Investigating the Impact of Polymer Functional Groups on the

Subscriber access provided by La Trobe University Library

Article

Investigating the Impact of Polymer Functional Groups on the Stability and Activity of Lysozyme-Polymer Conjugates Melissa Lucius, Rebecca Falatach, Cameron McGlone, Katherine Makaroff, Alex P. Danielson, Cameron Williams, Jay C. Nix, Dominik Konkolewicz, Richard Christopher Page, and Jason A. Berberich Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.5b01743 • Publication Date (Web): 11 Feb 2016 Downloaded from http://pubs.acs.org on February 24, 2016

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Biomacromolecules is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Investigating the Impact of Polymer Functional Groups on the Stability and Activity of Lysozyme-Polymer Conjugates Melissa Lucius1, Rebecca Falatach2, Cameron McGlone1, Katherine Makaroff1, Alex Danielson1, Cameron Williams1, Jay C. Nix,3 Dominik Konkolewicz1*, Richard C. Page1*, Jason A. Berberich2* 1

Miami University, Oxford, OH, Department of Chemistry and Biochemistry

2

Miami University, Oxford, OH, Department of Chemical, Paper and Biomedical

Engineering 3

Molecular Biology Consortium, Beamline 4.2.2, Advanced Light Source, Lawrence

Berkeley National Laboratory, Berkeley, California 94720, USA

*

Corresponding Author

[email protected], [email protected], [email protected]

Page 1

ACS Paragon Plus Environment

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 42

Abstract Polymers are often conjugated to proteins to improve stability; however, the impact of polymer chain length and functional groups on protein structure and function is not well understood. Here, we use RAFT polymerization to grow polymers of different lengths and functionality from a short acrylamide oligomer with a RAFT end group conjugated to lysozyme. We show by Xray crystallography that enzyme structure is minimally impacted by modification with the RAFT end group. Significant activity toward the negatively charged Micrococcus lysodeicticus cell wall was maintained when lysozyme was modified with cationic polymers. Thermal and chemical stability of the conjugates were characterized using differential scanning fluorimetry and tryptophan fluorescence. All conjugates had a lower melting temperature; however, conjugates containing ionic or substrate mimicking polymers were more resistant to denaturation by guanidine hydrochloride.

Our results demonstrate that tailoring polymer functionality can

improve conjugate activity and minimize enzymatic inactivation by denaturants.

Keywords: Protein-polymer conjugates, RAFT polymerization, Lysozyme, Enzymatic activity,

Page 2

Protein


stabilization

Page 3 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Introduction The conjugation of soluble polymers to proteins has been used impart new functions to proteins and to help stabilize them in non-native environments. This has led to the development of new protein-based technologies in areas such as drug delivery, bioremediation, and diagnostics.1,2

Perhaps one of the most successful applications of soluble protein-polymer

conjugates is for protein therapeutics. Attaching polymers to proteins for use in vivo stabilizes proteins by increasing the effective molecular size and creating steric hindrance which reduces access of proteolytic enzymes to the surface of the protein,3 limits glomerular filtration, and reduces antibody recognition by concealing antigenic epitopes on the modified protein.4,5 In addition, the attachment of polymers can also be used to add new functionality to the protein. For example, N-isopropylacrylamide (NiPAm) polymers attached to proteins lead to the creation of conjugates which change solubility with temperature providing new ways to separate and recycle biocatalysts.6–8 Polymer attachment can also be used to create protein conjugates that can be dissolved in organic solvents,9 processed in polymer melts,10 and incorporated into coatings.11–13 Perhaps one of the most promising benefits of polymer conjugation is the opportunity to improve the operational and storage stability of proteins.

A major limitation of proteins

compared to chemical catalysts is the difficulty in maintaining stability under the many diverse or extreme temperatures, pH ranges, solvent types, and interfaces that may be found in commercial enzyme formulations or industrial processing environments. There have been a number of reports demonstrating the protection of proteins against thermal denaturation.14 Maynard’s group reported stabilization of hen egg white lysozyme (HEWL) after conjugation with trehalose polymers synthesized using reversible addition-fragmentation chain transfer (RAFT) polymerization.15 In this case, the enzyme was reported to lose only 20% of its original

Page 3


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 42

activity after incubation at 90 °C for 1 hour compared to 80% of activity loss for the native enzyme.

Price and coworkers

demonstrated the possibility to select PEGylation sites on

proteins to provide optimal increases in conformational and proteolytic stability.14,16 Similarly, protein-polymer conjugates have been demonstrated to have improved activity and stability under extremes of pH,17–20 in mixtures of solvents,21 and at interfaces.22 Conjugation of novel polymers has even been shown to protect enzymes from inactivation due to exposure to UV and free radicals.23,24 There are many ways to synthesize protein-polymer conjugates. The traditional approach involves synthesizing the desired polymer in solvent and then creating an activated end group for attachment to the protein. This “grafting-to” approach has been the standard for almost 30 years and is the method by which most PEGylated protein therapeutics are produced.4,5,25

An

alternative approach, grafting-from, involves the covalent attachment of a small polymer initiator or chain transfer agent to the protein surface and the polymer chain is extended directly from this site.26 There are many advantages to this approach for synthesis of protein-polymer conjugates. First, conjugate clean-up is simplified significantly since any unreacted monomer can be removed from the final product by simple dialysis or ultrafiltration. With grafting-from it is much easier to synthesize protein-conjugates with a high density of polymers chains since the steric hindrance for attachment of the small molecule initiator or monomer is significantly smaller than for a full polymer. For synthesis of protein-polymer conjugates, biocompatible reversible deactivation radical polymerization methods such as atom transfer radical polymerization (ATRP) and RAFT are the most popular methods.8,27–33 These techniques offer good control over polymer chain length and polydispersity while being compatible with a wide range of monomers.

Page 4


Page 5 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

To date, there has been no comprehensive study to determine the impact of polymer chain lengths and functional groups on enzyme activity and stability. In this study, we use RAFT polymerization to synthesize a library of protein-polymer bioconjugates by using a grafting-to / grafting-from technique8 to produce active HEWL-polymer conjugates with polymers of varying length and functionality. We measure the activity of HEWL conjugates using a small fluorogenic substrate and the lyophylized cells of Micrococcus lysodeikticus. In addition, the thermal stability of the conjugates were characterized using a differential scanning fluorimetry assay and chemical stability was determined by measuring tryptophan fluorescence in increasing concentrations of guanadinium hydrochloride.

Materials and Methods Materials. Hen Egg White Lysozyme (HEWL) was purchased from MP Biomedicals. Acrylic acid (AA), dimethyl acrylamide (DMAm), and deuterated DMSO were obtained from Acros Organics. Acrylamide, N-hydroxysuccinimide (NHS) and 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) were purchased from Thermo-Fisher. methylumbelliferyl

β-D-N,N’,N”-triacetylchitotrioside,

Micrococcus lysodeikticus, 4azobisisobutyronitrile

(AIBN),

ethanethiol, carbon disulfide, 2-bromopropionic acid, phosphoroylcholine methacrylate (PCMA), acryloyl chloride, and oligo(ethylene oxide) methyl ether acrylate (OEOA) were purchased from Sigma Aldrich. Sodium nitrite and D-glucosamine hydrochloride were from Alfa Aesar. Dimethylaminoethoxy Methacrylate (DMAEMA) was obtained from TCI and potassium carbonate was purchased from MCB. VA-044 was obtained from Wako chemicals. All materials were used without further purification. SDS-poly(acrylamide) gel electrophoresis was performed using Bio-Rad Mini-PROTEAN TGX 4-20% gradient gels. Gels were stained with GelCode

