Design of Stomach Acid-Stable and Mucin-Binding Enzyme Polymer

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Design of stomach acid-stable and mucin binding-enzyme polymer conjugates Chad S. Cummings, Alan S Campbell, Stefanie L Baker, Sheiliza Carmali, Hironobu Murata, and Alan J Russell Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.6b01723 • Publication Date (Web): 12 Jan 2017 Downloaded from http://pubs.acs.org on January 18, 2017

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Biomacromolecules

Design of stomach acid-stable and mucin-

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binding enzyme polymer conjugates

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3 Chad S. Cummings1,2, Alan S. Campbell1,2, Stefanie Baker1,2, Sheiliza Carmali1,

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Hironobu Murata1, Alan J. Russell1,2,3,4*

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1

Center for Polymer-based Protein Engineering, Carnegie Mellon University, Pittsburgh,

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PA 15213 2

Department of Biomedical Engineering, Carnegie Mellon University, Pittsburgh, PA

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15213 3

Disruptive Health Technology Institute, Carnegie Mellon University, Pittsburgh, PA

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15213 4

Department of Chemistry, Carnegie Mellon University, Pittsburgh, PA 15213

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17 1

ACS Paragon Plus Environment

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ABSTRACT

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The reduced immunogenicity and increased stability of protein-polymer conjugates has

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made their use in therapeutic applications particularly attractive. However, the physicochemical

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interactions between polymer and protein, as well as the effect of this interaction on protein

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activity and stability, are still not fully understood. In this work, polymer-based protein

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engineering was used to examine the role of polymer physiochemical properties on the activity

7

and stability of the chymotrypsin-polymer conjugates and their degree of binding to intestinal

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mucin. Four different chymotrypsin-polymer conjugates, each with the same polymer density,

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were synthesized using “grafting-from” atom transfer radical polymerization. The influence of

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polymer charge on chymotrypsin-polymer conjugate mucin binding, bioactivity, and stability in

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stomach acid was determined. Cationic polymers covalently attached to chymotrypsin showed

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high mucin binding, while zwitterionic, uncharged, and anionic polymers showed no mucin

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binding. Cationic polymers also increased chymotrypsin activity from pH 6-8, while zwitterionic

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polymers had no effect, and uncharged and anionic polymers decreased enzyme activity. Lastly,

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cationic polymers decreased the tendency of chymotrypsin to structurally unfold at extremely

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low pH, while uncharged and anionic polymers induced unfolding more quickly. We

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hypothesized that when polymers are covalently attached to the surface of a protein, the degree

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to which those polymers interact with the protein surface is the predominant determinant of

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whether the polymer will stabilize or inactivate the protein. Preferential interactions between the

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polymer and the protein lead to removal of water from the surface of the protein and this, we

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believe, inactivates the enzyme.

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INTRODUCTION

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Protein-polymer conjugates have become central components of the biologic drug, synthetic

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and food industries. Chymotrypsin is one of the most commonly studied protein-polymer

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conjugates1-3 because there is a wealth of published information about the amino acid sequence,4

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crystal structure,5 and substrate preferences6,7 under a host of reaction conditions. Chymotrypsin

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is stable over a reasonably wide pH range8 and in many organic solvents.9,10 One of the most

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useful applications, and certainly the most therapeutically relevant application of chymotrypsin,

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is as an enzyme replacement therapy.11 In humans, chymotrypsin is secreted by the pancreas and

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is active in the small intestine, where it breaks down proteins. Specifically, chymotrypsin

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replacement therapy is used to treat diseases where low levels of the enzyme are a symptom.12

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During this therapy, exogenous chymotrypsin is delivered orally to the gastrointestinal (GI) tract.

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Unfortunately for exogenous chymotrypsin delivery, the stomach uses acid and proteases to

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rapidly breakdown proteins (including chymotrypsin) in order to facilitate amino acid nutrient

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absorption into the bloodstream. Unmodified chymotrypsin has been investigated as an enzyme

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replacement therapy to treat autism,13 cystic fibrosis,14-16 and exocrine pancreatic insufficiency.17

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Many other enzymes such as lactase,18-20 lipase,21,22 and amylase23 are used with varying

17

modification to treat GI tract disorders. In order for an enzyme replacement therapy to be

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optimally effective in the GI tract, it would need to remain stable from pH 1 to 8. In addition, the

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enzyme would ideally have a residence time that was extended compared to digested proteins.

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The mucosal innermost lining of the GI tract is replete with the glycosylated protein mucin,

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which is known to bind charged and hydrophilic polymers. One approach to keeping a

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therapeutic protein in the intestinal tract would be to combine the enzyme with mucoadhesive

23

molecules.24,25 3


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Unmodified chymotrypsin is active from pH 5-10,26 with its pH optimum at pH 8.27

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Interestingly, a cationic enzyme-polyamine conjugate was shown to be hyperactive at a wide

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range of pH.28 Additionally, polymer-encapsulated,29 protein engineered,30,31 or polymer

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conjugated32,33 enzymes exhibit some stability under harsh digestive tract conditions.

