Protein Immobilization Using Complex Coacervates and Complex Coacervate Thin Films Hursh Vardhan Sureka

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Protein Immobilization using Complex Coacervates and Complex

Coacervate Thin Films

Hursh Vardhan Sureka

B.S. Chemical and Biomolecular Engineering Georgia Institute of Technology, 2014

Submitted to the Department of Chemical Engineering in partial fulfillment of the requirements for the degree of

Doctor of Philosophy in Chemical Engineering

at the

MASSACHUSETTS INSTITUTE OF TECHNOLOGY

January 2021

Signature of Author: ______Department of Chemical Engineering January 14, 2021

Certified by: ______Bradley D. Olsen Professor of Chemical Engineering Thesis Supervisor

Accepted by: ______Patrick S. Doyle Robert T Haslam (1911) Professor Chairman, Committee for Graduate Students

Protein Immobilization using Complex Coacervates and Complex Coacervate Thin Films Hursh Vardhan Sureka

Submitted to the Department of Chemical Engineering on January 14, 2021 in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Chemical Engineering

Abstract

Enzymes can enable a wide and growing range of chemistries, often outperforming synthetic catalysts. However, enzymes must often be converted to heterogeneous catalysts. Protein immobilization enables this conversion and can enhance the stability of enzymes. Complex coacervates are highly effective at encapsulating and stabilizing enzymes. This thesis demonstrates the use of complex coacervate thin films for the immobilization of enzymes and systematically probes methods to enhance the performance of these materials. The first study presents a proof-of-concept demonstration of complex coacervate thin films for the synthesis of functional biomaterials. The immobilization method itself is all-aqueous, reducing the likelihood of enzyme denaturation, and facile, only requiring two steps: coating followed by crosslinking. A model biosensor was synthesized and demonstrated to have both high sensitivity and selectivity, and the immobilization method imparted increased thermal stability on the enzyme. From here, two directions were explored: how protein properties affect their coacervation behavior and optimizing the performance of the complex coacervate thin films. The second study aims to quantify the surface charge distribution or the “patchiness” of proteins and relate this to their complexation behavior. A patchiness parameter that averaged pair correlations between neighboring points on the protein surface was shown to correlate with the coacervation behavior of proteins with greater patchiness favoring the formation of complexes. Further work will enable this parameter to be incorporated with other protein properties in order to create robust predictive algorithms for protein-polymer coacervation. The third and fourth studies aimed to enhance the performance and properties of complex coacervate thin films. The third study probed whether the morphology of these composite materials could be controlled and found that morphologies varied greatly as a function of the polyelectrolyte strength and the loading of the encapsulated molecule. The strongest interactions led to precipitation, but weaker interactions led to micellization in both solution and the films. The fourth study aimed to understand how various polymer properties, including polyelectrolyte strength and monomer conformational freedom, affect the performance of complex coacervate thin films. Strong interactions were found to favor greater catalytic activity but lower stability, while weaker interactions favored greater stability.

Thesis Supervisor: Bradley D. Olsen, Professor of Chemical Engineering

Acknowledgements

The work presented in this thesis would not have been possible without the mentorship, support, and collaboration of my advisor, the Olsen group, my friends, and my family. My thesis advisor, Brad Olsen, has helped me become both a better researcher and a more collaborative and mindful colleague. We have had many interesting and insightful conversations about everything from science to management styles to Star Wars. He has challenged me to always produce high quality work by guiding me to ask interesting research questions and holding me to high standards for the data I produced. He has also supported my career goals as they evolved throughout my Ph.D. and actually allowed me to stick around to work on a very interesting research problem in conjunction with a start-up. Thank you Brad for creating an environment for scientists to thrive. I would also like to thank the Olsen group as a whole for making work more enjoyable, providing lots of interesting conversation, and creating an intellectually stimulating environment to conduct science. Thank you to my year-mates Helen and Irina for always being around to help keep me sane, provide moral support, discuss science, and accept fist-bumps of solidarity. Thank you to my scattering buddies Justin, Takuya, Sarah, and Aaron (and Helen even though we never went on a scattering trip together) for their support during those times of peak stress. Thank you to Allie, Aaron, Michelle S., and Carrie for all the training when I first started in the group. Thank you to Melody, Daphne, Danielle, Reggie, and Brian for being a calming influence on me and helping me keep my eye on the big picture. To my mini COVID lab pod, Melody, Haley, and Ameya, thank you for making the finishing stretch more bearable. Thank you to Melody, Haley, Emil, Patrick, Michelle C., Bruno, Jorge, everyone already mentioned, and many others for always creating a fun and interesting work environment and being excellent colleagues at work and friends outside of work. My friends at MIT have been a constant source of support and fun as well. Thank you to the “lunch train” for lots of fire bant, baseball and pong, TTS, late night conversations, and so much more over the years. It’s hard to believe we kept the chat alive for 5 years. You have all been instrumental in keeping me chugging on through grad school. Thank you to my year-mates for always providing moral support and lots of opportunities for fun and comradery like weekly trivia. Thank you to the tripod for being there since before we even started. Thank you to my COVID pod for helping me through the final push of grad school. Thank you to the ChemE department as a whole for welcoming me with open arms and allowing me to become part of this wonderful community. Finally, thank you to my family for being my rock through this arduous process. I can’t accurately express the amount of support my parents have provided these past 5 years (and for so many years before that). I didn’t always see it, but I know they set the path that got me here starting with our move to Jacksonville more than 20 years ago to get both me and my sister into Stanton for high school (and the feeder schools before). Thank you to my sister for helping keep me sane throughout. I know my extended family has kept me in their thoughts through all of this, and even though we haven’t been able to visit as much I know I had their support throughout. And thank you to the people who have become family over the years, the Jacksonville family friends, Deese, Cifredo, Ben, Sona, Sam, Teresa, and Nick, all of whom have always and will always have my back. I know this section does not do justice to the friendship, support, and mentorship that have gotten me here, but everyone listed here and anyone I have missed has my heartfelt gratitude.

Table of Contents Abstract ...... 3 Acknowledgements ...... 5 List of Figures ...... 11 List of Tables ...... 20 Chapter 1. Introduction ...... 21 1.1 Motivation ...... 21 1.2 Enzymes and Their Uses ...... 22 1.3 Functional Biomaterials from Enzymes...... 23 1.4 Complex Coacervates and Complex Coacervate Core Micelles (C3Ms) ...... 25 1.5 Protein Encapsulation in Complex Coacervates and C3Ms ...... 27 1.6 Ionic-Neutral Block Copolymer Self-Assembly ...... 29 1.7 Thesis Summary...... 31 1.8. References ...... 32 Chapter 2. Methods...... 39 2.1. Polymer Synthesis...... 39 2.1.1. BP monomer synthesis...... 39 2.1.2. Gel Permeation Chromatography (GPC)...... 43 2.1.3. OEGMA, 4VP, and DMAEMA purification...... 43 2.1.4. Test reaction setup...... 44 2.1.5. POEGMA and POEGMA-r-BP Syntheses...... 45 2.1.6. POEGMA-b-P4VP and (POEGMA-r-BP)-b-P4VP Synthesis...... 50 2.1.7. (POEGMA-r-BP)-b-PDMAEMA Synthesis...... 54 2.1.8. Quaternization of P4VP and PDMAEMA...... 57 2.1.9. RAFT Polymerization of Polyelectrolytes for Patchiness Study (Chapter 4)...... 59 2.2. Protein Preparation...... 66 2.2.1. Alkaline Phosphatase Cloning...... 66 2.2.2. Alkaline Phosphatase Expression and Purification...... 68 2.2.3. Preparation of Amylase Supercharged Variants...... 69 2.2.4. Expression and Purification of GFP Mutants...... 71 2.2.5. Nitroreductase Expression and Purification...... 73 2.3. Thin film preparation...... 75 2.3.1. PEG-Modification of Si surface...... 75 2.3.2. Flow coating...... 76 2.4. UV-Vis Absorption Methods...... 78 2.4.1. Turbidimetry ...... 78 2.4.2. Preparation of Standard Curve of 4-Methylumbelliferone (4MU)...... 79 2.4.3. Bulk Activity assays...... 81 2.4.4. Thin Film Activity and Metal Sensing Assays...... 82 2.5. Protein Surface Charge Analysis ...... 84 2.5.1. Design of GFP Mutants...... 84 2.5.2. Generating the Solvent Accessible Surface (SAS) and Surface Potential Map ...... 86 2.5.3. Search algorithm for patches...... 87 2.5.4. Algorithm for determining protein patchiness parameter...... 88 2.6. Solution and Bulk Characterization...... 88 2.6.1. Dynamic light scattering (DLS)...... 88

2.6.2. SANS...... 89 2.6.3. Optical Microscopy...... 89 2.7. Thin Film Characterization ...... 89 2.7.1. Atomic Force Microscopy (AFM)...... 89 2.7.2. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS)...... 90 2.8. References ...... 92 Chapter 3. Catalytic Biosensors from Complex Coacervate Core Micelle (C3M) Thin Films .... 93 3.1. Abstract ...... 93 3.2. Introduction ...... 94 3.3. Results and Discussion ...... 97 3.3.1. Materials Design...... 97 3.3.2. C3M Formation and Activity...... 100 3.3.3. Enzyme Immobilization and sensing within C3M Films...... 101 3.3.4. Film stability to thermal degradation ...... 111 3.4. Conclusions ...... 113 3.5. Methods...... 114 3.5.1. Benzophenone monomer synthesis (BP)...... 114 3.5.2. POEGMA-b-qP4VP Synthesis...... 114 3.5.3. Quaternization of (POEGMA-r-BP)-b-P4VP...... 116 3.5.4. Cloning of charge mutants of PhoA...... 116 3.5.5. Expression and purification of PhoA...... 116 3.5.6. Dynamic Light Scattering (DLS)...... 117 3.5.7. Grafting PEG to silicon wafers...... 117 3.5.8. Flow coating of coacervate solutions...... 118 3.5.9. Scanning probe microscopy (SPM)...... 118 3.5.10. Film height measurement...... 118 3.5.11. Grazing incidence small-angle X-ray scattering (GISAXS)...... 118 3.5.12. PhoA activity assays...... 119 3.6. References ...... 121 Chapter 4. The Effect of Protein Surface Charge Distribution on Protein–Polyelectrolyte Complexation ...... 125 4.1. Abstract ...... 125 4.2. Introduction ...... 126 4.3. Experimental ...... 129 4.3.1. Materials...... 129 4.3.2. Designing GFP mutants in silico...... 129 4.3.3. Algorithms for assessment of protein surface patchiness ...... 129 4.3.4. Search algorithm for patches...... 130 4.3.5. Algorithm for determining protein patchiness parameter...... 130 4.3.6. GFP mutant expression and purification ...... 131 4.3.7. Reversible addition-fragmentation transfer (RAFT) Polymerization ...... 132 4.3.8. Gel Permeation Chromatography...... 132 4.3.9. PAA polymerization...... 133 4.3.10. qP4VP polymerization...... 133 4.3.11. PSS polymerization...... 134 4.3.12. PDMAEA polymerization...... 134

4.3.13. Turbidity measurement...... 135 4.3.14. Zeta Potential and Dynamic Light Scattering (DLS) Measurements...... 135 4.3.15. Optical microscopy...... 136 4.4. Results and Discussion ...... 136 4.4.1. Protein design, patch calculation and expression ...... 136 4.4.2. Protein complexation with polyelectrolytes at various NaCl concentrations ...... 144 4.5. Conclusions ...... 156 4.6. References ...... 158 Chapter 5. Polyelectrolyte Complexation Driven Morphological Changes in Cationic-Neutral Block Copolymer Thin Films ...... 163 5.1. Abstract ...... 163 5.2. Introduction ...... 164 5.3. Results and Discussion...... 167 5.3.1. Materials Design...... 167 5.3.2. Complexation with PSS...... 168 5.3.3. Complexation with PAA...... 169 5.3.4. Complexation with Amylase...... 175 5.4. Conclusions...... 183 5.5. Materials and Methods...... 184 5.5.1. Materials...... 184 5.5.2. POEGMA Synthesis...... 184 5.5.3. POEGMA-b-P4VP Synthesis...... 185 5.5.4. POEGMA-b-P4VP Quaternization...... 185 5.5.5. P4VP Synthesis...... 185 5.5.6. P4VP Quaternization...... 186 5.5.7. PAA Polymerization...... 186 5.5.8. Preparation and Purification of α-Amylase and Supercharged α-Amylase...... 186 5.5.9. Turbidimetry...... 187 5.5.10. Dynamic Light Scattering (DLS)...... 188 5.5.11. Grafting PEG to Si Wafers...... 188 5.5.12. Film Casting...... 188 5.5.13. Atomic Force Microscopy (AFM)...... 188 5.5.14. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS)...... 188 5.6. References...... 189 Chapter 6. Enzyme Immobilization and Stabilization via Complex Coacervate Core Micelle (C3M) Thin Films ...... 193 6.1. Abstract ...... 193 6.2. Introduction ...... 193 6.3. Experimental...... 197 6.3.1. Materials...... 197 6.3.2. (POEGMA-r-BP) Synthesis...... 197 6.3.3. POEGMA-P4VP Synthesis...... 198 6.3.4. POEGMA-PDMAEMA Synthesis...... 198 6.3.5. Polymer Quaternization...... 198 6.3.6. NfsB Expression and purification...... 199 6.3.7. NfsB Supercharging...... 200

6.3.8. Preparation of Fused Silica...... 200 6.3.9. Polyethylene Glycol (PEG) Modification of Substrates...... 200 6.3.10. Film Casting...... 201 6.3.11. Dynamic Light Scattering...... 201 6.3.12. Atomic Force Microscopy (AFM)...... 201 6.3.13. Film Height Measurement...... 202 6.3.14. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS)...... 202 6.3.15. Film Aging...... 202 6.3.16. Activity Assays...... 202 6.4. Results and Discussion...... 204 6.4.1. Material Design...... 204 6.4.2. Morphology of NfsB-Block Copolymer Hybrid Materials...... 206 6.4.3. Film Performance...... 232 6.5. Conclusions...... 238 6.6. References ...... 241 Chapter 7. Conclusions ...... 245 7.1. Thesis Summary...... 245 7.2. Future Outlook ...... 248 7.3. References ...... 252 Appendix A. Supporting Information for Chapter 3...... 254 Appendix B. Supporting Information for Chapter 4...... 272 Appendix C. Supporting Information for Chapter 5 ...... 293 Appendix D. Supporting Information for Chapter 6 ...... 311

List of Figures

Figure 2.1. Elution profile of benzophenone methacrylate (BP) from Biotage. 41 Figure 2.2. 1H NMR of benzophenone methacrylate with labeled peaks in CDCl3 (400 MHz). 42 Figure 2.3. LRMS of BP Monomer. Expected mass was 266.1, found was 266.1. 42 Figure 2.4. Synthesis of POEGMA and POEGMA-r-BP via RAFT polymerization. 47 Figure 2.5. Representative GPC differential light scattering (solid) and light scattering (dashed) curves of POEGMA-r-BP. 48 1 Figure 2.6. Representative H NMR (400 MHz, CDCl3) of POEGMA-r-BP. 49 Figure 2.7. RAFT synthesis of (POEGMA-r-BP)-b-P4VP. 52 Figure 2.8. Sample 1H NMR of POEGMA-P4VP. 53 Figure 2.9. Sample GPC of POEGMA-P4VP. 54 Figure 2.10. RAFT synthesis of (POEGMA-r-BP)-b-PDMAEMA. 55 Figure 2.11. Sample 1H NMR of POEGMA-PDMAEMA. 56 Figure 2.12. Sample GPC of POEGMA-PDMAEMA. 57 Figure 2.13. Sample 1H NMR of POEGMA-qP4VP . 58 1 Figure 2.14. H NMR (500 MHz) in D2O of a) hyaluronic acid, b) sodium salt of PAA, c) qP4VP , d) PSS, and e) PDMAEA. 62 Figure 2.15. Gel permeation chromatography of synthesized charged polymers; PAA, qP4VP, PSS and PDMAEA. 65 -1 Figure 2.16. Titration data for PDMAEA. pKa was found to be 7.32 ± 0.05. 25 mL of 2 mg mL polymer was acidified to pH 2.56 and titrated with 50 mM NaOH. 66 Figure 2.17. a) FPLC chromatogram and b) paired SDS-PAGE analysis for the purification of PhoA M4. The red boxed regions correspond with one another. 69 Figure 2.18. SDS-PAGE of purified native α-amylase. 70 Figure 2.19. MALDI-TOF of native and supercharged α-amylase. 71 Figure 2.20. Properties of GFP mutants. (a) MALDI TOF of GFP variants (SP: 28.5 kDa, MP: 27.8 kDa, LP: 28.8 kDa, and XLP: 28.7kDa), (b) stained SDS-PAGE gel, (c) Circular dichroism spectra of GFP variants, and (d) UV-vis spectra of mutants at 1 mg mL-1, with inset image of UV fluorescence of purified GFP variants (2 mg mL-1). 73 Figure 2.21. a) SDS-PAGE analysis of Ni-NTA chromatography fractions. Elutions were pooled and dialyzed. b) SDS-PAGE analysis of dialyzed protein. 75 Figure 2.22. a) Standard curve in 96-well plates in MOPS buffer. b) Standard curve in 96-well plate in Tris buffer. c) Standard curve in 12-well plate in Tris buffer. 80 Figure 3.1. Synthesis of (POEGMA-rBP)-b-qP4VP via RAFT polymerization. POEGMA-r-BP was first polymerized via RAFT, then the polymer was used as a macro-RAFT agent to polymerize (POEGMA-r-BP)-b-P4VP. (POEGMA-r-BP)-b-P4VP was then quaternized with methyl iodide. 98 Figure 3.2. a) Structure of alkaline phosphatase (PDB: 3TG0)50 with mutations highlighted. b) Table of mutants and corresponding mutations and α-values. c) kcat and d)KM data for supercharged PhoA mutants determined by fluorimetric assay measuring conversion of 4-MUP to 4-MU. All mutants showed significantly greater kcat than the wild-type and a slight increase in KM. Mutant 4 was chosen for subsequent studies because it had sufficient negative charge ratio to enable complexation without a relative drop-off in activity. 99

Figure 3.3. a) Particle mass distributions determined by DLS for mixtures of PhoA and (POEGMA-r-BP)-b-qP4VP. Complexes began to form at 5% (wt/wt), with micellization occurring at 40% and 50%, and the formation of larger complexes occurring past this point. b) Effect of polymer addition on the activity of PhoA in solution. The polymer was added at 50% (wt/wt) with enzyme concentration remaining constant between the two cases. The effective kcat of the enzyme was reduced from 6.31 ± 0.01 s-1 to 3.84 ± 0.01 s-1 upon addition of the polymer and effective KM was found to decrease from 1.27 ± 0.03 μM to 0.94 ± 0.07 μM. 101 Figure 3.4. Method for immobilization of enzymes in C3M thin films. Polymer and protein are pre-mixed to form complex coacervate core micelles and then flow coated onto a solid substrate. UV irradiation is used to crosslink the soft block via benzophenone groups, rendering the final film insoluble. 102 Figure 3.5. a) Film activity for films of 50 wt.% enzyme at varying thickness. Activity increase with thickness is minimal beyond about 150 nm. b) Relative response to 5 ppm Zn2+ as a function of film thickness. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Relative response increase was minimal after 150 nm. 103 Figure 3.6. SPM height and phase images and GISAXS scattering patterns for films with 0% (a- c), 10% (d-f), 25% (g-i), and 50% (j-l) (wt/wt) protein loading. The pure block copolymer structures into disordered micelles under the conditions tested. With increasing protein concentration, this structure gives way to the formation of larger features. 105 Figure 3.7. a) Film activity and b) relative response to 5 ppm Zn2+ as functions of protein loading. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Both activity and response scaled with the protein loading. 106 Figure 3.8. SPM height and phase images for films as cast (a,b) and films annealed with water (c,d), acetone (e,f), DMSO (g,h), and DMF (i,j). Solvent annealing did not greatly affect the morphology of the films showing large complexes in all samples. 107 Figure 3.9. a) Film activity and b) relative response to 5 ppm Zn2+ after annealing with various solvents. All films are 50 wt.% protein. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Treatment with water had no effect on activity, but all other treatments led to a marked decrease in both activity and response. 108 Figure 3.10. a) Relative response of C3M films to treatments with various M2+ at 5 ppm. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with M2+ to the native film activity. Films showed high selectivity for zinc in all cases. The addition of other metals led to decreased response to Zn2+. b) Relative response of C3M film to varying Zn2+ concentration. Response was non-linear after 10 ppb Zn2+ and constant after 50 ppb. 109 Figure 3.11. The C3M films were tested under a variety of conditions and the samples were doped with 10 and 50 ppb Zn2+. This included a control in Millipore water, 10 ppb Cu2+, Co2+, and Ni2+, Charles River water (CRW), filtered Charles River water (CRWF), and 2 mixtures of metals (Mix 1: 5 ppb Cu2+, 1 ppb Co2+, 4 ppb Ni2+, and 7 ppb Zn2+; Mix 2: 2 ppb Cu2+, 6 ppb Co2+, 2 ppb Ni2+, and 4 ppb Zn2+). The response to 50 ppb Zn2+ was found to be statistically significant in all cases (**: p < 0.01, *: p < 0.05), except in CRWF (p = 0.052). The p-values are summarized in Table A.1. 110

Figure 3.12. Activity and response of films aged under ambient conditions (a,d), 40 °C and 50% RH (b,e), and 70 °C and 4% RH (c,f). Films retained their activity and response under ambient conditions and 40 °C and 50% RH, but were greatly inactivated at 70 °C and 4% RH leading to unreliable response. 112 Figure 4.1. Amino acid sequences of sfGFP mutants with mutation sites indicated in red. 139 -1 Figure 4.2. Different views of the electrostatic surface potentials (±2 kT ec ) of GFP mutants at the solvent accessible surface rendered from solutions of the linearized Poisson–Boltzmann equation using the Adaptive Poisson-Boltzmann Solver (APBS). The theoretical net charge was the same for all mutants (-4 at pH 8.0). 140 Figure 4.3. a) Charge patches identified by search algorithm for each GFP mutant at 0, 50, and 300 mM NaCl, with negative patches in and positive patches in blue. b) The average positive and negative patch areas of each mutant. c) The neutral area of each mutant where surface potential was less than |2 kT e-1|. 141 Figure 4.4. Patchiness parameter calculated for the panel of sfGFP mutants used in the study at pH 8.0 in varying salt concentrations (0, 50, and 300 mM). 143 Figure 4.5. Properties of GFP mutants. (a) MALDI TOF of GFP variants (SP: 28.5 kDa, MP: 27.8 kDa, LP: 28.8 kDa, and XLP: 28.7kDa), (b) stained SDS-PAGE gel, (c) Circular dichroism spectra of GFP variants, and (d) UV-vis spectra of mutants at 1 mg mL-1, with inset image of UV fluorescence of purified GFP variants (2 mg mL-1). 144 Figure 4.6. Zeta-potentials of a) the GFP mutants and b) the synthesized polymers in Tris buffer pH 8.0 with different concentrations of NaCl (0, 50 and 300 mM). 147 Figure 4.7. Turbidity profile of GFP mutants as a function of NaCl concentration and protein- polymer fraction. 149 Figure 4.8. Optical micrographs showing protein-polymer complexes resulting from mixing the GFP mutants with qP4VP at the maximum point of turbidity. 151 Figure 4.9. DLS data of GFP mutants with polyelectrolytes at different volume fractions and different concentrations of NaCl (0, 50 and 300 mM) in 20 mM Tris buffer (pH 8.0). 152

Figure 4.10. Small-angle neutron scattering (SANS) curves of XLP-PAA blends with fprot of a) 0.35 at 50 mM NaCl and b) 0.4 at 300 mM NaCl in Tris buffered D2O pH 8.0. SANS data are solvent corrected and fitted to a cylinder model62-64 with radii of 1.97 and 1.76 nm and lengths of 779 and 875 nm at 50 and 300 mM NaCl, respectively. 153 Figure 4.11. Phase behavior of GFP variants with a) qP4VP, b) PAA, and c) HA at the point of maximal complexation as a function of the patchiness of each protein at 0, 50, and 300 mM NaCl. 155 Figure 5.1. Turbidimetry data for the complex coacervation of P4VP 20q and 40q with PAA at 2 mg mL-1 in water. The point of maximal complexation is at f+ = 0.45-0.5. 170 Figure 5.2. AFM height and phase scans and GISAXS patterns for films of PAA and POEGMA-b-pqP4VP20 at various molar ratios of PAA to P4VP. The addition of PAA to the BC triggers the formation of a micellar phase. At the lowest loading of PAA (f+ = 0.67), the micelles are non-uniform as seen from both the defects in the AFM images and the broadness of the GISAXS peak. At higher loading of PAA (f+ = 0.5 and 0.33), near the point of charge balance, the micelles become more uniform as seen from both the AFM and the sharpening of the GISAXS peak. At the highest loading of PAA (f+ = 0.2), the micelles become swollen and non-uniform as seen from both the AFM and the broadening of the GISAXS peak. Figure C.4 provides 1-dimensional linecuts of the GISAXS data for further clarity. 171

Figure 5.3. AFM height and phase scans and GISAXS patterns for films of PAA and POEGMA-b-pqP4VP40 at various molar ratios of PAA to P4VP. The addition of PAA to the BC triggers the formation of a micellar phase. At the lowest loading of PAA (f+ = 0.67), the micelles are non-uniform as seen from both the defects in the AFM images and the broadness of the GISAXS peak. At the point of charge balance, the micelles become more uniform as seen from both the AFM and the sharpening of the GISAXS peak. At the highest loading of PAA (f+ = 0.3), the micelles become swollen and non-uniform as seen from both the AFM and the broadening of the GISAXS peak. Figure C.4 provides 1-dimensional linecuts of the GISAXS data for further clarity. 173 Figure 5.4. Turbidimetry data for the coacervation of pqP4VP20 and pqP4VP40 and the amylase variants at 2 mg mL-1 in water. The turbidity peak, which corresponds with the point of maximal complexation, is at f+ ≈ 0.6 for all variants. 176 Figure 5.5. AFM height and phase scans and GISAXS patterns for films of native amylase (A22) and POEGMA-b-pqP4VP40 at various mixing ratios. 178 Figure 5.6. AFM height and phase scans and GISAXS patterns for films of native (A22) and supercharged amylase (A30, A32, A37) and POEGMA-b-pqP4VP40 at various mixing ratios. 181 Figure 6.1. Activity of a) native and b) supercharged NfsB as a function of 4-NBS concentration at 0.5 mM NADH. Dashed lines represent Michaelis-Menten fits of the activity data. Thermally denaturing the enzyme variant at 70 °C for 12 h rendered each variant inactive. 206 Figure 6.2. Particle mass distributions determined by DLS for complexes formed between native NfsB and 0qPOEGMA-b-PDMAEMA at 1 mg mL-1 in water. Bimodal distributions of particle sizes give way to monomodal distributions with increasing protein loading for all three block copolymers. The mass fraction of complexes increases with increasing protein loading. 207 Figure 6.3. AFM height and phase scans of films of 0qPOEGMA36-PDMAEMA13 and native NfsB. The addition of protein led to the formation of micellar complexes, with the degree of complexation increasing with the protein loading. Similar behavior was seen with both 0qPOEGMA15-PDMAEMA5 and 0qPOEGMA55-PDMAEMA19 shown in Figures D.9 and D.10, respectively. Scale bars at 200 nm. 208 Figure 6.4. Particle mass distributions determined by DLS for complexes formed between native NfsB and 100qPOEGMA-b-PDMAEMA at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading. The mass fraction of complexes increases with increasing protein loading. 210 Figure 6.5. AFM height and phase scans of films of 100qPOEGMA-PDMAEMA and native NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two lower molecular weight encapsulants behave similarly, but the largest does not. At 50% protein loading, 100qPOEGMA15-PDMAEMA5 produced poor quality films due to apparent gelation of the coating solution (Figure D.12). Scale bars at 200 nm. 212 Figure 6.6. Particle mass distributions determined by DLS for complexes formed between native NfsB and 20qPOEGMA-b-P4VP at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading. The mass fraction of complexes increases with increasing protein loading. 213 Figure 6.7. AFM height and phase scans of films of 20qPOEGMA-P4VP and native NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two lower molecular weight encapsulants behave similarly at 10 and 25% protein loading, but not at 50%. At 50% protein loading the two higher molecular weight encapsulants behave similarly, while the smallest produced a poor quality film due to apparent gelation of the coating solution (Figure D.12). Scale bars at 200 nm. 215

Figure 6.8. Particle mass distributions determined by DLS for complexes formed between native NfsB and 100qPOEGMA-b-P4VP at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading for 100qPOEGMA15-P4VP5. Bimodal distributions gave way to monomodal distributions with greater protein loading for 100qPOEGMA36-P4VP11. For 100qPOEGMA55-P4VP17, the particle size distribution is only bimodal at 25%. The mass fraction of complexes increases with increasing protein loading. 216 Figure 6.9. AFM height and phase scans of films of 100qPOEGMA-P4VP and native NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two lower molecular weight encapsulants behave similarly at 10 and 25% protein loading, but not at 50%. At 50% protein loading the two higher molecular weight encapsulants behave similarly. Scale bars at 200 nm. 219 Figure 6.10. Particle mass distributions determined by DLS for complexes of SC NfsB and 0qPOEGMA-PDMAEMA. The mass fraction of material incorporated in the complexes increases with increased protein loading and the system forms small, monodisperse complexes. 220 Figure 6.11. AFM height and phase scans of films of 0qPOEGMA-PDMAEMA and SC NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two higher molecular weight encapsulants behave similarly, but not the lowest. Scale bars at 200 nm. 222 Figure 6.12. Particle mass distributions determined by DLS for complexes of SC NfsB and 100qPOEGMA-PDMAEMA. The mass fraction of material incorporated in the complexes increases with increased protein loading. 223 Figure 6.13. AFM height and phase scans of films of 100qPOEGMA-PDMAEMA and SC NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two higher molecular weight encapsulants behave similarly, but not the lowest. Scale bars at 200 nm. 225 Figure 6.14. Particle mass distributions determined by DLS for complexes of SC NfsB and 20qPOEGMA-P4VP. At 10% and 25% protein loading and at 50% for the two larger encapsulants, the system forms small, monodisperse complexes. With the smallest encapsulant at 50% protein loading, no apparent complexation occurs. The mass fraction of material incorporated in the complexes is maximized at 25% protein loading rather than 50% as seen with the other systems because the protein is in excess to the polymer (f+ < 0.5) for the 50% blends. 226 Figure 6.15. AFM height and phase scans of films of 20qPOEGMA-P4VP and SC NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two higher molecular weight encapsulants behave similarly, but not the lowest. Scale bars at 200 nm. 228 Figure 6.16. Particle mass distributions determined by DLS for complexes of SC NfsB and 100qPOEGMA-P4VP. Monomodal particle size distributions at 10% and 25% protein loading gave way to bimodal distributions at 50%/ The mass fraction of material incorporated in the complexes increases with increased protein loading for the larger encapsulants, but was nearly constant for the smallest. 229 Figure 6.17. AFM height and phase scans of films of 100qPOEGMA-P4VP and SC NfsB. Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two higher molecular weight encapsulants behave similarly, but not the lowest. Scale bars at 200 nm. 231

Figure 6.18. Activity data for films encapsulating native NfsB with initial activities shown in blue and activities after aging at 70 °C and 10% RH. For films encapsulating the native NfsB, choice of encapsulant had a much smaller effect on performance than with the supercharged variant. 234 Figure 6.19. Activity data for films encapsulating supercharged NfsB with initial activities shown in blue and activities after aging at 70 °C and 10% RH. For films encapsulating the supercharged NfsB, choice of encapsulant had a much greater effect on performance than with the native variant with the strongest polyelectrolytes showing the highest activity, but the weakest showing the greatest thermal stability. 236 1 Figure A.1. H NMR (400 MHz, CDCl3) of poly((oligo-ethylene glycol methacrylate)-r- (benzophenone methacrylate)) (POEGMA-r-BP). 253 1 Figure A.2. H NMR (400 MHz, CDCl3) of poly-[(oligo-ethylene glycol methacrylate)-r- (benzophenone methacrylate)]-b-(4-vinylpyridine)] ((POEGMA-r-BP)-b-P4VP). 254 1 Figure A.3. H NMR (400 MHz, D2O) of poly-[(oligo-ethylene glycol methacrylate)-r- (benzophenone methacrylate)]-b-(methyl-quaternized 4-vinylpyridine)] ((POEGMA-r-BP)-b- qP4VP). The polymer was quaternized > 95%. 255

Figure A.4. SEC analysis of POEGMA-r-BP. The polymer was found to have Mn of 31.8 kDa and Đ of 1.13. 256 Figure A.5. SEC analysis of (POEGMA-r-BP)-b-P4VP. Đ was 1.11. 256 Figure A.6. a) SDS-PAGE analysis of PhoA mutants before and after periplasmic purification and b) after FPLC purification. c) Representative Michaelis-Menten fits to enzyme activity. 257 Figure A.7. Turbidimetry data for qP4VP and PhoA. 258 Figure A.8. AFM height and phase images for C3M film before crosslinking (a,b), after crosslinking (c,d), and after soaking (e,f). Scale bars are 200 nm and z-scales are 15 nm and 15° for height and phase, respectively. The treatments did not lead to significant changes of the film morphology. 259 Figure A.9. Scanning probe microscope (SPM) height (a) and phase (b) images and and grazing incidence small-angle X-ray scatering (GISAXS) pattern (c) for film with 5% (wt/wt) protein loading. Scale bars are 200 nm and z-scales are 20 nm (a) and 10° (b). Scattering intensity is shown on a logarithmic scale. Film thickness was 120 ± 2 nm. SPM shows formation of large complexes; however, GISAXS indicates that the underlying structure is the same as that of the pure polymer (disordered micelles). 260

Figure A.10. The qy and qz linecuts of GISAXS data for films with increasing protein loading. The qy linecut was averaged over qz of 0.2 and 0.4 and the qz linecut was averaged over qy of 0.05 to 0.15. The structure showed diminishing peaks in both qy and qz with increasing protein concentration, indicating that the nanostructure of the polymer gives way to the formation of larger features with increased protein content. The dual peak in qz is observed due to internal reflections, while the peak at 0.30 is the position of the reflected beam. Based on both SPM and GISAXS the polymer appears to form a disordered micellar phase. 261 Figure A.11. GISAXS patterns for PhoA-polymer films annealed with different solvents. Intensity is shown on a logarithmic scale. The structure showed no significant change with annealing condition. 262

Figure A.12. Relative response to 5 ppm Zn2+ in the presence of various biological contaminants at relevant concentrations. No statistical difference was found for any of the contaminant tested (p > 0.05 vs. control). The concentration of glucose is 1 mM, based on the total concentration of carbohydrates reported in the sediment layer of rivers (1 mM)2, and much greater than that reported in river water (2.3 µM).3 The protein concentration is 10 mg L-1, based on the reported amino acid content in river water sediment (6.7 mg L-1),4 and significantly higher than reported amino acid content in the water column (less than 1 mg L-1).5-6 The E. coli concentration is 2300 ± 200 CFU per 100 mL, which is greater than the EPA standard for bacteria in recreational water (410 CFU per 100 mL).7 The mixture contains all of the contaminants at the previous concentrations. 264 Figure A.13. 1H NMR of BP Monomer. 265 Figure A.14. LRMS of BP Monomer. Expected mass was 266.1, found was 266.1. 266 Figure A.15. Sequence of PhoA. 266 Figure A.16. Primer Designs for engineering of PhoA. 267 Figure A.17. a) Standard curve in 96-well plates in MOPS buffer. b) Standard curve in 96-well plate in Tris buffer. c) Standard curve in 12-well plate in Tris buffer. 268 Figure A.18. Activity assay with and without additional DMSO to evaluate solvatochromic effects of DMSO concentration. The points at 50 μM are collected at the same conditions. The -1 -1 additional DMSO caused a negligible change in kcat from 7.45 ± 0.45 s to 7.17 ± 0.62 s ; however, it did cause a significant shift in KM from 1.7 ± 0.6 μM to 2.6 ± 0.6 μM. For the metals assay, this was not considered significant because the assays were conducted at a concentration such that the enzyme was expected to achieve its maximum reaction rate. 269 Figure A.19. Negative control with carbon tape (CT). Plotted against protein loading data for reference. Carbon tape had negligible activity. 270 Figure B.1. GFP variants model combined by EDTSurf and APBS. 272 Figure B.2. DNA sequences of GFP mutants 273 1 Figure B.3. H NMR (500 MHz) in D2O of a) hyaluronic acid, b) sodium salt of PAA, c) qP4VP (peak at 4.25 ppm confirms full quaternization, approx.. 5% (wt/wt) residual DMF), d) PSS, and e) PDMAEA (C′, D′, E′ correspond non-protonated PDMAEA, approx.. 5%). 275 Figure B.4. Gel permeation chromatography of synthesized charged polymers; PAA, qP4VP, PSS and PDMAEA. 278 Figure B.5. Positive (blue) and negative (red) electrostatic field isosurface contours for GFP -1 mutants (±2 kBT ec ) 280 Figure B.6. Purity analysis of GFP mutants after Ni-NTA purification. Main peaks are saturated for all mutants, but represent approximately 90% of all contents. Saturation prevents determination of true purity, which would be greater than 90%. 281 -1 Figure B.7. Titration data for PDMAEA. pKa was found to be 7.32 ± 0.05. 25 mL of 2 mg mL polymer was acidified to pH 2.56 and titrated with 50 mM NaOH. 282 1 Figure B.8. H NMR of synthesized qP4VP in D2O. The peak at 4.25 ppm shows fully quaternized methyl peak. Approximately 5% (wt/wt) residual DMF was found in the final polymer. 283 Figure C.1. Photographs of Precipitates from PSS with 20% quaternized P4VP. Positive charge fractions from left to right range from 0.1 to 0.9 with macroscopic precipitates forming at f+ of 0.3 to 0.7. 293 Figure C.2. Schematic of P4VP–PSS interaction showing the presence of induced charges on P4VP resulting from interaction with PSS. 293

Figure C.3. DLS data for PAA complexing with POEGMA-b-pqP4VP at f+ of 0.5. Data were taken in water at 1 mg mL-1. 294

Figure C.4. 1-D analysis of GISAXS data for PAA. Linecuts were averaged over qz of 0.25 to 0.3 nm-1. Peak micellization indicated by sharpest peaks occurs at the point of charge balance for both encapsulants. 295 Figure C.5. DLS of Amylase variants blended with POEGMA-b-pqP4VP20. Maximum radius of complexes occurred at f+ of 0.5 for all mutants. 296 Figure C.6. DLS of Amylase variants blended with POEGMA-b-pqP4VP40. 296 Figure C.5. AFM height and phase scans and GISAXS patterns for films of supercharged amylase (A30) and POEGMA-b-pqP4VP40 at various mixing ratios. 297 Figure C.6. AFM height and phase scans and GISAXS patterns for films of supercharged amylase (A32) and POEGMA-b-pqP4VP40 at various mixing ratios. 298 Figure C.7. AFM height and phase scans and GISAXS patterns for films of supercharged amylase (A37) and POEGMA-b-pqP4VP40 at various mixing ratios. 299 Figure C.8. SEC trace of POEGMA. 301 Figure C.9. 1H NMR of POEGMA-b-P4VP. 302 Figure C.10. SEC trace of POEGMA-b-P4VP. 303 Figure C.11. 1H NMR of POEGMA-b-20qP4VP. Comparison of initial P4VP aromatic peaks at 6.6 (2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) after quaternization confirm >95% conversion. 304 Figure C.12. 1H NMR POEGMA-b-40qP4VP. Comparison of initial P4VP aromatic peaks at 6.6 (2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) after quaternization confirm >95% conversion. 305 Figure C.13. SEC trace of P4VP homopolymer 306 Figure C.14. 1H NMR of 20qP4VP. Comparison of sum of peaks from 6.5 to 9.5 (aromatic P4VP peaks, 4H) to methyl peak at 4.4 (3H) confirms >95% conversion. 307 Figure C.15. 1H NMR of 40qP4VP. Comparison of sum of peaks from 6.5 to 9.5 (aromatic P4VP peaks, 4H) to methyl peak at 4.4 (3H) confirms >95% conversion. 308 Figure C.16. SDS-PAGE analysis of purified native amylase (A22). 309 Figure C.17. MALDI-TOF of native and supercharged amylase. 309 Figure D.1. Differential refractive index chromatographs from SEC of homopolymer and block copolymers of a) POEGMA-r-BP 15, b) POEGMA-r-BP 35, and c) POEGMA-r-BP 55. 311 Figure D.2. 1H NMR spectra of block copolymers prior to quaternization. 313 Figure D.3. 1H NMR spectra of quaternized block copolymers 314 Figure D.4. SDS-PAGE analysis of Native NfsB a) fractions from Ni-NTA chromatography and b) final dialyzed protein. 315 Figure D.5. MALDI-TOF analysis of native and supercharged NfsB. 316 Figure D.6. DLS of Native and supercharged NfsB in Water. 317 Figure D.7. Activity assay of Native and Supercharged NfsB plotted together. SC NfsB is 3 approximately 10 × less active than the native based on kcat/KM. 317 Figure D.8. AFM height and phase scans of 0qPOEGMA15-PDMAEMA5 with Native NfsB. 318 Figure D.9. AFM height and phase scans of 0qPOEGMA55-PDMAEMA19 with Native NfsB. 318

Figure D.10. GISAXS scattering patterns and qy linecuts for films of 0qPOEGMA-PDMAEMA. 319

Figure D.11. GISAXS scattering patterns and qy linecuts for films of 100qPOEGMA- PDMAEMA. 320 Figure D.12. Photograph of poor quality films (100qPOEGMA15-PDMAEMA5+50%N and 20qPOEGMA15-P4VP5+50%N) vs. regular (100qPOEGMA36-P4VP11+10%N). Poor quality films are cloudy and lack the color associated with the light refraction through the film because light is scattered instead, while the higher quality films have a mirror-like finish. 321

Figure D.13. GISAXS scattering patterns and qy linecuts for films of 20qPOEGMA-P4VP. 322 Figure D.14. 5 µm x 5 µm AFM images of 20qPOEGMA55-P4VP17 with 10% and 25% (wt/wt) protein loading of native NfsB. Large complexes increase in prevalence with the addition of protein over this range of loading. 323 Figure D.15. Enlarged AFM image of 100qPOEGMA15-P4VP5 Control film. a) Height and b) phase. 323

Figure D.16. GISAXS scattering patterns and qy linecuts for films of 100qPOEGMA-P4VP 324 Figure D.17. 5 µm x 5 µm AFM images of 20qPOEGMA55-P4VP17 with 0-50% (wt/wt) protein loading of SC NfsB. 325

List of Tables

Table 2.1. Synthesis conditions for POEGMA and POEGMA-r-BP presented in this thesis. The Mn and Đ were determined by GPC. Composition of POEGMA-r-BP was determined by NMR. 47 Table 2.2. Reaction conditions and data for POEGMA-b-P4VP and (POEGMA-r-BP)-b-P4VP. 52 Table 2.3. Reaction conditions and data for (POEGMA-r-BP)-b-PDMAEMA. 55 Table 2.4. Summary of Polymer Properties for Chapter 4. 61 Table 2.5. Summary and analysis of MALDI-TOF data for native and supercharged α-amylase. 71 Table 4.1. Summary of Polymer Properties 145 - Table 5.1. Structures, pKa/pI, and mass-to-charge ratios (m/z ) of model polyanions 168 Table 6.1. Molar mass and compositional data of diblock copolymer encapsulants. All polymers were synthesized with volume fraction of the cationic block of 22.5% ± 1.5%. 204 Table A.1. Contact angle of C3M Films before and after crosslinking and after soaking in buffer as determined on a goniometer. The films were found to be hydrophilic in all cases. However, soaking the films led to a marked increase in the contact angle, likely due to excess non crosslinked polymer and protein on the surface being washed away. 260 Table A.2. Statistical analysis (p-values for 1-tail T-test) of testing data 265 Table B.1. The number, size and potential of each patch in GFP mutants 279 Table B.2. Negative charge ratio (α) values of GFP mutants. 282 Table C.1. AFM image z-scales for Figure 5.7. 300 Table C.2. Summary and analysis of MALDI-TOF data. 310 Table D.1. Polymerization conditions for POEGMA-r-BP 311 Table D.2. Polymerization conditions for POEGMA-P4VP and POEGMA-PDMAEMA 312

Chapter 1. Introduction

1.1 Motivation

Enzymes are excellent biocatalysts that can enable complicated chemistries, such as carbon sequestration, biofuel production, complex sensing, and pharmaceuticals manufacturing, at very mild conditions.1-12 However, enzymes can be sensitive to environmental conditions, especially if they are far from the physiological range, such as high temperatures, non-neutral pH, and high solvent concentrations.1-2,5,13 In addition, enzymes can be both difficult and expensive to produce and purify.3,5-6,13-15 Because enzymes are typically homogeneous catalysts, i.e. dissolved in solution, recovery of these macromolecules from reaction mixtures can also prove difficult, making these processes economically infeasible without a method of retaining the enzymes.1-2,5,16

While methods like directed evolution show promise for engineering enzymes that are more robust against degradation,1,4,6,13-15,17 enzyme immobilization enables a simultaneous solution to two issues: the robustness of the enzymes and enzyme retention and recovery.1-2,5,13,18-19

Enzyme immobilization converts enzymes which are typically homogeneous biocatalysts into heterogeneous (or solid) catalysts.1-2,5,13,18-19 Immobilization has two main advantages: it enables easier retention and recovery of the enzyme and it can help safeguard the enzyme from degradation.1-2,5,13,18-19 While a number of strategies exist for achieving immobilization, many suffer from drawbacks such as low protein loading, transport limitations leading to poor kinetics, long processing times, or manufacturing processes that can damage the enzyme.1,13,20 This thesis aims to design and optimize a novel method for enzyme immobilization that achieves high protein loading, high activity, and high enzyme stability using simple, aqueous manufacturing techniques under ambient conditions. This chapter will provide an overview of the growing usage of enzymes

21 in industry and existing immobilization technologies, and then delve into the topics more specific to this work including complex coacervation and block copolymer morphology.

1.2 Enzymes and Their Uses

Enzymes are biocatalytic globular proteins that can be utilized to solve challenging problems in industry, medicine, defense, and other fields.1,3,5-6,14-15,19,21-23 Enzymes are active under aqueous and ambient conditions, unlike synthetic catalysts that often demand high temperature and pressure for ideal performance.1-3,5,18,24 Additionally, enzymes are selective, and can be both enantiospecific and stereospecific, allowing for minimization of undesired side reactions that can occur with synthetic catalysts.4,6,17 The superior properties of enzymes have led to the development of new technologies including enzymatic synthesis pathways in industry and enzyme-based sensors for medical, environmental, and defense applications, among others.1,3,5-6,14-15,19,21-23

The most common uses of enzymes in industry today include glucose-fructose isomerization for the production of high fructose corn syrup, animal feed, and detergents, but a growing variety of new enzymatic processes and applications are being developed and deployed.1,3,5,7,18-19,23 Enzymes have enabled complex chemistries in pharmaceuticals manufacturing, providing alternative syntheses for drugs, and have even been used as therapeutics.1,6,14-15,22,25-26 The expanding use of enzymatic processes has been largely driven by directed evolution and rational design driven by modeling which have enabled a wider range of chemistries and enhanced performance of enzymes for industrially relevant reactions.4,6,13,15,17

The incorporation of enzymes into biomaterials allows for further applications as well as potential enhancement of enzyme properties. For example, glucose sensors commonly used for diabetic patients rely on immobilized glucose oxidase to enable sensing and the encapsulation of therapeutic enzymes can aid in the prevention of proteolysis and aid in delivery of the

22 therapeutic.12,27-29 Size-exclusion mechanisms driven by material properties can also aid in the performance of protein-based biosensors by preventing diffusion of off-target macromolecules into the films, leading to increased sensitivity in biological fluids.30 Furthermore, the immobilization of enzymes enables their recovery and enhance their stability, both of which are important in the development of industrial enzymatic processes.1-2,5,13,18,31 The further development of encapsulation and immobilization technologies will enable the expansion of their potential uses.