Page 5


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Blue protein stain obtained from Thermo Scientific. The chain transfer agent 2-(((ethylthio)carbonothioyl)thio)propanoic acid (PAETC) was synthesized as outlined in our previous work.34 Synthesis of N-acryloyl-D-glucosamine (AGA). Synthesis of the AGA monomer was adapted from the procedure of Matsuda et al.35 D-glucosamine hydrochloride (8.6007 g, 39.88 mmol) and NaNO2 (0.1468 g, 2.13 mmol) were mixed in 20 mL of 2M aqueous K2CO3 before cooling to 0°C in a salt/ice bath. Acryloyl chloride (4.0208 g, 44.42 mmol) was added dropwise to the stirring reaction mixture. The temperature was maintained below 5°C for three hours and then warmed to room temperature and allowed to react for 21 hours. The reaction mixture was added to 200 mL of absolute ethanol and refrigerated overnight. The mixture was filtered to remove the precipitated salts and then concentrated under vacuum. The resulting product was recrystallized twice in methanol, followed by methanol and ethyl acetate at a ratio of 1:2 respectively. 1H NMR spectra showed peaks as described in literature, and ESI-MS confirmed the product.35,36 RAFT Polymerization of oligo-Acrylamide-chain transfer agent (O-Am-CTA) with PAETC. To synthesize oligo-acrylamide-CTA (O-Am-CTA), RAFT polymerization was used to grow a short oligomer of acrylamide from PAETC in methanol. AIBN (0.2340g, 1.425mmol), PAETC (2.9998 g, 14.27 mmol), and acrylamide (5.0864 g, 71.56 mmol) were added to a glass vial with 10 mL of methanol and mixed to dissolve. The reaction mixture was quantitatively transferred to a round bottom flask with 10 mL of water. The flask was purged with nitrogen gas for 10 minutes to remove oxygen from the system. The reaction was heated to 65 °C in an oil bath to initiate polymerization and the temperature was maintained for 15 hours. Samples taken initially and upon completion of the reaction were analyzed via 1H NMR to confirm at least 95% conversion of the monomer. The final reaction mixture was precipitated by dropwise addition to

Page 6


Page 6 of 42

Page 7 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

an ice cold solution of 200 mL THF and 50 mL ether. The resulting mixture was centrifuged at 6000 rpm for two minutes until all precipitate had been isolated. The collected product was left to dry in a fume hood. RAFT Polymerization of L-Am polymer (L-Am Polymer Only) . VA-044 (0.0065 g, 0.0201 mmol), O-Am-CTA (0.0578 g, 0.1022 mmol), and acrylamide (0.3597 g, 5.0605 mmol) were added to a 5 mL round bottom flask with 0.65 mL of water and mixed to dissolve. The flask was purged with nitrogen gas for 10 minutes to remove oxygen from the system. The reaction was heated to 45 °C in an oil bath to initiate polymerization and the temperature was maintained for 19 hours. Samples taken initially and upon completion of the reaction were analyzed via 1H NMR to confirm at least 95% conversion of the monomer. The final reaction mixture was precipitated by dropwise addition to an ice cold solution of 25 mL THF. The collected product was left to dry in a fume hood. A similar procedure was used for the synthesis of H-Am Polymer Only. RAFT Polymerization of L-DMAm polymer (L-Am Polymer Only) . AIBN (0.0037 g, 0.0226 mmol), O-Am-CTA (0.0574 g, 0.1015 mmol), and N,N-dimethylacrylamide (0.5195 g, 5.2406 mmol) were added to a 5 mL round bottom flask with 0.71 mL of ethanol and mixed; a few drops of water were added to the mixture to help dissolve. The flask was purged with nitrogen gas for 10 minutes to remove oxygen from the system. The reaction was heated to 65 °C in an oil bath to initiate polymerization and the temperature was maintained for 19 hours. Samples taken initially and upon completion of the reaction were analyzed via 1H NMR to confirm at least 95% conversion of the monomer. The final reaction mixture was precipitated by dropwise addition to an ice cold solution of 25 mL hexane. The collected product was left to dry in a fume hood. A similar procedure was used for the synthesis of H-DMAm Polymer Only.

Page 7


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 42

NHS/EDC Coupling of O-Am-CTA to HEWL. The O-Am-CTA was coupled to the reactive amines

of lysozyme

by NHS/EDC

coupling.

HEWL,

O-Am-CTA,

1-ethyl-3-(3-

dimethylaminopropyl)carbodiimide (EDC), and N-hydroxysuccinimide (NHS) were combined in 0.1 M sodium phosphate, pH 7.4 and allowed to react at room temperature for 2 hrs before quenching the reaction with glycine. HEWL was added at a final concentration of 1 mg/ml, NHS was kept at a final concentration 5 mM, and EDC was added at a molar ratio of 1 EDC to 1 CTA. A representative reaction is as follows: HEWL (5 mg, 0.35 µmol), O-Am-CTA (25.7 mg, 0.122 mmol), EDC (23.5 mg, 0.122 mmol), and NHS (5.4 mg, 25 µmol) were added to the reaction vial using stock solutions all prepared in the reaction buffer. Purification of the O-Am-CTA HEWL Conjugate. Ammonium sulfate precipitation was used to separate the O-Am-CTA HEWL conjugate from unreacted O-Am-CTA, NHS, and EDC. This was done by cooling the reaction mixture on ice while stirring. Ammonium sulfate was slowly added to the mixture until the final concentration was 63 % (w/v). The resulting mixture was allowed to stir on ice for 10 minutes and was centrifuged at 4 °C to collect the HEWL conjugate. The resulting pellet was resuspended in 25 mM sodium phosphate, pH 6.2. This process removed the majority of the unreacted O-Am-CTA but required purification by dialysis. Dialysis was performed using phosphate buffer in an Amicon stirred ultrafiltration cell with Ultracel 10 kDa ultrafiltration discs. The purified conjugate was concentrated to 10 mg/ml for RAFT polymerization. HEWL Activity Assays. Two different sized substrates were used to determine the activity of the HEWL at each stage of modification. The large substrate was lyophilized Micrococcus lysodeikticus. Enzyme catalyzed lysis of M. lysodeikticus cell wall was monitored by the change in optical density at 450 nm over time. Reactions were performed in 66 mM KH2PO4, pH 6.2

Page 8


Page 9 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

using and optical density values measured with a BioTek Synergy H1 microplate reader. The optical density was recorded every 15 seconds over a 4 minute time period and the slope of the optical density vs time represented the rate of reaction. The reaction rates for HEWL-polymer conjugates were compared to the rate of reaction for native HEWL to determine percent activity retention. The small HEWL substrate used in this study was 4-methylumbelliferyl β-D-N,N’,N”triacetylchitotrioside ((NAG)3-MUF), which is enzymatically hydrolyzed to produce a fluorescent product. This reaction was monitored using an excitation/emission setting of 360/460 using monochromators on a BioTek Synergy H1 microplate reader. The enzymatic hydrolysis reaction was performed in 50mM acetate, pH of 5.5 at 37 °C. Since the MUF product is only fluorescent at high pH the reaction was stopped using a glycine buffer at pH 10.8 prior to measuring the fluorescence at each time point. Typical Chain Extension from O-Am-CTA Modified HEWL by RAFT Polymerization. Acrylamide, AGA, DMAEMA, PCMA, and AA were each chain extended from the O-Am-CTA modified lysozyme. AGA (16 mg, 68.7 µmol), VA-044 (2.2 mg, 6.8 µmol) and O-Am-CTA lysozyme (0.95 mL, 0.62 µmol) were combined in a glass vial, mixed to dissolve, and transferred to a Schlenk flask. The system was deoxygenated for 10 minutes under a gentle flow of nitrogen gas. The polymerization was initiated by heating at 30 °C in an oil bath. The reaction was allowed to proceed for 14 hours and initial and final samples were analyzed via 1H NMR to determine percent conversion. Thermal Stability using Differential Scanning Fluorimetry. Native HEWL, polymer modified HEWL derivatives, and representative polymers only were analyzed in triplicate via differential scanning fluorimetry. Protein samples were diluted to a final concentrations of 5 µM with 20 mM HEPES, pH 7.5, 150 mM NaCl. Controls containing polymers only were diluted to

Page 9


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 42

a final concentration of 12.5 µM with 20 mM HEPES, pH 7.5, 150 mM NaCl, to match the polymer concentrations on the conjugates, due to the fact that typically 2.5 polymers attach to a single HEWL. Differential scanning fluorimetry was conducted using a CFX96 RT-PCR (BioRad). Fluorescence was monitored at 570 nm while temperature was ramped in 0.5 °C increments from 25 °C to 95 °C with a 5 second hold for equilibration at each temperature step. Melting temperatures of native HEWL and its conjugates (Tm) were determined by fitting data to a Boltzmann equation37 using Prism (GraphPad). Chemical Stability using Guanidine Hydrochloride. The chemical stability of HEWL conjugates was explored by using guanidine hydrochloride to chemically denature the protein and measuring the intrinsic fluorescence of tryptophan residues. Twelve guanidine hydrochloride solutions with concentrations ranging from 0 M to 6 M were prepared in 20 mM HEPES, 150 mM NaCl, pH 7.5. Native- and conjugated-HEWL samples were measured in triplicate at 27 µM final protein concentrations using a BioTek Synergy H1 microplate reader equipped with monochromators set to 280 nm excitation and an emission spectral window of 310 nm to 400 nm. Non-linear sigmoidal curves representing a two-state folding model were fit using Prism (GraphPad). Matrix Assisted Laser Desorption/Ionization-Mass Spectrometry (MALDI-MS). MALDITOF spectra were collected on a Bruker Autoflex III MALDI-TOF mass spectrometer (Billerica, MA). Mass spectra were calibrated using Bruker Protein Calibration Standard I (spanning 5.7 kDa to 16.9 kDa) as an external standard. Generally, 0.5 µL of sample (0.5 mg/ml) was mixed with 0.5 µL of saturated sinapinic acid (Fluka, WI) solution (0.1 % TFA, 40 % acetonitrile) directly on the target plate and allowed to dry at room temperature. Samples were analyzed in the positive ion linear mode to detect [M + H]+ ions.