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Chymotrypsin has a close to net-neutral surface charge and as a result native chymotrypsin is not

6

mucoadhesive. We hypothesized, therefore, that polymer-based protein engineering of

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chymotrypsin with charged polymers could generate enzyme variants that were mucin-binding

8

and stable at extremes of acidic pH.

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To further investigate the activity and mucin-binding of charged chymotrypsin-polymer

10

conjugates, we grew neutral, zwitterionic, and charged polymers directly from the surface of

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chymotrypsin using atom-transfer radical polymerization. The polymers, poly(carboxybetaine

12

acrylamide)

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poly(quaternary ammonium methacrylate) (pQA(+)), and poly(sulfonate methacrylate) (pSMA(-

14

)), were chosen to incorporate charged moieties (sulfonate anion, ammonium cation) generally

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considered to be kosmotropes (order-making/stabilizing) in the Hofmeister series.34,35 Both

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positively and negatively charged polymers have been shown to possess mucoadhesive

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properties as a result of binding to the strongly hydrophilic glycosylated polymers that cover

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mucin proteins.36,37 While uncharged and likely not mucoadhesive, pOEGMA has been shown to

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improve protein stability to different stressors such as temperature,38 protease degradation,39 and

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lyophilization.40 We hypothesized that each of the conjugates would have an impact on enzyme

21

stability at low pH while each polymer would have a distinct impact on mucin-binding of the

22

chymotrypsin-polymer conjugates. In this first study, our goal was to elucidate the relationship

(pCBAm(+/-)),

poly(oligoethylene

glycol

methacrylate)

(pOEGMA),

4


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between polymer physicochemical properties and the activity, stability, and mucin binding of the

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chymotrypsin-polymer conjugates.

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MATERIALS

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All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) and used as received

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unless otherwise indicated. Poly(ethylene glycol) methyl ether methacrylate (Mn = 475)

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(OEGMA475) was filtered through basic alumina column to remove inhibitor prior to use.

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Me6TREN,41 carboxybetaine acrylamide,42 and quaternary ammonium methacrylate43 were

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synthesized as described previously. Dialysis tubing (molecular weight cut off, 15kDa,

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Spectra/Por®, Spectrum Laboratories Inc., CA) for conjugate isolation were purchased from

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Thermo Fisher Scientific (Waltham, MA).

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METHODS

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Initiator Immobilization onto Chymotrypsin

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Synthesis of the ATRP initiating molecules was carried out as described previously.44

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Following synthesis, the initiator molecule (NHS-Br) (469 mg, 1.4 mmol) and CT (1.0 g, 0.04

15

mmol protein, 0.56 mmol -NH2 group in lysine residues) were dissolved in sodium phosphate

16

buffer (100 mL, 0.1 M NaPhos (pH 8)). The solution was stirred at 4 °C for 3 hours, and then

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dialyzed against deionized water, using dialysis tubing with a molecular weight cut off of 15

18

kDa, for 24 hours at 4 °C and then lyophilized. Initiator immobilization was quantified using

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matrix assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF-MS)

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on a PerSeptive Voyager STR MS with nitrogen laser (337 nm) and 20kV accelerating voltage

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located at the CMA, CMU, Pittsburgh, PA using sinapinic acid as the matrix and a gold sample

22

plate. MALDI-TOF MS instrumentation was supported by NSF grant CHE-9808188.

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Surface Initiated ATRP from CT-Br 5


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Chymotrypsin-pOEGMA and chymotrypsin-pSMA were synthesized using CuCl/CuCl2/bpy in

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deionized water.45 For CT-pOEGMA, 4.6 mL of a deoxygenated CuCl/CuCl2/bpy stock solution

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(5mM/45mM/110mM) in deionized water was added to 16.4 mL of a CT-Br (50 mg, 1.4 mM

4

initiator) and OEGMA475 (2415 mg, 330 mM, targeted degree of polymerization 225) solution in

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deoxygenated deionized water and allowed to react at 4 °C for 80 minutes. CT-pSMA was

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synthesized by adding 4.6 mL of stock CuCl/CuCl2/bpy (5 mM/45mM/110 mM) in

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deoxygenated deionized water to 16.4 mL of CT-Br (50 mg, 1.4 mM initiator) and SMA (1190

8

mg, 285 mM, targeted degree of polymerization 227) in deoxygenated 100 mM NaPhos (pH 7)

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and allowed to react for 65 minutes at 4 °C. CT-pQA was synthesized by adding 2 mL of CuBr

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(3.7 mg, 16 mM) and HMTETA (7.4 mg, 16 mM) in deoxygenated deionized water to 25 mL of

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CT-Br (50 mg, 1.4 mM initiator) and QA monomer (405 mg, 64 mM) in 64 mM deoxygenated

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NaSO4 solution and allowed to react at 25°C for 120 minutes.46 Lastly, CT-pCBAm was

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synthesized by adding 5 mL of CT-Br (50 mg, 1.4 mM initiator) and CBAm (348 mg, 332 mM)

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in deoxygenated 100 mM NaPhos (pH 7) buffer to 2 mL of CuCl (2.5 mg, 12 mM) and

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Me6TREN (5.5 mg, 12 mM) in deoxygenated deionized water and allowed to react for 120

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minutes at 4°C.47 All conjugates were purified using dialysis tubing (MWCO 25 kDa) against

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deionized water for 48 hours at 4°C. Samples were lyophilized and chymotrypsin weight percent

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in each conjugate was determined using BCA assay.