1.3 Functional Biomaterials from Enzymes

Functional biomaterials can be synthesized using a wide variety of methods, but the most common is surface immobilization to form an enzyme monolayer.3,5,7,13,15,19 Monolayer immobilization can be achieved using non-specific physical, electrostatic, or chemical adsorption or more specific methods, such as covalent bonding via coupling reactions of a chemically- modified surface to a specifically engineered site on the enzyme surface.1-2,5,13,16,18,31-34 Monolayer immobilization has been demonstrated to retain the enzymes, maintain—and even enhance, in some cases—enzyme activity, and stabilize enzymes against denaturation.1-2,5,13,16,18,31-35 A particular disadvantage of the more commonly used non-specifically bound monolayers is that there is no control over the orientation of the protein, which can lead to loss of function due to obstruction of the functional site.30,35-37 Binding to an engineered site on the protein can help alleviate this issue by allowing conformational control, and the use of a spacer between the enzyme and the surface has been demonstrated to improve the enzyme stability.35,37-38 Protein loading in monolayers is limited by the grafting density at the surface, and moving from a 2-dimensional to a 3-dimensional (3D) geometry can significantly increase specific protein loading.20,30,36

Immobilization can be achieved with a 3D geometry using numerous methods including encapsulation in polymer gels, direct self-assembly of protein-polymer block copolymers, and encapsulation in polyelectrolyte complexes.2,13,20,39-41 Protein can be encapsulated within both synthetic and natural materials such as hydrogels, metal-organic frameworks (MOFs), and even viral capsids, and entrapment within these nanoconfined environments has been demonstrated to stabilize enzymes, but transport and conformational limitations within these materials can decrease the effective activity of the enzymes within them.3,29,41-44 The addition of a semi-permeable membrane around a liquid or solid enzyme-containing material can enhance enzyme retention and stability but can cause loss of enzymatic function at the interface and introduce transport limitations.45 In both cases, the tuning of material properties such as particle and pore size is essential to maximizing functionality.

Layer-by-layer (LbL) methods utilize polyelectrolyte complexation to enable the aqueous processing of biofunctional materials.46 LbL films are based on the electrostatic adsorption of oppositely-charged polyelectrolytes to one another, and can be employed to immobilize charged enzymes with enzyme-loading directly controlled by the number of adsorption cycles.46-47 The method does require that the protein carries a moderately strong charge, but the effect of protein charge on the loading can be lowered by associating the protein with an oppositely-charged polymer first to form polyelectrolyte complexes (PECs), then forming LbL films with the PECs and a polymer of opposite charge to the net charge of the PECs.48 The LbL method has been demonstrated to be effective on simple surfaces as well as nanoparticles, implants, and other advanced materials.46-47

Incorporating a protein into a block copolymer (BC) by grafting it to a polymer provides a means of creating 3D microphase separated functional biomaterials.20,36,39,49-50 These protein-

24 polymer bioconjugates have been demonstrated for use in biosensors and biocatalysts, outperforming traditional 2D immobilization methods and some existing 3D methods.20,30,36 The protein-polymer BCs demonstrate phase behavior reminiscent of tradition BCs in both the solution and solid state, exhibiting known phases including cylindrical and lamellar morphologies.39,51

Selection of the polymer can have a significant effect on the morphology of the bioconjugates due to changes in the Flory-interaction parameter, χ, similar to traditional BCs.52 One difficulty is that the yield of bioconjugate can be relatively low (30-60%) even when using a 10-fold excess of the polymer.39 Similar phase behavior has been observed in protein-ELP fusions, where an ELP behaves like the polymer block of the bioconjugate, and the direct expression of the protein-ELP fusion avoids this synthetic loss and simplifies protein purification.53-55 While these methods and others have their advantages, polyelectrolyte complexation presents the potential to combine the self-assembly behavior of the BCs with the facile synthesis of the LbL methods.

1.4 Complex Coacervates and Complex Coacervate Core Micelles (C3Ms)

Complex coacervates are bio-inspired materials that have generated significant interest for use in adhesives, medicine, and protein immobilization.27,56-68 Complex coacervation is a liquid- liquid phase transition that can occur in solutions of oppositely-charged macromolecules in systems that are traditionally composed of two oppositely-charged polymers, salt, and water.69-71

The phase behavior in these systems is most strongly driven by the salt concentration, polymer concentration, and ratio of the interacting polymers with phase separation leading to the formation of a polymer dilute phase and a polymer dense phase, known as the complex coacervate.69-73 The phase separation occurs due to both the entropic effect of counterion release and the enthalpic effect of the polymer-polymer coulombic interactions.69,73-75

Complex coacervation is well-predicted for polymer-polymer systems by a number of models. These models have been discussed in great detail in recent work by Charles Sing and

Sarah Perry.73-74,76 Complex coacervation was first modeled by Voorn and Overbeek via an entropic mixing term with an added Debye-Hückel interaction term to account for electrostatic interactions, which was further built upon by Tainaka with the addition of the Flory-Huggins interaction to give the following equation:

3 퐹푉푂 휙푖 1 = ∑푖 ln 휙푖 + ∑푖푗 휒푖푗휙푖휙푗 − 훼[∑푖 𝜎푖휙푖]2 (1.1) 푉푘푏푇 푁푖 2

where 휙푖 is the volumetric fraction of each component, 푁푖 is the degree of polymerization of each component, 휒푖푗 is the Flory-Huggins interaction parameter, 𝜎푖 is the charge density, and 훼 is the dimensionless Bjerrum length.69,71,75 The Voorn-Overbeek model can provide excellent agreement with experimental data in terms of fitting the phase transition curve, but often requires

72,77 that 휒푖푗, 𝜎푖, and 훼 be used as fitting parameters. Additionally, the Voorn-Overbeek theory incorrectly predicts that the coacervate phase will contain a greater salt concentration than the dilute phase because it ignores both the effect of molecular connectivity on reducing the entropic penalty for charges on the same chain interacting with another interconnected chain and the entropic effect of counterion release on the complex coacervation phase transition.72-73,77-78 In fact, counterion release alone can be used to provide a model for this phase transition.79-80 In reality, the phase transition is likely driven by both enthalpic and entropic driving forces, with the relative importance of these forces depending on the specific system.79,81 A number of recent models have been able to more accurately capture the complex coacervation phase transition and more closely predict experimental results including phase compositions.72-74,76-78

When one or both of the interacting polymers are swapped for an ionic-neutral block copolymer, the charge interactions can lead to the microphase separation in solution, most

26 commonly forming complex coacervate core micelles (C3Ms), where a hydrophilic neutral block from the corona and the core of the micelle consists of the oppositely-charged polyelectrolytes.27,63,67,82-84 This architecture can be reversed by the use of a hydrophobic neutral block.82 These soluble complexes have been demonstrated to adopt vesicular, spherical, or cylindrical morphologies as a function of the salt concentration, neutral block length, and the charge ratio.85-86 The phase behavior of these complexes has been modeled, predicting shifts in morphology as a function of both the polymer properties and the salt concentrations; however, these results have not been fit to experimental results.86-87 Both complex coacervates and C3Ms have been demonstrated as effective methods for protein immobilization.56-57,64,67,82,84-85,88-89

1.5 Protein Encapsulation in Complex Coacervates and C3Ms

The high water content and densely populated nature of the complex coacervate phase make it an ideal candidate for the immobilization of proteins because of the similarity to the proteins’ natural environment.90-91 In fact, coacervation is considered to have been essential in the origins of life, and may play an important role in the segregation of proteins in the cytoplasm of cells, even allowing for the formation of membrane-less organelles via multiphase complex coacervation.90-91 Protein complex coacervates were first demonstrated by Bungenberg de Jong and Kruyt, who coined the concept in 1929,70 but the encapsulation of functional proteins has been a more recent development in the field.40,66 The encapsulation of proteins within coacervates and

C3Ms has since been proposed as a method for the stabilization and purification of proteins.26,88,92-

94 A wide variety of model proteins and enzymes have been immobilized in both complex coacervates and C3Ms, and both complex coacervates and C3Ms have been demonstrated to achieve high protein loading.26,58,66-67,82,88-93,95 Both complex coacervates and C3Ms can improve the activity and stability of proteins, and have been demonstrated to broaden the pH and

27 temperature range over which enzymes are stable and to provide protection against denaturants such as organic solvents and urea.21,66,93-94,96 This stabilization ability makes these materials interesting for use in biomaterials because biocatalysts and biosensors may be exposed to both harsh conditions and long storage times.

Encapsulation in both CCs and C3Ms can take place in two ways, either direct complex coacervation of the protein where the protein’s own charge can allow for complexation to occur with a polyelectrolyte or indirect encapsulation where the protein partitions into a coacervate of two oppositely charged polyelectrolytes.21,56-57,64,67 Typically, the direct method leads to higher protein loading, but may not always be feasible.21,64,67 Genetic or chemical alteration of the protein charge can enable the direct complex coacervation of the protein, but may lead to diminished performance of the enzyme depending on positioning of the active site and other properties of the protein.64,85,93 The partitioning method allows for proteins with weaker surface charge and proteins that cannot be modified without altering function to be incorporated into the coacervate.21 Recent work has provided some insight into how to better modify proteins for incorporation into complex coacervates and C3Ms. The use of localized charged tag, as opposed to isotropically distributed charge modifications on the protein surface, promotes complex coacervation over precipitation of the target protein, and provides a facile method for charge modification requiring only one terminus of the target protein to be modified instead of multiple sites.93 However, localizing charge interactions to a tag on the protein may diminish some of the thermal and chemical stabilization properties that have been demonstrated with complex coacervates, and will need to be further investigated. Despite the growing interest in the encapsulation of proteins within complex coacervates, predictions of protein coacervation behavior are very limited.

The models for polymer-polymer complex coacervation described previously do not work well for protein-polymer coacervation due to the folded morphology of the protein, the zwitterionic nature of the protein surface, the anisotropic distribution of charge on the protein surface, and the ability of amino acids to charge regulate.26,57,64,92,97 These properties of proteins can lead to proteins coacervating on the “wrong side” of their isoelectric point (pI), i.e. a protein with a net-negative charge coacervating with a negatively charged polymer or vice versa.57,92-93 Some work has begun to look at how to predict the complex coacervation of proteins. One heuristic predicts coacervation behavior based on the ratio of negative charges to positive charges of the protein (α) for the prediction of when a negatively charged protein would form complex coacervates and C3Ms, where α greater than approximately 1.2 and 1.4 was necessary for complex coacervate and C3M formation, respectively.64 A pair correlation function of the charged residues of proteins can serve as a metric to describe the charge distribution of proteins and patches seen by this metric have been shown to correlate with increased portioning of proteins into complex coacervates.57 Some modeling work exploring the effect of charge anisotropy on complexation of colloidal particles with polyelectrolytes has shown widened phase envelopes for the coacervation of patchy proteins as opposed to homogeneously charges ones, but further work is necessary to verify the results of the simulations.98 Further development of the theory of protein-polymer complex coacervation is necessary to enable predictive design of protein-polymer hybrid materials based on CCs and

C3Ms.

1.6 Ionic-Neutral Block Copolymer Self-Assembly

The ionic-neutral block copolymers that are used in the formation of C3Ms are also of interest in technologies such as new battery materials, selective membranes, and templated layer- by-layer deposition because the charged block can act as an ion-channel or functional template,

29 while the neutral block can provide structural integrity or serve as a channel for substrates.99-101

Two common forms of these block copolymers exist, the first, which is used in the C3Ms, is a charged-neutral BC, and the second is a neutral-neutral BC where one of the blocks has a significantly higher dielectric constant and so has a preferable solubility for salts.66,102-104

The self-assembly of these materials is determined by the block volume fraction, the molecular weight, and χ similar to regular block copolymers; however, the χ of ionic-neutral BCs is a function of the degree of ionization of the charged block or the salt loading.102-103,105-106 When an ionizable BC is used, the degree and method of ionization can alter the effective value of χ, allowing for tuning of the nanostructure in both thin films and solid-state.102-103 When driven by differences in salt solubility between the blocks, the shift in the phase behavior is driven by two competing factors, the entropically favorable uniform distribution of the ions between the blocks, which leads to increased miscibility of the blocks and an effective decrease in χ, and the energetically favorable solvation of the ions in the block with the higher dielectric constant, which leads to decreased miscibility of the blocks and an effective increase in χ.100,105,107-108 In polyethylene-b-poly(ε-caprolactone) (PEO-b-PCL), the addition of LiClO4 induces microphase separation, with χ increasing linearly with salt concentration at low concentrations, but decreasing at higher concentrations.105,108 In polystyrene-b-polyethylene oxide (PSS-b-PEO), the addition of lithium salts increases the effective χ, leading to a shift in the phase behavior.107 As previously discussed, the addition of both oppositely charged polymers and proteins to these materials in solution can trigger phase changes to various morphologies, but the solid-state behavior of such materials has not been studied.

1.7 Thesis Summary

The goal of this thesis is to demonstrate the viability of C3M thin films for creating nanopatterned enzymatic biosensors and biocatalysts and to explore how polymer and protein properties affect protein encapsulation and the film nanostructure and performance. The remainder of the thesis is organized as follows. Chapter 3 contains a proof-of-concept biosensor for enzyme- based sensing of transition metals using C3M thin films. In addition to the sensing efficacy, the film morphology, and the stability are studied. Chapter 4 aims to better understand the coacervation behavior of proteins by understanding and attempting to quantify how surface charge distribution of the protein affects the phase behavior of protein-polyelectrolyte systems. Chapter 5 explores how the charge density of the BC affects the complexation behavior and thin film morphology of the BC with oppositely-charged polyelectrolytes of varying polyelectrolyte strength, including a model protein. Chapter 6 expands on the work in Chapter 3 and 5 by exploring the structure-property relationship for C3M thin films. In addition to structural and activity analyses, the folding of the model enzyme is directly measured using optical analyses of dual- labeled enzymes.

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Chapter 2. Methods.

Reproduced (adapted) in part with permission from Sureka, et al. Catalytic Biosensors from

Complex Coacervate Core Micelle (C3M) Thin Films, ACS Appl. Mater. Interfaces 2019, 11, 35,

Reproduced (adapted) in part with permission from Kim,* Sureka,* et al. Effect of Protein Surface

Charge Distribution on Protein–Polyelectrolyte Complexation, Biomacromolecules 2020, 21, 8,

2.1. Polymer Synthesis.

2.1.1. BP monomer synthesis.

The benzophenone-based photo-crosslinking monomer, benzophenone methacrylate (BP), was synthesized per a previously described method.1 Triethylamine (10 mL, Sigma-Aldrich) was added to a solution of 4-hydroxybenzophenone (10 g, 50 mmol, Sigma-Aldrich) in dichloromethane (100 mL, VWR) in a 300 mL three-neck flask under stirring, and the solution was cooled to 0 °C in an ice water bath. A solution of methacryloyl chloride (5.9 mL, 60 mmol,

Sigma-Aldrich) in dichloromethane (50 mL) was prepared in a separate flask and transferred to a dropping funnel, and added dropwise at a rate of approximately 1 drop per 2 to 3 s while stirring.

Ensure there is ice present during the entirety of the addition, add more if necessary. The reaction mixture was then allowed to warm to room temperature and stirred overnight. The flask can be left in the ice bath, which will melt and allow the reaction to gradually get to room temperature.

The mixture was washed twice each with water (50 mL x 2), saturated NaHCO3 (50 mL x

2), and brine (50 mL x 2) in a separation funnel. The organic layer was then dried over Na2SO4 and filtered. The solvent was evaporated via rotovap. The resulting residue was purified by automated flash chromatography using a Biotage system with dichloromethane (DCM) and

39 methanol as a mobile phase using a 50 g Biotage® SNAP Ultra, KP-SIL column. The product was eluted from the column using 5 column volumes (CVs) of DCM followed by a 10 CV ramp to

10% methanol, with only one peak seen by UV, corresponding to the product (Figure 2.1). The tail of this peak was discarded. Please note that these columns have been discontinued, and the existing method in the Biotage system may need to be re-optimized for the new columns. The resulting solution of purified monomer was then dried by rotovap in a 250 or 500 mL flask. The solution will start to become very viscous as the product continues to dry, leading to higher risk of

“bumping” the mixture. If NMR indicated the presence of residual solvent, the monomer was further dried in the vacuum oven at room temperature overnight or until no further solvent was detected. 1 H NMR (400 MHz, CDCl3): δ 7.90 (m, 2 H), 7.83 (m, 2 H), 7.62 (m, 1 H), 7.52 (m, 2

H), 7.29 (m, 2 H), 6.42 (m, 1 H), 5.84 (m, 1 H), 2.12 (m, 3 H) (Figure 2.2). LRMS (electrospray

+ + ionization, ESI) calculated for C17H15O3 ([M H] ) 267.1, found 267.1 (Figure 2.3).

Figure 2.1. Elution profile of BP from Biotage, based on UV detector. Tail beyond 4 CV was discarded.

1 Figure 2.2. H NMR of benzophenone methacrylate with labeled peaks in CDCl3 (400 MHz).

Figure 2.3. LRMS of BP Monomer. Expected mass was 266.1, found was 266.1.

2.1.2. Gel Permeation Chromatography (GPC).

Unless otherwise noted, all GPC analyses were performed in dimethylformamide (DMF) on an Agilent 1260 LC system equipped with two columns (ResiPore, 300 × 7.5 mm, up to 500k

Da, Agilent Technologies, CA) in series. DMF with 0.02M LiBr was used as the eluent with a flow rate of 1 mL/min at 70 °C. The detector system consisted of a Wyatt miniDAWN TREOS multi-angle light scattering detector and a Wyatt Optilab T-rEX differential refractive index detector. Samples were prepared in DMF with 0.02M LiBr at a concentration of 1 to 5 mg mL-1 of polymer. 1 mL of the sample was drawn into a 1 mL syringe (Norm-Ject) and filtered through a

0.2 µm PTFE syringe filter into the appropriate vials. To avoid spillage, a gap should be left between the filter and the vial and a leur-lock syringe should be used.

2.1.3. OEGMA, 4VP, and DMAEMA purification.

Oligo(ethylene glycol methyl ether methacrylate) (OEGMA, Mn = 300, MilliporeSigma),

4-vinylpyridine (4VP, 98%, MilliporeSigma), and dimethylaminoethyl methacrylate (DMAEMA,

MilliporeSigma) were purified via basic alumina column. Columns were prepared with a small piece of cotton, an approximately 1 to 2 cm layer of washed sand, followed by basic alumina equal to approximately ½ the volume of monomer to be purified as described previously.2 The first several drops were discarded, and purified monomer was collected in an appropriately sized sealable container. The process was repeated once more. The monomer was then either used immediately or sealed further with parafilm and stored at -20 °C for no longer than 1 day. For volumes of 2-5 mL the columns were set up in either a pasture pipette or a 10 mL syringe. For less than 20 mL, the column was setup in a 20 mL syringe. For larger volumes, a traditional glass column was employed. 4VP changed from a yellow or brown color to nearly colorless upon removal of the inhibitor. OEGMA and DMAEMA did not change color and were clear both before

43 and after. If impurities were detected by NMR as was the case for one batch of 4VP, the monomer was vacuum distilled twice to remove the impurity. The 4VP was warmed to 80 °C over anhydrous

MgSO4, and distilled at approximately 500 mTorr. Distillate was collected in a flask chilled by a wet ice bath. The first several drops were discarded as a light fraction (~5% of the volume added) and the distillation was allowed to proceed until approximately 10-20% of the original volume remained in the bottoms. The monomer was sealed and stored at -20 °C for no longer than 1 day.

2.1.4. Test reaction setup.

To speed up testing of polymerization conditions, test reactions were setup such that 8-12 reactions could be run in parallel at the 2-10 mL scale. The reactions were done in 20 mL vials with septa caps for larger scale (> 4 mL) and 5 mL vials with septa caps for smaller scale in an appropriately-sized heat block (Chemglass CG-4904 and CG-4904 Dram). Reactions were degassed by N2-purging with needles connected to the N2 lines of the Schlenk line submerged below the liquid level (2-inch 21 gauge needles or 6-inch needles worked well) and outlet needles

(1.5-inch 21 gauge needles) inserted above the liquid level. Vials were purged for 20-25 min with

~3-5 bubbles per second and outlet needles were removed < 1 s before inlets. When more than 4 reactions were being run, the additional vials were purged in an available hood. Reactions were tested in varied solvents, temperatures, initiator ratios, and monomer concentrations. Conversion was monitored using NMR to track the disappearance of vinyl bonds and the ratio of monomer peaks to the shifted equivalent polymer peaks. Samples were taken every 2 or 3 h depending on the expected reaction rate for a total of 6 to 18 h. Sample needles and syringes were purged by drawing in and expelling N2 3 times from a purged, septum-sealed, N2-filled flask connected to the synthesis line.

2.1.5. POEGMA and POEGMA-r-BP Syntheses.

Both POEGMA and POEGMA-r-BP were synthesized by reversible addition- fragmentation chain transfer polymerization (RAFT) (Figure 2.4). Purified OEGMA was dissolved in dioxane at a 1:3 ratio by volume, and BP was added at a 1:20 ratio by weight to OEGMA for

POEGMA-r-BP polymerizations. This ratio was found to be sufficient for effective crosslinking by Dr. Allie Obermeyer. The chain transfer agent (CTA), 4-Cyano-4-(phenylcarbonothioylthio) pentanoic acid (CPP), and the initiator (I), azobisisobutyronitrile (AIBN), were added at a 5:1

(CTA:I) ratio at an appropriate ratio to the monomers to reach the target molecular weight. The reaction was carried out in a 500 mL Schlenk flask. It was degassed via 3 freeze-pump-thaw cycles and initiated by placing in a heated oil bath set to 65 °C. A freeze-pump-thaw cycle is a method to remove dissolved gases from the reagents used in the reaction, and enables air-free synthesis. Once all of the reagents for the RAFT reaction (and the stir-bar) have been added to the reaction flask, connect the flask to the Schlenk line and seal any openings with rubber septa. Pull gentle vacuum on the flask and flow in N2 a few times to remove most of the oxygen in the void space to avoid potentially condensing it, then seal the flask using the glass or Teflon stopper (you can flip the line to the flask to vacuum at this time to make sure the line is fully pumped down). Then, freeze the reaction mixture in LN2 (the LN2 will stop boiling hard when the reaction is fully frozen and the bottom of the Dewar will be easily visible), and once frozen pull vacuum on the reaction by opening the stopper until it reaches at maximum 100 mTorr, but ideally 40-50 mTorr (this may require some cleaning or maintenance of the line; the pump step is done while the vessel is in LN2).

After the vessel is pumped down, seal the flask and thaw in an isopropanol bath for 10-15 min then a water bath (using IPA initially prevents the formation of an ice cap that can slow down the thawing process, and replacing the water often can speed up the process). The thaw step is

45 substantially faster for small reactions (< 20 mL) and can be done in a single water bath. Repeat the freeze step and begin the next cycle (do not add N2 in between cycles). The vessel can filled with N2 after the final thaw step, but if being extra careful, add N2 after the last pump step but before thawing, pump out N2 and replace again to ensure pristine environment. After the reaction mixture is fully thawed, place on heat to start reaction. The reaction was terminated by removing from heat and exposing to air. The reaction was then precipitated three times into hexanes, using dichloromethane to dissolve the polymer after the first and second precipitations. The solvent was decanted off because the polymers do not solidify at room temperature. The polymer was then dried by rotovap. If there was still solvent detected by NMR, the polymer was furthered dried in the vacuum oven at room temperature or 40-50 °C for 1-2 days (be especially careful with heating the POEGMA-r-BP, and avoid if possible). Periodically stirring the polymer aided in the drying process. Molecular weight (Mn) and polydispersity (Đ) were determined by GPC. The monomer ratio for POEGMA-r-BP was determined by 1H NMR by comparison of the 2H and 3H peaks at

4.1 and 3.4, respectively, for OEGMA to the 3H and 4H peaks at 7.5-7.7 and 7.8-8, respectively, for BP. An example of the GPC and NMR characterization of POEGMA-r-BP is shown in Figure

2.5 and 2.6, respectively. GPC data do not include UV detection due to the possibility of crosslinking the benzophenone, and a GPC method in Breeze (“GPC DMF run no UV”) was used as opposed to the standard method with all settings but the wavelength of the UV detector (changed from 280 nm to 600 nm) kept the same. The reaction conditions and resulting Mn, Đ, and monomer ratios are summarized in Table 2.1. Supporting NMR and GPC data are included in the appendices of the chapters in which the materials were used.

Figure 2.4. Synthesis of POEGMA and POEGMA-r-BP via RAFT polymerization.

Table 2.1. Synthesis conditions for POEGMA and POEGMA-r-BP presented in this thesis. The

Mn and Đ were determined by GPC. Composition of POEGMA-r-BP was determined by NMR.

Vol. Vol. Mass Mass Mass Dioxane OEGMA BP CTA AIBN Temp. Time Mn %BP (mL) (mL) (g) (mg) (mg) (°C) (h) (kDa) Đ (mol) Chapter POEGMA-r- 58.0 19.05 1 133.0 15.6 65 7.5 22.3 1.13 8 3 BP22 (60 g) (20 g) POEGMA37 180 60 N/A 326.9 38.4 65 5 37.3 1.05 N/A 5 POEGMA- 180 60 3.20 609.6 71.7 65 6 15.5 1.06 6 6 BP15 POEGMA- 180 60 3.20 275.0 32.3 65 5.5 35.9 1.05 6 6 BP36 POEGMA- 180 60 3.20 195.6 23.0 65 5 55.0 1.10 6 6 BP55

Figure 2.5. Representative GPC differential light scattering (solid) and light scattering (dashed) curves of the POEGMA-r-BP of three molecular weights used in Chapter 6.

1 Figure 2.6. Representative H NMR (400 MHz, CDCl3) of POEGMA-r-BP from Chapter 3.

Comparison of the F (4H) and G (3H) peaks to the D (3H) peak was used to determine the ratio of

BP to OEGMA in the polymer.

2.1.6. POEGMA-b-P4VP and (POEGMA-r-BP)-b-P4VP Synthesis.

The POEGMA-r-BP-b-P4VP used in chapter 3 was synthesized as follows by Dr. Allie

Obermeyer. POEGMA-r-BP was used as a macromolecular chain transfer agent for RAFT polymerization of 4VP (Figure 2.7). The monomer (M), 4VP (3.68 g, 35 mmol), and AIBN (3.2 mg, 0.020 mmol) were added to a solution of the POEGMA-r-BP copolymer (3.24 g) in 12.5 g of a mixture of 1,4-dioxane and DMF in a molar ratio of 350:1.0:0.2 (M:CTA:I). The mixture was degassed by 3 freeze-pump-thaw cycles and backfilled with N2 then heated to 75 °C and terminated after 6 h by removal of heat and exposure to oxygen. The polymer was then precipitated in diethyl ether and dried under vacuum. The polymerization provided a well-defined (POEGMA-r-BP)-b-

P4VP diblock copolymer of molecular weight Mn = 29.3 kg mol–1 with a polydispersity of 1.11

1 (by SEC, Figure A1.5). DP4VP and Mn were determined by averaging the comparison of the H

NMR (400 MHz, CDCl3) peaks at δ 6.33 (m, 2 H) and 8.26 (m, 2 H) for 4VP and to the 3.31 (s, 3

H) peak for OEGMA (Figure A1.2).

Further reactions were designed to target approximately 20-40% conversion of 4VP per the test reactions and data from Stewart-Sloan’s work.2 Reactions contained equal parts dioxane and 4VP per Stewart-Sloan’s procedure to synthesize POEGMA-b-P2VP, but a small amount of

DMF was necessary due to the difference in P4VP and P2VP solubility (1:5 DMF:Dioxane by volume). For reactions targeting lower Mn P4VP, the solvent volume was doubled to enable dissolution of the POEGMA or POEGMA-r-BP macroCTA. POEGMA or POEGMA-r-BP macroCTA was dissolved in dioxane for 4 h to overnight. 4VP was then added, along with AIBN as a solid or in stock solution (10 mg mL-1 in dioxane, for < 4 mg), DMF, and balance dioxane.

The reactions were carried out in 20 mL septa-sealed vials and were degassed using N2 purging with 2-5 bubbles per second after which feed and outlet needles were removed. If reactions are not

50 working, there may be a leak, so the vials can be kept under constant N2 by leaving the feed line connected, but with the needle adjusted above the liquid level. Reactions were initiated by placing on heat at 75-80 °C in a heat block. After the prescribed reaction time, the reactions were terminated by rapidly cooling in ice and exposing to air. Reactions were purified by first precipitating in a 4:1 chilled mixture of hexanes and ether. After dissolving the polymer in DCM or methanol, the polymer was precipitated in ether twice more. Typically, a 10-fold or greater excess of the anti-solvent was used. The solvent was decanted off and the solid were rinsed with additional chilled ether, which was also decanted off. Decanting was preferred because significant losses had been seen by filtering especially with block copolymers containing lower fractions of

4VP. The resulting solids can be squeezed to remove additional solvent, but this actually made drying more difficult. The polymer was dried under flowing air for 1-2 h before being transferred into the vacuum oven where the polymer was dried at room temperature or 40-50 °C for 1-2 days until solvent was not detected by NMR (again heating is not advised for the BP containing

1 polymers, unless absolutely necessary). Monomer ratio and Mn were determined by H NMR by comparison of the δ 6.33 (m, 2 H) and 8.26 (m, 2 H) for 4VP and to the 3.31 (s, 3 H) peak for

OEGMA. Đ was determined by GPC. Table 2.2 contains all synthetic details and key characterization data for each polymer used. An example of the NMR and GPC analyses is provided in Figures 2.8 and 2.9.

Figure 2.7. RAFT synthesis of (POEGMA-r-BP)-b-P4VP. POEGMA-r-BP can be replaced with

POEGMA as well.

Table 2.2. Reaction conditions and data for POEGMA-b-P4VP and (POEGMA-r-BP)-b-P4VP.

Vol. Mass Mass Vol. Vol. Vol. Frac. CTA AIBN Dioxane DMF 4VP Temp. Time Mn P4VP MacroCTA (g) (mg) (mL) (µL) (mL) (°C) (h) (kDa) Đ (%) Ch. POEGMA22- POEGMA-r- 3.24 3.2 12.5 (g) 3.68 75 6 29.3 1.11 25.6 3 P4VP7 BP22 POEGMA37- P4VP11 POEGMA37 2 1.76 2.31 461.7 2.31 75 6 47.8 1.17 23.4 5

POEGMA15- POEGMA- 3 6.37 3.43 786.3 2.1 75 6 19.6 1.15 22.7 6 P4VP5 BP15 POEGMA36- POEGMA- 3 2.74 3.77 943.6 3.77 80 6 46.3 1.16 23.9 6 P4VP11 BP36 POEGMA55- POEGMA- 3 1.79 3.77 943.6 3.77 80 6 68.5 1.23 21.1 6 P4VP17 BP55

Figure 2.8. Sample 1H NMR of POEGMA-P4VP (POEGMA22-P4VP7 from Chapter 3).

Comparison of the I and J peaks to the D peak was used to determine the ratio of 4VP to OEGMA in the polymer.

Figure 2.9. Sample GPC of POEGMA-P4VP (POEGMA22-P4VP7 from Chapter 3)

2.1.7. (POEGMA-r-BP)-b-PDMAEMA Synthesis.

(POEGMA-r-BP)-b-PDMAEMA was synthesized via RAFT. POEGMA-r-BP was dissolved in dioxane in 40 mL septa-capped vials. DMAEMA and AIBN were added after dissolution, and the mixture was degassed via N2 purging for 25 min. Reactions were initiated by heating to 75 °C and terminated after a specified time by removing from heat and exposing to air.

Polymers were purified by precipitation in hexanes. Polymers were dissolved DCM and precipitated twice more in hexanes. Solvent was again decanted off, and the polymer was dried in a vacuum oven at room temperature for 1-2 days, with occasional stirring of the soft polymers to enable better drying. The polymer is soft and should not be filtered because it will stick to filter

54 paper and may go through it as well. The polymerization conditions and resulting polymer properties are given in Table 2.3. Final monomer ratios were determined by 1H NMR (Example in

Figure 2.11) and Đ was determined by GPC (Figure 2.12).

Figure 2.10. RAFT synthesis of (POEGMA-r-BP)-b-PDMAEMA.

Table 2.3. Reaction conditions and data for (POEGMA-r-BP)-b-PDMAEMA used in Chapter 6.

Mass Mass Vol. Vol. Vol. Frac. CTA AIBN Dioxane DMAEMA Temp. Time Mn PDMAEMA MacroCTA (g) (mg) (mL) (mL) (°C) (h) (kDa) Đ (%) POEGMA15- POEGMA- 6 3.18 11.27 5.64 75 4.5 20.5 1.11 23.5 PDMAEMA5 BP15 POEGMA36- POEGMA- 6 1.37 14.97 7.53 75 5.5 47.0 1.14 22.7 PDMAEMA13 BP36 POEGMA55- POEGMA- 6 0.896 14.97 7.53 75 7 71.6 1.18 22.4 PDMAEMA19 BP55

Figure 2.11. Sample 1H NMR of POEGMA-PDMAEMA (POEGMA15-PDMAEMA5).

Comparison of the K (6H) peak to the D (3H) peak was used to determine the ratio of DMAEMA to OEGMA in the polymer.

Figure 2.12. Sample GPC of POEGMA-PDMAEMA (POEGMA15-PDMAEMA5).

2.1.8. Quaternization of P4VP and PDMAEMA.

Full quaternization of the polymers was carried out using a 4x molar excess of methyl iodide compared to the monomer concentration of each polymer. Reactions were carried out in

DMF at 0.125 g mL-1 at room temperature overnight. The polymers were precipitated into ether twice to remove excess DMF and CH3I, polymer was re-dissolved in methanol between precipitations. Depending on block ratio, re-dissolving the polymer may be difficult in anything but water. In these cases, the solids can be thoroughly rinsed with ether, then dried under vacuum.

Both the solvents and the CH3I are sufficiently volatile to remove under vacuum and removal can be confirmed by 1H NMR. Alternatively, the polymers could also be dialyzed and lyophilized, enabling exchange of the anion from iodide to another if preferred. Quaternization was confirmed by 1H NMR (Figure 2.13 shows an example).

Partial quaternization of the POEGMA-P4VP block copolymers was also done with iodomethane in DMF. Polymers were dissolved in DMF at 0.125 g mL-1. Iodomethane was added in a 1:1 stoichiometric ratio with the target amount of the monomer to be quaternized and heated to 60 °C for 6 h. The polymer was then precipitated in diethyl ether twice and dried at room temperature or 60 °C for 18-24 h to remove residual DMF and ether. Conversion was measured by NMR and found to be >95%.

Figure 2.13. Sample 1H NMR of POEGMA-qP4VP (POEGMA22-qP4VP7 from Chapter 3). The total aromatic peaks are equal to the previously measured peaks. The total integral of the A (2H) and K (3H) peaks is expected to be 5 for complete quaternization of the polymer. The polymer was quaternized >95%.

2.1.9. RAFT Polymerization of Polyelectrolytes for Patchiness Study (Chapter 4).

These polymers were synthesized by Dr. Sieun Kim. The molecular weight, polydispersity

(Đ), and degree of polymerization (DP) of synthesized poly(acrylic acid) (PAA), quaternized poly(4-vinylpyidine) (qP4VP), poly(styrene sulfonate) (PSS), and poly(dimethylaminoethyl acrylate) (PDMAEA) were determined by GPC and are shown in Table 4.1 with traces shown in

Figure 2.15. HA was purchased from Lifecore Biomedical since HA is relatively difficult to obtain with narrow Đ (1H NMR provided in Figure 2.14a). Prior to polymerization, 4-vinylpyridine, acrylic acid and 2-dimethylaminoethyl acrylate were passed through a basic alumina column to remove hydroquinone or MEHQ inhibitor. The purified monomers were clear in color.

Polymerization of each of the monomers was performed by RAFT at 60-70 °C in a 200 mL Schlenk tube to obtain Đ < 1.4. Azobisisobutyronitrile (AIBN) was recrystallized in methanol to use as the radical initiator. Further details follow for each specific chemistry.

2.1.9.1. PAA polymerization.

2-(Dodecylthiocarbonothioylthio)-2-methylpropionic acid (72 mg, 0.2 mmol,

MilliporeSigma) and AIBN (6 mg, 0.037 mmol) were added to a solution of acrylic acid (10 g,

139 mmol) in water/DMF co-solvent (1:9 v/v) in the ratio of 700:1:0.2 (M:CTA:I). The solution was mixed until the CTA and radical initiator were completely dissolved in the solvent. Then, the mixture was degassed by three freeze-pump-thaw cycles in liquid nitrogen and purged with N2.

The polymerization was carried out in a sealed flask at 65 °C and terminated after 16 h by removal of heat and exposure to air. The polymer was purified by dialysis against water with a 3 kDa membrane for 7 cycles of 3 h each. The molar mass and dispersity of the obtained polymer were confirmed by aqueous GPC (Table 2.4, Figure 2.15). After confirmation of Mw, the polymer was stirred in 1 M NaOH for 3 hours to neutralize the polymer and convert it to the sodium salt of

PAA, dialyzed to exchange to water using the same procedure as before, and lyophilized. 1H NMR confirmed full deprotonation (Figure 2.14b).

2.1.9.2. qP4VP polymerization.

CPP (MilliporeSigma, 155 mg, 0.56 mmol) and AIBN (recrystallized twice from methanol,

18.2 mg, 0.11 mmol) were added to a solution of 4VP (10 g, 100 mmol) in 100 g 1,4-dioxane in the ratio of 500:1:0.2 (M:CTA:I). The solution was mixed in a 200 mL Schlenk reactor and then degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 60 °C and terminated after 12 h by removal of heat and exposure to air. The polymer was then precipitated three times in hexanes and dried under vacuum. The molar mass of the obtained polymer was confirmed by DMF GPC (Table 2.4, Figure 2.15). P4VP was quaternized with iodomethane (99%, MilliporeSigma, 13 mL) in N,N-dimethylformamide (DMF).

The reaction mixture was stirred at room temperature for 24 h, purified by precipitation in diethyl ether, and dried under vacuum. The degree of quaternization was 100% as determined by 1H NMR

(Figure 2.14c).

2.1.9.3. PDMAEA polymerization.

For PDMAEA polymerization, 2-ethylsulfanylthiocarbonyl-sulfanyl-2-methylpropionic acid (EMP) was used as the CTA and was synthesized following a previously reported procedure

(See Appendix B).3 EMP (31 mg, 0.14 mmol) and AIBN (4.6 mg, 0.028 mmol) were added to a solution of DMAEA (10 g, 70 mmol) in 100 mL 1,4-dioxane in the ratio of 500:1:0.2 (M:CTA:I).

The solution was degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 70 °C and terminated after 16 h by removal of heat and exposure to air. The polymer was then precipitated 3 times in hexanes and dried under

60 vacuum. Polymer molar mass was confirmed by GPC (Table 2.4, Figure 2.15). 1H NMR in acidified conditions provided in Figure 2.14e.

2.1.9.4. PSS polymerization.

4-Cyano-4-(phenylcarbonothioylthio) pentanoic acid (22.5 mg, 0.08 mmol) and AIBN (2 mg, 0.016 mmol) were mixed in 10 mL of DMF. Sodium 4-vinylbenzenesulfonate (10 g, 0.048 mmol) was dissolved in 90 mL of 1 M NaCl solution. CTA and initiator solution were mixed into the monomer solution in a 200 mL Schlenk reactor. Monomer:CTA:initiator were in the ratio of

600:1:0.2. The solution was degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 60 °C under N2 gas and terminated after 16 h by removal of heat and exposure to air. The polymer was then dialyzed against water for 7 cycles of 3 h each to get rid of monomer and dried under vacuum. Polymer molar mass was confirmed by GPC (Table 2.4, Figure 2.15). 1H NMR provided in Figure 2.14d.

Table 2.4. Summary of Polymer Properties for Chapter 4.

Polymers Mw (kDa) DP Đ Expected # of charge HA 190 473 1.20 473 PAA 33 454 1.34 454 qP4VP 64 411 1.07 411 PSS 70 340 1.07 340 PDMAEA 25 173 1.36 29

1 Figure 2.14. H NMR (500 MHz) in D2O of a) hyaluronic acid, b) sodium salt of PAA, c) qP4VP

(peak at 4.25 ppm confirms full quaternization, approx.. 5% (wt./wt.) residual DMF), d) PSS, and e) PDMAEA (C′, D′, E′ correspond non-protonated PDMAEA, approx. 5%).

Figure 2.15. Gel permeation chromatography of synthesized charged polymers; PAA, qP4VP,

PSS and PDMAEA.

2.1.9.5. Determination of pKa of PDMAEA.

PDMAEA did not have a reliable published pKa-value, so the polymer was titrated to

1 determine the pKa. A solution of PDMAEA in water (25 mL at 2 mg mL ) was acidified with 6 M

HCl to below the first equivalence point (in this case, pH 2.56). The solution was then titrated with

50 mM NaOH while pH was monitored with a Mettler Toledo pH probe. The pKa was found to be

7.32 (Figure 2.16).

-1 Figure 2.16. Titration data for PDMAEA. pKa was found to be 7.32 ± 0.05. 25 mL of 2 mg mL polymer was acidified to pH 2.56 and titrated with 50 mM NaOH.

2.2. Protein Preparation.

2.2.1. Alkaline Phosphatase Cloning.

This work was completed by Dr. Allie Obermeyer. The gene for PhoA in the pTrc99a plasmid was obtained as a generous gift from Dr. Jeff Glasgow. The QuikChange II Site Directed

Mutagenesis Kit (Agilent) was used to mutate the pTrc99a-PhoA plasmid to generate 5 mutants of alkaline phosphatase: M1) K92D + K96D, M2) K92D + K93D + K96D, M3) K352D + K353D,

M4) K92D + K96D + K352D + K353D, and M5) K92D + K93D + K96D + K352D +K353D. The plasmid for native PhoA was isolated from an overnight culture of TunerTM cells grown in lysogeny broth (LB) with ampicillin (0.2 mg mL-1) using a DNA mini-prep kit from Qiagen. The supplied protocol for the QuikChange kit was then followed.4 In brief, primers were designed with the aid of the primer design tool available online from Agilent (www.agilent.com/genomics/qcpd)

66 and purified primers were ordered from Integrated DNA Technologies, Inc. The plasmid was amplified by PCR in the presence of one of the designed primer pairs using the reaction conditions described in the protocol, after which the reactions were cooled to < 37 °C on ice. To each reaction,

1 µL of the provided Dpn I restriction enzyme was added and allowed to react for 1 h at 37 °C to degrade the parental DNA. This DNA could then be transformed into the provided XL1-Blue supercompetent cells (stored at -80 °C). The XL1-Blue cells were then thawed on ice and used immediately after thawing. As the cells were thawing, sterile 14 mL BD Falcon round bottom polypropylene tubes we pre-chilled on ice. The use of these tubes is necessary because the heat- shock step has been optimized for these tubes, and transformation may fail if they are not used.

After cells thawed, 50 µL of cells were aliquoted into each tube and 1 µL of the appropriate PCR product was added to each tube, pipetting up and down several times to ensure all DNA had been dispensed. Tubes were gently swirled or flicked to mix, then incubated on ice for 30 min. The tubes were then heat shocked for 45 s at 42 °C, then returned to ice for 2 min. To each tube, 0.5 mL of pre-warmed NZY+ broth (provided) was added to each tube and the cells were incubated at

37 °C for 1 h with 225 rpm shaking. Afterwards, 250 µL of the reactions were placed on LB-agar plates containing 0.2 mg mL-1 of ampicillin and spread using sterile glass beads. The plates were incubated at 37 °C for 16 h, then wrapped with parafilm and stored at 4°C until use. 5-8 colonies from each of the plates were selected for sequencing if the plates were not overgrown and the colonies were not in contact with other colonies. The colonies were picked up with a sterile P200 pipette tip and transferred to a sterile culture tube containing 5 mL LB and ampicillin (0.2 mg mL-

1), and grown overnight at 37 °C in the shaking incubator. DNA was extracted from these cultures by mini-prep and sequenced to ensure the correct mutations were present. DNA was stored at -20

°C until use. The gene sequence and primer sequences are shown in Figures A12 and A13,

67 respectively. The optimal PhoA mutant was found to be Mutant 4 based on activity and yield

(Chapter 3). This mutant was used for all subsequent studies.

2.2.2. Alkaline Phosphatase Expression and Purification.

Engineered genes were transformed into TunerTM cells following a standard protocol from

Novagen. A 5 mL overnight culture of the cells was prepared in LB supplemented with 0.2 mg mL-1 ampicillin and grown at 37 °C in a standard culture tube. The overnight cultures were added to 1 L of 2xYT media supplemented with 0.2 mg mL-1 ampicillin and grown at 37 °C. When the optical density at 600 nm reached 0.4, the cultures were induced with 1 mM IPTG, 1 mM MgSO4, and 0.1 mM ZnSO4, and allowed to grow overnight at 37 °C.

The cells were pelleted by centrifugation at 3,500 x G for 10 min and resuspended in 50 mM Tris buffer pH 8.0 after each centrifugation. The periplasm was isolated by adding solid sucrose (0.5 M), EDTA (2.5 mM), and lysozyme (0.6 mg mL-1) to the cells and incubating at 37

°C for 30 min. Sucrose was often difficult to dissolve, so several inversions were necessary; when not dissolved a clear or white layer was seen at the bottom of the bottle. EDTA was added from a

250 mM stock solution in water that was adjusted to pH 8.0 prior to the procedure. The sphaeroplasts were removed by centrifuging at 10,000 x G for 20 min. The resulting supernatant was the periplasmic fraction. The periplasmic fraction was precipitated by heating the solution for

10 min in an 80 °C water bath. The mixture was centrifuged at 24,000 x G to remove precipitates.

The supernatant was dialyzed against 20 mM Tris pH 8.0, and then purified via anion exchange chromatography using 2 5 mL Q columns (HiTrap Q HP, GE Healthcare) in series equilibrated with 20 mM Tris pH 8.0 and eluted with a 0-0.5 M NaCl gradient using the GE Akta FPLC.

Fractions containing PhoA were identified by UV absorbance at 280 nm and SDS-PAGE analysis.

Figure 2.17 provides an example of the PhoA M4 elution profile from FPLC and corresponding

SDS-PAGE analysis. Fractions containing PhoA were pooled and dialyzed against 20 mM Tris pH 8.0 with 1 mM MgSO4 and 0.1 mM ZnSO4 and then stored at 4 °C. Typical yield of PhoA M4 was approximately 40 mg L-1.

Figure 2.17. a) FPLC chromatogram and b) paired SDS-PAGE analysis for the purification of

PhoA M4. The red boxed regions correspond with one another.

2.2.3. Preparation of Amylase Supercharged Variants.

The α-amylase was purchased from MilliporeSigma. SDS-PAGE showed that the native protein was pure (Figure 2.18), but the protein made up only ~10% of the dry mass of the powder per A280 measurement. The small molecule contaminants were removed by ultrafiltration with a

15 mL 10 kDa MWCO Amicon Ultra centrifugal filtration unit. The protein was dissolved in phosphate buffer pH 8.0, spun down to 1.5 mL or less, and diluted by addition of buffer up to 15 mL (≥ 10x dilution). This was repeated 6 times, and the solution went from dark brown to near

69 clear with a yellow-brown tint. To reach a volume of 1.5 mL took approximately 15 min at 4000 g at 4 °C. Protein loss was ~10%. After purification, the supercharged α-amylase variants were made by reaction with succinic anhydride (SA). The previously purified native amylase was concentrated to 5 mg mL-1 in 10 mM phosphate buffer using 15 mL 10 kDa NMWL Amicon Ultra centrifugal filters. Three parallel reactions using 20 mL of the solution in 50 mL conical tubes were set up with 20, 50, and 200 molar equivalents of SA per unit protein by directly adding solid

SA to the tubes and allowing to stir at room temperature for 16-20 h, resulting in 3 variants named

A30, A32, and A37, respectively. A fourth reaction with 1000 molar equivalent resulted in the protein precipitating. Excess SA was removed via ultrafiltration with a 15 mL 10 kDa NMWL

Amicon Ultra centrifugal filters with 6 10-fold dilutions. The degree of supercharging was determined by MALDI-TOF (Figure 2.19, Table 2.5).