Page 10


Page 11 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Determination of Modification Site of O-Am-CTA modified HEWL. Trypsin digests to determine O-Am-CTA attachment sites were carried out using 20 µg native HEWL or acrylamide O-Am-CTA conjugated HEWL. Samples were digested using the In-Solution Tryptic Digestion and Guanidination Kit (Thermo Scientific # 89895). Digested samples were purified using Pierce C18 Spin Columns (Thermo Scientific # 89870). Native- and polymer-conjugated HEWL samples were used at 1.0 mg/mL concentrations. Samples were spotted on a Bruker MTP 384 ground steel target plate (Bruker Deltonik, Bremen, Germany). Saturated α-Cyano-4hydroxycinnamic acid in TF40 (0.1 % TFA, 40 % acetonitrile) was used as the matrix for trypsin digested samples. All samples were mixed 1:1 with the appropriate matrix on the MALDI plate. MALDI-TOF data acquisition was performed on a Bruker AutoFlexIII MALDI-TOF mass spectrometer at the Mass Spectrometry Facility, Miami University. Identification of carbamidomethyl- and guanidinyl-modified peptides from MALDI data was carried out using MASCOT (Matrix Science). Identified tryptic peptides are listed in Table S1. Crystallization, X-Ray Data Collection and Processing. A solution of 10 mg/ml HEWL was mixed 1:1 with crystallization condition consisting of 10 % (w/v) sodium chloride and 0.1 M sodium acetate, pH 4.6 to produce a final drop of 0.8 µl. Crystals were grown by vapor diffusion against a bath of 70 µl of crystallization condition. A solution of 10 mg/ml CTA-HEWL was mixed with a crystallization condition consisting of 2 M sodium chloride and 12 % (w/v) PEG 6000. CTA-HEWL crystals were grown by vapor diffusion against a bath of 70 µl of crystallization condition. Both CTA-modified and native HEWL crystals were harvested through LV cryo-oil (MiTeGen) for cryoprotection and immediately flash frozen in liquid nitrogen. Xray diffraction data were integrated and scaled using XDS38 and SCALA.39 The high resolution cutoff for each data set was chosen based on the performance of crystallographic statistics

Page 11


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 42

including CC1/2,40 I/σI, and completeness. Five percent of reflections were set aside for the test set for calculating Rfree. Structure Determination and Refinement. A structure of HEWL (PDB code: 1IEE)41 was used as molecular replacement search models for the native HEWL data set using PHASER.42 Iterative rounds of refinement in PHENIX43 and model building in Coot44 were used to build the native HEWL structure. The native HEWL structure was used as the molecular replacement model in PHASER42 for the CTA-HEWL data set. The final CTA-HEWL structure was built from iterative rounds of refinement in PHENIX43 and model building in Coot. 44 Geometric and sterochemical validation of both structures were performed using MolProbity v4.1.45 Crystallographic and refinement statistics for each structure and dataset are shown in Table S2. The native HEWL structure was deposited in the Protein Data Bank (PDB) under accession ID 5F14. The CTA-HEWL structure was deposited in the PDB under accession ID 5F16. Molecular Dynamics Simulations. All-atom simulations were performed using NAMD 2.1039 in an NPT ensemble at a constant pressure of 1 atm and temperature of 298 K utilizing the CHARMM22 force field,46 particle mesh Ewald technique for electrostatic interactions, and explicit TIP3P water. HEWL-polymer conjugate starting models were built using PDB ID 1IEE41 as the starting coordinates for the protein backbone and side chains. HEWL-DP5polyethylene glycol (O-PEG~HEWL) and HEWL-DP5-acrylamide (O-Am~HEWL) conjugates were each solvated in a water box with 15 Å padding and periodic boundary conditions in all three dimensions. A single simulation was run for each HEWL-polymer conjugate and each simulation was minimized for 30 ps with 1 fs time steps followed by a 10 ns simulation with 1 fs time steps.

Page 12


Page 13 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Results and Discussion Synthesis and Characterization of O-Am-CTA modified HEWL. Since direct modification of HEWL with the chain transfer agent PAETC reduces the water solubility of HEWL,34 a water soluble short chain of approximately five acrylamide (Am) units, termed an acrylamide oligomer (oligo-CTA), with an attached chain transfer agent was synthesized according to Scheme 1A. The oligomer of Am was synthesized by RAFT polymerization using 2-(propionic acid) ethyl trithiocarbonate (PAETC) as the chain transfer agent (CTA) to produce the oligo-CTA. These oligomers were coupled using EDC/NHS to the surface amines of HEWL using the grafting-to approach (Scheme 1B). The resulting acrylamide oligomer HEWL conjugates (O-Am) were soluble to at least 10 mg/mL at room temperature. Due to the overlap between the typical protein band at 280 nm and the trithiocarbonate peak at 310 nm,47 a protein concentration calibration curve at 270 nm was constructed. The protein concentration was determined by measuring the absorbance at 270 nm. Matrix assisted laserdesorption ionization mass spectrometry (MALDI-MS) traces for native HEWL and the HEWL conjugate modified with Am oligomers (O-Am) indicate that efficient modification of the protein was achieved as evidenced by the positive shift of the conjugate’s molecular weight after coupling, compared to the native protein (Figure S1). These results are comparable to those published earlier.34 A variety of monomers with different functionalities, including anionic, cationic and zwitterionic groups, were selected (Scheme 1C) for chain extension from the O-Am by the grafting from approach (Scheme 1B). The monomer N-acryloyl-D-glucosamine (AGA) was synthesized since it was a mimic of the native HEWL substrates and inhibitors composed of oligosaccharides of N-acetyl-D-glucosamine.48

Page 13


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 42

A)

B)

C

O Am

O NH2

O

OEOA

O 9

O HO O

N

DMA

AGA

N H

O O

N

DMAEMA

O

OH OH

OH

O O

O PCMA

AA OH

O P O O

O

N

Scheme 1. A) Synthesis of O-Am-CTA. B) The hybrid grafting-to and grafting-from method for protein-polymer conjugate synthesis. A preformed O-Am-CTA is attached to HEWL, followed by chain extension by RAFT. C) The various functional monomers used in this study for chain extension. The native HEWL and O-Am conjugate were analyzed using MALDI mass spectrometry of trypsin digests. Proteolytic cleavage by trypsin hydrolyzes the peptide backbone to the carboxyterminal side of lysine or arginine residues that have not been modified. Modification of lysine with the acrylamide oligomer precludes recognition by trypsin. By comparing peptides identified after trypsin digestion (Figure 1) we were able to identify the exact modification sites. Tryptic peptides with a characteristic “saw-tooth” pattern conclusively identify acrylamide oligomer modified peptides corresponding to modification at Lys1, Lys33, and Lys97 (Figure 1 and Table S1). The tryptic peptide data does not distinguish between modification of the amino-terminus or the ε-amino group of Lys1. However, these modification sites are consistent with our previous observation of ~2.5 oligo-CTA attachments per protein molecule34 and previously published

Page 14


Page 15 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

reports identifying Lys33, Lys97, and the amino-terminus as the most readily modifiable residues.49–51

Figure 1. Trypsin digestion of native and O-Am HEWL. (A) MALDI-TOF data for trypsin digested native HEWL identifies expected peptides with 87% sequence coverage. (B) MALDITOF for trypsin digested O-Am HEWL identifies expected peaks with 91% sequence coverage including peptides with O-Am modified lysine residues (K1, K33, and K97).

Page 15


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 42

Accommodation of Surface Amine Modifications. Although circular dichroism is conventionally used to probe structural effects of modification by polymers or chain transfer agents, we chose to use X-ray crystallography to provide highresolution determination of local and global structural changes. While the O-Am HEWL conjugate did not crystallize, we did obtain crystals for CTA-modified HEWL. We solved X-ray crystal structures of native and CTA-modified HEWL at resolutions of 1.148 Å and 1.200 Å, respectively. Globally the two structures are nearly identical with a 0.136 Å rmsd for all heavy atoms (Table S2). Compared to native HEWL, PAETC-modified HEWL exhibits an altered rotamer for Asn103 at the enzyme active site. This small change is likely the result of slight changes in unit cell size and intermolecular packing between the native and PAETC-modified HEWL crystals. Docking of a chitotriose substrate from a previously published HEWL structure41 indicates that this change would not preclude binding of substrates. Although the flexibility of the PAETC groups attached to Lys1, Lys33, and Lys97 resulted in electron densities that were too poor for reliable modeling of the PAETC, the local effects of PAETC modification are apparent when examining the orientation of lysine side chains (Figure 2 A-C and Figure S2). For native HEWL, both Lys97 (Figure 2C) and Lys1 (Figure S2) exhibit two distinct rotamers each; while their side chain positions in PAETC-modified HEWL are restricted to a single rotamer. This may be due to steric restriction added by the PAETC. In contrast, the orientation of Lys33 is identical in both structures (Figure 2B), likely due crystal packing that restrains allowed side chain rotamers. Thus, while minor differences in side chain positions are observed, modification by a PAETC does not result in substantial changes to the overall structure.

Page 16


Page 17 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Figure 2. Overlay of HEWL structures. (A) The structure of native HEWL (PDB ID 5F14, blue) is overlaid onto the structure of CTA-modified HEWL (PDB ID 5F16, green). For reference, a chitotriose substrate (grey sticks) from PDB ID 1HEW,41 is shown in the active site and active

Page 17


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 42

site residues within 3.5 Å of chitotriose are shown as sticks. Zoomed views of lysine 33 (B) and lysine 97 (C) show similarity across the native and PAETC-modified HEWL structures. Dynamic Structures of Polymer Conjugates. While limited changes in protein structure result from introduction of the small molecule PAETC, conjugation could have a profound influence on protein or polymer structure when a polymer is attached. We performed molecular dynamics (MD) simulations using all atom models for HEWL~polymer conjugates in explicit TI3P water for 10 ns each. Conjugates between HEWL and acrylamide oligomers (O-Am) as well as HEWL and ethylene glycol oligomers (OPEG) were studied by molecular dynamics. A family of snapshots at 100 ps intervals are shown in Figure 3 for O-Am and O-PEG conjugates with polymers containing 5 monomers each. Strikingly, the behavior of each conjugate is distinct for each type of monomer and for each conjugation site. At Lys33, both O-Am and O-PEG conjugates display interactions with HEWL surface residues. PEG chains preferentially cluster at a group of surface exposed hydrophobic residues consisting of two phenylalanine residues and one tryptophan. In contrast, the acrylamide chain occupies a separate region that enables interactions with glutamine, asparagine, and lysine residues at the protein surface. Surrounding Lys97, both hydrophobic and hydrophilic residues are more evenly distributed, thus the PEG chains exhibit no clear preference in location of interaction and the family of conformations adopts a mushroom shape observed in previous molecular dynamics simulations of peptide~PEG conjugates.52,53 In comparison, the acrylamide polymer exhibits a narrower “funnel” shape with protein surface interactions that are substantially more limited. In addition to preferential regions of interaction between each polymer at each location, it is immediately obvious that the current locations for polymer attachment, particularly Lys97, would produce substantial steric restrictions at the HEWL active

Page 18


Page 19 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

site. While small molecule substrates are likely to be readily accommodated by short polymer conjugates at Lys97, increased polymer lengths or the use of large substrates, such as M. lysodeikticus, are likely to result in diminished enzyme activity due to steric occlusion. Furthermore, while the short polymers are capable of interacting with the protein surface, these interactions tend to be limited. Near Lys33, each 5-monomer chain appears to be capable of forming approximately 2 hydrogens bonds total between the polymer and HEWL, predominantly via interactions with side chains. Near Lys97, similar degrees of hydrogen bonding are seen for the O-PEG conjugate and no hydrogen bonds are observed between the O-Am chain and HEWL. The MD simulations further suggest that the nature of interactions between the protein and conjugated polymer will depend upon the monomer composition of the polymer and the site of conjugation. Judicious choice of monomer composition may allow for protein-polymer interactions that stabilize the protein against thermal or chemical challenges. The behavior of polymer conjugates near Lys33 indicates that the polymer may offer increased resistance to proteolytic cleavage, stability against denaturants, or possibly thermal stabilization by creating a protein-polymer hydrogen bond network. However, the limited number of hydrogen bonds for the short polymers indicate that substantially longer polymers containing functional groups with considerably enhanced hydrogen bond propensity may be needed to enable stabilization against thermal or chemical denaturation.

Page 19


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 42

Figure 3. Molecular dynamics simulations of HEWL~O-Am and HEWL~O-PEG conjugates with

5

monomers

each.

(A)

Individual

structures represent a snapshot at 100 ps intervals over the course of the 10 ns MD simulations for O-Am (violet) and O-PEG (yellow). The chitotriose substrate from PDB ID 1HEW is shown to illustrate location of the HEWL active site.41 Zoomed views of the OAm and O-PEG conjugates to lysine 33 (B) and lysine 97 (C) show differential behavior of polymer chains at different sites.

Page 20


Page 21 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Chain Extension (Grafting From) O-Am HEWL Conjugate. Since the molecular dynamic simulations suggest that interactions between the acrylamide oligomer and the protein surface are likely to be minimal, a grafting from approach was used to chain extend the O-Am conjugates in order to enhance the interactions between the functional groups of the polymer chains and the protein surface. RAFT polymerization allowed straightforward synthesis of polymers with various functional groups and molecular weights from the well characterized O-Am conjugates. All grafting-from reactions were performed at 30 o

C to limit enzyme denaturation, and monomer conversions were in the range of 60-95%

conversion. The polymerization conditions closely follow those developed by Sumerlin and coworkers who showed excellent control over macromolecular architecture32, therefore the polymers are well controlled by the RAFT mechanism. The conjugates were grouped into two series. Those with relatively low molecular weight of ca. 30-50 repeat units, denoted as the “L” series, and those of much higher molecular weights containing 150-250 units, denoted as the “H” series. Note that the samples L-Am/AA* and L-Am/DMAEMA* have similar molecular weight to the L-Am/AA and L-Am/DMAEMA conjugates, although the L-Am/AA* and LAm/DMAEMA* constructs had approximately twice as many of the charged AA or DMAEMA groups. A summary of the polymerization data is given in Table 1. For conjugates of mean molecular weight 25,000 or lower, MALDI data could be obtained and these are given in the supporting information. For these lower molecular weight conjugates there is acceptable agreement between the theoretical molecular weight and the peak molecular weight of the MALDI, taken as a proxy for the conjugate’s average molecular weight. The MALDI-MS data for conjugates with non-baseline signal is given in the Supporting Information (Figures S3-S8). Differences between the molecular weight estimated by MALDI and the theoretical molecular

Page 21


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 42

weight could be due to differences in ionization efficiency between higher and lower molecular weight conjugates within the population. Additionally, due to the distribution of the number of attachments this can lead to further discrepancies between the theoretical and experimental molecular weights. Figure 4 shows the polyacrylamide gel-electrophoresis (PAGE) data for all conjugates studied. Native HEWL has a molecular weight of approximately 15 kDa, consistent with its reported size (14.3 kDa).54 The O-Am conjugate is at higher molecular weight than the native, and all other conjugates are at higher molecular weights than the native HEWL and the O-Am conjugate. Due to the complex nature of separation in PAGE, the apparent molecular weight is a complex factor involving the molecular charge, shape and mass, making direct evaluation of the true molecular weight almost impossible. However, all apparent molecular weights from PAGE show that the conjugates had higher molecular weight than the starting material, and higher ratios of monomer to enzyme tended to form higher molecular weight conjugates. Comparing the PAGE data in Figure 4, shows that the O-Am, L-Am and H-Am samples are clearly in different regions of the gel. Similarly, L-AGA and H-AGA are in different regions of the gel. This indicates that even within the limitations of PAGE, there is a clear distinction between the properties of the H and L series.