Figure 2.18. SDS-PAGE of purified native α-amylase.

Figure 2.19. MALDI-TOF of native and supercharged α-amylase.

Table 2.5. Summary and analysis of MALDI-TOF data for native and supercharged α-amylase.

Mean Modification Variant MW # Charge Alpha m/z- A22 54521 0 -22 1.77 2478 A30 54936 4.15 -30.3 2.20 1814 A32 55036 5.15 -32.3 2.33 1704 A37 55261 7.40 -36.8 2.64 1502 2.2.4. Expression and Purification of GFP Mutants.

The DNA and protein sequences for the GFP mutants are shown in Figures B1 and 4.1, respectively (PDB ID: 2B3P). All proteins were expressed in BL21(DE3) cells, which were transformed with plasmids encoding the GFP mutants. For each variant, a well-isolated colony was grown overnight in LB broth containing kanamycin at 37 °C. 5 mL of the overnight culture was diluted in 1 L of LB broth and incubated at 37 °C until OD600 0.8-1. Cultures were then induced with 1 mM IPTG, grown for 20 h at 37 °C, and harvested. Cells were collected by centrifugation at 4000x g for 30 min, and the supernatant was decanted. The collected wet cells were resuspended in lysis buffer (50 mM KH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0) and then sonicated in an ice bath for 10 min for two cycles. The resulting cell debris and the supernatant

71 were separated by centrifugation at 12000 rpm at 4 °C for 60 min. The presence of the expressed protein in the supernatant was confirmed by SDS-PAGE.

The crude proteins in supernatant were further purified using Ni-NTA metal affinity chromatography with 10 column volumes of wash buffer (50 mM KH2PO4, 300 mM NaCl, 20 mM imidazole, pH 8.0) to get rid of impurities. Using 3 column volumes of elution buffer (50 mM

KH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8.0), pure protein was obtained. The purified proteins were dialyzed into 20 mM Tris, 0-300 mM NaCl, pH 8.0 7 times with exchanges every 3 hours. The purified proteins were analyzed by SDS-PAGE, and molar mass was confirmed by

MALDI-TOF. If the proteins were not sufficiently purified, the Ni-NTA purification was repeated.

It was found that the purification of the GFP variants by Ni-NTA chromatography could be improved by the use of less resin to reduce off-target binding directly to the resin, the use of a reducing agent (5 mM β-mercaptoethanol) to prevent impurities from being carried through bound to the free cysteine on the GFP variants, and 1 M NaCl to screen charge interactions and prevent impurities that were electrostatically bound to the proteins from carrying through. The changes were especially important when trying to obtain high yields of the XLP variant. Further increases in salt concentration were not tested, but the QIAexpressionistTM handbook suggests that concentrations of up to 2 M may be used and may be beneficial.5 Additional additives such as glycerol were not tested, but may also provide some improvement of purification.

After purification, purity was confirmed by SDS-PAGE and the secondary structure of the mutants was confirmed by circular dichroism (CD) and UV fluorescence of purified GFP variants

(Figure 2.20). All CD spectroscopy was carried out on an Aviv model 202 CD spectrometer.

Proteins were measured in 20 mM Tris-Cl pH 8.0; samples were prepared at a concentration of 0.2

72 mg/mL and filtered using 0.2 µm syringe filters prior to measurement. Measurements were acquired in a 0.1 cm path length of the quartz cuvette at a scan rate of 6 nm/min.

Figure 2.20. Properties of GFP mutants. (a) MALDI TOF of GFP variants (SP: 28.5 kDa, MP:

27.8 kDa, LP: 28.8 kDa, and XLP: 28.7kDa), (b) stained SDS-PAGE gel, (c) Circular dichroism spectra of GFP variants, and (d) UV-vis spectra of mutants at 1 mg mL-1, with inset image of UV fluorescence of purified GFP variants (2 mg mL-1).

2.2.5. Nitroreductase Expression and Purification.

Plasmid DNA for a dimeric fusion of NfsB from E. coli, with mutations at residues 89 and

315 to p-azidophenylalanine (AzF) and cysteine, respectively, was provided by Neil Marsh

(University of Michigan). The plasmid was co-transformed with the pDule2 pCNF RS plasmid

(from Ryan Mehl, Oregon State University) into BL21 (DE3) E. coli cells. Overnight cultures were made in 50 mL LB supplemented with 40 mM spectinomycin and 50 mM kanamycin in a 250 mL baffled flask. The protein was expressed in 5 L LB supplemented with 40 mM spectinomycin, 50 mM kanamycin, and 0.7 mL of antifoaming agent (Antifoam 204, MilliporeSigma) in a bioreactor with an air flowrate of 5 L min-1. The bioreactor was equilibrated to 37 °C, and the overnight culture was added. When the optical density at 600 nm reached 0.6, 1 g AzF, dissolved in water by addition of NaOH, was added and the temperature was set to 18 °C. After 30 min, the culture was induced with 1 mM IPTG, and the expression was allowed to continue for 18-22 h. Cells were centrifuged down from the media at 4 °C at 4000 ×G for 15 min, and frozen at -80 °C and stored overnight or until needed.

Cells were thawed and resuspended in lysis buffer (~2-3 mL/g), after which 1 mg mL-1 of lysozyme (from chicken egg white, MilliporeSigma) was added and allowed to react for 30 min at

4 °C. The cells were then sonicated and the lysate was clarified by centrifugation at 10,000 ×G for

30 min at 4 °C. NfsB was then purified by metal affinity chromatography with Ni-NTA. The protein from one 5 L expression was allowed to bind to 6 mL of the resin overnight at 4 °C. The resin was then washed 4 times with 5 CVs of wash buffer containing 10% glycerol and then eluted with 5 CVs of elution buffer collected in 1 CV aliquots (Figure 2.21a). The protein was then dialyzed with 5 buffer changes into 50 mM phosphate buffer pH 8.0 with 100 mM NaCl. Purified enzyme was frozen in liquid N2 and stored at -80 °C. Before use, proteins were thawed at 4 °C overnight and stored at 4 °C for no longer than 5 days. Figure 2.21b shows an SDS-PAGE analysis of the purified enzyme.

Figure 2.21. a) SDS-PAGE analysis of Ni-NTA chromatography fractions. Elutions were pooled and dialyzed. b) SDS-PAGE analysis of dialyzed protein.

2.3. Thin film preparation.

2.3.1. PEG-Modification of Si surface.

In order to allow for adsorption of the POEGMA block to the Si surface, the surface had to be functionalized. A silicon wafer (WaferWorld, (100) orientation) was cleaned with acetone, methanol, and milliQ water then dried with filtered air from an air gun. The wafer was not allowed to dry between washing steps. The wafer was then cleaned with air plasma in a plasma cleaner

(Harrick) for 2-3 min. The smooth, shiny side of the wafer was then coated in a 0.25-0.5 cm layer of monomethyl ether polyethylene glycol (PEG, Mn = 750), and heated to 150 °C under vacuum for 18 h in a crystallization dish with diameter approximately 1 cm greater than the wafer. After removing from the oven and allowing to cool, acetone was added to the dish to allow the PEG to remain liquid. The wafer was stored in the crystallization dish for short term use (within 1 week),

75 or the wafer was cut using a diamond or SiC cutter and stored with the acetone-PEG mixture in a jar. When the wafers were ready for use, the wafers were quickly rinsed with acetone, methanol, and water and dried, then cut into rectangular shards for coating. The shards were then carefully individually cleaned with acetone, methanol, and water with care taken to remove any residual solvents with each subsequent wash. The shards were then dried with filtered air and stored in a fresh petri dish for no more than 1-2 days until coating.

2.3.2. Flow coating.

Flow coating was done on a system similar to that described by Stafford et al.6 Solutions were prepared at 15-20% (wt./wt.) in milliQ water, and diluted as necessary. To start, a standard microscope slide with no chips was cleaned with acetone, methanol, and water then dried with filtered air. The slide was attached to the angled, stationary upper platform with tape such that the long edge of the slide was flush with the far edge of the platform and the short edge was in line with the marking on the platform. The addition of a guiding groove would greatly improve this process. After the blade was in place, the chamber around the coater was adjusted to the right RH.

The make-shift humidity chamber around the flow coater was adjusted to the desired relative humidity using a number of methods depending on the season and weather. If the room was above the desired humidity (never occurred), the chamber could be dehumidified using the lab’s dehumidifier and tubing to connect to the chamber. If the room was at the desired humidity, the chamber was simply removed. If the room was slightly below the desired humidity (10-30%), a container(s) of water with a wet absorbent pad could be placed inside the chamber to reach the desired humidity passively. If the building heat was running on high, the humidity in the room could be 50-60% away from the target. If possible coating was avoided on these days, but if necessary, the room can be gently humidified with the cool, passive humidifier and the USB-

76 powered humidifier can be activated within the chamber. Care should be taken to minimize and wipe up condensation, and the humidifier should be turned off at least 2-3 min before coating to avoid any condensation or droplets on the films.

Once desired humidity was achieved, a rectangular shard of Si wafer (or another desired substrate) was taped such that it was centered on the alignment marking on the moving lower platform with the upper platform raised out of the way. The moving platform was then moved such that blade assembly covered the substrate and the edge of the blade was above the substrate and about 3-5 mm away from the tape. The blade platform angle was then adjusted to its lowest point

(making sure to avoid contacting the substrate and adjust the z-position up if necessary). The height of the blade was then adjusted to be 1-2 mm above the substrate (sufficiently wide for the liquid coating solution to be wicked in but not so wide that the solution just pools on the substrate), after which 2-4 µL of the coating solution was pipetted with the tip touching the gap between the substrate and the blade. Liquid should not be fully dispensed to avoid the formation of a bubble between the blade and the substrate. If a bubble forms oscillate the z-position or the angle of the blade to try to pop it (without breaking the capillary force between the blade and substrate). The z-position and the angle of the blade perpendicular to the angle of coating should now be adjusted.

The angle should be adjusted first. Lower the z-position of the blade such that the liquid is spread across the width of the substrate, then raise it until a droplet forms, and repeat several times. If the droplet is not centered or the liquid front when the blade is lowered is not parallel with the short edge of the blade, the angle needs to be adjusted using the appropriate knob. Adjust and repeat testing until the blade is flattened, then adjust the z-position of the blade so that an even 2-5 mm liquid front forms. This will need to be optimized by material and solution concentration as

(counterintuitively) smaller gaps lead to thicker films (to a point, after which nothing comes out).

Once everything is positioned, coating can start. If there was no prior knowledge, as a rule of thumb, the coating speed was initially tested at approximately 10 rpm for approximately 0.5 cm. Film color was used to determine if the desired thickness was achieved, with brown indicating

<50 nm thickness, dark blue 75-125 nm, light blue 125-150 nm, yellow 150-200 nm, and purple

>200 nm. Exact thicknesses were determined later by AFM. To start the coating, the substrate was first moved 2-5 mm to move off of the spot where adjustments were made, then a 0.5-1 cm patch was coated to determine if coating conditions were correct. If the films were too thick, slowing the sped by roughly the same factor by which the films were too thick worked well as a rule of thumb.

The z-position could also be adjusted up. If the films were too thick at a speed of 1-2 rpm, the solutions were diluted slightly (2-3 wt. %), and coating was attempted again. Coating conditions were highly non-linear with dilution, so if possible solutions that were found to work were not adjusted. After conditions were determined the desired number of test films were cast. Between each coating, excess coating solution was removed (while still wet) with a wet lab wipe, then the blade was cleaned with 70% ethanol or isopropanol twice and dried with a lab wipe and a lens wipe. If the blade could not be wiped clean or was damaged, it was replaced.

2.4. UV-Vis Absorption Methods.

2.4.1. Turbidimetry

Turbidimetry was used to determine whether pairs of polymers and/or proteins complexed and to what extent they complexed. The turbidity is defined as:

Turbidity = 100% − %T600 (2.1) where %T600 is the percent of light transmitted by the sample at 600 nm. This can be calculated from the absorbance at 600 nm as follows:

%T = 102−abs (2.2)

78 where the absorbance should be corrected for path-length and the blank. The turbidity measurements were taken on 200 µL samples in a 96-well plate in the TECAN Infinite M200Pro plate reader.

2.4.2. Preparation of Standard Curve of 4-Methylumbelliferone (4MU).

Standard curves were prepared for both the 96-well and 12-well (Figure 2.22) plates using

4-MU (Sigma-Aldrich) in the TECAN Infinite M200Pro plate reader. Individual 10 mM stock solutions of 4-MU were made in DMSO and diluted in water to 1 mM. For the 96-well plate standard, separate 1 mM stock solutions were further diluted to 500, 300, 100, 75, 50, 25, and 10

µM in water. 180 µL of 100 mM Tris pH 8.0 was added to 24 wells. To 8 wells, 20 µL of 500,

300, 100, 75, 50, 25, 10, and 0 µM 4-MU was added. The emission at 448 nm was measured after exciting at 362 nm. 20 µL of the wells’ contents were then added to the next well, providing a ten- fold dilution, and the excitation was measured again. This was repeated once more. For the 12- well plate, the 1 mM stock was diluted to 750, 500, and 250 µM. Data was collected at final concentrations of 100, 75, 50, 25, 10, 7.5, 5, 2.5, 1, 0.5, 0.25, and 0 µM and fitted to a line. High concentrations were excluded due to non-linearity and because experiments never reached that level of conversion.

Figure 2.22. a) Standard curve in 96-well plates in MOPS buffer. b) Standard curve in 96-well plate in Tris buffer. c) Standard curve in 12-well plate in Tris buffer.

2.4.3. Bulk Activity assays.

2.4.3.1. PhoA assay.

PhoA activity was measured fluorimetrically on a TECAN Infinite M200Pro plate reader.

4-methylumbelliferyl phosphate (4-MUP) (Aldrich) was converted to 4-methumbelliferone (4-

MU), which was excited at 362 nm and measured at 448 nm. 4-MUP was dissolved in dimethyl sulfoxide at 10 mM and diluted in water to the concentrations for the assays.

Assays on the enzyme and micelles were conducted in 96-well plates (Brand, #781602).

The mutants were compared in 100 mM 3-(N-morpholino)propanesulfonic acid (MOPS) buffer, pH 7.5, supplemented with 500 mM NaCl. The mutants were diluted to 0.5 µg of enzyme L-1 (0.5 ng mL-1) in MOPS buffer and stored on ice until use. 160 µL of MOPS buffer and 20 µL of substrate solution were added to each well. After taking an initial reading, 20 µL of diluted enzyme was injected into each well using the auto-injector on the spectrometer. The emission was measured for 2 min at 448 nm. The slope in terms of a.u. s-1 was converted to µM s-1 by dividing by the slope of the 4-MU standard curve (1376 a.u. mM-1).

Due to the high salinity of the MOPS buffer, the comparison of free enzyme activity to the micelle activity was conducted in 100 mM Tris at pH 8.0. A standard curve was prepared in this buffer. Micelles were prepared by adding an equivalent mass of POEGMA-b-qP4VP as protein in solution. The micelles and free enzyme were then diluted to 0.5 µg of enzyme L-1 in Tris buffer.

160 µL of Tris buffer and 20 µL of diluted enzyme or micelles were added to each well. After taking an initial reading, 20 µL of substrate solution was added to each well. The emission was measured for 2 min. The slope in terms of a.u. s-1 was converted to µM s-1 by dividing by the slope of the 4-MU standard curve (2338 a.u. mM-1).

2.4.3.2. NfsB Assay.

The activity of the NfsB was measured using an assay adapted from Schroeder, et al.7

Activity was measured in 100 mM NaPO4 buffer pH 8 for both the free enzymes and the films at

-1 -1 0.5 mM NADH. Oxidation of NADH (ε340 = 6220 M cm ) was monitored by plate reader. For all reactions, control samples with no enzymes/films were prepared in quadruplicate to monitor the basal conversion rate.

For the assay of free NfsB, assays were conducted with 0, 50, 100, 250, 500, 750, and 1000 mM 4-nitrobenzenesulfonamide (4-NBS). 4-NBS stock solution was prepared at 100 mM in

DMSO and diluted to 10x the assay concentrations with DMSO added such that all stock solutions had 1% DMSO (vol./vol.) to avoid solvatometric effects. NADH stock solution was prepared at

10 mM in 100 mM Tris pH 8. Assays were conducted at a volume of 200 µL in 96-well plates with 20 µL of 4-NBS solution, 10 µL of NADH solution, and 165 µL buffer with 5 µL of enzyme solution. For the native NfsB the enzyme stock solution was 0.5 µM and for the SC NfsB was 1 mg mL-1. For native NfsB the enzyme was added and the reaction was monitored for 2 min. For

SC NfsB the reaction was allowed to proceed for 2 h at room temperature under shaking during which the plate was sealed with an AeraSealTM film and the plate lid. The final conversion was then measured by plate reader. The thermally denatured protein was stored at 70 °C for 12 hours, the same condition used to screen the thermal stability of the film samples. The free enzyme reactions were run with 5 repeats.

2.4.4. Thin Film Activity and Metal Sensing Assays.

Assays on the films were conducted in 12-well plates (Falcon, #353043). Film areas were determined using photos of films on top of a calibration grid allowing for area determination with

ImageJ, an open-source image analysis software. Length scales was first calibrated by averaging

82 the number of pixels in 3 separate 6 cm lines and entering the conversion factor into ImageJ. The borders of the films were then traced using the polygon tool in ImageJ and ImageJ gave final areas in cm2.

2.4.4.1. PhoA Thin Film Assays.

Films were swollen overnight in 20 mM Tris pH 8.0 with 1 mM MgSO4 and 0.1 mM

ZnSO4. Prior to the assay, films were rinsed with water. 900 µL of 100 mM Tris pH 8.0 was then added to each well. After an initial background measurement, 100 µL of the substrate solution was added. The mixture was pipetted up and down a few times to ensure mixing. The 4-MU emission was then measured for 4 min. The activity data were converted from a.u. s-1 to µM s-1 by dividing rate by the slope of the calibration curve (530 a.u. µM-1). The resulting conversion rates were divided by the area of the film to give the final rates in terms of µM s-1 cm-2.

2.4.4.2. Metals sensing with PhoA thin films.

For metals sensing, the films were first assayed as described above with a final 4-MUP concentration of 100 µM, then were rinsed thoroughly with water (2-3 rinses with 2+ mL), and exposed to 100 mM EDTA in 100 mM Tris pH 8.0 for 25-30 min to convert the enzyme into apo- enzyme. Films were again rinsed thoroughly with water, and activity was measured again to ensure inactivation. After another water rinse, films were exposed to water containing the specified concentration of analyte and 1 mM MgSO4 for 20-25 min unless otherwise specified. Films were rinsed and assayed to determine the level of reactivation.

2.4.4.3. NfsB Thin Film Assays.

For the films assays, film samples of 0.3-0.5 cm2 were placed in 12-well plates. For each film condition, 4 samples were tested. Prior to the assay, the films were allowed to rehydrate in buffer (50 mM NaPO4 buffer pH 7.6 with 100 mM NaCl) overnight at 4 °C. A reaction mixture

83 with 1 mM 4-NBS and 0.5 mM NADH in 100 mM NaPO4 buffer was made using a 100 mM 4-

NBS stock in DMSO and a 10 mM stock of NADH in 100 mM Tris buffer pH 8. 1 mL of the reaction mixture was added to each well and the reactions were allowed to proceed for 30 min for the native NfsB and 12 h for SC NfsB under shaking. For the SC NfsB assays, the plates were sealed with both an AeraSealTM film and the plate lid to minimize evaporation. After the prescribed reaction time, 100 µL of the reaction mixture was transferred to a 96-well plate and the final conversion was measured.

2.4.4.4. Assessing Thermal Stability of Thin Films.

The thermal stability of the films was assessed by placing crosslinked films into an environmental chamber set to the prescribed conditions and stored under these conditions for the prescribed times. The films were then removed, chopped, and rehydrated as described in each of the film assays. For PhoA, the films were removed and rehydrated, then assayed the next day. For the extended aging assay of the NfsB films, the films were removed and stored dry at 4 °C until all time points were completed, then run simultaneously.

2.5. Protein Surface Charge Analysis

2.5.1. Design of GFP Mutants.

This work was completed by Dr. Sieun Kim. To create a series of GFP mutants having different patch sizes, the different amino acids within the protein that have previously been mutated successfully and are highly solvent exposed were determined based on analysis of super-charged

GFP proteins.8-9 To design the mutants, a base structure (.PDB) file for the protein was loaded into

PyMOL. These files are available on the Olsen lab server for some of the proteins the group has worked with previously or via the RCSB Protein Data Bank (www.rcsb.org).10 Note that for structures downloaded from PDB, several contain pairs of the proteins, and repeats should be

84 deleted before proceeding (with the exception of native multimers which should be kept intact).

For example, the structure for PhoA (PDB ID: 3TG0) contains a pair of the homodimers, so one of the two homodimers should be removed. If working with an existing plasmid, line up the

FASTA sequence of the protein structure with the plasmid sequence to avoid confusion about the positions of mutations. Mutation sites should be chosen based on literature if possible, as was done here, or to try to avoid disrupting structural and functional features of the protein such as β-sheets and the catalytic pocket.

After mutation sites have been selected and lined up with the FASTA sequence, the mutations can be made in PyMOL using the Mutagenesis wizard. Unfortunately, the wizard seems to require the user to click on the desired residue to modify it. To make this easier, change the color of the entire protein to white (or a color of your choosing), then use the commands “sele resi

##” (## should be replaced with the position of the amino acid in the FASTA sequence to be mutated) and “color red, sele” to change the color of the selected mutagenesis residue site to red, making it easy to identify and click on, and if mutating multiple sites, consider labeling them several different colors to ease the process. With the wizard open, click on the now identifiable target residue and in the mutagenesis panel click on the first row “No Mutation” and select the desired mutation. This will prompt PyMOL to generate several potential rotamers along with a prediction of the likelihood of each. The most likely rotamer was used for these calculations and should likely be used for all mutants in the absence of more data. After selecting the rotamer, click

“Apply” and “Done” to complete the mutation. If making multiple mutations, “apply” can be clicked after each and “done” after all mutations have been made to avoid having to re-open the wizard several times. After making all mutations, save the new structure as a new .pdb file with

85 the name or nickname of the mutant. In case of issues, the PyMOLWiki is an excellent resource available at: https://www.pymolwiki.org/index.php/Main_Page.

After mutating the amino acids in silico, the protein surface potential was visualized by using the Adaptive Poisson-Boltzmann Solver (APBS).11 The APBS method is a continuum model used to describe electrostatic interactions between solutes in aqueous salt solutions. After mutation of amino acids, PDB files of the mutated GFP proteins (XLP, LP, MP and SP) were generated by

PyMOL. The solver can be applied either on a personal machine or via the webserver at http://server.poissonboltzmann.org/, and for this work the webserver was used. The PDB files were first loaded into the PDB2PQR tool available at the webpage to generate a PQR file that stores data about the predicted atomic radii and charges within the protein as function of the pH and the chosen force field. For this work, the PARSE force field was used at pH 8. After the calculation is completed, the webserver allows users to adjust parameters such as the ionic strength and dielectric constant of the solvent prior to initiating the calculation of the electrostatic potential via the APBS tool. The data are output as a compressed .DZ file that contains electrostatic potential data over a

3D matrix. The file should be decompressed with 7zip or WinZip and saved with the same file name as the .PDB file.

2.5.2. Generating the Solvent Accessible Surface (SAS) and Surface Potential Map

The solvent accessible surface (SAS) of the proteins can be calculated using a Euclidean distance transform algorithm, EDTSurf, that uses the rolling-ball method to calculate the SAS of a given protein, and provides the vertices of a tessellated SAS for the proteins.12-13 The EDTSurf program is available at https://zhanglab.ccmb.med.umich.edu/EDTSurf/ as both source code and

Windows, Linux, and Apple executables. To use, place the executable and the .PDB files in the same folder, then open a command window. Type “cd ” then paste in the address of the folder and

86 hit enter to change the directory. Then type “EDTSurf.exe -i filename.pdb -s 2” where “filename” would be replaced with the name of the mutant and hit enter to run the EDTSurf program. This will generate a .PLY file containing the vertices of the SAS with the same file name as the .PDB file.

Using the code available in appendix B or the .M files available on the lab server at

\\olsenlab-old.mit.edu\Lab Member Files\Former Lab Members\Sieun Kim\GFP matlab, the SAS was then overlaid on the APBS potential map from the .DZ file to give the surface potential map,

[Q], showing the potential, q, as a function of position in 3 dimensions ([Q] = [푥̅ 푦̅ 푧̅ 푞̅]).

2.5.3. Search algorithm for patches.

The [Q] matrices were then used to calculate where charge patches occurred at the protein surface. All points in [Q] were first binned into positive (q ≥ 2 kT e-1; [Q+]), negative (q ≤ -2 kT e-1; [Q−]), and neutral matrices (-2 kT e-1 < q < 2 kT e-1; [Q0]). An arbitrary point (the first in the matrix) was chosen from [Q±] , and all its nearest neighbors within [Q±] were found (within 2 Å),

± ± removed from [Q ] , and placed in the matrix [Pi ]. The nearest neighbors of each subsequent

± ± ± ± point in [Pi ] within [Q ] were then deleted from [Q ], and added to [Pi ] until no further points were found in [Q±]. This process was repeated using the first remaining point in [Q±], until no

± ± points remained in [Q ]. For [Pi ] to be considered a patch, it had to have an area greater than 1 nm2. The area of the patches was approximated by assuming the points generated by EDTsurf were uniformly distributed and dividing the SAS by the number of points and multiplying the number of points in pi by this average area per point. After the positive and negative patches were found, the neutral area was calculated by subtracting the total area of the patches from the total area of the SAS. MATLAB code provided in Appendix B.

2.5.4. Algorithm for determining protein patchiness parameter.

For all points in [Q], all points within a radius of 2 Å were found, and each assigned a 1,

0, or -1 for the interaction between the two points. On average, each point had 6 neighbors within this radius, indicating that only the nearest neighbors were identified. A value of 1 was assigned if the potentials at the two points were both either greater than or equal 2 kT e-1 or less than or equal to -2 kT e-1, a value of 0 was assigned if the potential at either or both points was between the -2 and 2 kT e-1, and a value of -1 was assigned if the one point was greater than or equal to 2 kT e-1 and the other was less than or equal to -2 kT e-1. These values were summed and divided by the total number of interactions over the entire surface of the protein to give the patchiness correlation.

Under this algorithm, a completely positively or negatively charged surface would give a value of

1, a completely neutral surface or a perfectly alternating charge-neutral surface would give a value of 0, and a perfectly alternating positive-negative surface would give a value of -1. MATLAB code provided in Appendix B.

2.6. Solution and Bulk Characterization.

2.6.1. Dynamic light scattering (DLS).

DLS was used to detect the presence and size of soluble complexes such as complex coacervate core micelles (C3Ms). DLS was conducted either in water or buffer as noted for specific experiments. Unless otherwise noted, 10 measurements consisting of 10 acquisitions each were taken 3 times for each sample. Outliers that showed significant deviation from 1 at the baseline were excluded from averages because this indicates that a large piece of dust or other contaminant was in the field during the measurement. If more than 3 acquisitions had this issue, a new measurement was taken. Data were presented as mass distributions unless otherwise noted.

2.6.2. SANS.

SANS data were collected by Helen Yao and Haley Beech at Oak Ridge National

Laboratory spallation neutron source, beam line 6. As higher purity is necessary for these experiments, the GFP mutants were purified using buffers containing 1 M NaCl as opposed to 300 mM NaCl. SANS was attempted on samples of the patchy GFP mutants containing or thought to contain soluble complexes to determine the structure of these complexes. Of the blends tested,

-1 only the PAA-XLP mixture did not form solid precipitates at 10 mg mL in D2O (the minimum concentration to obtain sufficient signal). The scattering curves were fitted using SASview with a cylinder model as described in Chapter 4.

2.6.3. Optical Microscopy.

Optical microscopy was used to determine whether samples were precipitating or forming complex coacervates. Samples that were seen to macrophase separate by turbidimetry were prepared at 2 mg mL-1 at the blending ratio where the peak turbidity was observed. After the samples were blended, they were placed on a slide with a coverslip. The samples were imaged with a Zeiss Axioplan microscope equipped with a 100x objective and an Axiocam 503 Mono camera under bright field. The complex coacervates and precipitates look somewhat similar immediately after loading, but given 15-30 min, the complex coacervate droplets will coalesce into larger droplets, whereas precipitates will either continue to float around appears as small particles or will aggregate with individual particulates still visible.

2.7. Thin Film Characterization

2.7.1. Atomic Force Microscopy (AFM).

The NTEGRA system from NT-MDT was used to take height and phase scans using a

HQ:NSC16/Al BS tips from MikroMasch. Images were processed using the Nova PX software

89 from NT-MDT. Line-by-line linear corrections were applied to each image. Acquiring high-quality

AFM images requires fine-tuning of imaging conditions based on the properties of the materials being imaged. First, an appropriate AFM tip must be chosen based on the hardness of the sample.

The tips used here are typically good for “average” samples, but probes with lower spring constants should be screened if there is not sufficient contrast or if the tip appears to be damaging the sample.

Common issues that can arise are the emergence of a phantom pattern in the phase scan that appears as perfectly aligned diagonal lines and picking up debris on the tip that appears as perfectly repeating triangles in the image. In both cases, the tip should be replaced. If the diagonal lines persist there may be an issue with one of the internal components and NT-MDT should be contacted. Image samples at two appropriate lengths scales to obtain both long-range and short- range structural data as in some cases these may differ.

The film thicknesses were determined by scratch tests. Films were scratched using a 30 gauge needle (BD). An AFM scan was taken over the scratch and a planar flattening correction was applied to the exposed Si surface. The average height profile was then extracted from the data and analyzed to give the average film height. While this technique does work well, it highly time- consuming, and alternatives such as profilometry or ellipsometry should be used if possible. The main issues with this technique are the long imaging times, which can be reduced by taking slightly lower resolution images and picking up debris from the raised front of material from near the edge of the scratch. To minimize debris, use a fresh needle for each scratch.

2.7.2. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS).

GISAXS was conducted at beamline 8-ID-E at Argonne National Laboratory. The beamline scientists provide a detailed step-by-step procedure for the alignment and measurement of samples to all users. It is frequently updated to keep up with both software improvements and

90 capital improvements to the line. In brief, when the run starts, the beamline scientist will first create a parameters file that captures essential data like the energy of the beam and the sample to detector distance. Then work will begin by loading samples cast onto Si wafers (minimum of 1 cm x 1 cm) onto an 8-sample wheel by applying a scant amount of grease (provided at the beamline) to the back side of the sample and placing on one of the 8 spots on the sample wheel (positions are labeled and labels are visible via camera after the hutch is closed). After the samples are placed in the hutch and the hutch is closed, the beamline scientist will help to teach users (especially new users) how to properly align and image samples and will help to optimize instrument parameters like beamstop positioning. It is highly recommended to carry some sacrificial samples for this initial training and optimization as repeated imaging can (and will) destroy samples. If the critical angle of the samples is unknown, the beamline scientist can assist with determining this value as well.

An angle of 0.155° was used for this work because it exceeded the critical angle of the organic layer, but was less than the critical angle of the wafer. A rough alignment is accomplished by finding a maximum in intensity by varying the angle and finding the height at which the intensity is reduced by half compared to the maximum. A finer alignment based on a reflection at approximately 0.5° is then completed to correct both the angles in-line and perpendicular to the beam. After this the scattering patterns can be collected. Typically, data were collected at 3 angles using spots that were 2 beam-widths apart. If a spot produced significant flare, the position was adjusted to minimize flare. Data were analyzed using the GIXSGUI14 for MATLAB that was developed at this beamline available at: https://www.aps.anl.gov/Sector-8/8-ID/Operations-and-

Schedules/Useful-Links/Sector-8-GIXSGUI. Documentation of the software is available here as well. The beamline scientists can help in getting started with the tool.

2.8. References

1. Huang, J.; Cusick, B.; Pietrasik, J.; Wang, L.; Kowalewski, T.; Lin, Q.; Matyjaszewski, K., Synthesis and In Situ Atomic Force Microscopy Characterization of Temperature- Responsive Hydrogels Based on Poly(2-(dimethylamino)ethyl methacrylate) Prepared by Atom Transfer Radical Polymerization. Langmuir 2007, 23 (1), 241-249. DOI: 10.1021/la061683k 2. Stewart-Sloan, C. R. Understanding the effect of protonation on the self-assembly of a model polyelectrolyte-neutral block copolymer. Ph.D. Thesis, Massachusetts Institute of Technology, Cambridge, MA, 2016. 3. Thomas, C. S.; Glassman, M. J.; Olsen, B. D., Solid-State Nanostructured Materials from Self- Assembly of a Globular Protein–Polymer Diblock Copolymer. ACS Nano 2011, 5 (7), 5697-5707. DOI: 10.1021/nn2013673 4. Agilent, QuikChange II Site-Directed Mutagenesis Kit. 2015. https://www.agilent.com/cs/library/usermanuals/Public/200523.pdf (accessed 4/15/2018). 5. Qiagen, The QIAexpressionistTM: A handbook for high-level expression and purification of 6xHis-tagged proteins. Qiagen: Hilden, Germany, 2003. 6. Stafford, C. M.; Roskov, K. E.; III, T. H. E.; Fasolka, M. J., Generating thickness gradients of thin polymer films via flow coating. Rev. Sci. Instrum. 2006, 77 (2), 023908. DOI: 10.1063/1.2173072 7. Schroeder, M. M.; Wang, Q.; Badieyan, S.; Chen, Z.; Marsh, E. N. G., Effect of Surface Crowding and Surface Hydrophilicity on the Activity, Stability and Molecular Orientation of a Covalently Tethered Enzyme. Langmuir 2017, 33 (28), 7152-7159. DOI: 10.1021/acs.langmuir.7b00646 8. Lawrence, M. S.; Phillips, K. J.; Liu, D. R., Supercharging Proteins Can Impart Unusual Resilience. J. Am. Chem. Soc. 2007, 129 (33), 10110-10112. DOI: 10.1021/ja071641y 9. Lam, C. N.; Yao, H.; Olsen, B. D., The Effect of Protein Electrostatic Interactions on Globular Protein–Polymer Block Copolymer Self-Assembly. Biomacromolecules 2016, 17 (9), 2820-2829. DOI: 10.1021/acs.biomac.6b00522 10. Berman, H. M.; Westbrook, J.; Feng, Z.; Gilliland, G.; Bhat, T. N.; Weissig, H.; Shindyalov, I. N.; Bourne, P. E., The Protein Data Bank. Nucleic Acids Res. 2000, 28 (1), 235-242. DOI: 10.1093/nar/28.1.235 11. Jurrus, E.; Engel, D.; Star, K.; Monson, K.; Brandi, J.; Felberg, L. E.; Brookes, D. H.; Wilson, L.; Chen, J.; Liles, K.; Chun, M.; Li, P.; Gohara, D. W.; Dolinsky, T.; Konecny, R.; Koes, D. R.; Nielsen, J. E.; Head-Gordon, T.; Geng, W.; Krasny, R.; Wei, G.-W.; Holst, M. J.; McCammon, J. A.; Baker, N. A., Improvements to the APBS Biomolecular Solvation Software Suite. Protein Sci. 2018, 27 (1), 112-128. DOI: 10.1002/pro.3280 12. Xu, D.; Li, H.; Zhang, Y., Protein Depth Calculation and the Use for Improving Accuracy of Protein Fold Recognition. J. Comput. Biol. 2013, 20 (10), 805-816. DOI: 10.1089/cmb.2013.0071 13. Xu, D.; Zhang, Y., Generating Triangulated Macromolecular Surfaces by Euclidean Distance Transform. PLoS One 2009, 4 (12), e8140. DOI: 10.1371/journal.pone.0008140 14. Jiang, Z., GIXSGUI: a MATLAB toolbox for grazing-incidence X-ray scattering data visualization and reduction, and indexing of buried three-dimensional periodic nanostructured films. J. Appl. Crystallogr. 2015, 48 (3), 917-926. DOI: 10.1107/S1600576715004434

Chapter 3. Catalytic Biosensors from Complex Coacervate Core Micelle (C3M) Thin Films

Reproduced (adapted) with permission from Sureka, et al. Catalytic Biosensors from Complex

Coacervate Core Micelle (C3M) Thin Films, ACS Appl. Mater. Interfaces 2019, 11, 35, 32354–

3.1. Abstract

Enzymes have been applied to a variety of industrially and medically relevant chemistries as both catalysts and sensors. Incorporation of proteins and enzymes into complex coacervates has been demonstrated to improve the thermal, chemical, and temporal stability of enzymes in solution. In this work, a neutral-cationic block copolymer and an enzyme, alkaline phosphatase, are incorporated into complex coacervate core micelles (C3Ms) and coated onto a solid substrate to create a biocatalytic film from aqueous solution. The incorporation of photo-crosslinkable groups into the neutral block of the polymer allows the film to be crosslinked under ultraviolet light, rendering it insoluble. The morphology of the films is shown to depend most strongly on the protein loading within the films, while solvent annealing is shown to have minimal effect. These films are then demonstrated as specific sensors for Zn2+ in solution in the presence of other metals, a model reaction for ion-selective metal biosensing useful in environmental monitoring. They are shown to have low leaching and maintain sufficient activity and response for sensing for 1 month after aging under ambient conditions and at 40 °C and 50% relative humidity. The C3M immobilization method demonstrated can be applied to a wide variety of proteins with minimal chemical or genetic modification and could be used for immobilization of charged macromolecules in general to produce a wide variety of thin film devices.

3.2. Introduction

Enzymes are green, biodegradable catalysts that are typically selective, sensitive, and highly enantio- and stereo specific. They have been applied to a variety of industrially and medically relevant chemistries including glucose isomerization,1-2 pharmaceutical synthesis,3-6 biodiesel production,7-8 and carbon dioxide sequestration.9-10 Enzymes are most readily used as homogeneous catalysts, but heterogeneous catalysts are desirable in both biocatalysis and sensing applications because they allow for longer usage and easier purification without loss of the catalyst.1-2, 6-12 Enzyme immobilization presents a promising route to stabilize proteins against chemical and thermal stress, maximizing the useful lifetime of a protein-based device.1-2, 6-14

Surface immobilization is the most commonly used enzyme immobilization method, and it has been applied to make both biocatalysts and biosensors. Notable examples include glucose and ethanol sensors15 and ELISA assays,16 along with some industrially available glucose isomerase1-2 and lipase products.17 Surface immobilization can be achieved by either passively binding the protein to a substrate through van der Waals forces, electrostatic forces, etc. or by covalently binding the protein to a functionalized surface.9, 11, 15, 18 Despite its common usage, surface immobilization is known to have low protein loading and reduced function due to protein orientation defects.15-16 The use of spacers is thought to allow for greater translational and conformational freedom to try to overcome the issue of protein orientation defects.15 In order to increase the shelf-life of these sensors and catalyst, proteins can be crosslinked with glutaraldehyde or supplemented with polyelectrolytes or polyols.9, 11, 15, 18

Alternative immobilization methods try to increase protein loading and protein mobility by moving from a 2-dimensional surface to a 3-dimensional matrix. Entrapment, encapsulation, layer- by-layer (LbL) assembly, complex coacervation, and protein-polymer block copolymer self-

94 assembly are methods that have been demonstrated to create 3-dimensional enzymatic systems.9,

19-26 Among the many different methods, the use of LbL technologies and complex coacervates to achieve immobilization have garnered significant attention in recent years due to their relative ease.18, 21, 23-30 Complex coacervate based methods enable coating to be achieved in only one step, which is attractive for manufacturing.19, 23, 25, 27, 30

Both complex coacervates and C3Ms have been demonstrated to achieve high protein loading by mass and to improve the stability of encapsulated proteins to temporal, thermal, and chemical degradation.21, 26-27, 30-33 Encapsulation of GFP, lipase, organophosphate hydrolase, and many other proteins has been demonstrated.21, 26-27, 30-33 Supercharging of proteins has been demonstrated to aid in the incorporation of proteins into both complex coacervates and C3Ms, and has allowed for proteins to replace one of the polyelectrolytes in the system.25, 34-35 The formation of both phases is well-predicted by the negative charge ratio of the protein when combined with a strong polyelectrolyte.25 Thus, enzymes can be genetically engineered20 or chemically modified to form complex coacervates and C3Ms.25, 34-35 In cases where protein modification is difficult, complex coacervates and C3Ms incorporating enzymes can still be formed; however, overall protein loading is lower.27 This is achieved by combining an anionic polymer, a cationic polymer, and a protein in solution and allowing the protein to partition into the complex coacervate or C3M phase.26-27, 30 Charged-neutral diblock copolymers, like those used in C3Ms, have also been demonstrated to microphase separate,36 and templating of mCherry has been demonstrated in C3M films previously.19

The current standard for heavy metals analysis in water is inductively coupled plasma-mass spectrometry on a pre-concentrated sample, per the U.S. Environmental Protection Agency

(EPA).37 This technology is requires a trained technician to operate, a clean workspace, and

95 electricity. To overcome these limitations, technologies are under development. One household test that is available today uses a color-changing paper strip to estimate the concentration of heavy metals in water.38 While this works for civilian applications in the home, military and industrials applications, like detection of nuclear production and pollution levels, require specificity and a higher degree of precision.

Several amperometric, field effect transistor (FET)-based, and optical sensors have been developed.39 Forster resonance energy transfer (FRET) has been leveraged to develop a Hg2+ sensor, where the presence of the ions causes the formation of a thymine- Hg2+-thymine hairpin structure that brings a quencher into close proximity of an organic fluorophore, diminishing its signal.40 Newer devices use more stable species like quantum dots for fluorescence and Au- nanoparticles (NPs) to achieve a similar effect, nanometal surface energy transfer (NSET), with lower likelihood of photo-bleaching.41 Functionalized Au NPs have also been used to detect heavy metals colorimetrically, through a shift in the surface plasmonic resonance (SPR) peak upon aggregation induced by the presence of the metals.39, 42-44Amperometric devices utilizing cabon nanotubes have also been developed with high sensitivity and specificity; however, they show poor performance after day-to-day use.39

The use of enzymes to accomplish metals sensing is also under development. Satoh and coworkers have developed a flow sensor for Cu2+, Zn2+ and Co2+ using ascorbate oxidase (ASOD) and alkaline phosphatase (PhoA) based on apoenzyme reactivation.14 Urease, glucose oxidase

(GOx), and horseradish peroxidase (HRP) have been used for amperometric detection of heavy metals.45 A colorimetric assay for the detection of several different heavy metals using β- galactosidase deactivation on a paper strip has also been developed.46

This work demonstrates a method to create insoluble, biocatalytic films using a one-step aqueous coating process based on complex coacervation followed by photo-cross linking chemistry. The enzyme alkaline phosphatase is used as a model protein for selective sensing of transition metals in water. It is genetically engineered to increase negative charge density and drive

C3M formation with a photo-crosslinkable neutral-cationic block copolymer. The effects of film thickness, annealing conditions, and protein loading on film activity and morphology are evaluated with fluorimetric assays, scanning probe microscopy (SPM), and grazing-incidence small-angle

X-ray scattering (GISAXS). The films are then tested for sensitivity and selectivity for Zn2+ under a variety of conditions including in the presence of various mixtures of metals and in river water.

Finally, the films are tested for their thermal stability. The results demonstrate the viability of this immobilization method for the fabrication of a multitude of enzyme and macromolecule-based thin film devices.

3.3. Results and Discussion

3.3.1. Materials Design.

A polymer encapsulant for thin film formation was designed using a neutral-cationic block copolymer synthesized by reversible addition-fragmentation chain transfer (RAFT) polymerization. The synthesis of poly-[(oligo-ethylene glycol methacrylate)-r-(benzophenone methacrylate)]-b-(methyl-quaternized 4-vinylpyridine)] ((POEGMA-r-BP)-b-qP4VP) was performed according to Figure 3.1. This polymer is advantageous for enzyme encapsulation because proteins are weakly interacting with the uncharged POEGMA block,25, 27, 47 the POEGMA and qP4VP are sufficiently segregated to yield ordered nanostructures even in the absence of protein,36 and the rubbery-glassy contrast between the two blocks of the copolymer provides imaging contrast in SPM. First, the random copolymer of OEGMA and BP was synthesized via

RAFT. The resulting polymer was then used as a macro chain transfer agent in the polymerization of 4VP. The final product was quaternized by reaction with iodomethane. The benzophenone concentration was optimized to achieve successful photo-crosslinking while also maintaining water solubility. The final polymer had degrees of polymerization (DP) of 107 and 96 for

OEGMA/BP and 4VP blocks, respectively, and dispersity (Đ) of 1.11 as confirmed by 1H NMR and size-exclusion chromatography (Figures A.1-5). The OEGMA/BP block was 9 mol% BP monomer.

Figure 3.1. Synthesis of (POEGMA-r-BP)-b-qP4VP via RAFT polymerization. POEGMA-r-BP was first polymerized via RAFT, then the polymer was used as a macro-RAFT agent to polymerize

(POEGMA-r-BP)-b-P4VP. (POEGMA-r-BP)-b-P4VP was then quaternized with methyl iodide.

To prepare metal sensing catalysts, alkaline phosphatase (PhoA) was selected as a model enzyme. This enzyme can be used as a zinc sensor,14 a phosphate scavenger for bone scaffolds,13 and a detector for pesticides.48 It is known to be relatively easy to engineer and express in large quantities.49 In order to promote strong aggregation between the cationic polymer and protein,

PhoA was genetically modified to have increased anionic charge. Previous work has shown that a negative charge ratio (α) of approximately 1.4 or greater is necessary for the formation of C3Ms,25

98 so the modifications were designed to surpass this level. Wild-type PhoA has α of 1.26. Genetic modification was chosen over chemical charge modification in order to provide a single molecular species for coacervation experiments. Five mutants were expressed and purified by periplasmic extraction and fast protein liquid chromatography (FPLC) using anion exchange. The structure of the protein and a summary of the mutations are shown in Figures 3.2a,b. Polyacrylamide gel electrophoresis (PAGE) analysis of the purified mutants is shown in Figure A.6a,b. The enzyme activity was fit to Michaelis-Menten kinetics, and the negative mutants were found to have higher kcat than the native enzyme in solution, with slightly increased KM (Figure 3.2). A representative fit is shown in Figure A.6c. PhoA Mutant 4 (K92D + K96D + K352D + K353D) was chosen for the sensor because of its high negative charge ratio of 1.50, high expression yield, and high activity.

Figure 3.2. a) Structure of alkaline phosphatase (PDB: 3TG0)50 with mutations highlighted. b)

Table of mutants and corresponding mutations and α-values. c) kcat and d)KM data for supercharged

PhoA mutants determined by fluorimetric assay measuring conversion of 4-MUP to 4-MU. All mutants showed significantly greater kcat than the wild-type and a slight increase in KM. Mutant 4 was chosen for subsequent studies because it had sufficient negative charge ratio to enable complexation without a relative drop-off in activity.

3.3.2. C3M Formation and Activity.

Upon mixture of the protein and the block copolymer, the two components formed complex coacervate core micelles (C3Ms). Micelle formation was confirmed by dynamic light scattering

(DLS) as a function of polymer-protein blending ratio (Figure 3.3a) at 1 mg mL-1 in water. Both the pure polymer and pure protein showed minimal aggregation. Upon addition of 5% (wt/wt) protein to the polymer, a small amount of complexation was seen as peaks began to form above

10 nm. The level of complexation steadily increased until micelles were formed at 40% (Rh = 23.8

± 9.6 nm) and 50% (Rh = 39.9 ± 9.8 nm). Past this point, at 60% and 75% protein larger aggregates began to form. A solution at 50% protein was chosen for all further work as this mixture formed micelles and was the point of maximum coacervation with the qP4VP homopolymer per the method described by Obermeyer, et al.25 (Figure A.7).