Page 22


Page 23 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Table 1. Conditions and results for RAFT polymerizations. All polymerizations except the synthesis of the short Am oligomer were performed at pH=6 at 30 oC at 10 mg/mL enzyme. All conversions estimated by NMR.

[M1]:[M2]: [CTA]:[VA-044]

Time (h)

Conv. M1

Conv. M2

DP M1

DP M2

0

0

-

5

-

Mn Polymer (Th.)

Mn Conjugate (Th.)

Mp Conjugate (MALDI)

-

565

15678

15000

50

-

4239

22778

21000

34

-

4055

22410

-

50

-

24689

63678

0.6

0.9

24

9

5048

24396

18000

7

0.65

1

26

10

4105

22510

21000

40:20:1:5

13.5

0.5

1

20

20

5249

24798

21000

AA

40:10:1:5

7.5

0.5

0.5

20

5

2469

19238

18000

Am

AA

40:20:1:5

13.5

0.75

0.7

30

14

3827

21954

20000

L-AGA

AGA

-

50:0:1:5

14

0.95

47.5

-

11756.5

37813

H-Am

Am

-

250:0:1:5

14

0.95

-

237.5

-

17551.5

49403

H-Am/DMAEMA

Am

DMAEMA

200:50:1:5

28

0.5

1

100

50

15639

45578

H-Am/AA

Am

AA

200:50:1:5

14

0.8

0.75

160

37.5

14749

43798

H-AGA

AGA

-

250:0:1:5

14

0.9

-

225

-

53114

120528

Sample

M1

M2

HEWL

-

-

O-Am

Am

-

5:0:1:0.2

24

1

L-Am

Am

-

50:0:1:5

13.5

1

L-DMAm

DMAm

-

50:0:1:5

14

0.68

L-OEOA

OEOA

-

50:0:1:5

7.5

1

L-Am/PCMA

Am

PCMA

40:10:1:5

7.5

L-Am/DMAEMA

Am

DMAEMA

40:10:1:5

L-Am/DMAEMA*

Am

DMAEMA

L-Am/AA

Am

L-Am/AA*

Page 23


Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 42

Figure 4. PAGE data for HEWL conjugates with high molecular weight polymers. Panel A shows the PAGE data for the majority of the lower molecular weight (L) conjugates. Panel B shows the PAGE for the higher molecular weight conjugates (H) and the L-AGA sample.


24

Page 25 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

Effect of Polymer Molecular Weight and Functional Groups on Lysozyme Activity. To better understand the impact of polymer modification on enzyme kinetics, activity assays were performed on two different substrates. The first substrate, M. lysodeikticus, is a large (micron sized) lyophilized bacterial cell.

The ß-1,4 glycosidic linkages between N-

acetylmuramic acid and N-acetylglucosamine in the negatively charged cell walls are preferentially hydrolyzed by HEWL.54 The second substrate, 4-methylumbelliferyl β-DN,N’,N”-triacetylchitotrioside ((NAG)3-MUF), is a neutral small molecule containing a trimer of N-acetylglucosamine with a 4-methylumbelliferyl group. Upon hydrolysis of the β-1,4 linkages by HEWL the (NAG)3-MUF releases the 4-methylumbelliferyl group that is fluorescent at high pH. Figure 5 A investigates the influence of polymer molecular weight on the enzymatic activity. Acrylamide polymers were synthesized with degrees of polymerization (DP) in the order of 5, 50 and 250 units. The small molecule (NAG)3-MUF substrate showed relatively small variations in the enzymatic activity with increasing polymer molecular weight. The difference between the lowest (O-Am) and highest (H-Am) molecular weight acrylamide conjugates was only 40% of the native enzyme activity. Higher molecular weight attached polymers led to lower activity, consistent with the idea that the small (NAG)3-MUF substrate can relatively easily enter the enzyme’s active site regardless of the polymer size. In contrast, the M. lysodeikticus activity assay showed significant reduction as the polymer’s molecular weight was increased. The O-Am had approximately 60% of the native enzyme’s activity against M. lysodeikticus, while the enzymatic the activity decreased to approximately 20% of the native for L-Am, and the the HAm conjugate had essentially no enzymatic activity against M. lysodeikticus. Since the M. lysodeikticus substrate is large (1 µm in diameter), steric effects are important. Figure 5A


25

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 42

suggests that higher molecular weight polymers lead to more steric repulsions between the conjugate and the substrate, and lower activity. Figure 5B investigates the effect of functional group on enzymatic activity for relatively low molecular weight (L) conjugates. All conjugates shown in Figure 3B had attached polymers with the degree of polymerization on the order of 35-50 units. The functional groups considered were cationic (DMAEMA), anionic (AA), zwitterionic (PCMA) and neutral (OEOA, AGA, Am). With the exception of AGA, the activities against the (NAG)3-MUF substrate were in the order of 60-80% of the native enzyme’s activity. This indicates that, in general, the functional group on the polymer has minimal influence on the active site and leads to minimal decrease in the enzyme’s intrinsic ability to catalyze the degradation of β-1,4-glycosidic linkages. The exception to this is the L-AGA based conjugate which had a lower activity against the (NAG)3-MUF substrate. It is important to note that N-acetylglucosamine oligomers have been shown to noncovalently bind the active site of lysozyme efficiently,55–57 and the decrease in activity may be attributed to this binding. Figure 5B also investigates the influence of the functional group on HEWL activity against the larger anionic M. lysodeikticus substrate. In contrast, to the (NAG)3-MUF substrate where only minor variations were observed between functional groups, the M. lysodeikticus assay showed significant variation in the enzymatic activity for polymers with varying functional groups. In general the activity of the L- conjugates was typically approximately 20% of the activity of the native enzyme. This is most likely due to the steric repulsions caused by conjugating the polymer to the enzyme. The AGA based polymer showed only 1% activity against the substrate, likely due to a combination of the AGA polymer partially binding the active site and the steric repulsions caused by the polymer. The other non-ionic and zwitterionic


26

Page 27 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

polymers had activities in the order of 10-20% of the native against the M. lysodeikticus substrate. The charge on the polymer had the biggest influence on enzymatic activity against M. lysodeikticus. The data in Figure 5B shows that some of the lowest activities are observed for conjugates containing acrylic acid units. At the pH =6.2 used in the assay these groups will be significantly deprotonated leading to a negative charge on the polymer, potentially causing electrostatic repulsions between conjugate and the anionic M. lysodeikticus surface.58,59 In contrast, the DMAEMA based conjugates had apparent activities in the order of 80% of the native enzyme. At the pH values used for the M. lysodeikticus assay the amine groups on DMAEMA will be protonated,60 possibly leading to an electrostatic attraction between the conjugate and the anionic M. lysodeikticus surface, causing a sharp increase in the apparent rate of catalysis. A similar effect on enzymatic activity was observed by Satishkumar and Vertegel for HEWL covalently attached to polystyrene nanoparticles.61 HEWL coupled to nanoparticles with different surface charges had similar activities against a low molecular weight oligosaccharide substrate; however, electrostatic repulsions by negatively charged nanoparticles led to a reduction in enzymatic activity against M. lysodeikticus, while positively charged surfaces led to improved enzyme activity due to electrostatic attraction to the bacteria. Similar results were observed by Russell’s group and by Thayumanavan’s group for chymotrypsin where the binding of differently charged substrates and inhibitors were modulated by changing the charge of polymer chains attached to the enzyme.62,63 Figure 5C shows the apparent activity data for the high -molecular weight conjugates. For both the (NAG)3-MUF and M. lysodeikticus substrates the enzymatic activity is reduced for the H molecular weight polymers, compared their L counterparts. Against the (NAG)3-MUF


27

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 28 of 42

substrate, the Am, Am/AA and Am/DMAEMA showed activities in the order of 30-40% of the native. This suggests that there is steric crowding due to the high molecular weight polymer, but the small molecule substrate is still able to access the active site. In contrast, the H-AGA system showed no measureable activity against the (NAG)3-MUF substrate, which is consistent with occupancy of the HEWL active site by the high molecular weight polymer harboring Nacetylglucosamine, thereby excluding substrate and preventing catalysis.55–57 Against the larger anionic M. lysodeikticus substrate all conjugates except the H-Am/DMAEMA showed no measurable activity. This is likely due to steric effects. The H-Am/DMAEMA conjugate showed 20% of the native’s activity, which is consistent with an electrostatic attraction between the anionic M. lysodeikticus substrate and the H-Am/DMAEMA conjugate partially offsetting the loss of activity due to the attachment of the large polymer. The activity data are given in Table S3. In summary, the type and size of polymer attached to HEWL has a significant impact on enzymatic activity. For the small uncharged substrate (NAG)3-MUF, activity appears to be nearly independent of polymer type, but is reduced as the molecular weight of the attached polymer is increased. For the negatively charged M. lysodeikticus substrate, positively charged functional groups increase enzymatic activity; however, activity is more sensitive to increasing polymer molecular weight due to steric effects.


28

Page 29 of 42

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 ACS Paragon Plus Environment

29

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 30 of 42

Figure 5. Activity data for different HEWL-polymer conjugates. Panel A shows the activity of HEWL polymer conjugates, panel B shows the activity of low molecular weight (L) conjugates with different functional groups and panel C shows the activity of high molecular weight (H) conjugates. Effect of Polymer Molecular Weight and Functional Groups on Lysozyme Stability. In addition to enzymatic activity, the structural stability of each enzyme-conjugates was determined. There are several measures of protein stability, including ability to withstand thermally or chemically induced denaturation, in vivo half-life in circulation, and the capability to resist proteolytic degradation. Typically, conjugating a polymer to a protein will enhance its ability to withstand proteolytic degradation and improve the conjugate’s in vivo circulation time. However, due to the complex interactions between the solvent, the polymer and the protein, thermal and chemical stability are more difficult to predict. To date the majority of studies have focused on functional thermal stability (retained activity as a function of heating time), proteolytic resistance, and in vivo circulation times. In many cases, polymer modification of


30

Page 31 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

proteins has been shown to improve the functional thermal stability of proteins. For example, Maynard and coworkers showed that functional stability could be improved by conjugating trelahose polymers to HEWL, possibly due to the beneficial interactions between the polysaccharide nature of trelahose and the protein.15 Here we focus on structural stability against thermal and chemical denaturation for HEWL-polymer constructs with different polymer functionalities and molecular weights. Thermal stability was determined using differential scanning fluorimetry (DSF).37,64 This method measures the fluorescence of a solvatofluorochromic dye, SYPRO orange, as a function of temperature. When the protein is folded with hydrophobic residues buried in the core, the fluorescent signal of the SYPRO orange is weak. However, protein unfolding exposes hydrophobic residues that interact with SYPRO orange causing an increase in quantum yield of the fluorophore and a concomitant increase in fluorescence signal. By measuring fluorescence as a function of temperature, the melting temperature (Tm) is approximated by either measuring the inflection point, or fitting a Boltzmann function to the DSF data.37 A series of typical DSF curves and Boltzmann function fits are shown in Figure 6A. In general all DSF curves have similar profiles to those shown in Figure 6A. Below the melting temperature the fluorescence is very low, and the fluorescence signal increases as the protein unfolds. Figure 6A shows the DSF traces for the native HEWL, the O-Am, L-Am and H-Am conjugates. All conjugates are less thermally stable than the unmodified HEWL, as measured by the Tm. Interestingly the higher molecular weight conjugates exhibit lower Tm values. Figure S9A and Figure S9B highlight the effect of functional groups and polymer length on thermal stability, which is summarized in Table 2. Of particular interest is the comparison of the HEWL, L-OEOA, L-Am/AA, L-Am/DMAEMA, H-Am/AA and H-Am/DMAEMA samples. The non-


31

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 32 of 42

ionic L-OEOA samples unfolded at slightly lower temperatures compared to the anionic LAm/AA and L-Am/DMAEMA based polymers, and the unfolding transition was broader for the OEOA based conjugate than the other systems studied. Finally, Table 2 compares the effect of polymer molecular weight, with the lower molecular weight (L) conjugates typically having melting approximately 5 oC above the corresponding higher molecular weight (H) conjugates. Finally, to confirm that the DSF response measured was a direct consequence of the protein unfolding, rather than an interaction between the polymer backbone and the SYPRO orange dye, polymer only controls were performed. Four polymers were tested including poly(Am) with molecular weight similar to that of L-Am and H-Am, poly(DMAm) with molecular weight similar to that of L-DMAm, and a higher molecular weight DMAm polymer targeting 250 repeat units. DSF was performed on these polymer only samples at the same concentration as the polymer is on the lysozyme conjugates, although the data was comparable for samples with higher and lower concentrations of polymer. As indicated in Figure S9C, the signal is essentially baseline over the whole temperature range. The negligible change in the DSF signal for all 4 polymers studied indicates that there is negligible interaction between the dye and the polymer backbone. Figure 6B shows the stability against denaturation by guanidine hydrochloride (Gdn-HCl) as a chemical denaturant. Chemical stability was assessed by measuring the maximal wavelength (λmax) of tryptophan fluorescence. As the protein unfolds the λmax increases from ~330 nm in the fully folded state to ~342 nm in the fully unfolded state. Different concentrations of Gdn-HCl were used and the fluorescence maximum wavelength (λmax) was determined at each concentration. This can be fit to a two-state model for unfolding giving a measure of chemical stability with more stable constructs having inflection points at higher Gdn-HCl values. Figure


32

Page 33 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

6B shows the chemical denaturation data for the native HEWL, O-Am, L-Am and H-Am conjugates. Interestingly the inflection point initially decreases as the polymer molecular weight increases from HEWL to O-Am to L-Am, however the H-Am sample interestingly was more chemically stable than the HEWL to O-Am to L-Am samples. Figure S10 shows the results of chemical denaturation by Gdn-HCl for the remaining conjugates which is summarized in Table 2. Chemical stability is assessed using the concentration of Gdn-HCl denaturant needed to unfold 50% of the protein (D50). Several key conclusions can be drawn from this data. The highest molecular weight conjugates H-Am, HAm/AA and H-Am/DMAEMA and H-AGA had the best chemical stability, which exceeded the native HEWL. Additionally, the AGA monomer showed better chemical stabilization compared to the other non-ionic polymers, which can be explained by the AGA side-chains of the polymer hydrogen bonding with the protein domains near the active site and improving stability.48 In general, groups that can interact with the protein surface such as ionic functional groups or AGA tended to improve the stability compared to a non-ionic polymer of comparable molecular weight. In general, higher molecular weight polymers tended to lower the Tm, with the L series typically having higher Tm values than the H series. The functional group identity had very little influence on the Tm. The lower thermal stability observed with higher molecular weight polymers attached to the protein could be due to the larger polymer causing an entropic penalty for folding, which lowers thermal stability. The results from the chemical stability data suggests that polymer functionality and molecular weight both influence the ability to resist chemical denaturation. Unlike the thermal data, high molecular weight polymers (those from the H series) tended to have improved chemical stability compared to the lower molecular weight (L series)


33

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 34 of 42

polymer conjugates. This suggests that larger polymers are better able to interact with and tie together different segment of the protein and improve stability. Additionally, the ionic functional groups of DMAEMA and AA, and the AGA polymer had improved D50 values compared to their non-ionic counterparts. These data highlight that protein stability is a complex phenomenon, with some conjugates such as H-Am, H-AGA, H-Am/DMAEMA and H-Am/AA all being less stable than native HEWL respect to thermal denaturation, yet more stable than native HEWL with respect to chemical denaturation by Gdn-HCl.

Figure 6. Analyses of thermal and chemical stability. (A) Differential scanning fluorimetry of Native HEWL(black), O-Am (red), L-Am (cyan), and H-Am (blue) modified HEWL was used to


34

Page 35 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

examine thermal stability of each sample. A Boltzmann fit to each sample (dotted lines) enabled determination of Tm values reported in Table 2. (B) Tryptophan fluorescence plotted versus concentration of guanidine for Native HEWL(black), O-Am (red), L-Am (cyan), and H-Am (blue) modified HEWL. Non-linear sigmoidal fits using a two-state model (solid lines) allow for calculation of guanidine concentrations needed to reach 50% denaturation (D50) reported in Table 2.