Polymer addition caused a slight decrease in the effective activity of the enzyme (Figure

3.3b). Solutions of the enzyme alone and the enzyme and polymer together at a 50:50 ratio were prepared with equal concentrations of enzyme. The effective kcat of the enzyme was reduced from

-1 -1 6.31 ± 0.01 s to 3.84 ± 0.01 s upon mixing, and the effective KM was found to decrease from

1.27 ± 0.03 μM to 0.94 ± 0.07 μM. This could be due to noncompetitive inhibition of the enzyme,51 denaturation, or the effect of the local environment on activity.15

100

Figure 3.3. a) Particle mass distributions determined by DLS for mixtures of PhoA and

(POEGMA-r-BP)-b-qP4VP. Complexes began to form at 5% (wt/wt), with micellization occurring at 40% and 50%, and the formation of larger complexes occurring past this point. b) Effect of polymer addition on the activity of PhoA in solution. The polymer was added at 50% (wt/wt) with enzyme concentration remaining constant between the two cases. The effective kcat of the enzyme

-1 -1 was reduced from 6.31 ± 0.01 s to 3.84 ± 0.01 s upon addition of the polymer and effective KM was found to decrease from 1.27 ± 0.03 μM to 0.94 ± 0.07 μM.

3.3.3. Enzyme Immobilization and sensing within C3M Films.

Enzyme immobilization was achieved by flow coating a 15-20% (wt/wt) solution of

POEGMA-BP-b-qP4VP and PhoA M4 (50:50) in water onto a polyethylene glycol (PEG)-treated

Si surface and crosslinking under ultraviolet light (254 nm) (Figure 3.4). The protein is expected to segregate strongly into the charged domains as shown in Figure 3.4, per work done by Kim, et al.19 The PEG-treatment of the Si surface was necessary to allow for adhesion and wetting of the films to the surface. The films were photo-crosslinked for 5 minutes and soaked overnight in buffer prior to use. Photo-crosslinking and soaking had little effect on the structure of the C3M films 101

(Figure A.8). Based on film color, which went from light blue (ca. 150 nm) to a pale yellow (ca.

450 nm) or magenta (ca. 600 nm) after immersion, the films swelled by approximately 300 to

400% in solution. The films were found to be hydrophilic based on contact angle analysis (Table

A.1).

Figure 3.4. Method for immobilization of enzymes in C3M thin films. Polymer and protein are pre-mixed to form complex coacervate core micelles and then flow coated onto a solid substrate.

UV irradiation is used to crosslink the soft block via benzophenone groups, rendering the final film insoluble.

Reconstitution of the PhoA apo-enzyme in various transition metal solutions, followed by activity assays, provides a basis for the selective detection of low concentration transition metals using the coacervate thin films. The sensors can achieve high selectivity for zinc within metal mixtures because PhoA apo-enzyme becomes active only when it is reconstituted with zinc. In a typical sensing cycle, the coacervate films are first converted to the apo-enzyme form by treatment with EDTA to chelate any metal. The films are then incubated for a short period in a desired metal

102 solution, and subsequently the film activity is assayed. The ratio of the film activity after exposure to the sample and the initial film activity is referred to as the relative response and is used to quantify the film response to the metals.

Increasing film thickness was found to increase the film activity and response; however, this effect diminished beyond a certain thickness (Figure 3.5). Activity increased between the 53,

101 and 143 nm films but did not increase between the 143 and 175 nm films. Relative response also showed an increasing trend between the 53, 101 and 143 nm films, but did not increase between the 143 and 175 nm films. These phenomena could be the result of transport limitations.

Figure 3.5. a) Film activity for films of 50 wt.% enzyme at varying thickness. Activity increase with thickness is minimal beyond about 150 nm. b) Relative response to 5 ppm Zn2+ as a function of film thickness. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Relative response increase was minimal after 150 nm.

Protein loading was found to have a significant effect on the morphology, activity, and response of the C3M films. The polymer alone showed a disordered micelle phase in both scanning probe microscopy (SPM) and grazing-incidence small-angle X-ray scattering (GISAXS) (Figure

103

3.6a-c). The SPM images show that large structures of phase separated protein-polymer complexes begin to form at 5% (wt/wt) protein loading (Figure A.9), and increase in abundance at 10%

(Figure 3.6d-f) and 25% (Figure 3.6g-i), until they fully dominate the film structure at 50% (Figure

3.6j-l). The GISAXS patterns show a peak for the disordered micelles of the pure polymer that gradually weakens with increased protein loading up to 25% protein loading. The disordered micelles of the block copolymer alone were not visible by SPM past 0% protein loading, however the GISAXS data suggest that the subsurface structure is still partially similar to that of the pure polymer. At 50% protein loading, the peak in the GISAXS pattern no longer appears, suggesting that the larger structures fully dominate the film structure at this point. This was further confirmed by 1-D analysis of the GISAXS data (Figure A.10), with the diminishing peak in both qy and qz.

While the precise structure of the complexes in the films is unknown, the trend in complexation with protein loading is consistent with the DLS data in Figure 3.3a, which showed a gradual increase in structures with radii greater than 10 nm with increased protein loading until only the large structures were detected at 40% and 50% protein loading. The activity of the films is proportional to the protein loading (Figure 3.7a), as expected. The relative response, which should remain constant with protein loading (apart for the film with 0% protein), was maximized at the highest protein loading studied, which was 50% protein (Figure 3.7b). Film thickness was held relatively constant at 105-126 nm, apart from the 10% protein film because this solution had noticeably lower viscosity than the others, producing a thinner film.

104

Figure 3.6. SPM height and phase images and GISAXS scattering patterns for films with 0% (a- c), 10% (d-f), 25% (g-i), and 50% (j-l) (wt/wt) protein loading. Scale bars are 200 nm. Z-scales are 20 nm*, 40 nm†, 80 nm‡, and 10° for phase images. Scattering intensity is presented on a logarithmic scale. Film thicknesses were 117 ± 2, 77 ± 5, 105 ± 7, and 126 ± 3 nm for 0, 10, 25, and 50 wt.% protein loading, respectively. The pure block copolymer structures into disordered micelles under the conditions tested. With increasing protein concentration, this structure gives way to the formation of larger features.

105

Figure 3.7. a) Film activity and b) relative response to 5 ppm Zn2+ as functions of protein loading.

Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Both activity and response scaled with the protein loading.

Film thicknesses were 117 ± 2, 120 ± 2, 77 ± 5, 105 ± 7, and 126 ± 3 nm for 0, 5, 10, 25, and 50 wt.% protein loading.

Solvent annealing in polar and aqueous solvents was explored as a method to control morphology in the coacervate films. Because of the electrostatic nature of the coacervate interactions and the need to maintain enzymatic activity, solvents with a known ability to solubilize salt and aqueous-based formulations were the focus of annealing studies. Films were annealed with water, dimethyl sulfoxide (DMSO), dimethylformamide (DMF), acetone, and ethanol.

Solvents were mixed at a 50:50 volume ratio with methoxypolyethylene glycol (Mw=350) to reduce solvent activity, as described by Stewart-Sloan, et al.36 Under these conditions, film swelling of 30 to 50% was observed for water and acetone and less than 10% for DMSO and DMF based on color change in the films. Annealing led to little change in the height of the films after drying. Under all conditions tested, the films appear to form large complexes, with all solvents causing no significant change except DMSO which caused a shrinking of the complexes (Figure

106

3.8). Grazing incidence small angle X-ray scattering (GISAXS) confirmed that there was no significant difference between films annealed with the various solvents; however, the feature sizes observed in SPM would not be accessible by GISAXS (Figure A.11). Annealing with solvents other than water led to a decrease in both the film activity (Figure 3.9a) and the relative response of the films (Figure 3.9b). Based on these results, water annealing was utilized for subsequent studies.

Figure 3.8. SPM height and phase images for films as cast (a,b) and films annealed with water

(c,d), acetone (e,f), DMSO (g,h), and DMF (i,j). Scale bars are 200 nm. Z-scales are 35 nm for height images and 5° for phase images. All films are 50 wt.% protein. Film thicknesses were 89

± 2, 88 ± 3, 91 ± 2, 80 ± 4, and 83 ± 3 nm for the as cast, water annealed, acetone annealed, DMSO annealed, and DMF annealed films, respectively. Solvent annealing did not greatly affect the morphology of the films showing large complexes in all samples.

107

Figure 3.9. a) Film activity and b) relative response to 5 ppm Zn2+ after annealing with various solvents. All films are 50 wt.% protein. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with ZnSO4 to the native film activity. Treatment with water had no effect on activity, but all other treatments led to a marked decrease in both activity and response. Film thicknesses were 89 ± 2, 88 ± 3, 91 ± 2, 80 ± 4, and 83 ± 3 nm for the as cast, water annealed, acetone annealed, DMSO annealed, and DMF annealed films, respectively.

When tested against a panel of various different metals individually, PhoA M4 C3M thin films were capable of sensing zinc specifically when compared to Cu2+, Co2+, Ni2+ (Figure 3.10a), none of which showed signal above the background. However, in the presence of a multi-metal mixture, the overall response to zinc was reduced. Likely, this is due to competitive binding of the catalytic site by transition metals that are not catalytically active. Additionally, the presence of biological contaminants was found to have no statistically significant effect on the response of the films (Figure A.12). When tested with Zn2+ alone, the response follows Langmuir isotherm behavior (Figure 3.10b), and was fit to a scaled Langmuir isotherm,

푐 2+ 퐾 푀 푀 푀2+ 푅(푐푀2+) = (푅푚푎푥 − 푅푚푖푛) × 푐 2+ + 푅푚푖푛 (3.1) 1+퐾 푀 푀 푀2+

108

2+ where 푅 is the relative response, 푅푚푎푥 is the relative response at 5000 ppb Zn , 푅푚푖푛 is the relative

2+ -1 response at 0 ppb Zn , 푐푀2+is the concentration of the metal in solution in ppb (μg L ), 푀푀2+ is the molecular mass of the metal (g mol-1), and 퐾 is the equilibrium constant in μM-1. The equilibrium constant, K, was found to be 8.70 μM -1. This behavior is consistent with a set number of binding sites within the film, and the active site of the enzyme is expected to either be empty or have a bound metal center. In the absence of contaminants, the limit of detection (LOD) was found to be less the 5 ppb Zn2+.

Figure 3.10. a) Relative response of C3M films to treatments with various M2+ at 5 ppm. Relative response is the ratio of the film activity after deactivation with EDTA and reactivation with M2+ to the native film activity. Average film thickness was 124 ± 16 nm and all films were 50 wt.% protein. Films showed high selectivity for zinc in all cases. The addition of other metals led to decreased response to Zn2+. b) Relative response of C3M film to varying Zn2+ concentration.

Average film thickness was 97 ± 7 nm. Response was non-linear after 10 ppb Zn2+ and constant after 50 ppb.

109

The films were tested under a variety of conditions, including addition of contaminants, in water drawn from the Charles River before (CRW) and after filtering (CRWF), and in mixtures of metals (Figure 3.11). The samples were doped with 10 and 50 ppb of Zn2+. The two metal mixtures tested were 5 ppb Cu2+, 1 ppb Co2+, 4 ppb Ni2+, and 7 ppb Zn2+ (Mix 1) and 2 ppb Cu2+, 6 ppb

Co2+, 2 ppb Ni2+, and 4 ppb Zn2+ (Mix 2). The samples in Charles River water and in the two mixtures had a higher baseline response due to contaminants in the river water and the basal level of Zn2+ in the mixtures, but in all cases, the response increased with increasing Zn2+ concentration.

The response to 50 ppb Zn2+ was statistically significant compared to the control (p < 0.05) in all cases, except in filtered Charles River water (p = 0.052), which has a somewhat lower mean value at 50 ppb and higher variance than other samples. In the control, 10 ppb Cu2+, and 10 ppb Co2+ samples, there was a statistically significant response to 10 ppb Zn2+. The p-values for all conditions are summarized in Table A.1.

Figure 3.11. The C3M films were tested under a variety of conditions and the samples were doped with 10 and 50 ppb Zn2+. This included a control in Millipore water, 10 ppb Cu2+, Co2+, and Ni2+,

110

Charles River water (CRW), filtered Charles River water (CRWF), and 2 mixtures of metals (Mix

1: 5 ppb Cu2+, 1 ppb Co2+, 4 ppb Ni2+, and 7 ppb Zn2+; Mix 2: 2 ppb Cu2+, 6 ppb Co2+, 2 ppb Ni2+, and 4 ppb Zn2+). The response to 50 ppb Zn2+ was found to be statistically significant in all cases

(**: p < 0.01, *: p < 0.05), except in CRWF (p = 0.052). The p-values are summarized in Table

A.1. Films had an average thickness of 128 ± 16 nm, and all films were 50 wt.% protein.

3.3.4. Film stability to thermal degradation

Alkaline phosphatase has been shown to have high thermal stability, and can be held at higher than 80 °C for 5 to 30 minutes with minimal loss of activity in the presence of Mg2+, so the films were expected to have high thermal stability.52-54 Films were tested for thermal stability in 3 environments: ambient conditions (approximately 20 °C and 5-30% RH), 40 °C and 50% relative humidity (RH), and 70 °C and 4% RH. The physical appearance of the films did not change over time. The activity of the films decreased by 30.1 ± 5.7% under ambient conditions (Figure 3.12a) and by 24.0 ± 7.2% at 40 °C and 50% RH (Figure 3.12b) over 30 days. This decay occurred mostly within the first 3 to 7 days, which may be due to some initial enzyme inactivation, the film drying out over time, or equilibrating with the environment, but the films seem to be stable past that point.

The enzyme alone has been shown to have a similar activity decay profile at 20 °C in solution.55

The activity of the films at 70 °C and 4% RH decayed by 84.5 ± 20.6% over 30 days (Figure

3.12c). At 70 °C, the enzyme alone in solution was nearly fully deactivated after 3 days.55 This decay occurred rapidly with a 47.9 ± 9.6% decrease in activity happening in the first day. At both ambient conditions and 40 °C and 50% RH, the films retained sufficient activity to be usable.

The relative response of the films under ambient conditions (Figure 3.12d) and 40 °C and

50% RH (Figure 3.12e) decayed by 29.5 ± 13.4% and 27.9 ± 5.8%, respectively, similar to the decay seen in activity. If the decrease in activity were purely due to enzyme deactivation, the

111 relative response would be constant over time, since relative response is scaled by the activity of the film at the given time point. The response of the films aged at 70 °C and 4% RH (Figure 3.12f) initially decreased, but became unreliable because of the decay in the film activity. The decay in response at both ambient conditions and 40 °C and 50% RH mostly occurred within the first 3 days to 1 week after which there was an apparent steady state, and there was sufficient activity for sensing after 1 month. This suggests that after a brief curing period, the films may have an extended shelf-life.

Figure 3.12. Activity and response of films aged under ambient conditions (a,d), 40 °C and 50%

RH (b,e), and 70 °C and 4% RH (c,f). Films retained their activity and response under ambient conditions and 40 °C and 50% RH, but were greatly inactivated at 70 °C and 4% RH leading to unreliable response. Film thicknesses were 111 ± 2, 122 ± 2, and 142 ± 1 nm for ambient conditions, 40 °C and 50% RH, and 70 °C and 4% RH, respectively.

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3.4. Conclusions

Immobilization in C3M films has been demonstrated as an effective encapsulation and stabilization method for enzymes, and this work shows that this technology may be used to produce catalytically active polymer/protein hybrid thin films for biosensors. Micelle formation between charged block copolymers and proteins was confirmed via DLS and was found to maintain enzyme activity with a decrease in kcat likely caused by transport limitations or inhibition. These micelles were successfully cast into thin films, and the effects of film thickness, protein loading, and solvent annealing on morphology, activity, and response to Zn2+ were evaluated. Film thickness was found to have a diminishing effect on the activity and response of the films above a critical thickness.

SPM and GISAXS studies of the films showed that protein loading had a significant effect on the morphology of the films as the underlying micro-phase structure of the block copolymer gave way to formation of larger complexes with increased protein loading. Both the activity of the films and the response of the films increased with protein loading. Solvent annealing was found to have little effect on the film morphology, but to cause both the activity and response to decrease for all solvents tested except water.

The films were found to have both high selectivity and sensitivity to Zn2+. Selectivity was demonstrated in the presence of Cu2+, Co2+, and Ni2+, and response to Zn2+ was found to follow a

Langmuir isotherm. The response was tested under a variety of conditions and was found to be reliable for detection of 50 ppb Zn2+ in a range of aqueous samples. Aging under both ambient conditions and 40 °C and 50% RH led to slight decreases in both the activity and response; however, aging under 70 °C and 4% RH led to a sharp decrease in activity, leading to an unreliable response to Zn2+. The C3M film immobilization method can be easily employed on a range of enzymes with minimal genetic engineering or a simple chemical modification. Used with alkaline

113 phosphatase, this technology allows for selective sensing of zinc, the metal native to PhoA and serves as a model for generalized enzymatic sensing of metals. This immobilization technique could be employed on a variety of enzymes and macromolecules, and by tuning the polymer chemistry, immobilization could be achieved on a variety of substrates making this a general approach for biocatalytically active thin film fabrication.

3.5. Methods

3.5.1. Benzophenone monomer synthesis (BP).

Benzophenone methacrylate was synthesized following a previously reported protocol.56

Triethylamine (10 mL) was added to a solution of 4-hydroxybenzophenone (10 g, 50 mmol) in dichloromethane, and the solution was cooled to 0 °C in an ice water bath. A solution of methacroyl chloride (5.9 mL, 60 mmol) in dichloromethane was added dropwise while stirring.

The reaction mixture was subsequently allowed to warm and stirred at room temperature overnight. The mixture was washed twice each with water, saturated NaHCO3, and brine. The organic layers were dried over Na2SO4, filtered, and the solvent was evaporated. The resulting residue was purified by automated flash chromatography using a Biotage system with dichloromethane and methanol as a mobile phase. 1H NMR (400 MHz, CDCl3):  7.90 (m, 2 H),

7.83 (m, 2 H), 7.62 (m, 1 H), 7.52 (m, 2 H), 7.29 (m, 2 H), 6.42 (m, 1 H), 5.84 (m, 1 H), 2.12 (m,

3 H) (Figure A.13). LRMS (ESI) calculated for C17H15O3 ([M+H]+) 267.3, found 267.1 (Figure

A.14).

3.5.2. POEGMA-b-qP4VP Synthesis.

RAFT polymerization was used to synthesize a block copolymer from 4-vinylpyridine

(4VP) (95%, Aldrich) and oligo(ethylene glycol) methyl ether methacrylate (OEGMA, Mn = 300 g/mol) (Aldrich) with a small fraction of a benzoylphenyl methacrylate (BP) with a narrow

114 molecular weight distribution. OEGMA and 4VP were passed through basic alumina columns prior to polymerization to remove inhibitors. 4-cyano-4-(phenylcarbonothioylthio) pentanoic acid

(Aldrich, 133 mg, 0.50 mmol) and AIBN (recrystallized twice from methanol, 15.6 mg, 0.10 mmol) were added to a solution of OEGMA (20 g, 66.6 mmol) and BP (1 g, 3.8 mmol) in 60 g

1,4-dioxane in a molar ratio of 140:8:1:0.2 (OEGMA:BP:CTA:initiator). The solution was degassed by three freeze-pump-thaw cycles. The polymerization was carried out in a sealed flask at 65 °C and terminated after 7.5 h by removal of heat and exposure to oxygen. The polymer was then precipitated in hexanes and dried under vacuum.

The POEGMA-r-BP copolymer was found to have Mn = 31.8 kg/mol and Đ = 1.13 as determined by size exclusion chromatography (SEC) (Figure A.4). SEC analyses were performed on a Waters HPLC system equipped a Water 1515 Isocratic HPLC Pump with two columns

(ResiPore, 300 × 7.5 mm, up to 500k Da, Agilent Technologies, CA) in series. DMF with 0.02M

LiBr was used as the eluent with a flow rate of 1 mL/min at 70 °C. The detector system consisted of a Wyatt miniDAWN TREOS multi-angle light scattering detector and a Wyatt Optilab T-rEX differential refractive index detector. The ratio of BP to OEGMA was determined by 1H NMR

(400 MHz, CDCl3, Figure A.1) by comparison of δ 7.52 (m, 3 H) and 7.84 (m, 4H) for BP and

3.39 (s, 3 H) peak for OEGMA, respectively.

The POEGMA-r-BP was then used as a macromolecular chain transfer agent for RAFT polymerization of 4VP. 4VP (3.68 g, 35 mmol) and AIBN (3.2 mg, 0.020 mmol) were added to a solution of the POEGMA-BP copolymer (3.24 g) in 12.5 g of a mixture of 1,4-dioxane and DMF in a molar ratio of 350:1.0:0.2 (monomer:CTA:initator). The polymerization was carried out in a sealed flask at 75 °C and terminated after 6 h by removal of heat and exposure to oxygen. The polymer was then precipitated in diethyl ether and dried under vacuum. The polymerization

115 provided a well-defined (POEGMA-r-BP)-b-P4VP diblock copolymer of molecular weight Mn =

41.9 kg/mol with a dispersity of 1.11 (by SEC, Figure A.5). DP4VP and Mn were determined by

1 averaging the comparison of the H NMR (400 MHz, CDCl3) peaks at δ 6.33 (m, 2H) and 8.26 (m,

2H) for 4VP and to the 3.31 (s, 3H) peak for OEGMA, respectively (Figure A.2).

3.5.3. Quaternization of (POEGMA-r-BP)-b-P4VP.

The block copolymer was quaternized with a four-fold molar excess of iodomethane to

4VP monomer in minimal DMF to dissolve the polymer. The reaction mixture was stirred at room temperature for 24 h, and the modified polymer was precipitated in diethyl ether and dried under

1 vacuum. The degree of quaternization was > 95% as determined by H NMR (400 MHz, D2O) based on the emergence of a methyl peak at δ 4.24 (s, 3H) (Figure A.3).

3.5.4. Cloning of charge mutants of PhoA.

The gene for PhoA in the pTrc99a plasmid was obtained as a generous gift from Dr. Jeff

Glasgow. The QuikChange II Site Directed Mutagenesis Kit (Agilent) was used to mutate the pTrc99a-PhoA plasmid to generate 5 mutants of alkaline phosphatase: M1) K92D + K96D, M2)

K92D + K93D + K96D, M3) K352D + K353D, M4) K92D + K96D + K352D + K353D, and M5)

K92D + K93D + K96D + K352D +K353D. The supplied protocol was followed.57 The optimal

PhoA mutant was found to be Mutant 4. This mutant was used for all subsequent studies. The gene sequence and primer sequences are shown in Figure A.15 and A.16, respectively.

3.5.5. Expression and purification of PhoA.

Engineered genes were transformed into TunerTM cells following a standard protocol from

Novagen. A 5 mL overnight culture of the cells was prepared in LB media supplemented with 0.2 mg mL-1 ampicillin and grown at 37 °C. The overnight cultures were added to 1 L of 2xYT media supplemented with 0.2 mg mL-1 ampicillin and grown at 37 °C. When the optical density at 600

116 nm reached 0.4, the cultures were induced with 1 mM IPTG, 1 mM MgSO4, and 0.1 mM ZnSO4, and allowed to grow overnight at 37 °C. The cells were pelleted by centrifugation at 3,500 x G for

10 min and resuspended in 50 mM Tris buffer pH 8.0 after each centrifugation. The periplasm was isolated by adding solid sucrose (0.5 M), EDTA (2.5 mM), and lysozyme (0.6 mg mL-1) to the cells and incubating at 37 °C for 30 min. The sphaeroplasts were removed by centrifuging at

10,000 x G for 20 min. The resulting supernatant was the periplasmic fraction. The periplasmic fraction was precipitated by heating the solution for 10 min in an 80 °C water bath. The mixture was centrifuged at 24,000 x G to remove precipitates. The supernatant was dialyzed against 20 mM Tris pH 8.0, and then purified via anion exchange chromatography using 2 5 mL Q columns

(HiTrap Q HP, GE Healthcare) in series equilibrated with 20 mM Tris pH 8.0 and eluted with a 0-

0.5 M NaCl gradient using the GE Akta FPLC. Fractions containing PhoA were identified by UV absorbance at 280 nm and SDS-PAGE analysis. These fractions were pooled and dialyzed against

20 mM Tris pH 8.0 with 1 mM MgSO4 and 0.1 mM ZnSO4 and then stored at 4 °C.

3.5.6. Dynamic Light Scattering (DLS).

DLS was done on a Wyatt Möbius. PhoA was exchanged into water by ultrafiltration and diluted to 1 mg mL-1. (POEGMA-r-BP)-b-P4VP was dissolved in water at 1 mg mL-1 in water.

Samples were then made by blending the two solutions at the appropriate ratios. Samples were allowed to equilibrate overnight at 4 °C. Each sample was measured 3 times for 10 seconds.

3.5.7. Grafting PEG to silicon wafers.

Silicon wafers (WaferWorld, (100) orientation) were cleaned with an air plasma for 1 min.

An approximately 0.25 cm thick layer of poly(ethylene glycol) monomethyl ether (PEG, Mn=750)

(Aldrich, #202495) was applied to the wafer. The coated wafer was then heated to 150 °C for 18 h under vacuum. Wafers were stored in acetone prior to use.

117

3.5.8. Flow coating of coacervate solutions.

PEG-coated wafers were rinsed with acetone, methanol, and water and dried using filtered air. PhoA was exchanged into water 4 times, diluting by a factor of 10 each cycle, and concentrated to 15-20% (wt/wt) via ultrafiltration (Amicon Ultra 30k-15mL). Concentration was measured via absorption at 280 nm. A 25% solution of POEGMA-BP-b-qP4VP was prepared in water and mixed with the protein solution to achieve a 50:50 mass ratio. The final mixture was diluted to 15-20%.

The solution was then flow coated using a setup similar to that described by Stafford, et al.58 onto the substrate under 50-70% RH at room temperature. Films were then annealed with water vapor in a chamber containing 0.6M NaCl in water overnight and crosslinked under ultraviolet light for

5 minutes (UVP Model UVGL-15, 4 W, 254 nm), unless otherwise stated.

3.5.9. Scanning probe microscopy (SPM).

The NTEGRA system from NT-MDT was used to take height and phase scans using a

HQ:NSC16/Al BS probe from MikroMasch. Images were processed using the Nova PX software from NT-MDT. Planar or second order corrections were applied to all images.

3.5.10. Film height measurement.

Film thickness was determined by a scratch test. Films were scratched using a 30 gauge needle (BD). An SPM scan was taken over the scratch and a planar flattening correction was applied to the film surface. The average height profile was then extracted from the data and analyzed to give the average film height.

3.5.11. Grazing incidence small-angle X-ray scattering (GISAXS).

GISAXS experiments were conducted at Argonne National Laboratory at beamline 8-ID-

E using X-ray with energy 10.86 keV. Samples were measured under ambient conditions at 3 incident angles (i) 0.14°, 0.15°, and 0.155° between the critical angle of PEG-coated silicon and

118 the critical angle of the C3M film. The reported images are from an incident angle of 0.155°. Two images obtained at two different detector positions were combined to fill the gaps in the detector.

Samples were exposed for 5 seconds at each angle and detector position, keeping total exposure time to 30 seconds. Sample to detector distance (SDD) was 2165 mm. Data were analyzed using the GIXSGUI software package written by Dr. Zhang Jiang. Data were converted to q space by applying correction parameters (stored as .MAT file format) provided by the beamline scientists.

3.5.12. PhoA activity assays.

PhoA activity was measured fluorimetrically on a TECAN Infinite M200Pro plate reader.

4-methylumbelliferyl phosphate (4-MUP) (Aldrich) was converted to 4-methumbelliferone (4-

MU), which was excited at 362 nm and measured at 448 nm. 4-MUP was dissolved in dimethyl sulfoxide at 10 mM and diluted in water to the concentrations for the assays. Standard curves

(Figure A.17) for the assays were prepared as described in the SI. Solvatochromic effects due to changes in DMSO concentration were negligible (Figure A.18).

Assays on the enzyme and micelles were conducted in 96-well plates (Brand, #781602).

The mutants were compared in 100 mM 3-(N-morpholino)propanesulfonic acid (MOPS) buffer, pH 7.5, supplemented with 500 mM NaCl. The mutants were diluted to 0.5 µg of enzyme L-1 in

MOPS buffer. 160 µL of MOPS buffer and 20 µL of substrate solution were added to each well.

After taking an initial reading, 20 µL of diluted enzyme was injected into each well. The emission was measured for 2 min.

119

Assays on the films were conducted in 12-well plates (Falcon, #353043). The films were secured to the bottom of the wells using carbon tape; negative controls were conducted to ensure that the tape did not affect activity (Figure A.19). Films were swollen overnight in 20 mM Tris pH

8.0 with 1 mM MgSO4 and 0.1 mM ZnSO4. Prior to the assay, films were rinsed with water. 900

µL of 100 mM Tris pH 8.0 was then added to each well. After an initial background measurement,

100 µL of the substrate solution was added. The emission was then measured for 4 min.

For metals sensing, the films were first assayed as described above, then exposed to 100 mM EDTA in 100 mM Tris pH 8.0 for 25-30 min to convert the enzyme into apo-enzyme. Films were rinsed thoroughly with water, and activity was measured again to ensure inactivation. After another rinse, films were exposed to water containing the specified concentration of analyte and 1 mM MgSO4 for 20-25 min unless otherwise specified. Films were rinsed and assayed to determine the level of reactivation.

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47. Wang, Z.; Luan, Y.; Gan, T.; Gong, X.; Chen, H.; Ngai, T., Long-Range Interactions Between Protein-Coated Particles and POEGMA Brush Layers in a Serum Environment. Colloids and Surfaces B: Biointerfaces 2017, 150 (Supplement C), 279-287. DOI: 10.1016/j.colsurfb.2016.10.040 48. Mazzei, F.; Botrè, F.; Montilla, S.; Pilloton, R.; Podestà, E.; Botrè, C., Alkaline Phosphatase Inhibition Based Electrochemical Sensors for the Detection of Pesticides. Journal of Electroanalytical Chemistry 2004, 574 (1), 95-100. DOI: 10.1016/j.jelechem.2004.08.004 49. Glasgow, J. E.; Capehart, S. L.; Francis, M. B.; Tullman-Ercek, D., Osmolyte-Mediated Encapsulation of Proteins inside MS2 Viral Capsids. ACS Nano 2012, 6 (10), 8658-8664. DOI: 10.1021/nn302183h 50. Bobyr, E.; Lassila, J. K.; Wiersma-Koch, H. I.; Fenn, T. D.; Lee, J. J.; Nikolic-Hughes, I.; Hodgson, K. O.; Rees, D. C.; Hedman, B.; Herschlag, D., High-Resolution Analysis of Zn2+ Coordination in the Alkaline Phosphatase Superfamily by EXAFS and X-ray Crystallography. Journal of Molecular Biology 2012, 415 (1), 102-117. DOI: 10.1016/j.jmb.2011.10.040 51. Tomita, S.; Shiraki, K., Poly(Acrylic Acid) is a Common Noncompetitive Inhibitor for Cationic Enzymes with High Affinity and Reversibility. Journal of Polymer Science Part A: Polymer Chemistry 2011, 49 (17), 3835-3841. DOI: 10.1002/pola.24822 52. Muller, B. H.; Lamoure, C.; Le Du, M.-H.; Cattolico, L.; Lajeunesse, E.; Lemaître, F.; Pearson, A.; Ducancel, F.; Ménez, A.; Boulain, J.-C., Improving Escherichia coli Alkaline Phosphatase Efficacy by Additional Mutations inside and outside the Catalytic Pocket. ChemBioChem 2001, 2 (7‐8), 517-523. DOI: 10.1002/1439- 7633(20010803)2:7/8<517::AID-CBIC517>3.0.CO;2-H 53. Garen, A.; Levinthal, C., A Fine-Structure Genetic and Chemical Study of the Enzyme Alkaline Phosphatase of E. coli I. Purification and Characterization of Alkaline Phosphatase. Biochimica et Biophysica Acta 1960, 38, 470-483. DOI: 10.1016/0006- 3002(60)91282-8 54. Janeway, C. M. L.; Xu, X.; Murphy, J. E.; Chaidaroglou, A.; Kantrowitz, E. R., Magnesium in the Active Site of Escherichia coli Alkaline Phosphatase is Important for Both Structural Stabilization and Catalysis. Biochemistry 1993, 32 (6), 1601-1609. DOI: 10.1021/bi00057a026 55. Poltorak, O. M.; Chukhray, E. S.; Torshin, I. Y.; Atyaksheva, L. F.; Trevan, M. D.; Chaplin, M. F., Catalytic Properties, Stability and the Structure of the Conformational Lock in the Alkaline Phosphatase from Escherichia coli. Journal of Molecular Catalysis B: Enzymatic 1999, 7 (1), 165-172. DOI: 10.1016/S1381-1177(99)00039-9 56. Huang, J.; Cusick, B.; Pietrasik, J.; Wang, L.; Kowalewski, T.; Lin, Q.; Matyjaszewski, K., Synthesis and In Situ Atomic Force Microscopy Characterization of Temperature- Responsive Hydrogels Based on Poly(2-(dimethylamino)ethyl methacrylate) Prepared by Atom Transfer Radical Polymerization. Langmuir 2007, 23 (1), 241-249. DOI: 10.1021/la061683k 57. Agilent, QuikChange II Site-Directed Mutagenesis Kit. 2015. https://www.agilent.com/cs/library/usermanuals/Public/200523.pdf (accessed 4/15/2018). 58. Stafford, C. M.; Roskov, K. E.; III, T. H. E.; Fasolka, M. J., Generating thickness gradients of thin polymer films via flow coating. Review of Scientific Instruments 2006, 77 (2), 023908. DOI: 10.1063/1.2173072

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Chapter 4. The Effect of Protein Surface Charge Distribution on Protein–Polyelectrolyte

Complexation

Reproduced (adapted) in part with permission from Kim,* Sureka,* et al. Effect of Protein Surface

Charge Distribution on Protein–Polyelectrolyte Complexation, Biomacromolecules 2020, 21, 8,

4.1. Abstract

Charge anisotropy or the presence of charge patches at protein surfaces has long been thought to shift the coacervation curves of proteins and has been used to explain the ability of some proteins to coacervate on the “wrong side” of their pI. This work makes use of a panel of engineered sfGFP mutants with varying surface charge distributions but equivalent net charge and a suite of strong and weak polyelectrolytes to explore this concept. A patchiness parameter, which assessed the charge correlation between points on the surface of the protein, was used to quantify the patchiness of the designed mutants. Complexation between the polyelectrolytes and proteins showed that the mutant with largest patchiness parameter was the most likely to form complexes, while the smallest was the least likely to do so. The patchiness parameter was found to correlate well with the phase behavior of the protein-polymer mixtures, where both macrophase separation and the formation of soluble aggregates were promoted by increasing patchiness depending upon the polyelectrolyte with which the protein was mixed. Increasing total charge and increasing strength of the polyelectrolyte promote interaction for oppositely charged polyelectrolytes, while charge regulation is also key to interactions for similarly charged polyelectrolytes which must interact selectively with oppositely charged patches.

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4.2. Introduction

Protein immobilization has been widely used in the food, pharmaceuticals, textiles, and agriculture industries to stabilize proteins against degradation and enable easier integration of the protein into devices or industrial processes.1-12 In their native state, proteins are sensitive to chemical and physical conditions such as high temperature, extremes of pH, high ionic strength, and organic solvent, which makes the use of protein-based sensing and catalysis challenging in many industrial settings.1-2,6-10,13-16 Additionally, functional proteins are often used as homogeneous catalysts in industrial settings, making recovery of the protein difficult, which can increase the cost of bioprocesses by requiring greater amounts of enzyme, and complicating the design of devices such as biosensors. Protein immobilization can convert proteins into an active solid form that can be more easily incorporated into devices and industrial processes.1-2,6-10,13,16

Furthermore, immobilization can improve protein stability and help prevent protein aggregation.1-

2,6-10,13-16 Protein immobilization can be accomplished by a variety of methods including surface immobilization, encapsulation into a polymer matrix or gel, incorporation into a protein-polymer diblock copolymer with a protein block and a polymer block, layer-by-layer technologies, and complex coacervation.1-2,9,17-32 While several of these methods have been successfully demonstrated and deployed, complex coacervation avoids drawbacks such as low protein loading and misorientation as in the case of surface immobilization and losses from chemical degradation during the encapsulation process in polymer matrices and gels.17-18,22,28-29

Over about the past two decades, complex coacervation has garnered significant interest in the fields of biomaterials, biomedicine, and biosensing as an effective method for encapsulation because of its simplicity and ability to stabilize proteins.20,22,24,26-27,29,33-37 Complex coacervation is a liquid-liquid phase change in which electrostatically interacting polymers form a polymer-dense

126 phase (i.e. the complex coacervate), which is gel-like and can contain more than 70% water, and a polymer-dilute phase.11,22,25-27,29,34,38-44 Unlike the other methods, complex coacervation is easily accomplished by simply blending the polyelectrolytes and proteins in solution.11,22,26-27,29-30,32,34,38-

40 Furthermore, complex coacervates have been demonstrated to stabilize proteins against chemical, thermal, and temporal degradation and can maintain a high protein loading.22,29-30,32,34

Complex coacervation between synthetic polymers can be reasonably modeled by a range of different theories and computational methods ranging from the original Voorn-Overbeek model to more advanced computational methodologies.42-44 These theories make a range of assumptions that cannot be directly applied to the coacervation of folded proteins because of the rigidity of folded protein structures, the anisotropic charge distribution on the surface of the proteins, and the charge regulation that can occur on the protein surface.11,26-27,33,39,41-44 As such, protein-polymer coacervation has been mainly studied empirically. Various studies have confirmed that protein- polymer coacervates respond similarly to pH changes and salt concentration as synthetic polymer- only coacervates.11,26,29,33,39-41 Some studies have demonstrated systematic trends in protein coacervation, such as the work by Obermeyer, et al. that demonstrated that there is a cutoff point in the negative charge ratio for predicting whether or not a coacervate would form between a negatively charged protein and a strong polycation.26,39 However, in protein-polymer coacervates, there is often a shift in the point of maximal complexation away from the point of charge balance, as is predicted for synthetic polymers.26,33,45 This shift can be attributed to the presence of charge patches or charge regulation caused by polymer-protein interactions, which can shift the effective pKa of a residue by up to 2 pH units, depending on the specific protein and the polyelectrolyte strength of the complexing polyelectrolyte.11,26-27,33,39,41-44

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Several studies have specifically examined the role of charge anisotropy within the context of protein-polymer coacervation.33,35-37,39,45-46 A recent notable example is the study from Kapelner and Obermeyer which demonstrated that localizing a charged tag on a protein, as opposed to distributing the charge over the surface isotropically, can change the nature of the complexation that occurs between a protein and synthetic polymer with the tagged proteins being more likely to form complex coacervates and not solid flocculants.39 Additionally, recent work in the Perry lab has proposed the use of a pair correlation function between charged residues to quantify protein charges patches and hypothesized that the presence of larger patches leads to greater encapsulation efficiency of a protein into the coacervate phase.11 While charge anisotropy has been demonstrated to greatly affect the coacervation of proteins based on observed trends, these effects have not yet been quantified.

This study aims to elucidate and quantify the effects of charge anisotropy (patchiness) on the surface of proteins on the coacervation of proteins. A panel of mutants of a model protein, superfolder green fluorescent protein (sfGFP), were synthesized such that their net charges were approximately neutral and identical and isoelectric points were similar, but the distribution of charges on the surface varied from near perfectly isotropic to near perfectly anisotropic, providing a test sample set to demonstrate and quantify the effect of surface charge distribution on protein- polymer coacervation. The degree of charge patchiness at the surface of the proteins was confirmed visually by mapping out charge patches at the protein surface and numerically using a patchiness parameter. The coacervation behavior of the protein panel was determined with model strong and weak polycations and polyanions at various salt conditions to provide a detailed understanding of how charge patchiness affects coacervation under a wide range of classically studied conditions.

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4.3. Experimental

4.3.1. Materials.

Monomers (acrylic acid, 4-vinylpyridine, 4-styrenesulfonic acid sodium, and 2-

(dimethylamino)ethyl acrylate), 1,4-dioxane, hexane, dimethylformamide, ethanethiol, 4-cyano-

4-(phenylcarbonothioylthio) pentanoic acid and azobisisobutyronitrile (AIBN), K3PO4·H2O, carbon disulfide and 2-bromoisobutyric acid were purchased from Sigma Aldrich.

4.3.2. Designing GFP mutants in silico.

To create a series of GFP mutants having different patch sizes, the different amino acids within the protein that have already been mutated and are highly solvent exposed were determined based on analysis of super-charged GFP proteins.21,47 After mutating the amino acids in silico to invert or remove charge (changing K or R to D or E), the protein surface potential was visualized by using the Adaptive Poisson-Boltzman Solver (APBS).48 The APBS method is a continuum model used to describe electrostatic interactions between solutes in aqueous salt solutions. After mutation of amino acids, PDB files of the mutated GFP proteins (XLP, LP, MP and SP) were generated by

PyMOL. The PDB files were converted to PQR files using the PDB2PQR webserver prior to being loaded into APBS. The PQR files were generated at pH 8.0 at different ionic strengths using the

PARSE force field. After generating the PQR file, the surface potential file was obtained using

APBS.

4.3.3. Algorithms for assessment of protein surface patchiness

To enable quantitative analysis of the surface potential, a Euclidean distance transform algorithm

(EDTSurf) was used first to calculate the SAS utilizing the rolling ball method, providing the vertices of a tessellated SAS for each mutant.49-50 The SAS was then overlaid on the APBS

129 potential map that was previously calculated (Figure B.1) to give surface positions and potentials in three dimensions ([Q] = [푥̅ 푦̅ 푧̅ 푞̅]). MATLAB code in provided in SI.

4.3.4. Search algorithm for patches.

The [Q] matrices were then used to calculate where charge patches occurred at the protein surface.

All points in [Q] were first binned into positive (q ≥ 2 kT e-1; [Q+]), negative (q ≤ -2 kT e-1; [Q−] ), and neutral matrices (-2 kT e-1 < q < 2 kT e-1; [Q0]). An arbitrary point (the first in the matrix) was chosen from [Q±] , and all its nearest neighbors within [Q±] were found (within 2 Å), removed

± ± ± from [Q ] , and placed in the matrix [Pi ]. The nearest neighbors of each subsequent point in [Pi ]

± ± ± within [Q ] were then deleted from [Q ], and added to [Pi ] until no further points were found in [Q±]. This process was repeated using the first remaining point in [Q±], until no points remained

± ± 2 in [Q ]. For [Pi ] to be considered a patch, it had to have an area greater than 1 nm . The area of the patches was approximated by assuming the points generated by EDTsurf were uniformly distributed and dividing the SAS by the number of points and multiplying the number of points in pi by this average area per point. After the positive and negative patches were found, the neutral area was calculated by subtracting the total area of the patches from the total area of the SAS.

MATLAB code provided in SI.

4.3.5. Algorithm for determining protein patchiness parameter.

For all points in [Q], all points within a radius of 2 Å were found, and each assigned a 1, 0, or -1 for the interaction between the two points. On average, each point had 6 neighbors within this radius, indicating that only the nearest neighbors were identified. A value of 1 was assigned if the potentials at the two points were both either greater than or equal 2 kT e-1 or less than or equal to

-2 kT e-1, a value of 0 was assigned if the potential at either or both points was between the -2 and

130

2 kT e-1, and a value of -1 was assigned if the one point was greater than or equal to 2 kT e-1 and the other was less than or equal to -2 kT e-1. These values were summed and divided by the total number of interactions over the entire surface of the protein to give the patchiness correlation.

Under this algorithm, a completely positively or negatively charged surface would give a value of

4.3.6. GFP mutant expression and purification

The DNA and protein sequences for the GFP mutants are shown in Figure B.2 and 4.1, respectively

(PDB ID: 2B3P). All proteins were expressed in BL21(DE3) cells, which were transformed with plasmids encoding the GFP mutants. For each variant, a well-isolated colony was grown overnight in LB broth containing kanamycin at 37 °C. 5 mL of the overnight culture was diluted in 1 L of

LB broth and incubated at 37 °C until OD600 0.8-1. Cultures were then induced with 1 mM IPTG, grown for 20 h at 37 °C, and harvested. Cells were collected by centrifugation at 4000 g for 30 min, and the supernatant was decanted. The collected wet cells were homogeneously re-suspended in lysis buffer (50 mM KH2PO4, 300 mM NaCl, 10 mM imidazole, pH 8.0) and then sonicated in an ice bath for 10 min for two cycles. The resulting cell debris and the supernatant were separated by centrifugation at 12000 rpm at 4 °C for 60 min. The presence of the expressed protein in the supernatant was confirmed by SDS-PAGE. The crude proteins in supernatant were further purified using Ni-NTA metal affinity chromatography with 10 column volumes of wash buffer (50 mM

KH2PO4, 300 mM NaCl, 20 mM imidazole, pH 8.0) to get rid of impurities. Using 3 column volumes of elution buffer (50 mM KH2PO4, 300 mM NaCl, 250 mM imidazole, pH 8.0), pure protein was obtained. The purified proteins were dialyzed into 20 mM Tris, 0-300 mM NaCl, pH

131

8.0 with 7 times every 3 hours. The purified proteins were analyzed by SDS-PAGE, and molar mass was confirmed by MALDI-TOF. The secondary structure of the mutants was confirmed by

CD and UV fluorescence of purified GFP variants. All circular dichroism (CD) spectroscopy was carried out on an Aviv model 202 CD spectrometer. Proteins were measured in 20 mM Tris-Cl, pH = 8.0; samples were prepared at a concentration of 0.2 mg/mL and filtered using 0.2 µm syringe filters prior to measurement. Measurements were acquired in a 0.1 cm path length of the quartz cuvette at a scan rate of 6 nm/min.

4.3.7. Reversible addition-fragmentation transfer (RAFT) Polymerization

The molecular weight, polydispersity (Đ), and degree of polymerization (DP) of synthesized polyacrylic acid (PAA), quaternized poly(4-vinylpyidine) (qP4VP), polystyrene sulfonate (PSS), and poly(dimethyl aminoethyl acrylate) (PDMAEA) were determined by GPC and are shown in

Table 1 with traces shown in Figure B.3. HA was purchased from Lifecore Biomedical since HA is relatively difficult to obtain with narrow Đ (1H NMR provided in Figure B.3a). Prior to polymerization, 4-vinylpyridine, acrylic acid and 2-dimethylaminoethyl acrylate were passed through a basic alumina column to remove hydroquinone or MEHQ inhibitor. The purified monomers were clear in color. Polymerization of each of the monomers was performed by reversible addition fragmentation chain transfer (RAFT) polymerization to obtain dispersity Đ <

1.4 at 60-70 °C in a 200 mL Schlenk tube. Azobisisobutyronitrile (AIBN) was recrystallized in methanol to use as the radical initiator. Further details follow for each specific chemistry.

4.3.8. Gel Permeation Chromatography.

DMF GPC analyses were performed on a Waters HPLC system equipped with a Waters 1515

Isocratic HPLC Pump with two columns (ResiPore, 300 × 7.5 mm, up to 500k Da, Agilent

Technologies, CA) in series. DMF with 0.02M LiBr was used as the eluent with a flow rate of 1

132 mL min-1 at 70 °C. The detector system consisted of a Wyatt miniDAWN TREOS multi-angle light scattering detector and a Wyatt Optilab T-rEX differential refractive index detector.

Aqueous GPC analyses were run on an Agilent Technologies 1260 Infinity system using two

Aquagel columns in 0.5 M NaCl (aq) with 0.02% sodium azide as the mobile phase. The detector system consisted of a Wyatt miniDAWN TREOS multi-angle light scattering detector and a Wyatt

Optilab T-rEX differential refractive index detector.

4.3.9. PAA polymerization.

2-(Dodecylthiocarbonothioylthio)-2-methylpropionic acid (72 mg, 0.2 mmol) and AIBN (6 mg,

0.037 mmol) were added to a solution of acrylic acid (10 g, 139 mmol) in water/DMF co-solvent

(1:9 v/v) in the ratio of 700:1:0.2. The solution was mixed until the CTA and radical initiator were completely dissolved in the solvent. Then, the mixture was degassed by three freeze-pump-thaw cycles in liquid nitrogen and purged with N2. The polymerization was carried out in a sealed flask at 65 °C and terminated after 16 h by removal of heat and exposure to air. The polymer was purified by dialysis with a 3 kDa membrane. The molar mass and dispersity of the obtained polymer were confirmed by aqueous GPC (Table 4.1, Figure B.4). After confirmation of Mw, the polymer was stirred in 1 N NaOH for 3 hours to introduce sodium ions for neutralization, dialyzed to exchange to water, and lyophilized. 1H NMR confirmed full deprotonation (Figure B.3b).

4.3.10. qP4VP polymerization.

4-Cyano-4-(phenylcarbonothioylthio) pentanoic acid (Aldrich, 155 mg, 0.56 mmol) and AIBN

(recrystallized twice from methanol, 18.2 mg, 0.11 mmol) were added to a solution of 4- vinylpyridine (10 g, 100 mmol) in 100 g 1,4-dioxane in the ratio of 500:1:0.2. The solution was mixed in a 200 mL Schlenk reactor and then degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 60 °C and terminated after 12 h

133 by removal of heat and exposure to air. The polymer was then precipitated 3 times in hexanes and dried under vacuum. The molar mass of the obtained polymer was confirmed by DMF GPC (Table

1, Figure B.4). P4VP was quaternized with iodomethane (99%, Sigma-Aldrich, 13 mL) in N,N- dimethylformamide (DMF). The reaction mixture was stirred at room temperature for 24 h, purified by precipitation in diethyl ether, and dried under vacuum. The degree of quaternization was 100% as determined by 1H NMR (Figure B.3c).

4.3.11. PSS polymerization.

4-Cyano-4-(phenylcarbonothioylthio) pentanoic acid (22.5 mg, 0.08 mmol) and AIBN (2 mg,

0.016 mmol) were mixed in 10 mL of DMF. Sodium 4-vinylbenzenesulfonate (10 g, 0.048 mmol) was dissolved in 90 mL of 1 M NaCl solution. CTA and initiator solution was mixed in the monomer solution in a 200 mL Schlenk reactor. Monomer:CTA:initiator were in the ratio of

600:1:0.2. The solution was degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 60 °C under N2 gas and terminated after 16 h by removal of heat and exposure to air. The polymer was then dialyzed against water 7 times to get rid of monomer and dried under vacuum. Polymer molar mass was confirmed by GPC (Table

1, Figure B.4). 1H NMR provided in Figure B.3d.

4.3.12. PDMAEA polymerization.

For PDMAEA polymerization, 2-ethylsulfanylthiocarbonylsulfanyl-2-methylpropionic acid

(EMP) as a CTA was synthesized as following a previously reported procedure (details in SI).

EMP (31 mg, 0.14 mmol) and AIBN (4.6 mg, 0.028 mmol) were added to a solution of DMAEA

(10 g, 70 mmol) in 100 mL 1,4-dioxane in the ratio of 500:1:0.2. The solution was degassed by three freeze-pump-thaw cycles in liquid nitrogen. The polymerization was carried out in a sealed flask at 70 °C and terminated after 16 h by removal of heat and exposure to air. The polymer was

134 then precipitated 3 times in hexanes and dried under vacuum. Polymer molar mass was confirmed by GPC (Table 1, Figure B.4). 1H NMR in acidified conditions provided in Figure B.3e.

4.3.13. Turbidity measurement.

Protein and polymer samples were prepared at 2 mg mL-1 in 20 mM Tris buffer, pH 8.0, except for PDMAEA which was prepared at 6 mg mL-1 to maintain the same charge ratio. The concentration of protein was confirmed by absorption at 280 nm using an Implen P330 nanophotometer. Turbidity was used to measure the extent of complex formation as a function of charge stoichiometry and salt concentration. To vary the charge stoichiometry, the polymer solutions and the protein were mixed at ratios varying from 100% polymer to 100% protein in 10% increments of protein. Samples were prepared in triplicate in a 96 well plate with 200 µL as a total volume, and the percent absorbance was measured on a plate reader at 600 nm. Error bars on the plots represent the calculated standard deviation of the data. All combinations of the proteins and polymers were measured at 0, 50 and 300 mM of NaCl in 20 mM Tris buffer pH 8.0.

4.3.14. Zeta Potential and Dynamic Light Scattering (DLS) Measurements.

A Zetasizer nano series (Malvern Instruments) was used for the determination of zeta-potential (ζ) of GFP and polymer solutions as a function of different ionic strengths. A volume of 1 mL of protein and polymer solution in buffer solution (20 mM Tris pH8.0 and 0 to 300 mM of NaCl) was analyzed. The acquisition was for 4 seconds 5 times per sample and phase analysis light scattering

(PALS) frequency was 20 Hz. The proteins contained a single cysteine, so they were measured with 5 mM TCEP. The results were analyzed using the Smoluchowski model.

Dynamic light scattering (DLS) was performed on a Wyatt DynaPro Nanostar using a wavelength of 658 nm, beyond the optical absorbance of all proteins and polymers used. Data were acquired over 10 acquisitions of 5 s each. Samples with varying ratios of protein to polymer were prepared

135 at 2 mg mL-1 in Tris pH 8.0 with varying salt concentrations and filtered using 0.2 µm Whatman

Anotop 10 syringe filters prior to measurement to get rid of impurities. Samples were loaded into plastic 384 well plates for measurements. DYNAMICS V7 software was used for data analysis.

For LP and XLP at 20 mM Tris with 0 mM NaCl solution, the protein solutions precipitated prior to the measurement, so DLS was not performed. For the mixture of MP with qP4VP at 0 mM NaCl and the mixtures of MP, LP, and XLP with qP4VP at 50 mM NaCl, macrophase separation was observed, so DLS was not performed. SP had a weak turbidity signal with qP4VP, so DLS was performed on these mixtures.

4.3.15. Optical microscopy.

Samples that were seen to macrophase separate by turbidimetry were prepared at 2 mg mL-1 at the blending ratio where the peak turbidity was observed. After the samples were blended, they were placed on a slide with a coverslip. The samples were imaged with a Zeiss Axioplan microscope equipped with a 100x objective and an Axiocam 503 Mono camera under bright field.

4.4. Results and Discussion

4.4.1. Protein design, patch calculation and expression

To assess the effect of different charge patch size on protein interactions, superfolder green fluorescent protein (sfGFP; PDB ID: 2B3P) was chosen as a model protein because the protein is a robust folder, more resistant to denaturation than many proteins,51 has a known crystal structure,52 and has known plasticity at many amino acid (AA) positions that allows reconstruction of the protein surface.47,53 In this study, sfGFP protein was mutated to create a panel of mutants with differing electrostatic patch size on their surfaces. Mutation sites were selected such that they avoided alterations to structural features (beta barrel and alpha helix) of the proteins, and sites where mutations had previously been reported were preferred.21,47 Selected AAs were mutated to

136 be either positively (Lys and Arg) or negatively charged (Asp and Glu) to create or enlarge charge patches on sfGFP, and mutated to be neutral (but not hydrophobic) to decrease or remove charge patches. Then, the solvent-accessible surface (SAS) of the mutants was visualized via PyMOL.

The open-source Adaptive Poisson–Boltzmann Solver (APBS)48 was then applied to assess the charge distribution of each sfGFP mutant by allowing direct visualization of the electrostatic potential at the SAS of the mutants under various NaCl concentrations at pH 8.0. A pH of 8.0 was chosen for this study to prevent intermolecular interactions by cation pi-interaction, hydrophobic interaction, etc., and to avoid isoelectric point precipitation, while also minimizing the net charge of the protein to enable assessment of the effect of patchiness without convolution due to strong net charge.

Four mutants, small patch (SP), medium patch (MP), large patch (LP), and extra-large patch (XLP) were successfully designed such that their net charges were all between -1.5 and -1.8 at pH 8.0 and isoelectric points were between 7.3 and 7.6 per calculation with PROPKA3,54 with

SP being the least “patchy” and XLP the most. Native sfGFP was not tested because it did not match these strict criteria. Maintaining the same net charge across all the mutants was necessary to allow direct assessment of the effect of charge patch size. The amino acid sequences of the mutants are provided in Figure 4.1 and the mutation sites of AA are marked in red. DNA sequences of the mutants are provided in Figure B.2. Figure 4.2 shows the surface potential (q) at the SAS of the designed mutants, with positive potentials shown in blue and negative in red. Visually, the mutants have increasingly large patches as designed.

To quantitatively assess the patchiness of each mutant, an algorithm for identifying and characterizing the charge patches at the surface of the proteins was developed. Contiguous regions

-1 with the magnitude of the potential greater than 2 kT ec were considered patches if the area of the

137 region was greater than 1 nm2. The charge patches, average patch area, and total neutral area of each mutant at various NaCl concentrations and pH 8.0 are shown in Figure 4.3. Table S1 shows the number of patches found on each mutant, the area and average potential of each patch, and the neutral area on each mutant. As expected, increased NaCl concentration led to decreasing patch size in all mutants due to charge screening at the protein surface. The average patch area increased with increased expected patchiness, while the number of patches decreased. Also, the neutral area was found to decrease with increased charge patchiness, likely because of the reduced interfacial area between patches as the number of patches decreased. Visual inspection of the electrostatic field isosurfaces (± 2 kT e-1) of the mutants (Figure B.5) shows that the mutants with the largest patches have electrostatic fields that extend out further from the SAS than those with smaller patches, indicating that they have a greater potential for intra- and interspecies interactions. The fields shrink with increased ion concentration as expected suggesting that these interactions will change with salt concentration. These trends suggest that the panel of mutants was successfully designed to have varying charge patch sizes and patch distribution.

138

Figure 4.1. Amino acid sequences of sfGFP mutants with mutation sites indicated in red.

139

-1 Figure 4.2. Different views of the electrostatic surface potentials (±2 kT ec ) of GFP mutants at the solvent accessible surface rendered from solutions of the linearized Poisson–Boltzmann equation using the Adaptive Poisson-Boltzmann Solver (APBS). The theoretical net charge was the same for all mutants (-4 at pH 8.0).

140

Figure 4.3. a) Charge patches identified by search algorithm for each GFP mutant at 0, 50, and

300 mM NaCl, with negative patches in and positive patches in blue. b) The average positive and negative patch areas of each mutant. c) The neutral area of each mutant where surface potential was less than |2 kT e-1|.

141

To further quantify the charge patchiness of the protein surface, a patchiness correlation parameter was developed, inspired by Wang and Swan’s work with colloidal mixtures of random patchy spheres.55 The patchiness parameter represents the charge correlation between neighboring sites on the protein surface. The parameter was calculated by discretizing the protein surface and assigning each point on the grid to be positive, negative, or neutral based on a user-defined threshold value. Nearest neighbor interactions were then assigned a value of 1, 0, or -1. A value of 1 was assigned if both sites had a potential of the same sign, 0 was assigned if either site was neutral (between the positive and negative threshold values), and -1 was assigned if the sites had opposite potentials. The patchiness parameter is the average of this nearest neighbor interaction parameter. Under this algorithm, a completely positively or negatively charged surface would give a value of 1, a completely neutral surface or a perfectly alternating charge-neutral surface would give a value of 0, and a perfectly alternating positive-negative surface would give a value of -1.

This patchiness parameter was found to increase from SP to XLP as expected and decrease with increasing NaCl concentration again due to charge screening (Figure 4.4). The advantage of this methodology over using the relative positions of the amino acids is that here only the surface of the protein is considered, and the internal electrostatics are ignored, which allows for clearer determination of the likely interactions between the protein surface and surrounding species.

142

Figure 4.4. Patchiness parameter calculated for the panel of sfGFP mutants used in the study at pH 8.0 in varying salt concentrations (0, 50, and 300 mM).

All GFP mutants were successfully expressed and purified as shown in Figure 4.5. Molar masses of the expressed mutants were shown to match theoretical values by MALDI-TOF and

SDS-PAGE analysis (Figure 4.5a,b). The presence of large positively charged patches on XLP and

LP likely led to their decreased electrophoretic mobility compared to SP and MP, and the slightly larger positive patch on SP may have contributed to the slight difference in the mobility of SP compared to MP. Additionally, it has been demonstrated in silico that the concentrating particle charge to a patch as opposed to distributing it across a surface can lead to reduced electrophoretic mobility.56 Protein purity estimated from analysis of the SDS-PAGE results indicated a purity of approximately 90%, but saturation of the protein bands indicates this is likely an underestimation

(Figure B.6). Circular dichroism showed that all GFP variants exhibited similar secondary structure, suggesting that this structure was maintained across the mutations (Figure 4.5c). The mutants were shown to have active fluorophores by UV-vis spectroscopy (Figure 4.5d), with a major absorption peak at 491, 488, 485, and 490 nm for SP, MP, LP, and XLP, respectively, and

143 a minor peak at 396 nm for SP and XLP. The presence of the minor peak in the spectra of SP and

XLP is indicative of protonation of the active site.39

Figure 4.5. Properties of GFP mutants. (a) MALDI TOF of GFP variants (SP: 28.5 kDa, MP: 27.8 kDa, LP: 28.8 kDa, and XLP: 28.7kDa), (b) stained SDS-PAGE gel, (c) Circular dichroism spectra of GFP variants, and (d) UV-vis spectra of mutants at 1 mg mL-1, with inset image of UV fluorescence of purified GFP variants (2 mg mL-1).

4.4.2. Protein complexation with polyelectrolytes at various NaCl concentrations

To explore the effects of charge patchiness on coacervation, five polymers with varying polyelectrolyte strength and charge were chosen for complexation studies. Poly(4-

144 methylvinylpyridinium iodide) (qP4VP) was chosen as a model strong polycation, poly(styrene sodium sulfonate) (PSS) as a model strong polyanion, poly(2-dimethyl amino ethyl acrylate)

(PDMAEA) as a model weak polycation, poly(acrylic acid) (PAA) as a model weak polyanion, and hyaluronic acid (HA) as a model biopolyelectrolyte. These polymers were synthesized by reversible addition-fragmentation chain-transfer polymerization (RAFT), except HA which was purchased. Molar masses (Mw), degree of polymerization (DP), polydispersity (Đ), expected number of charges, and polymer structure data are shown in Table 1. Rg of hyaluronic acid was

0.596 57 estimated based on the empirical formula 푅푔 = 0.0275(푀 ), where M is the molar mass. Rg

2 퐶∞푁푙 of the other polymers was estimated as 푅 = √ , where C∞ is Flory’s characteristic ratio, N 푔 6 is the number of backbone bonds, and l is the bond length. The C∞ of qP4VP was estimated as

58 approximately 14, the C∞ of PSS was approximated as that of polystyrene (9.54), and the C∞ of

PAA and PDMAEA was approximated as that of poly(methyl acrylate) (7.91). For comparison,

59 the Rg of sfGFP is approximately 2 nm. The Rg of the polyelectrolytes is likely to change as a function of the salt concentration and as a result of complexation.60

145

Table 4.1. Summary of Polymer Properties

Polymers Mw DP Đ Expected # Approximate Polymer Structure (kDa) of charges Rg (nm)

HA 190 473 1.20 473 38.5

PAA 33 454 1.34 454 5.3

qP4VP 64 411 1.07 411 6.7

PSS 70 340 1.07 340 5.1

PDMAEA 25 173 1.36 29 3.3

The GFP mutants were all found to have negative ζ-potentials, except SP, which was near neutral (Figure 4.6a). The increasing patchiness from SP through XLP correlated with increasingly negative ζ-potential, and increasing salt concentration led to ζ-potential approaching neutrality for all mutants, as expected. The negative potential is expected due to the slightly negative net charge on the mutants. The polymers all showed appropriate zeta-potentials— PDMAEA and qP4VP were positive and PSS, PAA, and HA were all negative—however, there appears to be a negative bias on the measurements, as the potential of qP4VP would be expected to be highly positive, and the potential of PDMAEA at least slightly positive and not neutral (Figure 4.6b). This may be due to the nature of the measurement, which does not directly measure the potential of the molecule of interest, but also includes some layers of condensed ions and solvent. 146

Figure 4.6. Zeta-potentials of a) the GFP mutants and b) the synthesized polymers in Tris buffer pH 8.0 with different concentrations of NaCl (0, 50 and 300 mM).

Strong macrophase separation was observed between the GFP mutants and qP4VP by turbidimetry (Figure 4.7), with the strength of the complexation increasing with increased patchiness and more negative ζ-potential of the protein. The other polymers did not show strong macrophase separation. Complexation was measured as a function of both the concentration of

NaCl in the buffer and the mixing ratio of the protein and polymer (fprot), defined as:

푀푝푟표푡 푓푝푟표푡 = (4.1) 푀푝표푙푦+푀푝푟표푡

where Mprot in the volume of protein solution added and Mpoly is the volume of polymer solution added. This ratio was used in place of the more traditional metric, the positive charge fraction, because the protein was near net neutral. Protein and polymer solutions were all 2 mg mL-1, except

-1 PDMAEA which was at 6 mg mL to increase the charge ratio due to its proximity to pKa (pKa ≈

7.32 ± 0.05, Figure B.7). SP showed weak complexation at both 0 mM and 50 mM NaCl with the peaks at fprot of 0.9, and no complexation at 300 mM NaCl. MP showed strong complexation at both 0 mM and 50 mM NaCl with peaks at fprot of 0.8 and 0.9, respectively, and no complexation

147 at 300 mM NaCl. LP and XLP precipitated in the absence of NaCl, so data were not collected for these mutants at 0 mM NaCl. Both of these mutants showed strong complexation with qP4VP at

50 mM NaCl with peaks at fprot of 0.8 and 0.9, respectively, and no complexation at 300 mM NaCl.

The macrophase separation behavior of the mutants appeared to correlate with the ζ-potential and the patchiness because the three mutants with negative ζ-potential at 0 and 50 mM NaCl and the larger patchiness parameters, MP, LP, and XLP, were shown to strongly macrophase separate with qP4VP, while SP, which had a near neutral ζ-potential and the smallest patchiness parameter, only weakly phase separated. As predicted by coacervation theory, the complexation weakened with increasing salt concentration due to electrostatic screening effects, which led to no interactions occurring at 300 mM NaCl. All the mutants also apparently violate the empirical rule set by

Obermeyer, et al. that coacervation/complexation would not be expected below a negative charge ratio (α)—defined as the ratio of negatively charged residues on the protein to positively charged— of 1.2, suggesting that the patchiness had a strong influence on the complexation behavior of the mutants.26 The α-values of the mutants are shown in Table S2.

148

Figure 4.7. Turbidity profile of GFP mutants as a function of NaCl concentration and protein-

푀푝푟표푡 polymer fraction (푓푝푟표푡 = , where Mprot in the volume of protein solution added and 푀푝표푙푦+푀푝푟표푡

Mpoly is the volume of polymer solution added).

To better understand the nature of the macrophase separation with qP4VP, the samples that phase separated were observed by optical microscopy at the point of maximal complexation

(Figure 4.8). SP was found to only weakly complex with qP4VP at both 0 and 50 mM NaCl and fprot of 0.9, forming sparse precipitates as seen by both microscopy and the weak turbidity peak.

MP was found to strongly phase separate and form solid precipitates at both 0 and 50 mM NaCl at fprot of 0.8 and 0.9, respectively. Pure LP and XLP (fprot = 1) at 0 mM NaCl were shown to self-

149 complex and precipitate out. At 50 mM NaCl, both LP and XLP formed complex coacervates with qP4VP at fprot of 0.8 and 0.9, respectively, based on the formation of coalesced droplets observed by microscopy. At even the lowest polymer loading tested (fprot = 0.9), there are only sufficient negatively charged residues to associate with 60% of the P4VP monomers or approximately a 2- fold excess, with each polymer molecule interacting with approximately 6-7 proteins. The excess of both positively-charged monomers and residues indicates that counterions and charge screening play significant roles in achieving complexation in these systems. In the case of LP and XLP, the large negative patches likely allow for greater conformational freedom of the interacting chain or chains, leading to the liquid complex coacervates, whereas the smaller patches of SP and MP do not allow the conformational freedom, leading to the solid complexes. In the case of the liquid coacervates, multiple polymers or functionally independent stretches of the same polymer are likely interacting over short runs with the patches, whereas with the solid precipitates, a single polymer likely becomes conformationally locked interacting with each small patch.

While turbidimetry provides a measure of macrophase separation, the single phase regions may contain soluble protein-polymer complexes. Dynamic light scattering (DLS) was used to examine the presence of these complexes, illustrating regions of complexation beyond the regions of macrophase separation (Figure 4.9). SP was confirmed to complex with qP4VP at 0 and 50 mM

NaCl (Figure 4.9). XLP was shown to form soluble complexes with both weak polyanions, PAA and HA, over a wide window (fprot = 0.2-0.5) at 50 mM NaCl, and at a single point at 300 mM

NaCl (fprot = 0.4). The stronger polyanion, PSS, was never shown to complex with the mutants.

This difference may be because of the ability of PAA and HA to charge regulate, a phenomenon which can cause the pKa of functional groups to shift by up to 2 units depending on the local environment.26,33,61 No other polymer-protein combinations showed significant levels of

150 complexation. The inability of PDMAEA to complex with XLP was likely driven by its low net charge due to the proximity between the pH and the pKa. The low net charge likely made it both less entropically favorable, since there were fewer counterions that could be shed, and less enthalpically favorable, since the interaction strength would have been weaker, for complexation to occur.

Figure 4.8. Optical micrographs showing protein-polymer complexes resulting from mixing the

푀푝푟표푡 GFP mutants with qP4VP at the maximum point of turbidity. 푓푝푟표푡 = , where Mprot in 푀푝표푙푦+푀푝푟표푡 the volume of protein solution added and Mpoly is the volume of polymer solution added. Scale bar is 10 µm.

151

Figure 4.9. DLS data of GFP mutants with polyelectrolytes at different volume fractions and different concentrations of NaCl (0, 50 and 300 mM) in 20 mM Tris buffer (pH 8.0). 푓푝푟표푡 =

푀푝푟표푡 , where Mprot in the volume of protein solution added and Mpoly is the volume of 푀푝표푙푦+푀푝푟표푡 polymer solution added.

Small-angle neutron scattering (SANS) measurements of the XLP-PAA blends shown to form soluble complexes in the DLS experiment revealed that the complexes had a cylindrical morphology (Figure 4.10). Measurements were taken at the fprot value as the center of the peak

-1 observed by DLS at 10 mg mL in Tris buffered D2O (pH 8.0). The use of D2O may slightly change the behavior of the complexes, but it must be used to achieve sufficient signal given the dilute conditions. The data were fit to a cylinder model,62-64 shown below:

152

1 1 퐼(푞) = ∫ 퐹2(푞, 푢)푑푢 + 퐵 (4.2) 푉 0 where

1 sin( 푞퐿푢) 2 2 퐽1(푞푅√1−푢 ) 퐹(푞, 푢) = 2(∆𝜌)푉 1 (4.3) 푞퐿푢 푞푅√1−푢2 2 and V is the volume of the cylinder (πR2L), B is the background intensity, Δρ is the scattering length density difference between the scatterer and the solvent, and J1 is the first order Bessel function. The cylinder model was chosen because of the q-1 scaling of I(q) when q was less than the inverse of the Rg of the protein (~2 nm), and because rod-like polyelectrolyte-protein complexes have been observed previously in similar systems.65-66 The complexes were found to have radii (R) of 1.97 and 1.76 nm and lengths (L) of 779 and 875 nm at 50 and 300 mM NaCl,

-1 respectively. At the concentration required for SANS, 10 mg mL , SP-qP4VP blends at fprot of 0.9 in buffered D2O formed insoluble complexes, and a XLP-HA blend at fprot of 0.35 formed aggregates.

Figure 4.10. Small-angle neutron scattering (SANS) curves of XLP-PAA blends with fprot of a)

0.35 at 50 mM NaCl and b) 0.4 at 300 mM NaCl in Tris buffered D2O pH 8.0. SANS data are

153 solvent corrected and fitted to a cylinder model62-64 with radii of 1.97 and 1.76 nm and lengths of

779 and 875 nm at 50 and 300 mM NaCl, respectively.

The complexation behavior between the various polyelectrolytes and the GFP mutant panel correlated well with the patchiness parameter (Figure 4.11). At patchiness values greater than approximately 0.6, the mutants were found to be insoluble (LP and XLP at 0 mM NaCl). When complexing with qP4VP (Figure 4.11a), a patchiness value of approximately 0.45 to 0.55 led to the formation of complex coacervates (LP and XLP at 50 mM NaCl) and a value of approximately

0.2 to 0.35 led to the formation of solid precipitates (MP and SP at 0 and 50 mM NaCl). At 300 mM NaCl, LP and XLP had patchiness values of 0.25 and 0.3, respectively, but the high salt concentration likely led to screening effects the prevented complexation. Only XLP complexed with PAA (Figure 4.11b) with patchiness greater than approximately 0.3 (XLP at 50 and 300 mM

NaCl). LP at 50 mM NaCl had a patchiness parameter of 0.45, but did not complex with PAA.

This difference in behavior was likely driven by the fact that the average positive patch area of LP was nearly half that of XLP (Figure 4.9a). Complexation with HA required a patchiness of greater than approximately 0.5 (Figure 4.11c), so only XLP at 50 mM NaCl complexed with it.

Complexation was not observed with PDMAEA or PSS. The strongest polyanion, PSS, never complexed with the mutants, while both HA and PAA formed only soluble complexes. This difference may be because of the ability of PAA and HA to charge regulate, a phenomenon which can cause the pKa of functional groups to shift by up to 2 units depending on the local environment.26,33,61 Because PSS cannot charge regulate, the repulsion between the polymers themselves and the repulsion of the polymer with the negative patches on the protein may be too large for complexation to occur, whereas both PAA and HA can charge regulate allowing the polymers to modulate their charge near molecules or patches of the same charge, reducing the

154 repulsion and allowing complexation to occur. Additionally, the ability of carboxylic acid groups to form salt bridges with exposed amines may aid the complexation of PAA and HA with the model proteins.67 The inability of PDMAEA to complex with XLP was likely driven by its low net charge due to the proximity between the pH and the pKa. The low net charge likely made complexation both less entropically favorable, since there were fewer counterions that could be shed, and less enthalpically favorable, since the interaction strength would have been weaker.

Figure 4.11. Phase behavior of GFP variants with a) qP4VP, b) PAA, and c) HA at the point of maximal complexation as a function of the patchiness of each protein at 0, 50, and 300 mM NaCl.

155

4.5. Conclusions

Using a panel of four sfGFP mutants engineered with varying surface charge distributions, the effect of charge patchiness of the protein surface on coacervation was explored. The patchiness of the mutants was determined using potential mapping software and quantified using a patchiness parameter. The mutant with the highest patchiness, XLP, showed the strongest complexation behavior, and the one with the lowest, SP, showed the weakest. Complexation behavior correlated with the patchiness parameter.

The strongest complexation was observed with the strong polycation, qP4VP, which was the only system in which macrophase separation occurred. Three types of phase behavior were observed in the qP4VP-GFP system, precipitation of the protein (patchiness greater than approximately 0.6), complex coacervation (patchiness of approximately 0.45 to 0.55), and precipitation (patchiness of approximately 0.20 to 0.35) with no complexation occurring at 300 mM NaCl likely due to charge screening effects. The slight negative charge on the mutants also likely contributed to their complexation with qP4VP. Soluble complexes were found to form between the two weak polyanions (HA and PAA) and the mutant with the highest patchiness, XLP, but with no others. This was attributed to both the high patchiness and XLP’s large average positive patch area compared to the other mutants. The model strong polyanion, PSS, did not complex with any of the mutants. The difference in the behavior of the strong and weak polyanions is likely driven by the ability of the weak polyanions to charge regulate. The weak polycation, PDMAEA, did not complex with any of the mutants either, likely due to its low charge density compared with the other polymers.

Overall, patchiness is a predictor for whether the mutants would complex and of the nature of the complexes. Additionally, proteins were demonstrated to be a sequence-controlled model

156 system that enabled the study of charge patchiness, and could be used to study novel colloidal systems. This sequence control can be further leveraged to tune the complexation behavior of proteins in systems of interest. Further study through simulation or theory will help to enable further quantification of the effect of patchiness, and is necessary to create predictive algorithms for the effect patchiness will have on coacervation, especially when convoluted with the effects of other protein properties.

157

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Chapter 5. Polyelectrolyte Complexation Driven Morphological Changes in Cationic-

Neutral Block Copolymer Thin Films

5.1. Abstract

Despite the widespread use of block copolymers for the encapsulation of charged cargo, the effect of polyelectrolyte ionic complexation on block copolymer morphology, especially in solid-state materials, remains poorly understood. This study aims to understand how ionic complexation affects the morphology of solid-state BC thin films as a function of the charge density of the BC and the polyelectrolyte strength of the added macromolecules. The interactions of a model cationic-neutral block copolymer, poly((oligo(ethylene glycol) methacrylate)-b-(4-vinyl pyridine)) (POEGMA-b-P4VP) that was methyl quaternized to 20 and 40%, were studied with four model polyanions of varying strength: poly(styrene sulfonate) (PSS), poly(acrylic acid)

(PAA), and native and supercharged α-amylase. PSS caused precipitation of both the BCs and analogous P4VP homopolymers. PAA followed classic complex coacervation behavior with the

P4VP homopolymers and was found to induce micellization in the BCs. The addition of PAA past the point of charge balance led to a degradation of the nanostructure of the films due to swelling of the micellar cores. The PSS is thought to be able to cause a pKa shift in P4VP, leading to stronger interactions between PSS and P4VP thus causing the precipitation, unlike PAA which is a weaker polyanion and exhibited classic complex coacervation, forming complex coacervate core micelles

(C3Ms) as expected. The addition of amylase led to more complex microphase separation behavior likely driven by the ability of both blocks to strongly interact with the protein. While at low protein loading the protein appears to segregate into large complexes surrounded by excess block copolymer, additional protein loading leads to a microphase separated, disordered structure similar

163 to that of the block copolymers alone. If further protein is added, microphase separation is reduced.

The protein behaved differently than PAA because of the greater conformation freedom of PAA, the lower charge density, and the affinity of proteins to polyethylene glycol functionality.

5.2. Introduction

Block copolymers (BCs) have generated wide interest in fields including medicine, biomaterials, lithography, and energy due to their ability to self-assemble and their ability to combine functionality with favorable physical properties.1-6 BCs can serve as a template for the nanopatterning of proteins and other biomolecules or as an encapsulant.5,7 The grafting of a polymer directly to the protein or biomacromolecule of interest enables BC-like self-assembly of the materials allowing for the production of highly active biocatalysts and biosensors.2-4,8 Adding salt to a BC or directly incorporating an ionizable group into a BC to create an ionic-neutral BC enables the creation of specialty materials for applications including battery materials, selective membranes, home goods, functional biomaterials, and drug delivery.7,9-11 When the BC itself is charged, the charged block can act as a selective nanoscale pore in a membrane and can be used to encapsulate surfactants, proteins, drugs and other molecules via polyelectrolyte complexation.7,11-14

Polyelectrolyte complexation has been proposed as a facile method for the production of film materials, protein encapsulation, drug delivery, and adhesives.7,13-20 Polyelectrolyte complexes (PECs) encompass all phases produced by electrostatic interactions between macromolecules including precipitates, complex coacervates, and micelles, each of which have favorable properties. Precipitation has been used to create easily processible and recyclable solid materials with high bioresistivity, favorably low gas permittivity, and as a purification method for proteins.15 Complex coacervates and C3Ms have been widely used to enable technologies

164 including protein encapsulation and stabilization, the formulation of effective adhesives, and drug delivery.5,14,16-17,21-22 In order to produce C3Ms, a charged-neutral BC must be used in place of one or more of the macromolecules, and the properties of these BCs can influence the morphology of the PECs formed.12,23 Coating these PECs onto a surface enables the immobilization of proteins or other target molecules onto a surface.20,24 This can be advantageous, especially in the case of functional proteins and enzymes, where immobilization enables the conversion of the solvated macromolecule into a solid materials necessary for biosensors and biocatalysts.4,8,25-27 The structure of these materials in the solid-state is not well-understood, but structure–function relationships in block copolymer based materials suggest that the structure of such films may play a strong role in performance,4,8,28 and the structure of the charged-neutral block copolymers alone may provide insight into the self-assembly of these materials.

While typically the nanostructure of BCs is governed by the volume fractions of the blocks, the molecular weight, and the Flory-Huggins interaction parameter between the monomers (χ), the addition of salts or an ionizable block can cause a shift in the phase diagram of the BCs that depends on the salt loading or the degree of ionization.9-10,29-30 When one of the BC blocks is ionizable, the degree and method of ionization can alter the effective value of χ, allowing for much greater tuning of the nanostructure of these materials in both thin films and solid-state.9-10 In the case of salt addition, the shift in the phase behavior is driven by two competing factors, the entropically favorable uniform distribution of the ions between the blocks, which leads to increased miscibility of the blocks and an effective decrease in χ, and the energetically favorable solvation of the ions in the block with the higher dielectric constant, which leads to decreased miscibility of the blocks and an effective increase in χ.6,29,31-32 In polyethylene-b-poly(ε- caprolactone) (PEO-b-PCL), the inclusion of LiClO4 has been demonstrated to induce microphase

165 separation, with χ increasing linearly with salt concentration at low concentrations when the solvation effect dominates, but decreasing at higher concentrations as the entropic effect begins to dominate.29,32 In polystyrene-b-polyethylene oxide (PSS-b-PEO), the addition of lithium salts has been demonstrated to increase the effective χ between the blocks, leading to a shift in the phase behavior.31 The traditional BC phase diagram can be replotted for these systems replacing χ with the salt loading.32

In the case of PECs, when the additive is a polyelectrolyte or polyampholyte as opposed to a simple salt, the self-assembly behavior of the systems has been studied extensively in dilute solution12,33-34 but as mentioned previously is not well understood in the solid state. The nature and morphology of the PECs that form between two or more macromolecules is a function of the solution properties such as salt and macromolecule concentrations and pH and of the properties of the macromolecules themselves including molecular weight, block ratio, polyelectrolyte strength, and charge density.12,35 In soluble PECs, where at least one of the macromolecules is typically a charged-neutral BC, these morphologies include micelles, vesicles, and lamellae with transitions between these phases being dictated by the above variables.12,33 These soluble PECs can be coated to make insoluble, but solvent-permeable thin films in order to increase the utility of PECs to applications where surface immobilization of a target molecule is favorable, such as biosensing or biocatalysis.20 These C3M thin films have been demonstrated to stabilize proteins and can be synthesized with an all-aqueous process, but the phase behavior of these films is not fully understood.20

The goal of this study is to determine how polyelectrolyte complexation affects the thin film morphology of ionic-neutral block copolymers. Model polyanions of differing polyelectrolyte strength have been blended with BCs of differing charge density to understand how both the

166 macromolecule properties of all components and the loading of anion in the BC affect the morphologies of these films. The coacervation behavior of each polyanion was tested via turbidimetry with the homopolymer analog of the ionic block of the BC, and the solution-state phase behavior of the BC-polyanion blends was analyzed by dynamic light scattering. Thin films cast from the BCs and BC-polyanion blends were analyzed via X-ray scattering and microscopy methods to assess changes in nanostructure that occurred as a result of polyelectrolyte complexation.

5.3. Results and Discussion.

5.3.1. Materials Design.

To assess the effect of polyelectrolyte complexation on the morphology of ionic-neutral BC thin films, four model polyanions of varying polyelectrolyte strength were blended with two block polymer encapsulants of varying charge density. The model polymer encapsulant chosen for this study was partially quaternized poly(ethylene glycol methyl ether methacrylate)-block-poly(4- vinyl pyridine) (POEGMA-b-pqP4VP), and the model polyelectrolytes were poly(styrene sulfonate) (PSS), poly(acrylic acid) (PAA), and amylase, a model protein. The BC encapsulant was quaternized to 20 and 40 mol% of P4VP monomers by reaction with iodomethane (POEGMA- b-pqP4VP20 and POEGMA-b-pqP4VP40, respectively), which led to a disordered, microphase- separated morphology (Figures 5.1 and 5.2). A degree of quaternization near to the order-disorder transition was specifically selected to enable probing the effect of complexation on ordering. A summary of relevant properties of the model polyanions is given below in Table 5.1. PSS was considered to be the strongest polyanion due to its low pKa, while PAA and native and supercharged amylase variants were considered weak polyanions due to their relatively higher pKa/pI values. The model protein was expected to behave differently from the synthetic weak

167 polyanion because proteins tend to have a much higher mass-to-charge ratio that synthetic polyelectrolytes, which was hypothesized to have a strong effect on the coacervation behavior. To explore how the mass-to-charge ratio of the protein affects its coacervation behavior, the protein was supercharged to different degrees by reaction with succinic anhydride.

- Table 5.1. Structures, pKa/pI, and mass-to-charge ratios (m/z ) of model polyanions

- Polyanion Abbreviation Structure pKa (pI) m/z

Polystyrene PSS ~1 206.2 sulfonate

Polyacrylic acid PAA ~4.5 94.1

Native Amylase A22 (~4.7) 2478 A30 (~4.5) 1814 Supercharged A32 (~4.5) 1704 Amylase A37 (~4.4) 1502

5.3.2. Complexation with PSS.

Complexation of the POEGMA-b-pqP4VP with PSS led to at least partial precipitation across all concentrations tested. For this reason, films of POEGMA-b-pqP4VP and PSS could not be reliably cast. This result was further verified via bulk coacervation studies between partially quaternized

P4VP homopolymer and PSS that showed the formation of flocculants across a wide range of mixing ratios (Figure C.1). These flocculants likely transition to complex coacervates in the presence of salt;36 however, salt cannot be added to the mixtures being coated because it crystallizes at the surface of the films.

168

As both quaternized P4VP and PSS are strong polyelectrolytes, and precipitation has been observed in a similar system, the precipitation with the partially quaternized P4VP is not unexpected. Furthermore, electrostatic interactions between PSS and the non-quaternized 4VP monomers are enabled by the ability of strong polyelectrolytes to induce effective pKa shifts of up

13,37-38 to 2 pH units in polymers they interact with, leading to an induced charge. Thus, PSS could

39 shift the pKa of the non-quaternized 4VP monomers (approximately 5) to near neutral and induce a greater positive charge on the P4VP, as illustrated in Figure C.2. This would enable the PSS to interact with longer runs of the P4VP block, which has been demonstrated to enhance complexation behavior.

5.3.3. Complexation with PAA.

In dilute solution (2 mg mL-1), PAA and P4VP homopolymer that was quaternized to 20 and 40%

(pqP4VP20 and pqP4VP40, respectively) formed complex coacervates based on turbidity measurements (Figure 5.1). The blending ratios of the polymers are expressed in terms of the positive charge fraction (f+), where

푀+ 푓+ = (5.1) 푀++푀− and M+ and M− are the total positive and negative charge of the macromolecules in the system.

The point of maximal complexation was found to be at f+ of approximately 0.45 for both levels of quaternization indicating that P4VP-PAA coacervation behaves as expected with the point of maximal coacervation occurring near the theoretical point of charge balance (f+ = 0.5) based on the peak of the turbidity curves in Figure 5.1. Additionally, in solution, the POEGMA-b-pqP4VPs and PAA formed micelles at the theoretical point of charge balance (f+ = 0.5). Dynamic light scattering (DLS) measurements at this blending ratio showed that the majority of material in

169 solution was incorporated into the complexes of hydrodynamic radius greater than 10 nm (Figure

C.3).

Figure 5.1. Turbidimetry data for the complex coacervation of P4VP 20q and 40q with PAA at 2 mg mL-1 in water. The point of maximal complexation is at f+ = 0.45-0.5.

Thin films of POEGMA-b-pqP4VP20 showed a transition from a disordered, microphase- separated nanostructure in the pure BC film to a micellar nanostructure when blended with PAA, with the strongest ordering occurring at the point of charge balance between the polymers. Non- uniform micelles upon addition of 10% PAA relative to the P4VP on a molar basis (% PAA), or f+ = 0.67, based on both the AFM and GISAXS data in Figure 5.2. The emergence of the micelles is apparent from the AFM phase image and emergence of the peak in GISAXS. These micelles become more uniform at the point of charge balance, 20% PAA and f+ = 0.5, as seen in the AFM

+ data and by the sharper peak in GISAXS (Figure 5.2). Further addition of PAA to 40% PAA (f =

0.33) did not significantly change the nanostructure based on AFM but did lead to a stronger peak in GISAXS. At 80% PAA (f+ = 0.2) the micellar structure displayed a decrease in the degree of ordering as demonstrated by the broadening of the peak in the GISAXS data. 1-D analysis of the

GISAXS data confirms the emergence of a peak at 10% PAA, the sharpening of the peak at 20 and

40% PAA, and broadening at 80% PAA (Figure C.4).

170

Figure 5.2. AFM height and phase scans and GISAXS patterns for films of PAA and POEGMA- b-pqP4VP20 at various molar ratios of PAA to P4VP. The addition of PAA to the BC triggers the formation of a micellar phase. At the lowest loading of PAA (f+ = 0.67), the micelles are non- uniform as seen from both the defects in the AFM images and the broadness of the GISAXS peak.

At higher loading of PAA (f+ = 0.5 and 0.33), near the point of charge balance, the micelles become more uniform as seen from both the AFM and the sharpening of the GISAXS peak. At the highest loading of PAA (f+ = 0.2), the micelles become swollen and non-uniform as seen from both the

AFM and the broadening of the GISAXS peak. Figure C.4 provides 1-dimensional linecuts of the

푀+ GISAXS data for further clarity. The positive charge fraction, 푓+, is defined as 푓+ = and 푀++푀−

M+ and M− are the total positive and negative charge of the macromolecules in the system. Scale

-1 bars are 200 nm. GISAXS patterns’ x-axes range from qy of -0.5 to 0.5 nm and y-axes range from

171

-1 qz of 0 to 1 nm . Height z-scales are 7, 4, 5, 3.5, and 7 nm. Phase z-scales are 27°, 8°, 35°, 8°, and

30°.

Thin films of POEGMA-b-pqP4VP40 with PAA showed similar behavior to that of

POEGMA-b-pqP4VP20. A transition from the disordered, microphase-separated pure BC film to uniform micelles occurred upon addition of 20% PAA (f+ = 0.67) based on both the AFM and

GISAXS data in Figure 5.3. However, in the AFM scans, there are apparent defects where the BC has not fully transitioned to micelles indicated by the presence of the elongated structures observed in the phase image. The micelles became more uniform at the point of charge balance, 40% PAA

(f+ = 0.5), without the presence of the defects seen at 20% PAA. Further addition of PAA to 80%

+ PAA (f = 0.33) degraded the uniformity of micellar nanostructure as demonstrated by the broadening of the peak in the GISAXS data. 1-D analysis of the GISAXS data confirms the emergence of a peak at 20% PAA, the sharpening of the peak at 40% PAA, and the degradation at

80% PAA (Figure C.4).

172

At the point of charge balance, the micelles become more uniform as seen from both the AFM and the sharpening of the GISAXS peak. At the highest loading of PAA (f+ = 0.3), the micelles become swollen and non-uniform as seen from both the AFM and the broadening of the GISAXS peak.

Figure C.4 provides 1-dimensional linecuts of the GISAXS data for further clarity. The positive

푀+ charge fraction, 푓+, is defined as 푓+ = and M+ and M− are the total positive and negative 푀++푀− charge of the macromolecules in the system. Scale bars are 200 nm. GISAXS patterns’ x-axes

173

-1 -1 range from qy of -0.5 to 0.5 nm and y-axes range from qz of 0 to 1 nm . Height z-scales are 4.5,

6, 8, and 14 nm. Phase z-scales are 14°, 45°, 25°, and 30°.

The order-disorder transition (ODT) observed in the POEGMA-b-pqP4VP thin films when combined with PAA is reminiscent of ODTs observed in salt-doped BC systems and similar to

ODTs reported in PEC systems.12,32 In the salt-doped BC systems, the salt preferentially dissolves into the block with the higher dielectric constant causing the volume fraction of that block to increase and causing an increase in the effective χ of the system, triggering an ODT and even phase changes when sufficient χ is achieved.32 In PEC systems in solution, the addition of an oppositely- charged polymer or BC to an ionic-neutral BC can trigger a transition from a simple soluble state to soluble complexes, i.e. micelles or other nanostructures.12 At higher concentrations, these PECs can become ordered, and when charged-neutral-charged triblock copolymers of opposite charges are combined, can even lead to the formation of ordered gels.40 In the POEGMA-b-pqP4VP and

PAA system, the PAA is expected to preferentially interact with the P4VP block due to the charge interaction and the entropically favorable release of counterions, similar to what is seen in PEC systems. This in turn causes increased ordering within the system. The apparent reentrant ODT at high polyanion loading is also reminiscent of an ODT seen in salt-doped BC systems when salt loading is increased and the entropic drive to more evenly distribute salt across the two phases takes over, leading to increased miscibility between the blocks. This apparent transition may also be a phase boundary with another PEC morphology, which can occur as a function of the mixing ratio, or simply be indicative of the polyanion loading being too high causing the micellar cores to overswell. This behavior has been observed in uncharged block copolymer-homopolymer blends where at higher fractions of the homopolymer loading, a disordered phase emerges.41-42

174

While PSS formed precipitates with the BCs, PAA was found to micellize with the BCs both in solution and in the thin films. This difference in behavior is attributed to the weaker association of PAA with the BCs compared to PSS. Whereas PSS could induce a charge on non- quaternized 4VP monomers, the PAA likely cannot induce the same pKa shift in non-quaternized

4VP. Additionally, PAA as a weak polyanion is known to be able to charge regulate at near neutral pH enabling the PAA to adjust to its local environment as necessary, enabling a reduction in repulsions between PAA monomers.

5.3.4. Complexation with Amylase.

In solution, complexation of amylase with pqP4VP20 and pqP4VP40 led to complex coacervation. The turbidity curve was found to be slightly asymmetric with peak complexation occurring at f+ of approximately 0.6 for all amylase variants. This asymmetry has been previously observed in protein-polyelectrolyte PEC systems and is thought to arise from either charge regulation at the protein surface or from conformational effects due to the folding of the protein.13,34,38,43 The pqP4VP20 showed higher turbidity at lower mixing ratios than the pqP4VP40

(Figure 5.4). The onset of coacervation shifts as a function of the supercharging of the protein for pqP4VP40, with the onset occurring at lower protein loading (closer to f+ = 1) with increased supercharging, while the onset of coacervation for pqP4VP20 was not dependent on the supercharging. The difference in the behavior between the two levels of quaternization is likely caused in part by the greater hydrophobicity of the pqP4VP20 compared to the pqP4VP40 at neutral pH, an effect that has been demonstrated to promote phase separation.44 In addition, it has been shown that the distribution of charges along a polymer plays a key role in determining its coacervation behavior because continuous stretches of charged monomers enable the formation of longer regions of interactions, or runs.45 The lower charge density of pqP4VP20 interrupts the

175 formation of longer runs of interactions regardless of the distribution of charges on the protein surface. The pqP4VP40 can form longer runs of interactions, and, as charges become more concentrated on the protein surface as a result of supercharging, the onset of coacervation occurs at lower protein loading because the proteins are able to interact more strongly with the polymers.

Figure 5.4. Turbidimetry data for the coacervation of pqP4VP20 and pqP4VP40 and the amylase variants at 2 mg mL-1 in water. The turbidity peak, which corresponds with the point of maximal complexation, is at f+ ≈ 0.6 for all variants.

In solution, complexes of amylase with POEGMA-b-pqP4VP20 showed slight increases in the hydrodynamic radii (RH) as a function of increasing protein loading for all variants by DLS, with the maximum occurring at f+ of 0.5 for all amylase variants (Figure C.5). In the case of A37, the average RH was found to increase dramatically owing to the emergence of peaks at 50-100 nm.