Table 2. Thermal and Chemical stability data for all lysozyme polymer conjugates. Tm (oC)

∆Tm (oC)

D50 (M)

∆D50 (M)

HEWL

67.5±0.2

0

4.1±0.5

0

O-Am

62±0.2

-5.5

3±0.7

-1.1

L-Am

64.1±0.2

-3.4

2.5±0.9

-1.6

H-Am

57.4±0.2

-10.1

4.6±0.4

0.5

L-OEOA

60.1±0.5

-7.4

2.8±0.5

-1.3

L-Am/PCMA

64.9±0.2

-2.6

2.9±0.5

-1.2

L-Am/AA

63.9±0.2

-3.6

3.7±0.4

-0.4

L-Am/AA*

63.6±0.2

-3.9

3.3±0.6

-0.8

H-Am/AA

57.8±0.2

-9.7

4.6±0.4

0.5

L-Am/DMAEMA

63.5±0.2

-4

2.9±0.4

-1.2

L-Am/DMAEMA*

63.8±0.2

-3.7

3.2±0.4

-0.9

H-Am/DMAEMA

57.2±0.2

-10.3

4.7±0.5

0.6

Sample


35

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 36 of 42

L-AGA

58.6±0.2

-8.9

4.8±0.4

0.7

H-AGA

59±0.2

-8.5

5±0.4

0.9

These stability data were used to estimate thermodynamic parameters for conjugate folding. The tryptophan fluorescence in the Gdn-HCl denaturation study was used to estimate the Gibbs free energy of protein folding (∆G) for each construct, and the Tm was used to extrapolate an enthalpy (∆H) and entropy (∆S) of folding, using the fact that at Tm, the ∆G of folding is zero. These thermodynamic parameters and their associated standard errors are given in Table S4. Due to the relatively large standard errors, typically 20-30% of the thermodynamic parameter, it is important to note that all thermodynamic conclusions are qualitative. Several key trends can be seen in the thermodynamic data. As anticipated at ambient temperature the folded state is thermodynamically favored. As the polymer molecular weight increases, the enthalpy of folding (∆H) becomes more negative, although the entropy of folding (∆S) also becomes more negative. This supports the hypothesis that higher molecular weight polymers offer more opportunities for the polymer to interact with and non-covalently tie different segments of the protein together, leading to enthalpic stabilization. However, the presence of a higher molecular weight polymer requires a more specific set of configurations for the conjugate to be in the folded state, leading to an entropic penalty for folding with a higher molecular weight polymer. Finally, the polymers containing charged groups DMAEMA and AA, as well as AGA tended to have a more negative enthalpy of folding compared to other polymers of similar molecular weight. This suggests that the cationic and anionic groups in the Am/AA and Am/DMAEMA polymers interact with complementary charges on the protein. This allows the polymer to supramolecularly bind and wrap previously independent segments of the protein. Similarly the AGA groups have been


36

Page 37 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

shown to bind the active site of the enzyme,55–57 potentially leading to stabilization by supramolecularly binding two segments of the protein together through hydrogen bonds at the active site. This is likely to make the entropy of folding more negative and less entropically favorable.

Conclusions In this study, we demonstrate that modification of HEWL with polymers of varying functional groups and lengths can significantly impact the activity and the stability of the resulting conjugates. In general, enzyme activity was reduced with increasing length of the attached polymer chain, most likely due to reduction in accessibility of the substrate to the active site. However, for the negatively charged M. lysodeikticus, enzyme activity was further reduced when the enzyme was modified with anionic polymers due to charge repulsion between the polymer chains and the anionic cells. Modification of HEWL with cationic polymers showed an increase in HEWL activity, most likely due to electrostatic attraction between the negatively charged cell wall and the positively charged polymer-conjugate. Polymer modification of HEWL had a complex effect on enzyme stability. The thermal stability of all conjugates was reduced as determined by the Tm of the conjugates. However, an increase in chemical stability against guanadine HCl was observed for many of the polymer conjugates. This increase in stabilization is most likely due to increased hydrogen bonding and ionic interactions between the functional groups on the polymer chains and the residues on the enzyme surface. Finally, molecular dynamics simulation of PEG and acrylamide oligomers attached to HEWL suggest that the functionality of the polymer chains, the protein attachment site, and residues adjacent to the attachment site each influence interactions between the polymer chain and the surface of the


37

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 38 of 42

protein. Future studies using site-specific modification will be important for determining the importance of modification site on the stability of protein-polymer conjugates.

Supporting Information Available Additional information including MALDI-MS data for lysozyme and the polymer modified conjugates, MALDI-TOF-MS data for the trypsin digested lysozyme and conjugates, details on X-ray crystal structure data collection and refinement, activity and stability data for the polymer modified conjugates, and thermodynamic parameters for the resulting lysozyme conjugates are available in the Supporting Information. This material is available free of charge via the Internet at http://pubs.acs.org.

Acknowledgements We are grateful to M. Sameer Al-Abdul-Wahid for assistance with the mass spectrometry analysis. DK, RCP and JAB acknowledge institutional startup funds from Miami University. JAB acknowledges the Miami University Committee on Faculty Research for support. The Advanced Light Source is supported by the US Department of Energy under contract number DE-AC03-76SF00098 at Lawrence Berkeley National Laboratory.

Corresponding Author Information *Jason A. Berberich, Ph.D. Email: [email protected] *Richard C. Page, Ph.D. Email: [email protected] *Dominik Konkolewicz, Ph.D. Email: [email protected]


38

Page 39 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

References (1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24) (25) (26) (27) (28)

Obermeyer, A. C.; Olsen, B. D. ACS Macro Lett. 2015, 4 (1), 101–110. Shu, J. Y.; Panganiban, B.; Xu, T. Annu. Rev. Phys. Chem. 2013, 64 (1), 631–657. Falatach, R.; Li, S.; Sloane, S.; McGlone, C.; Berberich, J. A.; Page, R. C.; Averick, S.; Konkolewicz, D. Polymer 2015, 72, 382–386. Veronese, F. M.; Mero, A. BioDrugs Clin. Immunother. Biopharm. Gene Ther. 2008, 22 (5), 315–329. Pelegri-O’Day, E. M.; Lin, E.-W.; Maynard, H. D. J. Am. Chem. Soc. 2014, 136 (41), 14323–14332. Hoffman, A. S. Adv. Drug Deliv. Rev. 2013, 65 (1), 10–16. Moatsou, D.; Li, J.; Ranji, A.; Pitto-Barry, A.; Ntai, I.; Jewett, M. C.; O’Reilly, R. K. Bioconjug. Chem. 2015, 26 (9), 1890–1899. Cobo, I.; Li, M.; Sumerlin, B. S.; Perrier, S. Nat. Mater. 2015, 14 (2), 143–159. Yang, Z.; Mesiano, A. J.; Venkatasubramanian, S.; Gross, S. H.; Harris, J. M.; Russell, A. J. J. Am. Chem. Soc. 1995, 117 (17), 4843–4850. Lee, P.; Towslee, J.; Maia, J.; Pokorski, J. Macromol. Biosci. 2015, 15 (10), 1332–1337. Drevon, G. F.; Danielmeier, K.; Federspiel, W.; Stolz, D. B.; Wicks, D. A.; Yu, P. C.; Russell, A. J. Biotechnol. Bioeng. 2002, 79 (7), 785–794. Zhang, L.; Wu, S.; Buthe, A.; Zhao, X.; Jia, H.; Zhang, S.; Wang, P. ACS Appl. Mater. Interfaces 2012, 4 (11), 5981–5987. Huang, A.; Qin, G.; Olsen, B. D. ACS Appl. Mater. Interfaces 2015, 7 (27), 14660–14669. Pandey, B. K.; Smith, M. S.; Torgerson, C.; Lawrence, P. B.; Matthews, S. S.; Watkins, E.; Groves, M. L.; Prigozhin, M. B.; Price, J. L. Bioconjug. Chem. 2013, 24 (5), 796–802. Mancini, R. J.; Lee, J.; Maynard, H. D. J. Am. Chem. Soc. 2012, 134 (20), 8474–8479. Chao, S.-H.; Matthews, S. S.; Paxman, R.; Aksimentiev, A.; Gruebele, M.; Price, J. L. J. Phys. Chem. B 2014, 118 (28), 8388–8395. Cummings, C.; Murata, H.; Koepsel, R.; Russell, A. J. Biomacromolecules 2014, 15 (3), 763–771. Murata, H.; Cummings, C. S.; Koepsel, R. R.; Russell, A. J. Biomacromolecules 2013, 14 (6), 1919–1926. Cummings, C.; Murata, H.; Koepsel, R.; Russell, A. J. Biomaterials 2013, 34 (30), 7437– 7443. Gauthier, M. A.; Klok, H.-A. Polym. Chem. 2010, 1 (9), 1352. López-Cruz, J. I.; Viniegra-González, G.; Hernández-Arana, A. Bioconjug. Chem. 2006, 17 (4), 1093–1098. Pai, S. S.; Przybycien, T. M.; Tilton, R. D. Langmuir 2010, 26 (23), 18231–18238. Lele, B. S.; Russell, A. J. Biomacromolecules 2005, 6 (1), 475–482. Lele, B. S.; Russell, A. J. Biomacromolecules 2004, 5 (5), 1947–1955. Harris, J. M.; Chess, R. B. Nat. Rev. Drug Discov. 2003, 2 (3), 214–221. Averick, S.; Mehl, R. A.; Das, S. R.; Matyjaszewski, K. J. Controlled Release 2015, 205, 45–57. Averick, S.; Simakova, A.; Park, S.; Konkolewicz, D.; Magenau, A. J. D.; Mehl, R. A.; Matyjaszewski, K. ACS Macro Lett. 2012, 1 (1), 6–10. Sumerlin, B. S. ACS Macro Lett. 2012, 1 (1), 141–145.