With POEGMA-b-pqP4VP40, the supercharged variants behaved differently from native amylase

(Figure C.6). In all cases, at the lowest protein loading, the hydrodynamic radii remained similar to that of the block copolymer alone. For native amylase, this remained true at f+ of 0.7, but for the supercharged variants the average hydrodynamic radii were found to reach their maxima here

176 due to the emergence of small peaks at 50-100 nm. At f+ of 0.5, the native amylase reached a maximum RH due to the emergence of similar peaks at 50-100 nm, but the supercharged variants showed a disappearance of this higher molecular weight peak but maintained RH greater than that of the BC and protein.

Thin films of POEGMA-b-pqP4VP20—amylase PECS showed changes in nanostructure as a function of f+, but not as a function of the supercharging of the proteins. Figure 5.5 shows the

AFM height and phase images and GISAXS patterns for native amylase (A22) as a function of f+.

The AFM and GISAXS data for the supercharged variants is shown in Figures C.7-9. At the lowest protein loading—20% (wt/wt) for A22, A30, and A32, and f+ = 0.8 for A37 (12.9 wt. %)—the films showed the presence of both a disordered, microphase separated nanostructure similar to the

BC alone and larger complexes likely composed of the protein and the BC. At higher protein loading, f+ of 0.7, the films showed a similar nanostructure, but with greater prevalence of the complexes. This led to a peak associated with the nanostructure of the BC fading by GISAXS and with this morphology being less prevalent in the AFM phase scans. At the highest protein loading, f+ = 0.5, the films show a microphase separated, disordered structure similar to the BC alone, but with both phases showing significant swelling compared to the BC alone. Fast Fourier transforms

(FFTs) of the AFM phase images at f+ = 0.5 indicated a characteristic wavelength of 110 to 150 nm, corresponding to a q of 0.04 to 0.06 nm-1, which is outside the q-range accessible by GISAXS.

177

Figure 5.5. AFM height and phase scans and GISAXS patterns for films of native amylase (A22) and POEGMA-b-pqP4VP40 at various mixing ratios. The positive charge fraction, 푓+, is defined

푀+ as 푓+ = and M+ and M− are the total positive and negative charge of the macromolecules 푀++푀−

-1 in the system. Scale bars are 200 nm. GISAXS patterns’ x-axes range from qy of -0.5 to 0.5 nm

-1 and y-axes range from qz of 0 to 1 nm . Height z-scales are 5, 55, 45, and 35 nm. Phase z-scales are 30°, 45°, 100°, and 35°.

Thin films of POEGMA-b-pqP4VP40—amylase PECs showed transitioning nanostructures as a function of both f+ and the supercharging of the proteins. The native amylase

(A22) formed similar nanostructures to the POEGMA-b-pqP4VP20 and amylase films (Figure

5.6). At 20% (wt/wt), the films showed the presence of both a disordered, microphase separated nanostructure similar to the BC alone and large complexes (with diameter of ~200 nm and greater)

178 likely composed of the protein and the BC. GISAXS showed a weak peak similar to that of the BC alone. At the corresponding point on the coacervation curve (Figure 5.4b), f+ = 0.88, the amylase was not found to complex with the homopolymer, which likely caused the macrophase separation observed. At f+ = 0.7, the film showed a similar nanostructure, but with some fading of the BC structure and shrinking of the macrophase separated regions, and peaks were no longer visible by

GISAXS. At f+ = 0.5, the films show a microphase separated, disordered structure similar to the

BC alone, but with both phases showing significant swelling. No structures were apparent by

GISAXS, and the FFT of the phase image showed no strong peaks, but did have a wide series of peaks between 140 and 1000 nm.

The first supercharged variant of amylase (A30) showed a shift in the nanostructures of the films with POEGMA-b-P4VP40 relative to A22 (Figure 5.6). At 20% (wt/wt), the film had a disordered, microphase separated nanostructure similar to the BC alone, and the GISAXS pattern showed a peak similar to that of the BC alone. At the corresponding point on the coacervation curve (Figure 5.4b), f+ = 0.84, the amylase was found to weakly complex with the homopolymer.

At f+ = 0.7, the film had a microphase separated, disordered structure similar to the BC alone, but with both phases showing significant swelling. No structures were apparent by GISAXS, but the

FFT of the phase image showed a peak at 384 nm. At f+ = 0.5, the nanostructure of the film degraded, and no structures were apparent by GISAXS.

The second supercharged variant of amylase (A32) showed a shift in the nanostructures of the films with POEGMA-b-pqP4VP40 relative to A30 (Figure 5.6). At 20% (wt/wt), the film had both a disordered, microphase separated nanostructure similar to the BC alone and the emergence of the swollen disordered, microphase separated structure observed at f+ of 0.7 with A30. At the corresponding point on the coacervation curve (Figure 5.4b), f+ = 0.83, the amylase was found to

179 complex with the homopolymer. At f+ = 0.7, the film had a microphase separated, disordered structure similar to the BC alone, but with both phases showing significant swelling, similar to

A30. No structures were apparent by GISAXS, but the FFT of the phase image showed a peak at

204 nm. At f+ = 0.5, the nanostructure of the film degraded, and no structures were apparent by

GISAXS.

The final supercharged variant of amylase (A37) showed similar nanostructure to the films with POEGMA-b-pqP4VP40 relative to A32 (Figure 5.6). At 20% (wt/wt), the film had both a disordered, microphase separated nanostructure similar to the BC alone and the emergence of the swollen disordered, microphase separated structure observed at f+ of 0.7 with A30 and 20% with

A32. At the corresponding point on the coacervation curve (Figure 5.4b), f+ = 0.82, the amylase was found to complex with the homopolymer. At f+ = 0.7, the film had a microphase separated, disordered structure similar to the BC alone, but with both phases showing significant swelling, similar to A30. No structures were apparent by GISAXS, but the FFT of the phase image showed a peak at 204 nm. No structures were apparent by GISAXS.

180

181

Figure 5.6. AFM height and phase scans and GISAXS patterns for films of native (A22) and supercharged amylase (A30, A32, A37) and POEGMA-b-pqP4VP40 at various mixing ratios. The

푀+ positive charge fraction, 푓+, is defined as 푓+ = and M+ and M− are the total positive and 푀++푀− negative charge of the macromolecules in the system. Scale bars are 200 nm. GISAXS patterns’

-1 -1 x-axes range from qy of -0.5 to 0.5 nm and y-axes range from qz of 0 to 1 nm . Height and phase z-scales are located in Table C.1.

Overall, the films of the amylase variants and the two BCs behaved similarly, but the greater charge density of the POEGMA-b-pqP4VP40 led to the formation of an additional morphology not observed with the POEGMA-b-pqP4VP20. For the films of POEGMA-b- pqP4VP20 and amylase, the films underwent two transitions, the first to a macrophase separated structure consisting of large complexes and a disordered, microphase separated phase similar to the BC alone, and the second from that phase to a single disordered, microphase separated structure resembling a swollen version of the BC alone. The films of POEGMA-b-pqP4VP40 and supercharged amylase exhibited a third transition to a disordered phase when the protein loading exceeded the loading at the peak of the turbidity curve. The proteins are expected to interact preferentially with the cationic block of the BCs likely leading to the formation of the complexes observed at lower protein loading. Proteins, however, have also been demonstrated to have an affinity for polyethylene glycol (PEG).46-47 As the protein loading is increased, the protein likely distributes into both blocks causing the formation of the swollen microphase separated structure observed, and as loading in further increased, the amphiphilic nature of the protein may cause it to behave as a compatibilizer between the two blocks leading to the disordered structure observed with the 40% quaternized BC and the supercharged amylase.

182

The differentiated behavior of PAA and the amylase variants emerges from three factors: the difference in charge density of the two species, the ability of PAA to more independently explore conformational space compared to the folded protein, and the affinity of proteins to PEG functionalities. The conformational freedom of PAA allows the BC-PAA systems to achieve charge neutrality at the theoretical point of charge balance as opposed to the proteins, which due to their folded state are not able to interact as freely with the BCs thus requiring an excess of P4VP to achieve neutrality. Together, the higher charge density and greater conformational freedom of

PAA also lead to stronger interactions between PAA and P4VP compared to the protein and P4VP causing greater divergence from the BC nanostructure and inducing the micellization seen in the films. The affinity of the protein to both block of the BC likely allows the protein to transition into the BC without significantly disrupting the nanostructure of the BC until high protein loading.

5.4. Conclusions.

The morphology of ionic-neutral BC thin films is altered by polyelectrolyte complexation when the BC is used as an encapsulant for an oppositely charge macromolecule. The nature of the complexation was found to be dependent on the strength and charge density of the polyanions. The strongest polyanion studied, PSS, caused the formation of solid flocculants across a wide range of mixing ratios. PAA, the weak, synthetic polyanion studied caused micellization with the BCs, triggering an ODT. High loading of the PAA, significantly past the point of charge balance caused either a re-entrant ODT or a phase transition as the micelles became less regular. The amylase and

P4VP showed shifted coacervation curves as have been previously reported in literature. In the films, when the protein loading corresponded to a point below the coacervation peak, large aggregates are observed in a sea of BC. This structure transitions to a swollen microphase separated structure near the coacervation peak. Beyond this peak, the films become disordered.

183

The differences in the behavior of PSS and PAA is attributed to the ability of strong polyelectrolytes to induce charge, which caused a stronger interaction between PSS and the BCs than between PAA and the BC. The difference in the behavior of the PAA and protein is attributed to the higher charge density of PAA, the greater conformational freedom of PAA, and the lower affinity of PAA towards PEG functionalities compared to the protein, which led to the more regular nanostructure observed in the BC-PAA films compared to the BC-protein films. This work will allow for better control of the nanostructure of the BC-protein films and facilitate testing of the structure-function relationship in these films.

5.5. Materials and Methods.

5.5.1. Materials.

All solvents and monomers were purchased from Millipore Sigma. Iodomethane, α-amylase, and

PSS (~70 kDa) were also purchased from Millipore Sigma. Succinic Anhydride was purchased from Fisher. 4-Cyano-4-(phenylcarbonothioylthio)pentanoic acid (CPP) was purchased from

Strem Chemicals.

5.5.2. POEGMA Synthesis.

The poly(oligo(ethylene glycol) methacrylate) (POEGMA) homopolymer was polymerized using reversible addition-fragmentation transfer polymerization (RAFT). Oligo(ethylene glycol) methacrylate (OEGMA, Mn = 300 Da) was purified by passing over a basic alumina column twice.

Azobisisobutyronitrile (AIBN) was recrystallized from methanol twice. CPP (326.9 mg, 1.17 mmol), AIBN (38.4 mg, 0.234 mmol), and OEGMA (Mn = 300 Da, 60 mL, 210 mmol) were dissolved in dioxane (180 mL) and degassed three times by freeze-pump-thaw. The vessel was backfilled with N2 and heated to 65 °C for 5 h. The reaction was terminated by taking off heat and exposing to air. The product was precipitated three times into hexanes and dried overnight under

184 vacuum at 50 °C to remove residual hexanes. Mn was found to be 37.3 kDa and Đ was found to be

1.05 by DMF GPC (Figure C.8).

5.5.3. POEGMA-b-P4VP Synthesis.

The POEGMA was used as a macro-RAFT agent to polymerize P4VP. 4VP was purified by passing over a basic alumina column twice. AIBN was recrystallized from methanol twice.

POEGMA (2 g, 0.054 mmol), AIBN (1.76 mg, 0.011 mmol), and 4VP (2.31 mL, 21.43 mmol) were dissolved in a mixture of 5:1 DMF and dioxane (2.77 mL) and degassed by N2 purging for

30 min. The mixture was heated to 75 °C and stirred for 6 h. The polymer was purified by precipitation in diethyl ether. The polymer was dried under vacuum at 50 °C for 20 h. The composition and Mn were found to be 44.4 mol% 4VP (23.4 vol% P4VP) and 47.8 kDa, respectively, as determined by NMR (Figure C.9). The Đ was found to be 1.17 by DMF GPC

Figure C.10).

5.5.4. POEGMA-b-P4VP Quaternization.

The BCs were quaternized with iodomethane in DMF. POEGMA-b-P4VP (200 mg) was dissolved in DMF (1600 µL). Iodomethane was added in a 1:1 stoichiometric ratio with the target amount of

4VP monomer to be quaternized, 20 and 40% (5.19 µL and 10.39 µL), and heated to 60 °C for 6 h. The polymer was then precipitated in diethyl ether twice and dried at 60 °C for 18-24 h to remove residual DMF and ether. Conversion was measured by NMR and found to be >95%

(Figures C.11 and C.12).

5.5.5. P4VP Synthesis.

P4VP was polymerized by RAFT at 70 °C for 16 h. 4VP was purified by basic alumina column.

4VP (60 mL, 556 mmol) was dissolved in DMF (180 mL) with 2-(dodecylthiocarbonothioylthio)-

2-methylpropionic acid (338.1 mg, 0.93 mmol) and AIBN (30.46 mg, 0.19 mmol). The reaction 185 mixture was degassed by 3 freeze-pump-thaw cycles, backfilled with N2, then placed on heat. The reaction was terminated by exposure to air. Mn and Đ were found to be 33.3kDa and 1.05 based on

DMF GPC (Figure C.13).

5.5.6. P4VP Quaternization.

P4VP was quaternized with iodomethane in DMF. P4VP (100 mg) was dissolved in DMF (1000

µL). Iodomethane was added assuming full conversion of the iodomethane, 20 and 40% on a molar basis to the 4VP monomer (11.8 µL and 23.7 µL) and heated to 60 °C for 6 h. The polymer was then precipitated in diethyl ether twice and dried at 60 °C for 18-24 h to remove residual DMF and ether. Conversion was measured by NMR and found to be >95% (Figures C.14 and C.15).

5.5.7. PAA Polymerization.

PAA was synthesized by RAFT polymerization as described by Kim, et al. Briefly, acrylic acid was passed over basic alumina to remove inhibitor. 2-(Dodecylthiocarbonothioylthio)-2- methylpropionic acid (72 mg, 0.2 mmol) and AIBN (6 mg, 0.037 mmol) were added to a solution of acrylic acid (10 g, 139 mmol) in water/DMF (1:9 v/v). The mixture was degassed by 3 freeze- pump-thaw cycles and backfilled with N2, placed on heat at 65 °C for 16 h, and terminated by removal from heat and exposure to air. The polymer was purified by dialysis with a 3 kDa MWCO membrane and converted to the sodium salt by stirring with 1 N NaOH, and dialyzed again. Mn was 33 kDa and Đ was 1.34 by aqueous GPC.35

5.5.8. Preparation and Purification of α-Amylase and Supercharged α-Amylase.

SDS-PAGE showed that the protein was pure as received, but the protein made up only approximately 10% of the dry mass of the powder per A280 measurement. The powder was dissolved at 20 mg mL-1, and the small molecule contaminants were removed by ultrafiltration with 15 mL 10 kDa NMWL Amicon Ultra centrifugal filters. The protein was dissolved in 10 mM

186 phosphate buffer pH 8.0 and subjected to 6 10-fold dilutions. The solution went from dark brown to near clear with a brown tint. SDS-PAGE analysis of the final protein is shown in Figure C.16.

Supercharging was accomplished with succinic anhydride (SA). The previously purified native amylase was concentrated to 5 mg mL-1 in 10 mM phosphate buffer using 15 mL 10 kDa NMWL

Amicon Ultra centrifugal filters. 20 mL of the solution was reacted with 20, 50, and 200 molar equivalents of SA per unit protein resulting in A30, A32, and A37, respectively. Excess SA was removed via ultrafiltration with a 15 mL 10 kDa NMWL Amicon Ultra centrifugal filters with 6

10-fold dilutions. The degree of supercharging was determined by MALDI-TOF (Figure C.17,

Table C.2).

To remove salts from the protein solutions prior to both film coating and solution measurements, the buffer was exchanged to MilliQ water by concentrating the protein in a 0.5 or 4 mL 10 kDa

NMWL Amicon Ultra centrifugal filter by at least 10-fold then diluting with water to the original volume 6 times.

5.5.9. Turbidimetry.

Measurements were done in MilliQ water at 2 mg mL-1 in a TECAN Infinite M200Pro plate reader in 96-well plates at 600 nm. Absorption was converted to turbidity by converting absorption to % transmission and subtracting this value from 100% using the following equation:

Turbidity (100 − %T) = 100 − 102−abs (5.2)

Samples were blended from 2 mg mL-1 samples of the BCs and polyanions at the appropriate charge ratios. 200 µL of sample was prepared in each well and mixed by repeatedly pipetting.

Turbidity was measured within 1 minute of mixing to minimize settling of flocculants/coacervate droplets.

187

5.5.10. Dynamic Light Scattering (DLS).

Measurements were completed on a Wyatt Mobius in MilliQ water at 1 mg mL-1. Samples were filtered with a 0.2 µm PTFE syringe filter to remove dust. 10 measurement of 10 s each were taken for each sample.

5.5.11. Grafting PEG to Si Wafers.

Silicon wafers (Wafer World, (100) orientation) were cleaned with an air plasma for 2 min using a Harrick PDC-001 Plasma Cleaner. The wafers were then coated with an approximately 0.25 cm thick layer of hydroxyl-terminated poly(ethylene glycol) methyl ether (750 Da, Sigma) and placed in a vacuum oven at 150 °C for 18 h. Wafers were stored in acetone prior to use.

5.5.12. Film Casting.

Films were cast as described previously.20 All films were approximately 150-200 nm thick based on light blue to yellow film color. Previous study showed little effect of thickness on C3M film activity and morphology at these thicknesses.20

5.5.13. Atomic Force Microscopy (AFM).

The NTEGRA system from NT-MDT was used to take height and phase scans using a

HQ:NSC16/Al BS probe from MikroMasch. Images were processed using the Nova PX software from NT-MDT. First order linear corrections were applied to all images.

5.5.14. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS).

GISAXS experiments were conducted at Argonne National Laboratory at beamline 8-ID-E using

X-ray with energy of 10.915 keV. Samples were measured under ambient conditions at three incident angles (αi) 0.14, 0.15, and 0.155° between the critical angle of PEG-coated silicon and the critical angle of the C3M film. The reported images are from an incident angle of 0.155°. Two

188 images obtained at two different detector positions were combined to fill the gaps in the detector.

Samples were exposed for 1 to 10 s at each angle and detector position, keeping the total exposure time under 60 s. Exposure time was modulated to accumulate sufficient detector counts. The sample-to-detector distance was 2185 mm. Data were analyzed using the GIXSGUI software package written by Dr. Zhang Jiang. Data were converted to q space by applying correction parameters (stored as .MAT file format) provided by the beamline scientists.

5.6. References.

1. Bates, C. M.; Maher, M. J.; Janes, D. W.; Ellison, C. J.; Willson, C. G., Block Copolymer Lithography. Macromolecules 2014, 47 (1), 2-12. DOI: 10.1021/ma401762n 2. Thomas, C. S.; Glassman, M. J.; Olsen, B. D., Solid-State Nanostructured Materials from Self-Assembly of a Globular Protein–Polymer Diblock Copolymer. ACS Nano 2011, 5 (7), 5697-5707. DOI: 10.1021/nn2013673 3. Huang, A.; Qin, G.; Olsen, B. D., Highly Active Biocatalytic Coatings from Protein–Polymer Diblock Copolymers. ACS Appl. Mater. Interfaces 2015, 7 (27), 14660-14669. DOI: 10.1021/acsami.5b01884 4. Dong, X.-H.; Obermeyer, A. C.; Olsen, B. D., Three-Dimensional Ordered Antibody Arrays Through Self-Assembly of Antibody–Polymer Conjugates. Angew. Chem. Int. Ed. 2017, 56 (5), 1273-1277. DOI: 10.1002/anie.201607085 5. Kataoka, K.; Harada, A.; Nagasaki, Y., Block copolymer micelles for drug delivery: Design, characterization and biological significance. Adv. Drug Del. Rev. 2012, 64, 37-48. DOI: https://doi.org/10.1016/j.addr.2012.09.013 6. Young, W.-S.; Kuan, W.-F.; Epps, I. I. I. T. H., Block copolymer electrolytes for rechargeable lithium batteries. J. Polym. Sci., Part B: Polym. Phys. 2014, 52 (1), 1-16. DOI: 10.1002/polb.23404 7. Kawamura, A.; Harada, A.; Kono, K.; Kataoka, K., Self-Assembled Nano-Bioreactor from Block Ionomers with Elevated and Stabilized Enzymatic Function. Bioconj. Chem. 2007, 18 (5), 1555- 1559. DOI: 10.1021/bc070029t 8. Paloni, J. M.; Dong, X.-H.; Olsen, B. D., Protein–Polymer Block Copolymer Thin Films for Highly Sensitive Detection of Small Proteins in Biological Fluids. ACS Sensors 2019, 4 (11), 2869-2878. DOI: 10.1021/acssensors.9b01020 9. Stewart-Sloan, C. R.; Olsen, B. D., Protonation-Induced Microphase Separation in Thin Films of a Polyelectrolyte-Hydrophilic Diblock Copolymer. ACS Macro Lett. 2014, 3 (5), 410-414. DOI: 10.1021/mz400650q 10. Stewart-Sloan, C. R.; Wang, R.; Sing, M. K.; Olsen, B. D., Self-Assembly of Poly(vinylpyridine-b- oligo(ethylene glycol) methyl ether methacrylate) Diblock Copolymers. J. Polym. Sci., Part B: Polym. Phys. 2017, 55 (15), 1181-1190. DOI: 10.1002/polb.24369 11. Annaka, M.; Morishita, K.; Okabe, S., Electrostatic Self-Assembly of Neutral and Polyelectrolyte Block Copolymers and Oppositely Charged Surfactant. The Journal of Physical Chemistry B 2007, 111 (40), 11700-11707. DOI: 10.1021/jp074404q 12. Horn, J. M.; Kapelner, R. A.; Obermeyer, A. C., Macro- and Microphase Separated Protein- Polyelectrolyte Complexes: Design Parameters and Current Progress. Polymers 2019, 11 (4), 578.

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13. Obermeyer, A. C.; Mills, C. E.; Dong, X.-H.; Flores, R. J.; Olsen, B. D., Complex Coacervation of Supercharged Proteins with Polyelectrolytes. Soft Matter 2016, 12 (15), 3570-3581. DOI: 10.1039/C6SM00002A 14. Blocher, W. C.; Perry, S. L., Complex Coacervate-Based Materials for Biomedicine. Wiley Interdiscip. Rev.: Nanomed. and Nanobiotechnol. 2017, 9 (4), e1442. DOI: 10.1002/wnan.1442 15. Kurtz, I. S.; Sui, S.; Hao, X.; Huang, M.; Perry, S. L.; Schiffman, J. D., Bacteria-Resistant, Transparent, Free-Standing Films Prepared from Complex Coacervates. ACS Applied Bio Materials 2019, 2 (9), 3926-3933. DOI: 10.1021/acsabm.9b00502 16. Dompé, M.; Cedano-Serrano, F. J.; Vahdati, M.; van Westerveld, L.; Hourdet, D.; Creton, C.; van der Gucht, J.; Kodger, T.; Kamperman, M., Underwater Adhesion of Multiresponsive Complex Coacervates. Advanced Materials Interfaces 2020, 7 (4), 1901785. DOI: 10.1002/admi.201901785 17. Black, K. A.; Priftis, D.; Perry, S. L.; Yip, J.; Byun, W. Y.; Tirrell, M., Protein Encapsulation via Polypeptide Complex Coacervation. ACS Macro Lett. 2014, 3 (10), 1088-1091. DOI: 10.1021/mz500529v 18. Lee, Y.; Ishii, T.; Cabral, H.; Kim, H. J.; Seo, J. H.; Nishiyama, N.; Oshima, H.; Osada, K.; Kataoka, K., Charge‐Conversional Polyionic Complex Micelles—Efficient Nanocarriers for Protein Delivery into Cytoplasm. Angew. Chem. Int. Ed. 2009, 48 (29), 5309-5312. DOI: doi:10.1002/anie.200900064 19. Mills, C. E.; Obermeyer, A.; Dong, X.; Walker, J.; Olsen, B. D., Complex Coacervate Core Micelles for the Dispersion and Stabilization of Organophosphate Hydrolase in Organic Solvents. Langmuir 2016, 32 (50), 13367-13376. DOI: 10.1021/acs.langmuir.6b02350 20. Sureka, H. V.; Obermeyer, A. C.; Flores, R. J.; Olsen, B. D., Catalytic Biosensors from Complex Coacervate Core Micelle (C3M) Thin Films. ACS Appl. Mater. Interfaces 2019, 11 (35), 32354- 32365. DOI: 10.1021/acsami.9b08478 21. Sing, C. E.; Perry, S. L., Recent progress in the science of complex coacervation. Soft Matter 2020, 16 (12), 2885-2914. DOI: 10.1039/D0SM00001A 22. Lindhoud, S.; Norde, W.; Cohen Stuart, M. A., Effects of Polyelectrolyte Complex Micelles and Their Components on the Enzymatic Activity of Lipase. Langmuir 2010, 26 (12), 9802-9808. DOI: 10.1021/la1000705 23. Rumyantsev, A. M.; Zhulina, E. B.; Borisov, O. V., Scaling Theory of Complex Coacervate Core Micelles. ACS Macro Lett. 2018, 7 (7), 811-816. DOI: 10.1021/acsmacrolett.8b00316 24. Kim, B.; Lam, C. N.; Olsen, B. D., Nanopatterned Protein Films Directed by Ionic Complexation with Water-Soluble Diblock Copolymers. Macromolecules 2012, 45 (11), 4572-4580. DOI: 10.1021/ma2024914 25. Brena, B. M.; Batista-Viera, F., Immobilization of Enzymes. In Immobilization of Enzymes and Cells, Guisan, J. M., Ed. Humana Press: Totowa, NJ, 2006; pp 15-30. 26. Basso, A.; Serban, S., Industrial applications of immobilized enzymes—A review. Molecular Catalysis 2019, 479, 110607. DOI: https://doi.org/10.1016/j.mcat.2019.110607 27. DiCosimo, R.; McAuliffe, J.; Poulose, A. J.; Bohlmann, G., Industrial Use of Immobilized Enzymes. Chem. Soc. Rev. 2013, 42 (15), 6437-6474. DOI: 10.1039/C3CS35506C 28. Young, W.-S.; Epps, T. H., Ionic Conductivities of Block Copolymer Electrolytes with Various Conducting Pathways: Sample Preparation and Processing Considerations. Macromolecules 2012, 45 (11), 4689-4697. DOI: 10.1021/ma300362f 29. Huang, J.; Tong, Z.-Z.; Zhou, B.; Xu, J.-T.; Fan, Z.-Q., Salt-induced microphase separation in poly(ε- caprolactone)-b-poly(ethylene oxide) block copolymer. Polymer 2013, 54 (12), 3098-3106. DOI: https://doi.org/10.1016/j.polymer.2013.03.070 30. Wang, J.-Y.; Chen, W.; Roy, C.; Sievert, J. D.; Russell, T. P., Influence of Ionic Complexes on Phase Behavior of Polystyrene-b-poly(methyl methacrylate) Copolymers. Macromolecules 2008, 41 (3), 963-969. DOI: 10.1021/ma071908d 31. Young, W.-S.; Epps, T. H., Salt Doping in PEO-Containing Block Copolymers: Counterion and Concentration Effects. Macromolecules 2009, 42 (7), 2672-2678. DOI: 10.1021/ma802799p

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32. Loo, W. S.; Galluzzo, M. D.; Li, X.; Maslyn, J. A.; Oh, H. J.; Mongcopa, K. I.; Zhu, C.; Wang, A. A.; Wang, X.; Garetz, B. A.; Balsara, N. P., Phase Behavior of Mixtures of Block Copolymers and a Lithium Salt. The Journal of Physical Chemistry B 2018, 122 (33), 8065-8074. DOI: 10.1021/acs.jpcb.8b04189 33. Voets, I. K.; de Keizer, A.; Cohen Stuart, M. A., Complex Coacervate Core Micelles. Adv. Colloid Interface Sci. 2009, 147–148, 300-318. DOI: 10.1016/j.cis.2008.09.012 34. Cooper, C. L.; Dubin, P. L.; Kayitmazer, A. B.; Turksen, S., Polyelectrolyte–protein complexes. Curr. Opin. Colloid Interface Sci. 2005, 10 (1), 52-78. DOI: https://doi.org/10.1016/j.cocis.2005.05.007 35. Kim, S.; Sureka, H. V.; Kayitmazer, A. B.; Wang, G.; Swan, J. W.; Olsen, B. D., Effect of Protein Surface Charge Distribution on Protein–Polyelectrolyte Complexation. Biomacromolecules 2020, 21 (8), 3026-3037. DOI: 10.1021/acs.biomac.0c00346 36. Sadman, K.; Delgado, D. E.; Won, Y.; Wang, Q.; Gray, K. A.; Shull, K. R., Versatile and High- Throughput Polyelectrolyte Complex Membranes via Phase Inversion. ACS Appl. Mater. Interfaces 2019, 11 (17), 16018-16026. DOI: 10.1021/acsami.9b02115 37. Cooper, C. L.; Dubin, P. L.; Kayitmazer, A. B.; Turksen, S., Polyelectrolyte–Protein Complexes. Curr. Opin. Colloid Interface Sci. 2005, 10 (1–2), 52-78. DOI: 10.1016/j.cocis.2005.05.007 38. Lund, M.; Jönsson, B., On the Charge Regulation of Proteins. Biochemistry 2005, 44 (15), 5722-5727. 39. Mahltig, B.; Gohy, J.-F.; Antoun, S.; Jérôme, R.; Stamm, M., Adsorption and structure formation of the weak polyelectrolytic diblock copolymer, PVP-b-PDMAEMA. Colloid. Polym. Sci. 2002, 280 (6), 495-502. DOI: 10.1007/s00396-001-0628-1 40. Srivastava, S.; Andreev, M.; Levi, A. E.; Goldfeld, D. J.; Mao, J.; Heller, W. T.; Prabhu, V. M.; de Pablo, J. J.; Tirrell, M. V., Gel phase formation in dilute triblock copolyelectrolyte complexes. Nature Communications 2017, 8 (1), 14131. DOI: 10.1038/ncomms14131 41. Hillmyer, M. A.; Maurer, W. W.; Lodge, T. P.; Bates, F. S.; Almdal, K., Model Bicontinuous Microemulsions in Ternary Homopolymer/Block Copolymer Blends. The Journal of Physical Chemistry B 1999, 103 (23), 4814-4824. DOI: 10.1021/jp990089z 42. Matsen, M. W., Phase Behavior of Block Copolymer/Homopolymer Blends. Macromolecules 1995, 28 (17), 5765-5773. DOI: 10.1021/ma00121a011 43. Cummings, C. S.; Obermeyer, A. C., Phase Separation Behavior of Supercharged Proteins and Polyelectrolytes. Biochemistry 2018, 57 (3), 314-323. DOI: 10.1021/acs.biochem.7b00990 44. Kayitmazer, A. B., Thermodynamics of complex coacervation. Adv. Colloid Interface Sci. 2017, 239, 169-177. DOI: https://doi.org/10.1016/j.cis.2016.07.006 45. Chang, L.-W.; Lytle, T. K.; Radhakrishna, M.; Madinya, J. J.; Vélez, J.; Sing, C. E.; Perry, S. L., Sequence and entropy-based control of complex coacervates. Nature Communications 2017, 8 (1), 1273. DOI: 10.1038/s41467-017-01249-1 46. Huang, A.; Yao, H.; Olsen, B. D., SANS partial structure factor analysis for determining protein– polymer interactions in semidilute solution. Soft Matter 2019, 15 (37), 7350-7359. DOI: 10.1039/C9SM00766K 47. Bekale, L.; Agudelo, D.; Tajmir-Riahi, H. A., The role of polymer size and hydrophobic end-group in PEG–protein interaction. Colloids Surf. B. Biointerfaces 2015, 130, 141-148. DOI: https://doi.org/10.1016/j.colsurfb.2015.03.045

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Chapter 6. Enzyme Immobilization and Stabilization via Complex Coacervate Core Micelle

(C3M) Thin Films

6.1. Abstract

Enzyme immobilization enables the conversion of enzymes from homogeneous to heterogeneous catalysts and the stabilization of enzymes against denaturation. Complex coacervation facilitates the encapsulation of enzymes within a polymer matrix in an aqueous process, and the coating and crosslinking of thin films of these complexes onto a substrate allows for the immobilization of enzymes via complex coacervation. This study aims to understand how the properties of polymer encapsulants—in this case polycationic-neutral diblock copolymers—can be tuned to optimize the activity and thermal stability of complex coacervate core micelle (C3M) thin films, and how these optima vary as a function of the net charge of the encapsulated protein. A library of 12 polymer encapsulants with varying polycationic chemistries and molecular weights was synthesized and combined with a model protein used in a native, low net-charge form or a supercharged form..

Morphologies of the resulting C3M thin films typically varied as a function of the protein loading, the molecular weight of the encapsulants, and the charge of the protein. The performance of the thin films, in terms of both activity and thermal stability, seemed to be a stronger function of the choice of polymer chemistry than of the molecular weight or the protein loading, with stronger protein-polymer interactions favoring higher activity and weaker polymer-protein interaction and greater conformational freedom of polymer side chains favoring greater thermal stability.

6.2. Introduction

Protein-based biocatalysts have been used in many manufacturing sectors including textiles, pharmaceuticals, food products, and biodiesel.1-5 The advantages of these biocatalysts

193 over synthetic catalysts include increased stereospecificity, milder processing conditions, and faster kinetics.4-7 The development of novel biocatalysts has been greatly accelerated through directed evolution, allowing for the synthesis of designer enzymes for specific processes.4,7-8 For example, lipases have been evolved to provide enhanced stereoselectivity and activity,9 cytochrome P450 enzymes have been evolved to both improve stereoselectivity and activity and enable new chemistries,10 and even enzyme cascades have been evolved to achieve one-pot multi- step conversions.8,11-12 Despite the great strides made in the development of novel stable biocatalysts, most natural enzymes and many engineered enzymes are susceptible to denaturation due to changes in pH, temperature, and solvent conditions, and most are homogeneous catalysts making enzyme retention challenging.4,13-14 Enzyme immobilization can address both these issues by converting enzymes to heterogeneous catalyst and helping to stabilize enzymes against denaturation.4,13-16

One method that has been proposed for the encapsulation and immobilization of enzymes is complex coacervation.17-25 Complex coacervation is a liquid-liquid phase transition that can occur in aqueous mixtures of electrostatically-interacting macromolecules that results in a dense phase (the complex coacervate) and a dilute phase.26 When one of the macromolecules used is a charged-neutral block copolymer, this phase change can also form complex coacervate core micelles (C3Ms). Complex coacervates and C3Ms have been demonstrated to achieve high protein loading with a wide range of proteins including alkaline phosphatase, green fluorescent protein, and organophosphate hydrolase, and have been shown to improve the stability of encapsulated proteins to temporal, thermal, and chemical degradation.21-22,24,27-31 While both C3Ms and complex coacervates are still wet, protein-loaded C3Ms can be coated onto surface to enable protein immobilization onto surfaces.24-25 Complex coacervates have also been dried to form solid

194 materials via phase inversion and via electrospinning, presenting a potential alternative method for the immobilization of enzymes within these systems.32-33 While the behavior of complex coacervates consisting of polymers has been modeled extensively with high accuracy,34 the use of a protein complicates modeling for various reasons including the ability of proteins to charge regulate based on local environments, the zwitterionic nature of the protein surface, and the effects that localized charge patches can have on the complexation of the proteins.35-42

Despite difficulties in modeling, several studies have begun to systematically probe the behavior of protein-polyelectrolyte complexation. In systems with a strong polycation and a negatively-charged protein, the ratio of negative charges to positive charges on the protein has been demonstrated to be predictive of whether a protein will form complex coacervates and C3Ms, with negative supercharging of the proteins favoring the formation of complexes.42 Additionally, the nature of complexes formed from negatively-supercharged enzymes and strong polycations is dependent on the distribution of charges on the protein, with isotropically distributed charges favoring precipitation and localization of charges to a tag favoring liquid-liquid separation.36 Thus, enzymes can be genetically engineered24,36,43 or chemically modified to form complex coacervates and C3Ms.19-20,42 Efforts have also been made to quantify the charge distribution on protein surfaces, i.e. the patchiness of these surfaces, and determine whether these values can be predictive of complexation behavior with some success.35,40

While factors affecting the incorporation of enzymes into C3Ms and complex coacervates have been widely explored, structure-function relationships, which for these systems are often convoluted with composition-function relations, have not been widely studied. One study did find that for a particular protein, lipase, the activity of the enzyme was maximized at a particular blending ratio of the components—notably not at the point of charge balance but rather when an

195 excess of the cationic-neutral block copolymer encapsulant was in excess—but did not explore how the morphologies of complexes evolved as a function of this blending ratio.21 The structure of C3Ms in solution has been shown to vary with the block ratios of the block copolymer(s) used, the salt concentration and pH, and the ratio of the two (or more) complexing species, but may also be influenced by a number of other variables including total concentration and temperature.44 The relationship of these structures to the function of encapsulated proteins has not been reported. In

C3M thin films, our recent work has demonstrated that the blending ratios and polyelectrolyte strengths of components can have a strong effect on the morphologies of the thin films, indicating that in these films a structure-composition relationship occurs, but structure-function relationships have not yet been explored.45 Structure-function relationships have been demonstrated to be essential to the performance of many classes of materials.46-47 In the case of biosensors made of protein-polymer fusion block copolymers, tuning domain sizes has enabled greater sensitivity of biosensors in biological fluids via a size-exclusion effect.46-47 In conductive block copolymer-salt systems, changes in block copolymer morphology can greatly affect conductivity.48 Thus, understanding both the composition-structure and resulting structure-function relationships is essential to optimizing the performance of C3M thin films.

This work aims to understand how the properties of cationic-neutral block copolymer encapsulants and the properties of enzymes affect the performance of enzymes encapsulated in

C3M thin films. The effects of the blending ratio of the polymer and protein, the polymer chemistry, length, and polyelectrolyte strength, and the effect of enzyme charge density on the morphology, activity, and thermal stability of C3M thin films are examined using a library of 12 polymer encapsulant and native and supercharged variants of a model protein. Thermal stability is

196 found to be a strong function of polymer chemistry while activity varies both as a function of chemistry and morphology.

6.3. Experimental.

6.3.1. Materials.

All solvents were purchased from VWR International. Monomers and azobisisobutyronitrile

(AIBN) were purchased from MilliporeSigma. 4-Cyano-4-(phenylcarbonothioylthio) pentanoic acid (CPP) was purchased from Strem Chemical.

6.3.2. (POEGMA-r-BP) Synthesis.

Reversible addition-fragmentation chain transfer polymerization (RAFT) was used to synthesize

3 random copolymers of oligo(ethylene glycol) methyl ether methacrylate (OEGMA, Mn = 300 g mol–1) and benzoylphenyl methacrylate (BP) with narrow molecular weight distributions as previously described.24 OEGMA was passed through 2 basic alumina columns prior to polymerization to remove inhibitors. CPP and AIBN (recrystallized twice from methanol) were added to a solution of OEGMA (20 g, 66.6 mmol) and BP (1 g, 3.8 mmol) in 60 g of 1,4-dioxane.

The solutions were degassed by three freeze–pump–thaw cycles. The polymerization was carried out in a sealed flask at 65 °C and terminated by removal of heat and exposure to oxygen. The polymer was then precipitated in hexane and dried under vacuum. Ratios and masses of OEGMA,

CPP, and AIBN are given in Table S1.

The Mn and of the POEGMA-r-BP copolymers are listed in Table 1 as determined by size- exclusion chromatography (SEC) (Figure D.1). SEC analyses were performed on a Waters high- performance liquid chromatography (HPLC) system equipped with a Waters 1515 Isocratic HPLC

Pump with two columns (ResiPore, 300 × 7.5 mm, up to 500 kDa, Agilent Technologies, CA) in

197 series. DMF with 0.02 M LiBr was used as the eluent with a flow rate of 1 mL min–1 at 70 °C.

The detector system consisted of a Wyatt miniDAWN TREOS multiangle light scattering detector and a Wyatt Optilab T-rEX differential refractive index detector.

6.3.3. POEGMA-P4VP Synthesis.

4-Vinylpyridine (4VP) was passed through 2 basic alumina columns prior to polymerization to remove inhibitors. POEGMA-r-BP was dissolved in a 5:1 mix of dioxane and dimethyl formamide

(DMF). 4VP and AIBN were added after dissolution, and the mixture was degassed via N2 purging for 25 min. Reactions were initiated by heating to 75-80 °C and terminated after a specified time by removing from heat and exposing to air. The exact polymerization conditions are given in Table

S2. Final monomer ratios were determined by 1H NMR (Table 6.1 and Figure D.2) and Đ was determined by SEC (Figure D.1).

6.3.4. POEGMA-PDMAEMA Synthesis.

2-(Dimethylamino)ethyl methacrylate (DMAEMA) was passed through 2 basic alumina columns prior to polymerization to remove inhibitors. POEGMA-r-BP was dissolved dioxane. DMAEMA and AIBN were added after dissolution, and the mixture was degassed via N2 purging for 25 min.

Reactions were initiated by heating to 75 °C and terminated after a specified time by removing from heat and exposing to air. The exact polymerization conditions are given in Table S2. Final monomer ratios were determined by 1H NMR (Table 6.1 and Figure D.2) and Đ was determined by SEC (Figure D.1).

6.3.5. Polymer Quaternization.

For 100% quaternization, the polymers (0.5 g) were dissolved in DMF (4 mL) and stirred with a

4x molar excess of methyl iodide to cationic monomer overnight at room temperature. For 20% quaternization, the POEGMA-P4VPs (0.5 g) were dissolved DMF (4 mL), and stirred with a 1:1

198 stoichiometric ratio of methyl iodide with the target amount of 4VP monomer to be quaternized,

20%. Polymers were precipitated in ether twice and dried at 40 °C under vacuum for 1-2 days.

Degree of quaternization and removal of solvents was confirmed by NMR (Figure D.3).

6.3.6. NfsB Expression and purification.

Plasmid DNA for a dimeric fusion of NfsB from E. coli, with mutations at residues 89 and 315 to p-azidophenylalanine (AzF) and cysteine, respectively, was provided by Neil Marsh (University of Michigan). The plasmid was cotransformed with the pDule2 pCNF RS plasmid (from Ryan

Mehl, Oregon State University) into BL21 (DE3) E. coli cells. Overnight cultures were made in

50 mL LB supplemented with 40 mM spectinomycin and 50 mM kanamycin in a 250 mL baffled flask. The protein was expressed in 5 L LB supplemented with 40 mM spectinomycin, 50 mM kanamycin, and 0.7 mL of antifoaming agent (Antifoam 204, MilliporeSigma) in a bioreactor with an air flowrate of 5 L min-1. The bioreactor was equilibrated to 37 °C, and the overnight culture was added. When the optical density at 600 nm reached 0.6, 1 g AzF, dissolved in water by addition of NaOH, was added and the temperature was set to 18 °C. After 30 min, the culture was induced with 1 mM IPTG, and the expression was allowed to continue for 18-22 h. Cells were centrifuged down from the media at 4 °C at 4000 xG for 15 min, and frozen at -80 °C and stored overnight or until needed.

Cells were thawed and resuspended in lysis buffer (~2-3 mL/g, [formulation]), after which 1 mg mL-1 of lysozyme (from chicken egg white, MilliporeSigma) was added and allowed to react for

30 min at 4 °C. The cells were then sonicated and the lysate was clarified by centrifugation at

10,000 xG for 30 min at 4 °C. NfsB was then purified by metal affinity chromatography with Ni-

NTA. Purified enzyme was frozen in liquid N2 and stored at -80 °C. Before use, protein were

199 thawed at 4 °C overnight and stored at 4 °C for no longer than 5 days. Figure D.4 shows an SDS-

PAGE analysis of the purified enzyme.

6.3.7. NfsB Supercharging.

NfsB was concentrated via an Amicon Ultra 30 kDa centrifugal filter and adjusted to 5 mg mL-1 in 100 mM NaPO4 buffer pH 8.0. 200 molar equivalents of succinic anhydride per mole of protein were added to the solution and allowed to react at room temperature overnight. The protein was the dialyzed against 50 mM NaPO4 buffer pH 7.6 with 100 mM NaCl in a 12-14 kDa MWCO dialysis membrane to remove excess succinic anhydride. The change in molar mass was determined to be 1.73 kDa corresponding to 17.3 modifications per protein by MALDI-TOF

(Figure D.5). The modification led to a change of the net charge from -7 to -41.6. Purified supercharged (SC) NfsB was frozen in liquid N2 and stored at -80 °C. Before use, the protein was thawed at 4 °C overnight and stored at 4 °C for no longer than 5 days.

6.3.8. Preparation of Fused Silica.

Fused silica substrates (Mark Optics) were sonicated in 2% Hellmanex solution for 30 min under heating, then thoroughly rinsed with water. The wafers were then cleaned in piranha solution (3:1

H2SO4:30% H2O2) for 1 h and then rinsed several times with MilliQ water. The wafers were then dried with filtered air and stored in a sterile petri dish until use.

6.3.9. Polyethylene Glycol (PEG) Modification of Substrates.

Silicon wafers (WaferWorld) and piranha-cleaned fused silica substrates were cleaned with an air plasma in a Harrick plasma cleaner for 2 min. Substrate were covered with sufficient monomethyl-

PEG (MilliporeSigma, Mn = 750) to submerge them in a crystallization dish and were placed in a

150 °C vacuum oven for 16 h. Substrates were rinsed and stored in acetone until use.

200

6.3.10. Film Casting.

PEG-coated substrates were rinsed with acetone, methanol, and water and dried using filtered air.

NfsB was exchanged into water six times, diluting by a factor of 30 each cycle, and concentrated to 15–20% (wt/wt) via ultrafiltration (Amicon Ultra 30K-15 mL). The concentration was measured via absorption at 280 nm. Polymer solutions were prepared at 25% in water and mixed with the protein solutions to achieve appropriate mass ratios. The final mixtures were diluted to 20% and adjusted down if necessary to achieve target film thickness. The solutions were then flow coated using a setup similar to that described by Stafford et al.49 onto the substrate under 68-72% relative humidity (RH) at room temperature. Films were then cross-linked under ultraviolet light for 7 min

(280 nm), and rinsed with water to remove excess polymer.

6.3.11. Dynamic Light Scattering.

Protein was exchanged into water by ultrafiltration and diluted to 1 mg mL-1 in water. Polymers were dissolved at 1 mg mL-1 in water. Protein-polymer blends were then made at 10%, 25%, and

50% protein and allowed to equilibrate overnight at 4 °C. The DLS data for the blends and the pure polymers and proteins were collected on a Wyatt Mobius in a 45 µL microcuvette with 10 measurements consisting of 10 acquisitions of 10 s each. Protein samples were measured in both

10 mM sodium phosphate buffer with 100 mM NaCl pH 7.6 and water. Data shown for the pure proteins are in buffer as the native protein aggregated and the supercharged protein unfolded in water (Figure D.6).

6.3.12. Atomic Force Microscopy (AFM).

The NTEGRA system from NT-MDT was used to take height and phase scans using a

HQ:NSC16/Al BS probe from MikroMasch. Images were processed using the Nova PX software from NT-MDT. Line-by-line linear corrections were applied to each image.

201

6.3.13. Film Height Measurement.

The film thickness was determined by a scratch test. Films were scratched using a 30 gauge needle

(BD). An AFM scan was taken over the scratch and a planar flattening correction was applied to the exposed Si surface. The average height profile was then extracted from the data and analyzed to give the average film height.

6.3.14. Grazing-Incidence Small-Angle X-ray Scattering (GISAXS).

GISAXS experiments were conducted at Argonne National Laboratory at beamline 8-ID-E using

X-ray with energy of 10.915 keV. Samples were measured under ambient conditions at incident angles (αi) of 0.14, 0.15, and 0.155° between the critical angle of PEG-coated silicon and the critical angle of the C3M film. The reported images are from an incident angle of 0.155°. Samples were exposed for 0.5 to 10 s at each angle with exposure time adjusted to achieve sufficient detector counts without damaging the samples. The sample-to-detector distance was 2185 mm.

Data were analyzed using the GIXSGUI software package written by Dr. Zhang Jiang. Data were converted to q space by applying correction parameters (stored as .MAT file format) provided by the beamline scientists.

6.3.15. Film Aging.

Thermal aging of the films was conducted in a Memmert environmental chamber set to 70 °C and

10% RH for the prescribed times.

6.3.16. Activity Assays.

The activity of the NfsB was measured using an assay adapted from Schroeder, et al.50 Activity was measured in 100 mM NaPO4 buffer pH 8 for both the free enzymes and the films at 0.5 mM

-1 -1 NADH. Oxidation of NADH (ε340 = 6220 M cm ) was monitored by plate reader. For all

202 reactions, control samples with no enzymes/films were prepared in quadruplicate to monitor the basal conversion rate.

For the assay of free NfsB, assays were conducted with 0, 50, 100, 250, 500, 750, and 1000 mM 4-nitrobenzenesulfonamide (4-NBS). 4-NBS stock solution was prepared at 100 mM in

DMSO and diluted to 10x the assay concentrations with DMSO added such that all stock solutions had 1% DMSO (vol./vol.) to avoid solvatometric effects. NADH stock solution was prepared at

203 reaction time, 100 µL of the reaction mixture was transferred to a 96-well plate and the final conversion was measured.

6.4. Results and Discussion.

6.4.1. Material Design.

A library of diblock copolymer encapsulants was designed with varying charge density, molar mass, and monomer conformational freedom to assess the effects of these properties on the activity and stability of protein-polymer C3M thin films. A corona block consisting of neutral POEGMA and a small amount of photocrosslinker, benzophenone methacrylate, was synthesized by RAFT at three molar masses (Table 1). These POEGMA-r-BP polymers then used as macroCTAs for the polymerizations of the cationic core blocks consisting of P4VP or PDMAEMA, yielding

POEGMA-P4VP and POEGMA-PDMAEMA. The block copolymers were synthesized such that the volume percent of the cationic block was near 25%, consistent with the block ratio typically used for the formation of C3Ms.31 Polymer data are provided in Table 1.

Table 1. Molar mass and compositional data of diblock copolymer encapsulants. All polymers were synthesized with volume fraction of the cationic block of 22.5% ± 1.5%.

Mn Mn Vol % Polymer POEGMA-BP DPOEGMA DPBP ĐPOEGMA-BP Polycation DPCation Mn BC ĐBC Polycation POEGMA15- 15470 45 3 1.06 4980 32 20451 1.11 22.7% PDMAEMA5 POEGMA36- 35920 105 7 1.05 11040 70 46961 1.14 21.9% PDMAEMA13 POEGMA55- 54960 161 11 1.10 16680 106 71642 1.18 21.7% PDMAEMA19 POEGMA15- 15470 45 3 1.06 4170 40 19644 1.15 22.7% P4VP5 POEGMA36- 35920 105 7 1.05 10360 99 46277 1.16 23.9% P4VP11 POEGMA55- 54960 161 11 1.10 13560 129 68519 1.23 21.1% P4VP17

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In order to probe the effects of polymer charge density on the complex coacervation of the protein-polymer systems, the polymers were modified to create low and high charge density variants. The POEGMA-PDMAEMAs were left non-quaternized for the low charge-density variant of the POEGMA-PDMAEMAs (0qPOEGMA-PDMAEMA) because the pKa of

PDMAEMA is 7.4-7.8,51 near neutral pH and likely to charge modulate in the presence of the

52 model protein. Because the pKa of P4VP is approximately 5, far enough from neutral pH to be unlikely to charge modulate in the presence of the model protein, the POEGMA-P4VPs were partially quaternized to 20% to create low charge-density POEGMA-P4VPs (20qPOEGMA-

P4VP). Both sets of block copolymers were fully quaternized to create the high-charge density variants of these polymers (100qPOEGMA-PDMAEMA and 100qPOEGMA-P4VP).

A monomeric NfsB construct with A89AzF and A315C mutations that has been previously described was used as the model protein to enable the use of FRET and single molecule tracking to understand dynamics within the C3M thin films studied.15 A supercharged variant of the protein, made by reaction with succinic anhydride, was also study the dependence of optimal polymer chemistry on the charge density of the encapsulated protein. The native and supercharged proteins had net charges of -7 and -42, respectively. The native protein and a supercharged variant were

-1 -1 found to have kcat of 136 ± 7 s and 0.011 ± 0.001 s , respectively, and Km of 1800 ± 120 µM and

180 ± 50 µM, respectively, at 0.5 mM NADH, respectively (Figure 6.1). Due to the vastly different kinetics, with kcat/KM being 3 orders of magnitude smaller for the surpercharged than the native variant, the activities are plotted on different axes, but Figure D.7 shows the activities of the two variants plotted on the same axes. The significantly lower activity of the supercharged enzyme is likely driven by increased electrostatic repulsion between negatively charged NADH, negatively

205

53 charged 4-NBS (pKa = 9.38) , and the negatively charged enzyme. Exposure of both variants to

70 °C for 12 h rendered them completely inactive.

Figure 6.1. Activity of a) native and b) supercharged NfsB as a function of 4-NBS concentration at 0.5 mM NADH. Dashed lines represent Michaelis-Menten fits of the activity data. Thermally denaturing the enzyme variant at 70 °C for 12 h rendered each variant inactive.

6.4.2. Morphology of NfsB-Block Copolymer Hybrid Materials.

In dilute solution, all 3 0qPOEGMA-PDMAEMAs show complexation with native NfsB with the mass fraction of complexes increasing with increased protein loading based on DLS analysis

(Figure 6.2). At 10% (wt./wt.) protein loading, particle mass distributions appear bimodal, suggesting the formation of micelles and of large aggregates or of morphologies with two diffusive modes such as a cylindrical micelles or a beads-on-a-string morphology. This secondary diffusive mode diminished with increased protein loading leading to a monomodal peak with the shoulder disappearing at 25% protein for 0qPOEGMA36-PDMAEMA13 and 0qPOEGMA55-

PDMAEMA17, and at 50% protein for 0qPOEGMA15-PDMAEMA5. The disappearance of this secondary diffusive mode is likely caused by the increased loading allowing the system to approach charge balance and decreasing the amount of free polymer in the system, leading to the formation of spherical micelles. The theoretical positive charge fraction, f+—defined as:

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푀+ 푓+ = (1) 푀++푀− and M+ and M- are the total positive and negative charges in the system, respectively—of the system at 50% protein loading is approximately 0.8, indicating that charge balance is in theory not achieved. However, due to the ability of both the protein and polymer to modulate charge and the proximity of the pH to the pKa of the polymer, the system may effectively achieve charge balance.

Figure 6.2. Particle mass distributions determined by DLS for complexes formed between native

NfsB and 0qPOEGMA-b-PDMAEMA at 1 mg mL-1 in water. Bimodal distributions of particle sizes give way to monomodal distributions with increasing protein loading for all three block copolymers. The mass fraction of complexes increases with increasing protein loading.

Thin films of 0qPOEGMA-PDMAEMA of all 3 molar masses showed similar nanostructure with the addition of native NfsB leading to a gradual transition from a disordered

207 morphology with no microphase separation, to micellar protein-polymer complexes. Figure 6.3 shows the AFM data for thin films of 0qPOEGMA36-PDMAEMA13 and native NfsB, and Figures

D.8 and D.9 show the AFM data for 0qPOEGMA15-PDMAEMA5 and 0qPOEGMA55-

PDMAEMA17, respectively. The polymers alone were neither ordered nor microphase separated.

At 10% protein loading, a small amount of complexation occurs in all three polymers indicated by the peaks in the AFM height scans and corresponding valleys in the AFM phase scans, with both larger complexes with diameter of approximately 100 nm and smaller complexes with diameter of approximately 20 nm forming. The presence of a bimodal distribution is consistent with the DLS measurements. However, the sizes of the complexes are inconsistent with DLS, likely driven by the drying of the micelles. At 25% and 50% protein loading, the films show only the smaller complexes, again consistent with the DLS. GISAXS analysis shows no significant features for the films except 25 and 50% protein loading with 0qPOEGMA55-PDMAEMA17, where a shoulder emerges (Figure D.10).

Figure 6.3. AFM height and phase scans of films of 0qPOEGMA36-PDMAEMA13 and native

NfsB. The addition of protein led to the formation of micellar complexes, with the degree of complexation increasing with the protein loading. Similar behavior was seen with both

208

0qPOEGMA15-PDMAEMA5 and 0qPOEGMA55-PDMAEMA19 shown in Figures D.9 and

D.10, respectively. Scale bars at 200 nm.

When blended with the native NfsB in solution, 100qPOEGMA-PDMAEMA also showed the formation of complexes at low protein loading and an increasing mass fraction of complexes with increased protein loading based on DLS (Figure 6.4). With 100qPOEGMA15-PDMAEMA5, the particle size distribution is monomodal at 10% protein loading and bimodal both 25 and 50% protein loading, again suggesting that larger aggregates or elongated nanostructures are forming.

With 100qPOEGMA36-PDMAEMA13, the particle size distribution is bimodal at all 3 protein loadings with the secondary diffusive mode becoming more prevalent with higher protein loading.

With 100qPOEGMA55-PDMAEMA19, the particle size distribution is bimodal at all 3 protein loadings with the secondary diffusive mode becoming more prevalent with higher protein loading.

The difference in behavior between 0qPOEGMA-PDMAEMA and 100qPOEGMA-PDMAEMA arises from the significantly higher charge density of 100qPOEGMA-PDMAEMA. Even at 50% protein loading, f+ exceeds 0.85, and unlike 0qPOEGMA-PDMAEMA, 100qPOEGMA-

PDMAEMA cannot modulate charge. This indicates that there is always an excess of polymer encapsulant compared to protein in these systems, and the nanostructure of the complexes could evolve as a function of the protein loading and adopt morphologies such as cylindrical micelles or beads on a string that have been previously reported in systems of polyelectrolyte complexes.

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Figure 6.4. Particle mass distributions determined by DLS for complexes formed between native

NfsB and 100qPOEGMA-b-PDMAEMA at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading. The mass fraction of complexes increases with increasing protein loading.

In thin films, the nanostructure of the 100qPOEGMA-PDMAEMA and native NfsB films varied strongly with the protein loading, and showed slight differences as a function of the encapsulants’ molecular weights. Figure 6.5 shows the AFM height and phase scans of all 3

100qPOEGMA-PDMAEMA encapsulants with native NfsB. The two lower molecular weight polymers, 100qPOEGMA15-PDMAEMA5 and 100qPOEGMA36-PDMAEMA13 alone appear to adopt a parallel cylinder morphology, which is consistent with both the block fraction and the use of a non-neutral coating surface (PEG-functionalized). However, GISAXS (Figure D.11) only shows a single peak for these samples, so the morphology may alternatively be lamellar. AFM imaging of the 100qPOEGMA55-PDMAEMA19 alone shows a disordered, microphase separated

210 structure, but GISAXS shows a peak at q* of 0.2 nm-1 (Figure D.11) and 3q*, again suggesting either a lamellar or cylindrical morphology. Upon the addition of 10% protein, both

100qPOEGMA15-PDMAEMA5 and 100qPOEGMA36-PDMAEMA13 show similar structure to the BC alone, but with defects which are likely protein-polymer complexes, as seen by the AFM scans. These defects varied in size, in agreement with the bimodal distribution of particle sizes observed by DLS at the protein loading for 100qPOEGMA36-PDMAEMA13 and the tail seen in the distribution for 100qPOEGMA15-PDMAEMA5. In the case of 100qPOEGMA55-

PDMAEMA19, the addition of 10% protein led to a swelling of one of the phases, likely the cationic block, based on the AFM scans. At 25% protein loading, all three encapsulants form apparent beads-on-a-string complexes, but only 100qPOEGMA15-PDMAEMA5 has apparent excess block copolymer based on the AFM scans. However, the GISAXS of all the 25% films still show peaks equivalent to those of the BCs alone, indicating that all three likely have free BC

(Figure D.11). The beads-on-a-string morphology is also consistent with the bimodal distributions observed by DLS. At the highest protein loading, 50%, the two lower molecular weight encapsulants show disordered micellar complexes, while the largest still shows a beads-on-a-string morphology. This is inconsistent with the bimodal distributions observed by DLS for the lower molecular weight encapsulants, but drying of complexes and coating from concentrated solution

(150 to 200-fold higher) likely led to morphological changes in films. 100qPOEGMA15-

PDMAEMA5 also produces poor quality films at 50% loading with visible defects because the coating solution becomes gel-like. The differentiated behavior between the higher molecular weight encapsulants and the lower likely arises from the ability of the larger encapsulant to interact with a greater number of the protein molecules, leading to alternate nanostructures.

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Figure 6.5. AFM height and phase scans of films of 100qPOEGMA-PDMAEMA and native NfsB.

Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two lower molecular weight encapsulants behave similarly, but the largest does not.

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At 50% protein loading, 100qPOEGMA15-PDMAEMA5 produced poor quality films due to apparent gelation of the coating solution (Figure D.12). Scale bars at 200 nm.

Like the previous two types of encapsulants, the 20qPOEGMA-P4VP also showed the formation of complexes in solution at all blending ratios tested with increasing complexation occurring with greater protein loading (Figure 6.6). At all the blending conditions, except 10% with 20qPOEGMA15-P4VP5, the distributions are bimodal suggesting the formation of larger complexes or non-spherical morphologies.

Figure 6.6. Particle mass distributions determined by DLS for complexes formed between native

NfsB and 20qPOEGMA-b-P4VP at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading. The mass fraction of complexes increases with increasing protein loading.

The morphologies of the thin films of 20qPOEGMA-P4VP and native NfsB showed a strong dependence on both the molecular weight of the polymer and the protein loading (Figure

213

6.7). The lowest molecular weight one, 20qPOEGMA15-P4VP5, was disordered with no protein and showed the gradual increase in formation of complexes as 10 and 25% protein loading.

GISAXS shows the emergence of a slight peak at 10 and 25%, which may indicate the formation of a disordered micellar phase (Figure D.13) that was not observed by AFM likely due to insufficient mechanical contrast. At 50% protein loading these complexes dominated and led to the formation of precipitates in the coating solution, and, like the 100qPOEGMA15-PDMAEMA5 with 50% protein, this blend produced poor quality films (Figure D.12). The middle encapsulant,

20qPOEGMA36-P4VP11, was disordered at both 0 and 10% protein loading. At 25%, it showed the formation of both smaller complexes with diameter of approximately 10 nm and 50 nm at 25%, consistent with the bimodal distribution observed by DLS (Figure 6.5), but with different length scales as observed previously. Like the smallest encapsulant, this one also shows the emergence of a slight peak at 10 and 25% by GISAXS, which may indicate the formation of a disordered micellar phase (Figure D.13) that again was not observed by AFM likely due to insufficient mechanical contrast. At 50%, the structure transitioned to a microphase separated, disordered phase as has been observed with a similar encapsulant, 20% quaternized POEGMA-b-P4VP (24 vol% P4VP, with no crosslinker), with 50% protein loading of native α-amylase.45 The largest encapsulant, 20qPOEGMA55-P4VP17, shows a disordered micellar phase at 0% protein loading, observed both by AFM and GISAXS (Figure D.13). At 10% and 25% protein loading, the disordered micellar structure of the polymer alone is observed alongside smaller complexes with

10-20 nm diameter, beads-on-a-string structures consisting of 2 or more of the smaller complexes, and larger spherical complexes with diameter greater than 200 nm. For clarity, Figure D.14 provides AFM scans of larger areas of these films. Both the beads-on-a-string54-55 and the larger spherical complexes45,54 have been previously reported in polyelectrolyte complexes. At 50%, the

214 structure transitioned to a microphase separated, disordered phase similar the 20qPOEGMA36-

P4VP11.

Figure 6.7. AFM height and phase scans of films of 20qPOEGMA-P4VP and native NfsB.

Morphologies varied as a function of both the protein loading and the molecular weight of the

215 polymer. The two lower molecular weight encapsulants behave similarly at 10 and 25% protein loading, but not at 50%. At 50% protein loading the two higher molecular weight encapsulants behave similarly, while the smallest produced a poor quality film due to apparent gelation of the coating solution (Figure D.12). Scale bars at 200 nm.

Like the previous three encapsulants, when blended with the native NfsB in solution,

100qPOEGMA-P4VP also showed the formation of complexes at low protein loading and an increasing mass fraction of complexes with increased protein loading based on DLS (Figure 6.8).

With 100qPOEGMA15-P4VP5, the particle size distribution is monomodal at 10% protein loading and bimodal both 25 and 50% protein loading, again suggesting that larger aggregates or elongated nanostructures are forming. With 100qPOEGMA36-P4VP11, the particle size distribution is bimodal at 10% protein loading but transitions to a monomodal distribution at higher protein loading. With 100qPOEGMA55-P4VP17, the particle size distribution is monomodal at 10% protein, becomes bimodal at 25% with the secondary peak diminishing at 50%.

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Figure 6.8. Particle mass distributions determined by DLS for complexes formed between native

NfsB and 100qPOEGMA-b-P4VP at 1 mg mL-1 in water. Bimodal distributions of particle sizes emerged with greater protein loading for 100qPOEGMA15-P4VP5. Bimodal distributions gave way to monomodal distributions with greater protein loading for 100qPOEGMA36-P4VP11. For

100qPOEGMA55-P4VP17, the particle size distribution is only bimodal at 25%. The mass fraction of complexes increases with increasing protein loading.

The morphologies of the thin films of 100qPOEGMA-P4VP and native NfsB showed a strong dependence on both the molecular weight of the polymer and the protein loading as well

(Figure 6.9). For all 3 molecular weights, films could not be cast at 50% protein loading because the solutions became gel-like. The 100qPOEGMA15-P4VP5 appears to be lamellar or cylindrical in the absence of protein (Figure D.15 provides an enlarged view of the AFM images for clarity), but is expected to be cylindrical based on the degree of quaternization and the block ratio.56

GISAXS shows a single peak implying weak self-assembly. Addition of 10% protein led to no significant change in the morphology seen by both AFM and GISAXS (Figure D.16), which is inconsistent with the DLS. At 25%, the morphology appears to shift to micellar with the presence of some larger complexes in agreement with the DLS data. There was also a shift in the GISAXS peak from q of 0.376 nm-1 to 0.407 nm-1 corresponding to a shrinking of the length scale from 16.7 nm to 15.4 nm upon the addition of 25% protein, which is likely caused by the complexation of the polymer with the protein allowing the cationic domain to relax. Both the 100qPOEGMA36-

P4VP11 and 100qPOEGMA55-P4VP17 show similar morphologies across the three loading conditions. The pure polymers appear to show a morphology suggesting that the polymers may sit on the cusp of the body centered cubic micellar phase and the hexagonal cylinder phase, but again are expected to be cylindrical based on block ratio and degree of charge.56 GISAXS of

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100qPOEGMA36-P4VP11 again only shows a primary peak implying weak self-assembly. The

GISAXS of 100qPOEGMA55-P4VP17 shows a weak secondary peak at approximately √5q*, further suggesting that the polymer lies near the BCC-cylindrical transition. In both cases, the addition of 10% protein led to no significant change in the morphology observed by both AFM and GISAXS, as with 100qPOEGMA15-P4VP5. At 25%, both formed rod-like complexes in the presence of non-complexed block copolymer. Rod-like complexes have been previously reported in polyelectrolyte-protein complex systems, and are thought to arise when the polycations are sufficiently large to bridge between multiple protein molecules,40,57-58 consistent with the observation that this morphology did not occur with the smallest of the 100qPOEGMA-P4VPs.

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Figure 6.9. AFM height and phase scans of films of 100qPOEGMA-P4VP and native NfsB.

Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two lower molecular weight encapsulants behave similarly at 10 and 25% protein

219 loading, but not at 50%. At 50% protein loading the two higher molecular weight encapsulants behave similarly. Scale bars at 200 nm.

Complexation of the 0qPOEGMA-PDMAEMA with SC NfsB in solution led to the formation of monodisperse micelles with peak radii of 10-25 nm, with the mass fraction of complexes increasing as a function of protein loading (Figure 6.10). This behavior differs from that of the native NfsB and 0qPOEGMA-PDMAEMA, which had much greater radii and dispersity by comparison (Figure 6.2), because of the greater charge density of the supercharged protein that enables it to more easily induce charge on the non-quaternized polymer.

Figure 6.10. Particle mass distributions determined by DLS for complexes of SC NfsB and

0qPOEGMA-PDMAEMA. The mass fraction of material incorporated in the complexes increases with increased protein loading and the system forms small, monodisperse complexes.

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In thin films, the addition of SC NfsB led to a transition from the disordered polymers to the formation of micelles for all molecular weights of 0qPOEGMA-PDMAEMA at 50% protein loading, but the intermediate morphologies varied with molecular weight. The 0qPOEGMA15-

PDMAEMA5 remained disordered showed slight complexation at 10% protein loading and transitioned to disordered C3Ms at 25%. Both 0qPOEGMA36-PDMAEMA13 and

0qPOEGMA55-PDMAEMA19 form rod-like complexes at 10% protein loading with the remaining polymer remaining disordered. At 25%, both appear to transition to a microphase separated, disordered morphologies based on AFM and, in the case of 0qPOEGMA36-

PDMAEMA13, the emergence of a peak in the GISAXS pattern (Figure D.10). At 50% protein loading all 3 encapsulants appear to form disordered micellar complexes by AFM and for

0qPOEGMA36-PDMAEMA13, this is accompanied with a broadening of the peak seen in

GISAXS (Figure D.10).

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Figure 6.11. AFM height and phase scans of films of 0qPOEGMA-PDMAEMA and SC NfsB.

Morphologies varied as a function of both the protein loading and the molecular weight of the polymer. The two higher molecular weight encapsulants behave similarly, but not the lowest. Scale bars at 200 nm.

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In solution, the 100qPOEGMA-PDMAEMA behaves similarly to the previously discussed conditions with the mass fraction of complexes increasing with protein loading (Figure 6.12). The smallest encapsulant, 100qPOEGMA15-PDMAEMA5, appears to behave differently than the two larger ones and shows bimodal size distributions at 10% and 25% protein loading, while the larger ones appear monomodal. At 50% protein loading, the distributions all appear monomodal. In all cases the peak complex radii were approximately 40-45 nm, larger than with the 0qPOEGMA-

PDMAEMA. The difference in behavior between the 0qPOEGMA-PDMAEMA and

100qPOEGMA-PDMAEMA arises from the greater charge density of 100qPOEGMA-

PDMAEMA which requires interactions with greater numbers of proteins per polymer to achieve charge neutrality.

Figure 6.12. Particle mass distributions determined by DLS for complexes of SC NfsB and

100qPOEGMA-PDMAEMA. The mass fraction of material incorporated in the complexes increases with increased protein loading.

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In thin films of 100qPOEGMA-PDMAEMA and SC NfsB, morphologies differed as both a function of protein loading and the molecular weight of the polymer encapsulant. The

100qPOEGMA15-PDMAEMA5 transitions from the hexagonal or lamellar structure of the block copolymer alone to a mixture of spherical and worm-like micellar complexes observed in the AFM imaging at 10% protein loading, which is consistent with the bimodal distribution observed by

DLS. This is accompanied by a shrinking of the peak observed by GISAXS (Figure D.11), but not a disappearance which indicates some of the polymer remains non-complexed, consistent with the

DLS. At 25% protein loading, the morphology changes to disordered, microphase separated structure. At 50%, this microphase separated structure remains but with a reduction in the apparent spacing between the domains and with the presence of spherical complexes. The

100qPOEGMA36-PDMAEMA13 and 100qPOEGMA55-PDMAEMA19 behave similarly to one another. At 10%, the AFM height scans appear similar to the polymer alone, but the phase scans appear to indicate the presence of a third phase other than the cationic and neutral blocks, likely the protein partitioned into the cationic phase. At 25%, both the AFM height and phase scans show a two-phase microphase separated structure for these encapsulants, where the additional protein is likely partitioned into the cationic block. At 50%, these domains appear to swell, making the other, presumably POEGMA, domain drastically shrink.

224

Figure 6.13. AFM height and phase scans of films of 100qPOEGMA-PDMAEMA and SC NfsB.

225

Unlike the other protein-encapsulant systems discussed thus far, complexation of SC NfsB with 20qPOEGMA-P4VP did not increase with protein loading, but was instead maximized at

25% protein loading (Figure 6.14). This is because at 50% protein loading the f+ is approximately

0.38 meaning the protein is in charge excess to the polymer and cannot be fully complexed with the protein. Although in theory, the 0qPOEGMA-PDMAEMA carries a similar charge density to the 20qPOEGMA-P4VP, the PDMAEMA functionality is able to charge regulate at pH 7, while the P4VP functionality cannot. This allows the 0qPOEGMA-PDMAEMA to still effectively complex all of the protein in the system despite there being a theoretical excess while

20qPOEGMA-P4VP cannot. Apart from this behavior at 50% protein loading, the 20qPOEGMA-

P4VP system produces monodisperse micelles similar to 0qPOEGMA-PDMAEMA.

Figure 6.14. Particle mass distributions determined by DLS for complexes of SC NfsB and

20qPOEGMA-P4VP. At 10% and 25% protein loading and at 50% for the two larger encapsulants, the system forms small, monodisperse complexes. With the smallest encapsulant at 50% protein

226 loading, no apparent complexation occurs. The mass fraction of material incorporated in the complexes is maximized at 25% protein loading rather than 50% as seen with the other systems because the protein is in excess to the polymer (f+ < 0.5) for the 50% blends.

In the thin films, the three 20qPOEGMA-P4VPs each behaved differently from one another when complexed with SC NfsB (Figure 6.15). The 20qPOEGMA15-P4VP5 transitions from a disordered morphology at 0% protein loading to a disordered, microphase separated morphology when 10% protein is added, but transitions back to a disordered phase when more protein is added. The addition of 10% protein to the 20qPOEGMA36-P4VP11 led to the formation of a disordered, microphase separated morphology, which was enhanced at 25% protein loading as apparent by both the AFM and the emergence and strengthening of the peak seen by GISAXS

(Figure D.13). At 50%, the system appears more disordered, and the peak seen by GISAXS disappears (Figure D.13). At 10% protein loading, the 20qPOEGMA55-P4VP17 remains micellar as with the pure block copolymer, but with some swelling of the micelles and the emergence of some large spherical complexes with diameters of approximately 200-1000 nm (see Figure D.17).

At 25%, these spherical complexes persist, but the micelles transition to a disordered, microphase separated morphology. At 50%, fewer of the large complexes were observed (see Figure D.17) and the disordered, microphase separated morphology persists.

227

Figure 6.15. AFM height and phase scans of films of 20qPOEGMA-P4VP and SC NfsB.

228

In solution, 100qPOEGMA15-P4VP formed nearly identical mass fractions of complexes across the three mixing ratios, while 100qPOEGMA36-P4VP11 and 100qPOEGMA55-P4VP17 showed increasing mass fractions of complexes with increased protein loading (Figure 6.16). The particle mass distributions of the complexes are monomodal for all the molecular weights at 10% and 25% protein loading, while at 50%, the distributions all become bimodal. Compared to native

NfsB and 100qPOEGMA-P4VP, the behavior in solution of the SC NfsB complexes is very similar, with the bimodality emerging at greater protein loading.

Figure 6.16. Particle mass distributions determined by DLS for complexes of SC NfsB and

100qPOEGMA-P4VP. Monomodal particle size distributions at 10% and 25% protein loading gave way to bimodal distributions at 50%/ The mass fraction of material incorporated in the complexes increases with increased protein loading for the larger encapsulants, but was nearly constant for the smallest.

229

In the thin films encapsulating SC NfsB, the 100qPOEGMA-P4VP had very similar morphology to the 100qPOEGMA-PDMAEMA with the morphologies differing as both a function of protein loading and the molecular weight of the polymer encapsulant. The

100qPOEGMA15-P4VP5 showed the same morphology as 100qPOEGMA15-PDMAEMA5 at

25% and 50% protein loading, but at 10% showed structure similar to 100qPOEGMA36-

PDMAEMA13 and 100qPOEGMA55-PDMAEMA19 at 10% protein loading with an apparent third microphase observed in the AFM phase scan. The 100qPOEGMA36-P4VP11 and

100qPOEGMA55-P4VP17 behave similarly to 100qPOEGMA36-PDMAEMA13 and

100qPOEGMA55-PDMAEMA19, except at 10% protein loading. At 10%, there appear to be wormlike micellar complexes forming likely with cores consisting of the cationic block and the protein. At 25%, the microphase separated structure shows very similar morphology to

100qPOEGMA36-PDMAEMA13 and 100qPOEGMA55-PDMAEMA19, but the spacing between the neighboring cationic block-protein domains is smaller. At 50%, they are nearly identical with 100qPOEGMA36-PDMAEMA13 and 100qPOEGMA55-PDMAEMA19.

230

Figure 6.17. AFM height and phase scans of films of 100qPOEGMA-P4VP and SC NfsB.

231

6.4.3. Film Performance.

When encapsulating the native NfsB, the polymer encapsulants showed fairly similar performance in terms of initial activity with all conditions producing films with an activity of approximately 25 µM min-1 cm-2 (Figure 6.18), with one outlier, 100qPOEGMA15-PDMAEMA5 with 50% protein loading, which as previously stated produced poor quality films. Figure 6.18 shows the activities of the films encapsulating native NfsB as determined by assaying with 0.5 mM NADH and 1 mM 4-NBS before and after aging at 70 °C and 10% RH for 12 h. After crosslinking and soaking overnight to rehydrate the films, several of the film conditions with lower protein loading fully delaminated likely due to insufficient complexation or crosslinker density to achieve mechanical stability in the relatively high salinity buffer, so these data are not included.

Some films only partially delaminated and were measured, and are indicated in Figure 6.18. The films of 0qPOEGMA-PDMAEMA and native NfsB show no significant difference in both activity and thermal stability as a function of the molecular weight and no severe nonlinearity in the activity as a function of the protein loading. These behaviors are expected as the film morphologies are similar for 25% and 50% protein loading for all molecular weights of the polymer. At 50% protein loading, all 3 molecular weights showed 80% or more (within error) activity retention after aging.

The films of 100qPOEGMA-PDMAEMA showed some molecular weight dependence with activity decreasing with molecular weight, but with the highest molecular weight encapsulant showing the greatest thermal stability. The 100qPOEGMA55-PDMAEMA19 adopted a rod-like or beads-on-a-string morphology at 50% protein loading as opposed to a disordered micellar morphology seen in the two lower molecular weight encapsulants, which appears to impart greater thermal stability to the enzymes possibly by providing the enzymes an additional degree of freedom in their mobility within the films. All three encapsulants showed 80% or more (within

232 error) activity retention after aging, but 100qPOEGMA15-PDMAEMA5 was excluded from further study due to the poor quality of films produced. Films of native NfsB and both 20q- and

100qPOEGMA-P4VP did not show significant differences in performance as a function of molecular weight. The 100qPOEGMA-P4VP did show higher activity than the 20qPOEGMA-

P4VP likely due to greater enzyme retention due to the stronger charge density. The

20qPOEGMA36-P4VP11 and 20qPOEGMA55-P4VP17 did show nonlinearity in film activity as a function of protein loading with the 25% and 50% protein loading films showing similar activities and the effect being more pronounced for the larger encapsulant. This nonlinearity likely arises from the morphological shift from the disordered micellar structure similar to that of the block copolymer to the disordered, microphase separated morphology observed at 50% protein loading for these two polymer encapsulants. This phase change may arise from the increased entropic drive of the enzyme to evenly distribute across the film leading to decreased segregation of the enzyme into the cationic block,45 leading to enzyme complexing with the POEGMA block and hindering transport of the substrates through the film. Only the 100qPOEGMA55-P4VP17 at

25% protein loading and the 20qPOEGMA15-P4VP at 50% protein loading showed greater than

80% activity retention after aging, but 20qPOEGMA15-P4VP with 50% protein was excluded from further study due to the poor quality of films produced.

233

*Partially delaminated films. **Poor quality films (see Figure D.12).

The performance, both in terms of activity and thermal stability, of the films encapsulating the SC NfsB had a much stronger dependence on choice of encapsulant than with the native NfsB

(Figure 6.19). In terms of activity, the two types of encapsulants with the largest polyelectrolyte strength, 100qPOEGMA-P4VP and 100qPOEGMA-PDMAEMA, outperformed both the weaker encapsulants. 0qPOEGMA-PDMAEMA outperformed 20qPOEGMA-P4VP in terms of activity likely because the PDMAEMA domain can modulate its charge in response to the protein loading whereas P4VP cannot, allowing the 0qPOEGMA-PDMAEMA to retain a greater amount of the

234 enzymes. This is supported by the DLS data for these encapsulants with SC NfsB which showed that 0qPOEGMA-PDMAEMA was able to complex with all of the SC NfsB even at 50% loading, while the 20qPOEGMA-P4VP could not leading to the presence of free protein. As the proteins are not directly crosslinked into the films and are held in the films through electrostatics, this poor complexation likely led to release some of the enzyme while the films were rinsed after crosslinking or while they were rehydrated prior to the assay. 100qPOEGMA-PDMAEMA shows nonlinearity between protein loading and activity, with the nonlinearity becoming more pronounced with higher molecular weight. At 25% protein loading, all three 100qPOEGMA-

PDMAEMAs showed a microphase separated, disordered morphology, but the length scales observed for the 100qPOEGMA15-PDMAEMA5 were much smaller than with the two larger encapsulants. At 50% loading, the cationic domain appears to swell significantly in response to the increased protein loading, causing the POEGMA domain to shrink. This behavior was more pronounced in the two higher molecular weight encapsulants, and suggests that the shrinking of the POEGMA domain hindered transport of the substrates through previously available “channels” through the POEGMA domain. Similar nonlinearities were observed in both 0qPOEGMA-

PDMAEMA and 20qPOEGMA-P4VP, but not with 100qPOEGMA-P4VP, where the change in the channel size was less pronounced between 25 and 50% loading. In terms of thermal stability, only the 0qPOEGMA-PDMAEMA performed well with SC NfsB, with only the 0qPOEGMA15-

PDMAEMA5 with 50% protein loading showing less than 80% activity retention and with several conditions showing near 100% activity retention.

235

Figure 6.19. Activity data for films encapsulating supercharged NfsB with initial activities shown in blue and activities after aging at 70 °C and 10% RH. For films encapsulating the supercharged

NfsB, choice of encapsulant had a much greater effect on performance than with the native variant with the strongest polyelectrolytes showing the highest activity, but the weakest showing the greatest thermal stability.

Comparing the cases of the native versus the supercharged NfsB, the performance of the films encapsulating native NfsB, both in terms of the stability and the initial activity, varies less as a function of the encapsulant than with the supercharged. This difference likely arises from the relative strength of the interactions between the two encapsulated NfsB variants and the encapsulants. Due to the low net charge of native NfsB, long series of interactions involving multiple sequential monomers are unlikely to form, whereas with the supercharged variant, the greater density of negatively charged sites on the protein surface likely enables the formation of

236 these longer runs. This difference in the two variants of NfsB leads to two phenomena: relatively weak electrostatic interaction between the native NfsB and the polymer encapsulants and relatively strong electrostatic interactions between the SC NfsB and the quaternized encapsulants. In the case of native NfsB, the relatively weak interactions (resulting from the protein itself being a weaker polyelectrolyte) led to all of the polymer encapsulants having relatively similar performance.

However, in the case of the SC NfsB, strong polymer-protein interaction, especially with the 100% quaternized polymers led to high initial activity but poor thermal stability, while weaker interactions with the 0qPOEGMA-PDMAEMAs led to much greater thermal stability at the expense of some activity.

So, there seems to be an optimum between activity and stability in the C3M thin films, where tuning the interaction strength between the polymer and protein can allow for maximization of protein stability without significant sacrifices in activity. The reason this optimum may arise is that when the polymers interact strongly with the proteins, the electrostatic bonds formed are more difficult to break, and as the temperature rises and thermal fluctuations in the protein’s structure begin to occur, the bonds are not easily rearranged in response to the fluctuations. However, when these interactions are weaker, these electrostatic bonds can more easily rearrange, allowing the thermal fluctuations to occur without “pulling” on the protein. In the case of 20qPOEGMA-P4VP with SC NfsB, the encapsulant falls victim to two issues, poor activity due to low enzyme retention driven by the low charge density of the polymer (and unlike 0qPOEGMA-PDMAEMA, an inability to charge regulate to make up for this at neutral pH), and poor thermal stability because the quaternized monomer-protein bonds are relatively strong. In the case of native NfsB, these effects are more subtle, again because of the protein itself behaving as a weak polyelectrolyte, but can be seen in the slight difference between the thermal stabilities of the POEGMA-P4VP films

237 and the POEGMA-PDMAEMA films where the POEGMA-P4VP may interact slightly more strongly with the protein due to both cation-π interactions and hydrophobic interactions leading to the slightly lower thermal stabilities observed. This difference in the thermal stability may also be driven by the greater conformational freedom of the allyl side chain of PDMAEMA compared to the aromatic side chain of P4VP.

Additionally, the nonlinearities observed between protein loading and film activity for some of the polymer encapsulants suggest that there is an optimum level of protein loading beyond which the additional protein gives diminishing returns and in some cases even decreases overall activity. There are also conditions which produced poor quality films or films with insufficient mechanical properties. This optimum varied as a function of both the charge of the encapsulated protein and the choice of polymer encapsulant because it appears to be driven by morphology rather than by chemistry. However, the thermal stability of the films seems to be less influenced by the morphology. This suggests that the optimization of polymer encapsulant can be done step- wise, finding an optimal encapsulant for thermal stability

6.5. Conclusions.

Enzymatic biocatalysis plays a growing role in industry, but as progress is made towards the design and deployment of designer enzymes, systems to retain enzymes within reactors must still be deployed. Enzyme immobilization enables both the retention of the enzyme by converting the enzymes from homogeneous to heterogeneous catalysts and the stabilization of the enzyme against denaturation. Complex coacervation facilitates the encapsulation of enzymes within a polymer matrix in an aqueous process, and the coating of thin films of these complexes onto a substrate and crosslinking allows for the immobilization of enzymes via complex coacervation. This study aimed to understand how the properties of polymer encapsulants—in this case polycationic-neutral

238 diblock copolymers—can be tuned to optimize the activity and thermal stability of complex coacervate core micelle (C3M) thin films, and how these optima vary as a function of the net charge of the encapsulated protein. To explore these questions a library of 12 model encapsulants consisting of a photocrosslinkable neutral block was synthesized at three molecular weights each coupled with four model polycation chemistries all with the same volumetric block ratio was synthesized. These were then combined with a model protein, NfsB, in wither its native charge state or a supercharged state.

In general, the morphologies of the C3M thin films varied strongly as a function of protein loading, with greater protein loading typically giving rise to greater complexation. However, the specific morphologies assumed by the films varied greatly with the chemistry and molecular weight of the polymers and with the net charge of the protein. The larger molecular weight encapsulants, unsurprisingly, gave rise to larger domain sizes, which proved important to the performance of the films because these larger spacing appeared to aid in transport of the enzyme substrates through the films.

The performance of the thin films, in terms of both activity and thermal stability, seemed to be a stronger function of the choice of polymer chemistry than of the molecular weight or the protein loading. For the native enzyme, which carried a very low net charge, the performance of the encapsulants was relatively similar, whereas large differences in performance were seen with the supercharged enzyme. This difference arose from the strength of electrostatic interactions between the enzymes and the encapsulants. Due to its low net charge, the native enzyme formed only weak interactions with all of the encapsulants so factors such as the conformational freedom of the side chains and hydrophobicity of the encapsulants started to play a role in the performance of the films with greater conformational freedom favoring greater thermal stability. The

239 supercharged enzyme experienced changes in the strength of interactions as a function of the encapsulant chemistries with the two model strong polyelectrolytes showing the highest activities, but poor thermal stability, and the weak polyelectrolyte showing lower activity, but near perfect thermal stability. Nonlinearities in the activity of the films and the protein loading arose from morphological differences, with increased loading sometimes causing the closing of “channels” of the neutral block, likely negatively affecting transport of the enzyme substrates within the films.

These data suggest that two optimizations should be done for new systems, first choosing an encapsulant with the desired ratio of activity and stability, which may be highly application- dependent, and second optimizing protein loading to achieve the greatest activity per unit enzyme.

A wide range of potential encapsulants exist in literature, and using random copolymers there of further expands this parameter space, so finding a global optimum for every protein of interest may not be feasible, but these data suggest that certain design principles can be applied to the choice of encapsulant. The goal of immobilization is typically to extend the longevity of enzymes and help protect enzymes from denaturation. The data suggest that this is best accomplished by the pairing of strong and weak polyelectrolytes which produces films with the highest thermal stabilities, while possessing moderate to high activity.

240

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Chapter 7. Conclusions

7.1. Thesis Summary

At its outset, this thesis aimed to demonstrate the feasibility of using thin films of Complex coacervates to immobilize enzymes facilitating the creation of biofunctional materials. These materials were of interest because the ionic-neutral block copolymers used to construct them are known to self-assemble into a variety of nanostructures and protein loading in polyelectrolyte complex films had been previously demonstrated,1-4 so it was hypothesized that they could enable the facile synthesis of nanostructured biofunctional materials. The first study provided a proof-of- concept for the plug-and-play immobilization of enzymes via complex coacervate thin films. A model enzyme, alkaline phosphatase, was successfully immobilized in a complex coacervate thin film and was demonstrated to retain its activity in the films. The immobilized enzyme was demonstrated to be an effective biosensor for a model metal ion, Zn2+, and to have improved thermal stability compared to the solvated enzyme. The complex coacervate thin films, however, did not self-assemble into the nanoscale morphologies expected for block copolymer, forming larger kinetically-trapped structures instead likely driven by the strong interaction between the strong polycationic encapsulant and the supercharged enzyme. Overall, the method performed well, so work turned to two directions: developing a deeper understanding of protein-polymer complex coacervation and optimizing the complex coacervate thin film immobilization method.

It is widely accepted that the surface charge distribution of proteins can play a strong role on their complex coacervation behavior, but until recently a method to quantify this distribution had not been developed and the resulting effect on coacervation had not been developed.5-7 The second study aimed to quantify the surface charge distribution, or “patchiness,” of proteins and correlate this to their coacervation behavior. To systematically study the effect of charge

245 distribution, a model protein was genetically modified into 4 mutants with the same net charge but varying charge distributions. A patchiness parameter, which assessed the charge correlation between points on the surface of the protein, was used to quantify the patchiness of the designed mutants. The complexation of the model proteins was studied against a library of strong and weak polyelectrolytes, and the behavior was found to correlate with the behavior observed. Both macrophase separation and the formation of soluble aggregates were promoted by increasing patchiness depending upon the polyelectrolyte with which the protein was mixed. Increasing total charge and increasing strength of the polyelectrolyte promotes interaction for oppositely charged polyelectrolytes, while, with polyelectrolyte of like net charge, charge regulation is key to interactions because the electrolytes must interact selectively with oppositely charged patches.

Further study to understand how patchiness can be convoluted with other protein parameters such as the net charge, charge ratio, and hydrophobicity are necessary to generate robust predictive algorithms for protein coacervation.

Optimizing the performance of protein-polymer complex coacervate thin films requires an understanding of both the self-assembly behavior of the films and the effect of the self-assembly and polymer chemistry on the performance of the films. The third study aimed to understand whether complex coacervate thin films could self-assemble at a nanoscale or whether kinetics would prevent these morphologies from occurring. The complexation of a model polycationic- neutral block copolymer (BC) encapsulant was studied in solution and thin films with a series of polyanions of increasing polyelectrolyte strength. The model encapsulant was chosen such that it weakly self-assembled and would more weakly interact with the encapsulants to avoid the formation of kinetically trapped complexes. The strength of the polyanion proved to be important to the complexation with the strongest, poly(styrene sulfonate) (PSS), precipitating in solution (and

246 so not being coatable), and the weaker ones, poly(acrylic acid) (PAA) and α-amylase, both forming soluble complexes in solution, but adopting very different morphologies in thin films. PAA formed complex coacervate core micelles (C3Ms) in solution with the encapsulant, as excomplex coacervateted, and this structure translated to the films as well, with micelles being most uniform at the point of charge balance. The model proteins, native α-amylase and supercharged variants, adopted three morphologies as a function of the charge density of the encapsulant and protein and the protein loading. All of the encapsulant-protein combinations underwent two transitions, the first to a macrophase separated structure consisting of large complexes and a disordered, microphase separated phase similar to the BC alone at lower protein loading, and the second from that phase to a single disordered, microphase separated structure resembling a swollen version of the BC alone at higher protein loading. The films of the higher charge density encapsulant and supercharged amylase exhibited a third transition to a disordered phase when the protein loading exceeded the loading at the peak of the turbidity curve. The differences in the behavior of PSS and

PAA was attributed to the ability of strong polyelectrolytes to induce charge, which caused a stronger interaction between PSS and the encapsulants than between PAA and the BC. The difference in the behavior of the PAA and protein was attributed to the higher charge density of

PAA, the greater conformational freedom of PAA, and the lower affinity of PAA towards PEG functionalities compared to the protein, which led to the more regular nanostructure observed in the BC-PAA films compared to the BC-protein films.

The final study expanded on the previous one to explore how polymer chemistry

(specifically monomer conformational freedom and polycationic strength) and molecular weight and protein charge density affected the catalytic activity and thermal stability of complex coacervate thin films and how this relates to the morphologies of the films. The model protein was

247 used in its native form, with low net charge, and a supercharged form, with much higher net charge, and combined with a library of 12 neutral-polycationic block copolymers consisting of 3 molecular weight variants of 4 polymer chemistries each with the same volumetric block ratio. Some key observations about the performance of the films emerged. The first was the stronger interactions between the protein and polymer lead to greater initial activity of the films likely due to greater enzyme retention. The second was that, contrarily, weaker interaction strength between the polymer and protein leads to greater thermal stability. The third was that in cases where nonlinearities in activity and protein loading are observed, morphological changes such as swelling of the cationic domain caused by increased protein loading can dramatically affect transport through the film. Finally, when the interaction strength is held constant, more flexible monomers impart greater thermal stability on the proteins. Optimization of polymer encapsulants for target proteins will still be required until further predictive tools are developed, but the above observations provide guidelines for the deployment of this technology and help to convert it to a truly plug-and-play method in the future.

This thesis has demonstrated the use of complex coacervate thin films to immobilize enzymes and convert them into heterogeneous biocatalysts. The method is fast, aqueous, and can result in nanostructured films with high thermal stability. For most proteins, this method is essentially plug-and-play, but activity of biomaterials could be enhanced by slight optimization of cationic chemistry and protein loading to optimize stability and activity.

7.2. Future Outlook

While this thesis explored several variables to begin optimizing the performance of these films, several have not yet been studied, such as the effect of the neutral block chemistry, the block ratio between the neutral and ionic block, and wide set of potential cationic block chemistries. This

248 variable space grows further when considering the benefits of replacing a homogeneous core cationic block with a random copolymer of varying properties such as polyelectrolyte strength, hydrophobicity, and even charge to better conform to the zwitterionic protein surface. Systematic studies of not only whether systems form complexes but of how this complexation affects the performance of encapsulated enzymes is necessary to achieve full optimization of complex coacervation technology.

While this variable space is large, the advent of cheaper high-throughput technologies coupled with advances in machine-learning (ML) is making solving problems like this much more tenable. Using metrics like the patchiness parameter described in the thesis and other protein properties (charge density, hydrophobicity, negative charge ratio,8 etc.), a vector representation of proteins capturing properties relevant to their polyelectrolyte complexation behavior could be developed and used alongside existing systems to describe polymer properties to enable the formulation of predictive algorithms for protein-polymer complexation. This can be further coupled with performance metrics to enable optimization of biofunctional material design. In fact, recent work has leveraged both these technologies to begin optimizing polymer design for drug delivery of ribonucleopeptides encapsulated within polyelectrolyte complexes.9 In this case, the target protein was held constant, so machine learning was only utilized on the design of the encapsulants, but importantly machine learning enabled the optimization of polymer properties for a highly nonlinear performance metric, drug delivery.

Another interesting avenue for the further development of both this technology and biomaterials in general would be the directed evolution of materials rather than just of a protein.