39

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 40 of 42

(29) Lele, B. S.; Murata, H.; Matyjaszewski, K.; Russell, A. J. Biomacromolecules 2005, 6 (6), 3380–3387. (30) Boyer, C.; Bulmus, V.; Liu, J.; Davis, T. P.; Stenzel, M. H.; Barner-Kowollik, C. J. Am. Chem. Soc. 2007, 129 (22), 7145–7154. (31) Li, H.; Bapat, A. P.; Li, M.; Sumerlin, B. S. Polym. Chem. 2011, 2 (2), 323–327. (32) Li, H.; Li, M.; Yu, X.; Bapat, A. P.; Sumerlin, B. S. Polym. Chem. 2011, 2 (7), 1531– 1535. (33) De, P.; Li, M.; Gondi, S. R.; Sumerlin, B. S. J. Am. Chem. Soc. 2008, 130 (34), 11288– 11289. (34) Falatach, R.; McGlone, C.; Al-Abdul-Wahid, M. S.; Averick, S.; Page, R. C.; Berberich, J. A.; Konkolewicz, D. Chem Commun 2015, 51 (25), 5343–5346. (35) Matsuda, T.; Sugawara, T. Macromolecules 1996, 29 (16), 5375–5383. (36) Cook, M. T.; Smith, S. L.; Khutoryanskiy, V. V. Chem. Commun. 2015, 51 (77), 14447– 14450. (37) Niesen, F. H.; Berglund, H.; Vedadi, M. Nat. Protoc. 2007, 2 (9), 2212–2221. (38) Kabsch, W. Acta Crystallogr. D Biol. Crystallogr. 2010, 66 (Pt 2), 125–132. (39) Phillips, J. C.; Braun, R.; Wang, W.; Gumbart, J.; Tajkhorshid, E.; Villa, E.; Chipot, C.; Skeel, R. D.; Kalé, L.; Schulten, K. J. Comput. Chem. 2005, 26 (16), 1781–1802. (40) Diederichs, K.; Karplus, P. A. Acta Crystallogr. D Biol. Crystallogr. 2013, 69 (Pt 7), 1215–1222. (41) Sauter, C.; Otálora, F.; Gavira, J.-A.; Vidal, O.; Giegé, R.; García-Ruiz, J. M. Acta Crystallogr. D Biol. Crystallogr. 2001, 57 (8), 1119–1126. (42) McCoy, A. J.; Grosse-Kunstleve, R. W.; Adams, P. D.; Winn, M. D.; Storoni, L. C.; Read, R. J. J. Appl. Crystallogr. 2007, 40 (4), 658–674. (43) Adams, P. D.; Afonine, P. V.; Bunkóczi, G.; Chen, V. B.; Davis, I. W.; Echols, N.; Headd, J. J.; Hung, L.-W.; Kapral, G. J.; Grosse-Kunstleve, R. W.; McCoy, A. J.; Moriarty, N. W.; Oeffner, R.; Read, R. J.; Richardson, D. C.; Richardson, J. S.; Terwilliger, T. C.; Zwart, P. H. Acta Crystallogr. D Biol. Crystallogr. 2010, 66 (2), 213–221. (44) Emsley, P.; Lohkamp, B.; Scott, W. G.; Cowtan, K. Acta Crystallogr. D Biol. Crystallogr. 2010, 66 (4), 486–501. (45) Chen, V. B.; Arendall, W. B.; Headd, J. J.; Keedy, D. A.; Immormino, R. M.; Kapral, G. J.; Murray, L. W.; Richardson, J. S.; Richardson, D. C. Acta Crystallogr. D Biol. Crystallogr. 2010, 66 (Pt 1), 12–21. (46) MacKerell, A. D.; Bashford, D.; Bellott, M.; Dunbrack, R. L.; Evanseck, J. D.; Field, M. J.; Fischer, S.; Gao, J.; Guo, H.; Ha, S.; Joseph-McCarthy, D.; Kuchnir, L.; Kuczera, K.; Lau, F. T. K.; Mattos, C.; Michnick, S.; Ngo, T.; Nguyen, D. T.; Prodhom, B.; Reiher, W. E.; Roux, B.; Schlenkrich, M.; Smith, J. C.; Stote, R.; Straub, J.; Watanabe, M.; Wiórkiewicz-Kuczera, J.; Yin, D.; Karplus, M. J. Phys. Chem. B 1998, 102 (18), 3586– 3616. (47) Skrabania, K.; Miasnikova, A.; Bivigou-Koumba, A. M.; Zehm, D.; Laschewsky, A. Polym. Chem. 2011, 2 (9), 2074–2083. (48) Pace, C. N.; McGrath, T. J. Biol. Chem. 1980, 255 (9), 3862–3865. (49) Suckau, D.; Mak, M.; Przybylski, M. Proc. Natl. Acad. Sci. 1992, 89 (12), 5630–5634. (50) da Silva Freitas, D.; Abrahão-Neto, J. Int. J. Pharm. 2010, 392 (1–2), 111–117. (51) Lee, H.; Park, T. G. J. Pharm. Sci. 2003, 92 (1), 97–103. (52) Hamed, E.; Ma, D.; Keten, S. ACS Biomater. Sci. Eng. 2015, 1 (2), 79–84.


40

Page 41 of 42

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Biomacromolecules

(53) Lam, C. N.; Chang, D.; Wang, M.; Chen, W.-R.; Olsen, B. D. J. Polym. Sci. Part A Polym. Chem. 2016, 54 (2), 292–302. (54) Lysozyme Worthington Enzyme Manual http://www.worthingtonbiochem.com/ly/default.html (accessed Dec 21, 2015). (55) Rupley, J. A.; Butler, L.; Gerring, M.; Hartdegen, F. J.; Pecoraro, R. Proc. Natl. Acad. Sci. U. S. A. 1967, 57 (4), 1088–1095. (56) Von Dreele, R. B. Acta Crystallogr. D Biol. Crystallogr. 2001, 57 (Pt 12), 1836–1842. (57) Ikeda, K.; Hamaguchi, K. J. Biochem. (Tokyo) 1969, 66 (4), 513–520. (58) Park, H.; Robinson, J. R. Pharm. Res. 1987, 4 (6), 457–464. (59) Högfeldt, E.; Miyajima, T.; Marinsky, J. A.; Muhammed, M. Acta Chem Scand 1989, 43, 496–499. (60) Van de Wetering, P.; Zuidam, N. J.; Van Steenbergen, M. J.; Van der Houwen, O.; Underberg, W. J. M.; Hennink, W. E. Macromolecules 1998, 31 (23), 8063–8068. (61) Satishkumar, R.; Vertegel, A. Biotechnol. Bioeng. 2008, 100 (3), 403–412. (62) Murata, H.; Cummings, C. S.; Koepsel, R. R.; Russell, A. J. Biomacromolecules 2014, 15 (7), 2817–2823. (63) Sandanaraj, B. S.; Vutukuri, D. R.; Simard, J. M.; Klaikherd, A.; Hong, R.; Rotello, V. M.; Thayumanavan, S. J. Am. Chem. Soc. 2005, 127 (30), 10693–10698. (64) Vollrath, F.; Hawkins, N.; Porter, D.; Holland, C.; Boulet-Audet, M. Sci. Rep. 2014, 4, 5625.


41

Biomacromolecules

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 42 of 42

Table of Contents Graphic:


42

Investigating the Impact of Polymer Functional Groups on the

Recommend Documents