Both complex coacervate systems and protein-ELP fusion materials are interesting targets for the development of this technology. In the case of coacervates, a polymer solution simply has to be

249 added prior to evaluation. In the case of the ELP-fusions nothing must be added, and the ELP itself can also be evolved for optimal performance. This technology could enable the rapid development of novel biomaterials.

In addition to optimization of encapsulation, complex coacervation technologies can be leveraged from co-encapsulation or co-immobilization of multi-enzyme cascades. Enzyme cascades can accomplish a wider variety of chemistries than singles enzymes. Co-immobilization can enable shuttling of the substrates between the various enzymes. Complex coacervates could enable the facile co-encapsulation of multiple enzymes with high-protein loading by simply blending solutions of the components together. The high-protein loading may help to enhance the shuttling of substrates between the enzymes. Additionally, the polymer chemistry and synthetic route may be optimized to encourage partitioning of the substrates into the coacervate while also encouraging partitioning of the product out of the coacervate allowing for “equilibrium-breaking” as well. The recent discovery of multiphase coacervates may enable shuttling of substrates along a synthetic route as well, but this requires a deeper understanding of the properties of these materials.

As understanding of the complex coacervation of proteins has evolved in recent years, the field is ripe for new work focusing on the function of these materials as opposed to only the form.

Wider studies of how properties like the morphologies of complex coacervates and C3Ms and the polymer chemistry affect various performance metrics like thermal and chemical stability and catalytic activity will enable conversion of this technology from an academic curiosity to a commercially deployable technique. Early work in this direction, including the work presented in this thesis, show great promise for the future of these materials.

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7.3. References

1. Stewart-Sloan, C. R.; Olsen, B. D., Protonation-Induced Microphase Separation in Thin Films of a Polyelectrolyte-Hydrophilic Diblock Copolymer. ACS Macro Lett. 2014, 3 (5), 410- 414. DOI: 10.1021/mz400650q 2. Stewart-Sloan, C. R.; Wang, R.; Sing, M. K.; Olsen, B. D., Self-Assembly of Poly(vinylpyridine-b-oligo(ethylene glycol) methyl ether methacrylate) Diblock Copolymers. J. Polym. Sci., Part B: Polym. Phys. 2017, 55 (15), 1181-1190. DOI: 10.1002/polb.24369 3. Stewart-Sloan, C. R. Understanding the effect of protonation on the self-assembly of a model polyelectrolyte-neutral block copolymer. Ph.D. Thesis, Massachusetts Institute of Technology, Cambridge, MA, 2016. 4. Kim, B.; Lam, C. N.; Olsen, B. D., Nanopatterned Protein Films Directed by Ionic Complexation with Water-Soluble Diblock Copolymers. Macromolecules 2012, 45 (11), 4572-4580. DOI: 10.1021/ma2024914 5. Blocher McTigue, W. C.; Perry, S. L., Design Rules for Encapsulating Proteins into Complex Coacervates. Soft Matter 2019, 15 (15), 3089-3103. DOI: 10.1039/C9SM00372J 6. Cooper, C. L.; Dubin, P. L.; Kayitmazer, A. B.; Turksen, S., Polyelectrolyte–Protein Complexes. Curr. Opin. Colloid Interface Sci. 2005, 10 (1–2), 52-78. DOI: 10.1016/j.cocis.2005.05.007 7. Xu, Y.; Mazzawi, M.; Chen, K.; Sun, L.; Dubin, P. L., Protein Purification by Polyelectrolyte Coacervation: Influence of Protein Charge Anisotropy on Selectivity. Biomacromolecules 2011, 12 (5), 1512-1522. DOI: 10.1021/bm101465y 8. Obermeyer, A. C.; Mills, C. E.; Dong, X.-H.; Flores, R. J.; Olsen, B. D., Complex Coacervation of Supercharged Proteins with Polyelectrolytes. Soft Matter 2016, 12 (15), 3570-3581. DOI: 10.1039/C6SM00002A 9. Kumar, R.; Le, N.; Tan, Z.; Brown, M. E.; Jiang, S.; Reineke, T. M., Efficient Polymer-Mediated Delivery of Gene-Editing Ribonucleoprotein Payloads through Combinatorial Design, Parallelized Experimentation, and Machine Learning. ACS Nano 2020, DOI: 10.1021/acsnano.0c08549

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253

Appendix A. Supporting Information for Chapter 3.

Reproduced (adapted) with permission from Sureka, et al. Catalytic Biosensors from Complex

Coacervate Core Micelle (C3M) Thin Films, ACS Appl. Mater. Interfaces 2019, 11, 35, 32354–

1 Figure A.1. H NMR (400 MHz, CDCl3) of poly((oligo-ethylene glycol methacrylate)-r-

(benzophenone methacrylate)) (POEGMA-r-BP). Comparison of the F and G peaks to the D peak was used to determine the ratio of BP to OEGMA in the polymer.

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1 Figure A.2. H NMR (400 MHz, CDCl3) of poly-[(oligo-ethylene glycol methacrylate)-r-

(benzophenone methacrylate)]-b-(4-vinylpyridine)] ((POEGMA-r-BP)-b-P4VP). Comparison of the I and J peaks to the D peak was used to determine the ratio of 4VP to OEGMA in the polymer.

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1 Figure A.3. H NMR (400 MHz, D2O) of poly-[(oligo-ethylene glycol methacrylate)-r-

(benzophenone methacrylate)]-b-(methyl-quaternized 4-vinylpyridine)] ((POEGMA-r-BP)-b- qP4VP). The total aromatic peaks are equal to the previously measured peaks. The total integral of the A (2H) and K (3H) peaks is expected to be 4.94 for complete quaternization of the polymer because the ratio of 4VP to OEGMA is 0.98 and for peak A is expected to be 2, while the new methyl peak is expected to be 2.94. Peaks are normalized against peak D (3H). The polymer was quaternized > 95%.

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Figure A.4. SEC analysis of POEGMA-r-BP. The polymer was found to have Mn of 31.8 kDa and Đ of 1.13.

Figure A.5. SEC analysis of (POEGMA-r-BP)-b-P4VP. Đ was 1.11.

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Figure A.6. a) SDS-PAGE analysis of PhoA mutants before and after periplasmic purification and b) after FPLC purification. c) Representative Michaelis-Menten fits to enzyme activity.

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Figure A.7. Turbidimetry data for qP4VP and PhoA.

Determination of PhoA coacervation. The method described by Obermeyer, et al.1 was used to characterize the coacervation behavior of the PhoA with qP4VP in order to determine the best mixing ratio for the copolymer with PhoA. The point of maximum turbidity was determined to

+ be at f of 0.82 ± 0.05. The block copolymer was determined to have a final Mn of 55.5 kDa based on full quaternization as follows:

푀푛,푓푖푛푎푙 = 푀푛,푖푛푖푡푖푎푙 + 퐷푃4푉푃 × 푀푊퐶퐻3퐼 × % 푞푢푎푡푒푟푛푖푧푎푡푖표푛 = 41100 + 96 × 141.94 × 100% (1)

Based on full quaternization, each polymer molecule carries a charge of +96. At pH 7, PhoA M4 carries a net charge of -16.7.

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Figure A.8. AFM height and phase images for C3M film before crosslinking (a,b), after crosslinking (c,d), and after soaking (e,f). Scale bars are 200 nm and z-scales are 15 nm and 15° for height and phase, respectively. The treatments did not lead to significant changes of the film morphology.

Table A.1. Contact angle of C3M Films before and after crosslinking and after soaking in buffer as determined on a goniometer. The films were found to be hydrophilic in all cases. However, soaking the films led to a marked increase in the contact angle, likely due to excess non crosslinked polymer and protein on the surface being washed away.

Film condition Contact Angle As cast 49.4 ± 5.8° Crosslinked 51.5 ± 1.3° Crosslinked and soaked 81.4 ± 0.5°

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Figure A.9. Scanning probe microscope (SPM) height (a) and phase (b) images and and grazing incidence small-angle X-ray scatering (GISAXS) pattern (c) for film with 5% (wt/wt) protein loading. Scale bars are 200 nm and z-scales are 20 nm (a) and 10° (b). Scattering intensity is shown on a logarithmic scale. Film thickness was 120 ± 2 nm. SPM shows formation of large complexes; however, GISAXS indicates that the underlying structure is the same as that of the pure polymer (disordered micelles).

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Horizontal Vertical

Figure A.10. The qy and qz linecuts of GISAXS data for films with increasing protein loading.

The qy linecut was averaged over qz of 0.2 and 0.4 and the qz linecut was averaged over qy of

0.05 to 0.15. The structure showed diminishing peaks in both qy and qz with increasing protein concentration, indicating that the nanostructure of the polymer gives way to the formation of larger features with increased protein content. The dual peak in qz is observed due to internal reflections, while the peak at 0.30 is the position of the reflected beam. Based on both SPM and

GISAXS the polymer appears to form a disordered micellar phase.

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Figure A.11. GISAXS patterns for PhoA-polymer films annealed with different solvents.

Intensity is shown on a logarithmic scale. The structure showed no significant change with annealing condition.

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Figure A.12. Relative response to 5 ppm Zn2+ in the presence of various biological contaminants at relevant concentrations. No statistical difference was found for any of the contaminant tested

(p > 0.05 vs. control). The concentration of glucose is 1 mM, based on the total concentration of carbohydrates reported in the sediment layer of rivers (1 mM)2, and much greater than that reported in river water (2.3 µM).3 The protein concentration is 10 mg L-1, based on the reported amino acid content in river water sediment (6.7 mg L-1),4 and significantly higher than reported amino acid content in the water column (less than 1 mg L-1).5-6 The E. coli concentration is 2300

± 200 CFU per 100 mL, which is greater than the EPA standard for bacteria in recreational water

(410 CFU per 100 mL).7 The mixture contains all of the contaminants at the previous concentrations.

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Table A.2. Statistical analysis (p-values for 1-tail T-test) of testing data

Condition 10 ppb Zn 50 ppb Zn Control 0.01 0.002 10 ppb Ni 0.287 0.001 10 ppb Co 0.016 0.000 10 ppb Cu 0.009 0.002 CRW 0.058 0.009 CRWF 0.123 0.052 Mix 1 0.085 0.020 Mix 2 0.136 0.018

Figure A.13. 1H NMR of BP Monomer.

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Figure A.14. LRMS of BP Monomer. Expected mass was 266.1, found was 266.1.

5’- GTGAAACAAAGCACTATTGCACTGGCACTCTTACCGTTACTGTTTACCCCTGTGACAAAA 60 GCCCGGACACCAGAAATGCCTGTTCTGGAAAACCGGGCTGCTCAGGGCGATATTACTGCA 120 CCCGGCGGTGCTCGCCGTTTAACGGGTGATCAGACTGCCGCTCTGCGTGATTCTCTTAGC 180 GATAAACCTGCAAAAAATATTATTTTGCTGATTGGCGATGGGATGGGGGACTCGGAAATT 240 ACTGCCGCACGTAATTATGCCGAAGGTGCGGGCGGCTTTTTTAAAGGTATAGATGCCTTA 360 CCGCTTACCGGGCAATACACTCACTATGCGCTGAATAAAAAAACCGGCAAACCGGACTAC 420 GTCACCGACTCGGCTGCATCAGCAACCGCCTGGTCAACCGGTGTCAAAACCTATAACGGC 480 GCGCTGGGCGTCGATATTCACGAAAAAGATCACCCAACGATTCTGGAAATGGCAAAAGCC 540 GCAGGTCTGGCGACCGGTAACGTTTCTACCGCAGAGTTGCAGGATGCCACGCCCGCTGCG 600 CTGGTGGCACATGTGACCTCGCGCAAATGCTACGGTCCGAGCGCGACCAGTGAAAAATGT 660 CCGGGTAACGCTCTGGAAAAAGGCGGAAAAGGATCGATTACCGAACAGCTGCTTAACGCT 720 CGTGCCGACGTTACGCTTGGCGGCGGCGCAAAAACCTTTGCTGAAACGGCAACCGCTGGT 780 GAATGGCAGGGAAAAACGCTGCGTGAACAGGCACAGGCGCGTGGTTATCAGTTGGTGAGC 840 GATGCTGCCTCACTGAATTCGGTGACGGAAGCGAATCAGCAAAAACCCCTGCTTGGCCTG 900 TTTGCTGACGGCAATATGCCAGTGCGCTGGCTAGGACCGAAAGCAACGTACCATGGCAAT 960 ATCGATAAGCCCGCAGTCACCTGTACGCCAAATCCGCAACGTAATGACAGTGTACCAACC 1020 CTGGCGCAGATGACCGACAAAGCCATTGAATTGTTGAGTAAAAATGAGAAAGGCTTTTTC 1080 CTGCAAGTTGAAGGTGCGTCAATCGATAAACAGGATCATGCTGCGAATCCTTGTGGGCAA 1140 ATTGGCGAGACGGTCGATCTCGATGAAGCCGTACAACGGGCGCTGGAATTCGCTAAAAAG 1200 GAGGGTAACACGCTGGTCATAGTCACCGCTGATCACGCCCACGCCAGCCAGATTGTTGCG 1260 CCGGATACCAAAGCTCCGGGCCTCACCCAGGCGCTAAATACCAAAGATGGCGCAGTGATG 1320 GTGATGAGTTACGGGAACTCCGAAGAGGATTCACAAGAACATACCGGCAGTCAGTTGCGT 1380 ATTGCGGCGTATGGCCCGCATGCCGCCAATGTTGTTGGACTGACCGACCAGACCGATCTC 1440 TTCTACACCATGAAAGCCGCTCTGGGGCTGAAATAA -3’ 1476

Figure A.15. Sequence of PhoA.

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(a) K92D + K96D sense: 5’-TACACTCACTATGCGCTGAATGATAAAACCGGCGATCCGGACTACGTCACCG-3’

antisense: 5’-GTAGTCCGGATCGCCGGTTTTATCATTCAGCGCATAGTGAGTGTATTGCCCG-3’ (b) K92D + K93D + K96D sense: 5’-GCAATACACTCACTATGCGCTGAATGATGATACCGGCGATCCGGACTACGTCACCGAC-3’

antisense: 5’-ACGTAGTCCGGATCGCCGGTATCATCATTCAGCGCATAGTGAGTGTATTGCCCGGTAA-3’ (c) K352D + K353D sense: 5’-CGCTGGAATTCGCTGATGATGAGGGTAACACGCTGGTCATA-3’

antisense: 5’-AGCGTGTTACCCTCATCATCAGCGAATTCCAGCGCCCGTTG-3’

Figure A.16. Primer Designs for engineering of PhoA.

Preparation of standard curves. Standard curves were prepared for both the 96-well (Figure

S14) and 12-well (Figure S15) plates using 4-methylumbelliferone (4-MU) (Aldrich). Individual

10 mM stock solutions of 4-MU were made in dimethyl sulfoxide (DMSO) and diluted in water to 1 mM. For the 96-well plate standard, separate 1 mM stock solutions were further diluted to

500, 300, 100, 75, 50, 25, and 10 µM in water. 180 µL of 100 mM Tris pH 8.0 was added to 24 wells. To 8 wells, 20 µL of 500, 300, 100, 75, 50, 25, 10, and 0 µM 4-MU was added. The emission at 448 nm was measured after exciting at 362 nm. 20 µL of the wells’ contents were then added to the next well, providing a ten-fold dilution, and the excitation was measured again. This was repeated once more. For the 12-well plate, the 1 mM stock was diluted to 750, 500, and 250 µM.

Data was collected at final concentrations of 100, 75, 50, 25, 10, 7.5, 5, 2.5, 1, 0.5, 0.25, and 0 µM and fitted to a line. High concentrations were excluded due to non-linearity and because experiments never reached that level of conversion.

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Figure A.17. a) Standard curve in 96-well plates in MOPS buffer. b) Standard curve in 96-well plate in Tris buffer. c) Standard curve in 12-well plate in Tris buffer.

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Figure A.18. Activity assay with and without additional DMSO to evaluate solvatochromic effects of DMSO concentration. The points at 50 μM are collected at the same conditions. The

-1 -1 additional DMSO caused a negligible change in kcat from 7.45 ± 0.45 s to 7.17 ± 0.62 s ; however, it did cause a significant shift in KM from 1.7 ± 0.6 μM to 2.6 ± 0.6 μM. For the metals assay, this was not considered significant because the assays were conducted at a concentration such that the enzyme was expected to achieve its maximum reaction rate.

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Figure A.19. Negative control with carbon tape (CT). Plotted against protein loading data for reference. Carbon tape had negligible activity.

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References

1. Obermeyer, A. C.; Mills, C. E.; Dong, X.-H.; Flores, R. J.; Olsen, B. D., Complex Coacervation of Supercharged Proteins with Polyelectrolytes. Soft Matter 2016, 12 (15), 3570-3581. DOI: 10.1039/C6SM00002A 2. Westrich, B.; Förstner, U., Sediment Dynamics and Pollutant Mobility in Rivers: An Interdisciplinary Approach. Springer Science & Business Media: 2007. 3. Wicks, R. J.; Moran, M. A.; Pittman, L. J.; Hodson, R. E., Carbohydrate Signatures of Aquatic Macrophytes and Their Dissolved Degradation Products as Determined by a Sensitive High- Performance Ion Chromatography Method. Applied and Environmental Microbiology 1991, 57 (11), 3135-3143. 4. Thomas, J. D., The Role of Dissolved Organic Matter, Particularly Free Amino Acids and Humic Substances, in Freshwater Ecosystems. Freshwater Biology 1997, 38 (1), 1-36. DOI: doi:10.1046/j.1365-2427.1997.00206.x 5. Lytle, C. R.; Perdue, E. M., Free, Proteinaceous, and Humic-Bound Amino Acids in River Water Containing High Concentrations of Aquatic Humus. Environmental Science & Technology 1981, 15 (2), 224-228. DOI: 10.1021/es00084a009 6. Chinn, R.; Barrett, S. E., Occurrence of Amino Acids in Two Drinking Water Sources. In Natural Organic Matter and Disinfection By-Products, American Chemical Society: 2000; Vol. 761, pp 96-108. 7. Environmental Protection Agency. Recreational Water Quality Criteria; 2012.

271

Appendix B. Supporting Information for Chapter 4.

Reproduced (adapted) in part with permission from Kim,* Sureka,* et al. Effect of Protein Surface

Charge Distribution on Protein–Polyelectrolyte Complexation, Biomacromolecules 2020, 21, 8,

Figure B.1. GFP variants model combined by EDTSurf and APBS.

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XLP GFP DNA sequence: ATGGGTCACCACCACCACCACCACGGTAGCGCGTGCGAACTGATGGTTAGCAAGGGCGAGG AACTGTTCACCGGTGACGTGCCGATCCTGGTTGAGCTGGACGGCGATGTGAACGGTCACGAC TTTAGCGTTCGTGGTGAGGGCGAAGGTGATGCGACCAACGGCGAACTGACCCTGAAATTCAT TTGCACCACCGGTGAACTGCCGGTGCCGTGGCCGACCCTGGTTACCACCCTGACCTACGGTG TGCAGTGCTTTAGCCGTTATCCGAAGCACATGAAACGTCACGACTTCTTTAAGAGCGCGATG CCGAAAGGCTACGTTCAAGAACGTACCATCAGCTTCAAGAAAGATGGCAAGTATAAAACCC GTGCGGAAGTGAAGTTTAAAGGCCGTACCCTGGTTAACCGTATCGAGCTGAAGGGTGAAGA CTTCAAAGAGGATGGCAACATTCTGGGTCACAAGCTGGAATACAACTTTAACAGCCACGAC GTGTATATCACCGCGGATAAGGAGAAGAACGGCATCAAGGCGGAGTTTAAAATTCGTCACA ACGTGGAGGACGGTAGCGTTCAGCTGGCGGATCACTACCAGCAAAACACCCCGATTGGCAA GGGTCCGGTTCTGCTGCCGCGTCGTCACTATCTGAGCACCCGTAGCAAGCTGAGCAAAGACC CGAACGAGGAACGTGATCACATGGTGCTGCTGGAATTCGTTACCGCGGCGGGCATTGACCA CGGTATGGATGAGCTGTACAAA LP GFP DNA sequence: ATGGGTCACCACCACCACCACCACGGTAGCGCGTGCGAGCTGATGGTTAGCAAAGGCGAGG AACTGTTCGAGGGTGACGTGCCGATCCTGGTTGAACTGGACGGCGATGTGAACGGTCACGA ATTTAGCGTTCGTGGTGAGGGCGAAGGTGATGCGACCAAGGGCGAGCTGACCCTGAAATTC ATTTGCACCACCGGTGAACTGCCGGTGCCGTGGCCGACCCTGGTTACCACCCTGACCTACGG TGTGCAGTGCTTTAGCCGTTATCCGAAGCACATGAAACAACACGACTTCTTTAAGAGCGCGA TGCCGGAGGGCTACGTTCAGGAACGTACCATCAGCTTCAAGGACGATGGTACCTATAAAAC CCGTGCGGAAGTGAAGTTTGAAGGCGACACCCTGGTTAACCGTATCGAGCTGAAGGGTAAA GATTTCAAGGAAAAAGGCAACATTCTGGGTCACAAACTGGAGTACAACTTTAACAGCCACC GTGTGTATATCACCGCGGATAAGCGTAAAAACGGCATCAAGGCGGAATTTAAAATTCGTCA CAACGTGAAGGACGGTAGCGTTCAACTGGCGGATCACTACCAGCAAAACACCCCGATTGGT CGTGGTCCGGTTCTGCTGCCGCGTCGTCACTATCTGAGCACCCGTAGCGCGCTGAGCAAGGA CCCGAAAGAGGAACGTGATCACATGGTGCTGCTGGAGTTCGTTACCGCGGCGGGCATTGAC CACGGTATGGATGAACTGTACAAA MP GFP DNA sequence: ATGGGTCACCACCACCACCACCACGGTAGCGCGTGCGAGCTGATGGTTAGCAAAGGCGAAA CCCTGTTCACCGGTGTGGTTCCGATCCTGGTGCAGCTGAACGGCGACGTTAACGGTCACGAG TTTAGCGTTCGTGGTAGCGGCACCGGTGATGCGACCAACGGCCAACTGACCCTGAAGTTCAT TTGCACCACCGGTGAACTGCCGGTGCCGTGGCCGACCCTGGTTACCACCCTGACCTACGGTG TGCAGTGCTTTAGCGCGTATCCGAACCACATGAAGGCGCACGACTTCTTTAAAAGCGCGATG CCGAACGGCTACGTTCAAGAGCGTACCATCAGCTTCAAGAACGACGGTACCTATAAAACCC GTGCGGAAGTGAAGTTTGCGGGCGATACCCTGGTTAACCGTATCGCGCTGAAGGGTATTGAC TTCAAAGAGGATGGCAACATCCTGGGTCACAAACTGGAATACAACTTTAACAGCCACAACG TGTATATTACCGCGGACAAACAGGCGAACGGCATCAAGGCGAACTTTGCGATTCGTCACAA CGTGCAGGATGGTAGCGTTCAACTGGCGACCCACTACCAGCAAAACACCCCGATCGGCAAC GGTCCGGTTCTGCTGCCGGATAACCACTATCTGAGCACCCAAAGCGCGCTGAGCAAAGACCC GAACGAGACCCGTGATCACATGGTGCTGCTGGAATTCGTTACCGCGGCGGGCATTACCCACG GTATGAACGAGGTGTACAAG SP GFP DNA sequence: ATGGGTCACCACCACCACCACCACGGTAGCGCGTGCGAGCTGATGGTTAGCAAGGGCGCGG AACTGTTCGACGGTAAAGTGCCGATCCTGGTTGAACTGAAGGGCGATGTGAACGGTCACAA

273

ATTTAGCGTTCGTGGTGAGGGTAACGGTGATGCGACCCGTGGCGATCTGACCCTGAAGTTCA TTTGCACCACCGGTGAACTGCCGGTGCCGTGGCCGACCCTGGTTACCACCCTGACCTACGGT GTGCAGTGCTTTAGCCGTTATCCGGACCACATGAAGCAACACGATTTCTTTAAAAGCGCGAT GCCGGAGGGCTACGTTCAGGAACGTACCATCAGCTTCAAGGACGATGGCAAGTATAAAACC CGTGCGGAAGTGAAATTTGAAGGCGACACCCTGGTTAACCGTATCGAGCTGAAGGGTATTG ATTTCAAGGAAAAAGGCAACATCCTGGGTCACAAACTGGAGTACAACTTTAACAGCCACGA CGTTTATATTACCGCGGATAAGGAGAAAAACGGCATCAAGGCGGAATTTAAAATTCGTCAC AACGTGCGTGACGGTAGCGTTCAACTGGCGGATCACTACCAGCAAAACACCCCGATTGGCA AGGGTCCGGTGCTGCTGCCGGACAAACACTATCTGAGCACCGAGAGCGTTCTGAGCAAAGA CCCGCGTGAAGCGCGTGATCACATGGTGCTGCTGGAGTTCGTTACCGCGGCGGGCATTACCC ACGGTGACAAGGAAGATTACAAA Figure B.2. DNA sequences of GFP mutants

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275

276

1 Figure B.3. H NMR (500 MHz) in D2O of a) hyaluronic acid, b) sodium salt of PAA, c) qP4VP (peak at

4.25 ppm confirms full quaternization, approx.. 5% (wt/wt) residual DMF), d) PSS, and e) PDMAEA (C′,

D′, E′ correspond non-protonated PDMAEA, approx.. 5%).

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Figure B.4. Gel permeation chromatography of synthesized charged polymers; PAA, qP4VP, PSS and

PDMAEA.

278

Table B.1. The number, size and potential of each patch in GFP mutants

Size of Pair # of Potential # of Size of - Potential Neutral Protein +Patch correlation Patch (KT/ec) Patch Patch (Å) (KT/ec) area(Å) (Å) f XLP_0 1 3672.45 4.50 1 3935.28 -4.14 3692.97 0.66 XLP_50 1 3116.22 4.07 1 2774.18 -3.46 5410.29 0.51 XLP_300 1 1571.63 3.33 1 349.08 -3.93 7820.16 0.30 2 154.65 12.38 2 428.62 -3.03 3 402.33 2.64 3 574.23 -2.78 LP_0 1 3244.40 4.13 1 3350.75 -4.25 4599.10 0.59 2 148.65 2.71 LP_50 1 2648.62 3.84 1 2391.87 -3.69 6135.94 0.45 2 166.48 2.74 1 831.21 3.31 1 685.72 -3.66 8566.78 0.25 LP300 2 243.18 9.33 2 461.95 -3.01 3 236.35 2.81 3 194.12 -3.02 4 123.59 2.70 MP_0 1 350.67 3.10 1 105.13 -2.32 7255.71 0.32 2 326.42 2.71 2 1753.86 -3.25 3 638.95 2.93 3 184.16 -3.18 MP_50 1 334.28 3.03 1 165.01 -3.07 8070.29 0.25 2 346.82 2.70 2 124.11 -2.79 3 686.21 2.92 3 888.18 -2.85 MP_300 1 267.46 2.73 1 136.49 -2.91 9713.76 0.13 2 166.18 2.84 2 204.31 -2.97 3 126.70 3.01 SP_0 1 840.73 5.26 1 406.93 -2.62 8662.14 0.25 2 265.09 2.65 2 399.01 -3.26 3 468.43 -3.46 4 241.47 -3.00 SP_50 1 773.31 5.17 1 119.07 -2.42 9328.24 0.20 2 245.23 2.59 2 191.83 -2.98 3 325.83 -3.29 4 103.30 -3.40 5 197.00 -2.94 SP_300 1 589.16 5.30 1 136.59 -2.77 10052.74 0.14 2 136.17 2.49 2 229.37 -2.96 3 139.76 -2.85

279

Figure B.5. Positive (blue) and negative (red) electrostatic field isosurface contours for GFP mutants (±2

-1 kBT ec )

280

Figure B.6. Purity analysis of GFP mutants after Ni-NTA purification. Main peaks are saturated for all mutants, but represent approximately 90% of all contents. Saturation prevents determination of true purity, which would be greater than 90%.

281

-1 Figure B.7. Titration data for PDMAEA. pKa was found to be 7.32 ± 0.05. 25 mL of 2 mg mL polymer was acidified to pH 2.56 and titrated with 50 mM NaOH.

Table B.2. Negative charge ratio (α) values of GFP mutants. α value = (# of D + # of E + N-terminal) / (#of

K + # of R + C terminal)

282

1 Figure B.8. H NMR of synthesized qP4VP in D2O. The peak at 4.25 ppm shows fully quaternized methyl peak. Approximately 5% (wt/wt) residual DMF was found in the final polymer.

Synthesis of 2-ethylsulfanylthiocarbonylsulfanyl-2-methylpropionic acid (EMP). The synthesis

1 method of the CTA was adapted from a previous study. K3PO4·H2O (5.71 g, 1.1 eq) was placed in 100 mL acetone and stirred for 20 min at ambient temperature. Ethanethiol (2 g, 1.2 eq) was added to a suspension of K3PO4·H2O. After 1hour, carbon disulfide (4.53 mL, 3 eq) was added, and the solution turned bright yellow. 2-bromoisobutyric acid (8.35 g, 1 eq) was slowly added, and then the mixture was stirred overnight. 300mL of 1M HCl was added to the mixture on an ice bath, and then the crude material was extracted with DCM (150 mL) and washed two times with HCl. The extracted material was again washed with deionized water and brine separately, and the material was dried over Na2SO4, and filtered. After

283 drying the crude product, silica gel (hexanes:ethyl acetate = 2:1) was utilized to purify EMP. After collect fraction, the material was dried under vacuum and a yellow solid was obtained (yield, 80%).1H NMR

(CDCl3, δ): 1.33 (t, 3H, -SCH2CH3), 1.72 (s, 6H, -(C=O)C(CH3)2S-), 3.30 (q, 2H, -SCH2CH3).

284

MATLAB Code Function to convert protein data files to usable matrix and run all subsequent analyses. function output=runall()

[BASE, fold]=uigetfile('*.pdb'); %[BASE, fold]=uigetfile('*.pdb');

SUB=uigetfile('*.dx');

BASE=[BASE(1:end-4)]; SUB=SUB(length(BASE)+1:end-3);

BASE=[fold,BASE]; getSurface_AB(BASE,SUB); output1=testScript1_AB(BASE,SUB);

SA=inputdlg("input SA value"); SA=str2num(SA{1});

[ppatchdata,ppatches,npatchdata,npatches,neuarea]... =protpatch2(output1(:,1), output1(:,2:4), 2, -2,1,100,SA); output={ppatchdata,ppatches,npatchdata,npatches,neuarea}; output{end+1}=patchcorr(output1(:,1), output1(:,2:4), 2, -2, 2); savefig([BASE,SUB,'_Fig_4.fig']); save([BASE,SUB],'output');

Function to convert .ply file (from EDTsurf) and .dx file (from APBS) to a [x, y, z, q] matrix at the SAS. function output1 = testScript1_AB(PDB_ID,PDB_SUB) %% Post-processing of APBS output file % % Input files include: % ply file: PDB#.ply % OpenDX file from APBS: PDB#DX.txt % % This scrips reads the ply file and potential values from the APBS output, which can % be then further processed to get useful information

%clear;clc;

%PDB_ID = '2p3b'; % The DX output seems to be rotated %PDB_ID = 'Janus';

% Read the ply file: surface mesh generated from EDTSurf % Ref: https://zhanglab.ccmb.med.umich.edu/EDTSurf/ PlyData = pcread([PDB_ID,'.ply']); PlyCoor = PlyData.Location;

285

% Read the APBS output OpenDX file % Ref: http://apbs- pdb2pqr.readthedocs.io/en/latest/formats/opendx.html % Get grid information dxFileId = fopen([PDB_ID,PDB_SUB,'DX.txt'], 'r'); for i = 1:4 fgetl(dxFileId); % Irrelevant lines end gridCountsLine = fgetl(dxFileId); nxyz = sscanf(gridCountsLine, 'object 1 class gridpositions counts %d %d %d'); nx = nxyz(1); ny = nxyz(2); nz = nxyz(3); originLine = fgetl(dxFileId); origin = sscanf(originLine, 'origin %f %f %f'); dxLine = fgetl(dxFileId); dx = sscanf(dxLine, 'delta %f %f %f'); dx = dx(1); dyLine = fgetl(dxFileId); dy = sscanf(dyLine, 'delta %f %f %f'); dy = dy(2); dzLine = fgetl(dxFileId); dz = sscanf(dzLine, 'delta %f %f %f'); dz = dz(3); fclose(dxFileId);

% Get potential information dxFileObj = importdata([PDB_ID,PDB_SUB,'DX.txt'], ' ', 11); dxFileData = dxFileObj.data'; dxFileData = reshape(dxFileData, size(dxFileData,2)*3, 1); dxFileData(isnan(dxFileData)) = []; % Remove NaN entries potDX = reshape(dxFileData, nz, ny, nx); % nz * ny * nx potDX = permute(potDX, [2,3,1]); % ny * nx * nz -> more natural plotting

% The grid on x,y,z xGrid = origin(1) + (0:nx-1)*dx; yGrid = origin(2) + (0:ny-1)*dy; zGrid = origin(3) + (0:nz-1)*dz;

% Plot the results figure('Color', [1 1 1]); isosurface(xGrid, yGrid, zGrid, potDX, 2); hold on; isosurface(xGrid, yGrid, zGrid, potDX, -2); plot3(PlyCoor(:,1), PlyCoor(:,2), PlyCoor(:,3), 'k.');

surfPot = interp3(xGrid, yGrid, zGrid, potDX, PlyCoor(:,1), PlyCoor(:,2), PlyCoor(:,3)); savefig([PDB_ID,PDB_SUB,'_Fig_1.fig']);

figure('Color', [1 1 1]); isosurface(xGrid, yGrid, zGrid, potDX, 2); hold on; isosurface(xGrid, yGrid, zGrid, potDX, -2); colormap(rwbColormap(50)); 286 savefig([PDB_ID,PDB_SUB,'_Fig_2.fig']); figure('Color', [1 1 1]); scatter3(PlyCoor(:,1), PlyCoor(:,2), PlyCoor(:,3), 40, surfPot); caxis([-5,5]); colormap(rwbColormap(50)); savefig([PDB_ID,PDB_SUB,'_Fig_3.fig']); output1=[surfPot,PlyCoor];

Function to locate and evaluate surface charge patches. function [ppatchdata,ppatches,npatchdata,npatches,neuarea]=protpatch2(q,X,pcuto ff,ncutoff,r,minsize,SA) %ppatchdata includes area and average potential of all positive patches

%npatchdata includes area and average potential of all negative patches

%p&npatches contain in a cell array the coordinates and potentials of all %points on the patches

%neuarea is the total area in sq. angstroms that is not included in patches

%q is the vector of potentials

%X is the matrix of coordinates on the surface to which q is mapped

%pcutoff and ncutoff are the positive and negative cutoff potentials for evaluation

%r is the minimum separation between patches in angstroms. %Note: this is not calculated as a distance along the surface but rather an %absolute distance

%minsize is the minimum size of a patch in terms of sq. angstroms

%SA is the surface area of the protein in square angstroms.

numpt=length(q); %stores the number of total points X(:,4)=q;%joins coord and q into one matrix ptarea=SA/size(X,1);%average area per point p=[]; parea=0;

%for loop to find isosurface

287

%the vector p contains coordinates and potentials of all points on the %isosurface for i=1:numpt if q(i)>=pcutoff p(end+1,:)=X(i,:); %find all points with positive pot>pcutoff

end end k=1; del=[]; patchpos=[]; ppatches={}; piso=[];

% Breadth first search for patches %only finds patches with size>minsize. minsize in terms of sq angstroms. while ~isempty(p) j=1; patchpos(1,:)=p(1,:); p(1,:)=[]; if size(p,1)>1 while j<=size(patchpos,1) for i=1:size(p,1) dist=((patchpos(j,1)-p(i,1))^2+(patchpos(j,2)... -p(i,2))^2+(patchpos(j,3)-p(i,3))^2); if dist<=r^2 del(end+1)=i; patchpos(end+1,:)=p(i,:); end end p(del,:)=[]; j=j+1; del=[]; end elseif size(p,1)<=1 p=[]; end if (length(patchpos)*ptarea)5 piso(i,4)=5; end end parea=parea*ptarea; pnumpatch=length(ppatches); avgppatch=parea/pnumpatch; pcoord=piso; figure if ~isempty(piso) scatter3(piso(:,1),piso(:,2),piso(:,3),1,piso(:,4)) colorbar hold on end

n=[]; narea=0; for i=1:numpt if q(i)<=ncutoff n(end+1,:)=X(i,:); end end length(n); k=1; del=[]; patchpos=[]; npatches={}; niso=[]; while ~isempty(n) j=1; patchpos(1,:)=n(1,:); n(1,:)=[]; if size(n,1)>1 while j<=size(patchpos,1) for i=1:size(n,1) dist=((patchpos(j,1)-n(i,1))^2+(patchpos(j,2)... -n(i,2))^2+(patchpos(j,3)-n(i,3))^2); if dist<=r^2 del(end+1)=i; patchpos(end+1,:)=n(i,:); end end n(del,:)=[]; j=j+1; del=[]; end 289

elseif size(n,1)<=1 n=[]; end if (length(patchpos)*ptarea)

Function to evaluate Protein Patchiness. function [patchiness]=patchcorr(q,X,pcutoff,ncutoff,correl_l) %numpatch is the integer number of patches %area is a vector of the areas of each of the patches %q is the vector of potentials %X is the matrix of coordinates on the surface to which q is mapped %cutoff is the cutoff potential below which points are considered neutral %correl_l is the search radius X(:,4)=q;%joins coord and q into one matrix

%code for patchiness parameter. count=0; %counts the number of interactions analyzed sum=0; %sums the correlation param. %Loop evaluates pairwise correlation function using neighbors within

290

%correl_L for i=1:size(X,1) for j=1:size(X,1) dist=((X(j,1)-X(i,1))^2+(X(j,2)-X(i,2))^2+(X(j,3)-X(i,3))^2); if i==j %skips self-correlation sum=sum+0; elseif dist<=correl_l^2 %evaluates whether point j is within correl_l of i count=count+1; %counts # of interactions with neighbors within correl_l if (X(j,4)>=pcutoff && X(i,4)>=pcutoff)... ||(X(j,4)<=ncutoff && X(i,4)<=ncutoff) sum=sum+1; %adds 1 to the correlation sum if the neighbors are of the same charge elseif (X(j,4)>=pcutoff && X(i,4)<=ncutoff) ... ||(X(j,4)<=ncutoff && X(i,4)>=pcutoff) sum=sum-1; %subtracts 1 from the sum if the neighbors are of the opposite charge else sum=sum+0; %sum does not change if |q_i| or |q_j| < |cutoff| end end end end patchiness=sum/count; %Patchiness parameter is the sum of the interactions divided by total number of interactions

References 1. Thomas, C. S.; Glassman, M. J.; Olsen, B. D., Solid-State Nanostructured Materials from Self- Assembly of a Globular Protein–Polymer Diblock Copolymer. ACS Nano 2011, 5 (7), 5697-5707. DOI: 10.1021/nn2013673

291

292

Appendix C. Supporting Information for Chapter 5

Figure C.1. Photographs of Precipitates from PSS with 20% quaternized P4VP. Positive charge fractions from left to right range from 0.1 to 0.9 with macroscopic precipitates forming at f+ of 0.3 to 0.7.

Figure C.2. Schematic of P4VP–PSS interaction showing the presence of induced charges on

P4VP resulting from interaction with PSS.

293

Figure C.3. DLS data for PAA complexing with POEGMA-b-pqP4VP at f+ of 0.5. Data were taken in water at 1 mg mL-1.

294

Figure C.4. 1-D analysis of GISAXS data for PAA. Linecuts were averaged over qz of 0.25 to

0.3 nm-1. Peak micellization indicated by sharpest peaks occurs at the point of charge balance for both encapsulants.

295

Figure C.5. DLS of Amylase variants blended with POEGMA-b-pqP4VP20. The positive

푀+ charge fraction, 푓+, is defined as 푓+ = and M+ and M− are the total positive and negative 푀++푀− charge of the macromolecules in the system. Maximum radius of complexes occurred at f+ of 0.5 for all mutants.

Figure C.6. DLS of Amylase variants blended with POEGMA-b-pqP4VP40. The positive

푀+ charge fraction, 푓+, is defined as 푓+ = and M+ and M− are the total positive and negative 푀++푀− charge of the macromolecules in the system.

296

Figure C.5. AFM height and phase scans and GISAXS patterns for films of supercharged amylase

(A30) and POEGMA-b-pqP4VP40 at various mixing ratios. The positive charge fraction, 푓+, is

푀+ defined as 푓+ = and M+ and M− are the total positive and negative charge of the 푀++푀− macromolecules in the system. Scale bars are 200 nm. GISAXS patterns’ x-axes range from qy of

-1 -1 -0.5 to 0.5 nm and y-axes range from qz of 0 to 1 nm . Height z-scales are 5, 60, 45, and 20 nm.

Phase z-scales are 30°, 30°, 45°, and 50°.

297

Figure C.6. AFM height and phase scans and GISAXS patterns for films of supercharged amylase

(A32) and POEGMA-b-pqP4VP40 at various mixing ratios. The positive charge fraction, 푓+, is

푀+ defined as 푓+ = and M+ and M− are the total positive and negative charge of the 푀++푀− macromolecules in the system. Scale bars are 200 nm. GISAXS patterns’ x-axes range from qy of

-1 -1 -0.5 to 0.5 nm and y-axes range from qz of 0 to 1 nm . Height z-scales are 5, 35, 13, and 16 nm.

Phase z-scales are 30°, 35°, 25°, and 40°.

298

Figure C.7. AFM height and phase scans and GISAXS patterns for films of supercharged amylase

(A37) and POEGMA-b-pqP4VP40 at various mixing ratios. The positive charge fraction, 푓+, is

푀+ defined as 푓+ = and M+ and M− are the total positive and negative charge of the 푀++푀− macromolecules in the system. Scale bars are 200 nm. GISAXS patterns’ x-axes range from qy of

-1 -1 -0.5 to 0.5 nm and y-axes range from qz of 0 to 1 nm . Height z-scales are 5, 20, 30, and 15 nm.

Phase z-scales are 30°, 12°, 30°, and 40°.

299

Table C.1. AFM image z-scales for Figure 5.7.

Protein Loading z-scale unit Control A22 A30 A32 A37

20 wt.% Height nm 6 50 4 45 35

20 wt.% Phase ° 35 25 7 50 35

f+ = 0.7 Height nm 6 35 35 20 20

f+ = 0.7 Phase ° 35 50 35 35 75

f+ = 0.5 Height nm 6 30 28 13 25

f+ = 0.5 Phase ° 35 60 25 30 27

300

Figure C.8. SEC trace of POEGMA.

301

Figure C.9. 1H NMR of POEGMA-b-P4VP. Comparison of OEGMA peak at 4 (2H) to 4VP aromatic peaks (2H) enables determination of final Mn and block ratio.

302

Figure C.10. SEC trace of POEGMA-b-P4VP.

303

Figure C.11. 1H NMR of POEGMA-b-20qP4VP. Comparison of initial P4VP aromatic peaks at

6.6 (2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) after quaternization confirm

>95% conversion.

304

Figure C.12. 1H NMR POEGMA-b-40qP4VP. Comparison of initial P4VP aromatic peaks at 6.6

(2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) after quaternization confirm >95% conversion.

305

Figure C.13. SEC trace of P4VP homopolymer

306

Figure C.14. 1H NMR of 20qP4VP. Comparison of sum of peaks from 6.5 to 9.5 (aromatic

P4VP peaks, 4H) to methyl peak at 4.4 (3H) confirms >95% conversion. Comparison of initial

P4VP aromatic peaks at 6.6 (2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) agree.

307

Figure C.15. 1H NMR of 40qP4VP. Comparison of sum of peaks from 6.5 to 9.5 (aromatic

P4VP peaks, 4H) to methyl peak at 4.4 (3H) confirms >95% conversion. Comparison of initial

P4VP aromatic peaks at 6.6 (2H) and 8.4 (2H) to shifted peaks at 7.8 (2H) and 8.8 (2H) agree.

308

Figure C.16. SDS-PAGE analysis of purified native amylase (A22).

Figure C.17. MALDI-TOF of native and supercharged amylase.

309

Table C.2. Summary and analysis of MALDI-TOF data.

Mean Modification Variant MW # Charge Alpha m/z- A22 54521 0 -22 1.77 2478 A30 54936 4.15 -30.3 2.20 1814 A32 55036 5.15 -32.3 2.33 1704 A37 55261 7.40 -36.8 2.64 1502

310

Appendix D. Supporting Information for Chapter 6

Table D.1. Polymerization conditions for POEGMA-r-BP

mL mL g BP mg CTA mg Temperature Time (h) Dioxane OEGMA AIBN (°C) POEGMA15 180 60 3.20 609.6 71.7 65 6 POEGMA36 180 60 3.20 275.0 32.3 65 5.5 POEGMA55 180 60 3.20 195.6 23.0 65 5

Figure D.1. Differential refractive index chromatographs from SEC of homopolymer and block copolymers of a) POEGMA-r-BP 15, b) POEGMA-r-BP 35, and c) POEGMA-r-BP 55.

311

Table D.2. Polymerization conditions for POEGMA-P4VP and POEGMA-PDMAEMA

g mg µL 1% AIBN mL mL µL Time T (°C) POEGMA AIBN in Dioxane Monomer Dioxane DMF (h)

POEGMA15- 6 3.184 318.4 5.64 10.97 0 4.5 75 PDMAEMA5 POEGMA36- 6 1.371 137.1 7.53 14.92 0 5.5 75 PDMAEMA1 3 POEGMA55- 6 0.896 89.6 7.53 14.96 0 7 75 PDMAEMA1 9 POEGMA15- 3 6.369 636.9 2.10 2.79 786.3 6 75 P4VP5 POEGMA36- 3 2.743 274.3 3.77 3.50 943.6 6 80 P4VP11 POEGMA55- 3 1.793 179.3 3.77 3.59 943.6 6 80 P4VP17

312

Figure D.2. 1H NMR spectra of block copolymers prior to quaternization.

313

Figure D.3. 1H NMR spectra of quaternized block copolymers.

314

Figure D.4. SDS-PAGE analysis of Native NfsB a) fractions from Ni-NTA chromatography and b) final dialyzed protein.

315

Figure D.5. MALDI-TOF analysis of native and supercharged NfsB.

316

Figure D.6. DLS of Native and supercharged NfsB in Water.

Figure D.7. Activity assay of Native and Supercharged NfsB plotted together. SC NfsB is

3 approximately 10 × less active than the native based on kcat/KM.

317

Figure D.8. AFM height and phase scans of 0qPOEGMA15-PDMAEMA5 with Native NfsB.

Figure D.9. AFM height and phase scans of 0qPOEGMA55-PDMAEMA19 with Native NfsB.

318

Figure D.10. GISAXS scattering patterns and qy linecuts for films of 0qPOEGMA-PDMAEMA.

319

Figure D.11. GISAXS scattering patterns and qy linecuts for films of 100qPOEGMA-

PDMAEMA.

320

Figure D.12. Photograph of poor quality films (100qPOEGMA15-PDMAEMA5+50%N and

20qPOEGMA15-P4VP5+50%N) vs. regular (100qPOEGMA36-P4VP11+10%N). Poor quality films are cloudy and lack the color associated with the light refraction through the film because light is scattered instead, while the higher quality films have a mirror-like finish.

321

Figure D.13. GISAXS scattering patterns and qy linecuts for films of 20qPOEGMA-P4VP.

322

Figure D.14. 5 µm x 5 µm AFM images of 20qPOEGMA55-P4VP17 with 10% and 25%

(wt/wt) protein loading of native NfsB. Large complexes increase in prevalence with the addition of protein over this range of loading.

Figure D.15. Enlarged AFM image of 100qPOEGMA15-P4VP5 Control film. a) Height and b) phase. 323

Figure D.16. GISAXS scattering patterns and qy linecuts for films of 100qPOEGMA-P4VP

324

Figure D.17. 5 µm x 5 µm AFM images of 20qPOEGMA55-P4VP17 with 0-50% (wt/wt) protein loading of SC NfsB.

325