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Molecular Characterization of Conjugates and Polymers

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All polymers were cleaved from the surface of CT-polymer conjugates using acid hydrolysis.

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CT conjugates (15 mg/mL) were incubated in 6N HCl at 110 °C under vacuum for 24 hours.

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Following incubation, cleaved polymers were isolated from CT using dialysis tubing (MwCO 1K 6


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Da) for 48 hours and then lyophilized. Number and weight average molecular weights (Mn and

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Mw) and the polydispersity index (Mw / Mn) were estimated by gel permeation chromatography

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(GPC) for polymers cleaved from CT. Analysis was conducted on a Waters 2695 Series with a

4

data processor, using 0.1 M sodium phosphate buffer (pH 7.0) with 0.01 volume % NaN3

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(pOEGMA, pCBAM), 0.1 M sodium phosphate (pH 2.0) with 0.5 % TFA (pQA), or 80%

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sodium phosphate (pH 9.0)/20% acetonitrile (pSMA) as eluent at a flow rate 1 mL/min, with

7

detection by a refractive index (RI) detector, and PEG (pOEGMA, pCBAm, pQA) or polystyrene

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sulfonate (pSMA) narrow standards for calibration.

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A Micromeritics (Norcross, GA) NanoPlus 3 dynamic light scattering (DLS) instrument was

10

used to measure the intensity average hydrodynamic diameter (Dh) of each of the chymotrypsin

11

conjugates at 2 mg/mL in 50 mM NaPhos (pH 7) buffer at 25 °C. Histograms of results were

12

plotted after 70 accumulation times, and average Dh values were calculated from these runs.

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In Vitro Polymer Mucoadhesion

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Mucoadhesion of free polymers was evaluated using mucin in different buffer systems. Free

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polymers were synthesized by the same protocol as for CT conjugates, but with a small molecule

16

initiator instead of the chymotrypsin macroinitiator. Polymers were dissolved at 1 mg/mL in

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different buffers (167 mM HCl (pH 1), 50 mM ammonium acetate (pH 4.5), 50 mM NaPhos (pH

18

8)) and mixed with mucin protein (3 mg/mL in deionized water) at different weight ratios. After

19

mixing, solutions were incubated for 30 minutes at 37 °C and absorbance at 400 nm (turbidity)

20

was recorded. Turbidity measurements were plotted as relative ratios to the turbidity

21

measurement at w/w ratio 0.0. For experiments with NaCl and ethanol, polymers were dissolved

22

in buffer solutions with either 0.2 M NaCl, 0.5 M NaCl, or 10% v/v ethanol and then mixed with

23

mucin. 7


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Free polymer zeta potential (ζ) values were measured on a Micromeritics (NanoPlus 3)

2

zetasizer instrument. Free polymers were dissolved at 2 mg/mL in specified buffer solution. Zeta

3

potential values were averages of 4 repeat runs.

4

CT Conjugate Biocatalytic Activity

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N-succinyl-L-Ala-L-Ala-L-Pro-L-Phe-p-nitroanilide was used as a substrate for enzyme

6

bioactivity assays. In a cuvette, 0.1 M sodium phosphate buffer (930-990 µL, pH 6,7, or 8),

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substrate (0-60 µL, 6 mg/mL in DMSO), and enzyme (10 µL, 0.1 mg enzyme/mL 0.1 M pH 8.0

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sodium phosphate buffer (4 µM)) were mixed at 37 °C using a circulating water bath. The rate of

9

the hydrolysis was determined by recording the increase in absorbance at 412 nm for the first 30

10

seconds after mixing. KM and kcat values were calculated using Graphpad software with

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Michaelis-Menten curve fit when plotting substrate concentration versus the initial rate for

12

substrate hydrolysis.

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In Vitro Gastric Acid Stability

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Native CT and CT-conjugates were incubated at 4 µM in 167 mM HCl at 37 ° C in 50 µL

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aliquots. Aliquots were removed at specified time points and residual activity was measured at

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37 °C in 0.1 M sodium phosphate buffer (pH 8.0) with Suc-AAPF-pNA as substrate (288 µM).

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Each time point was measured in triplicate and residual activity was calculated as the ratio of

18

activity remaining from time zero.

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Intrinsic Tryptophan Fluorescence of CT Conjugates

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CT conjugates were incubated at 37 °C in 167 mM HCl (pH 1) at 12 µM CT in 100 µL

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aliquots for each time point. At the specified time point, samples were diluted to 4 µM using 0.1

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M NaPhos buffer (pH 8) and the intrinsic fluorescence was measured in triplicate at 37 °C.

23

Spectrum emission from 300-400 nm was measured for each sample after excitation at 270 nm. 8


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The wavelength values corresponding to the maximum emission intensity for each measurement

2

were calculated and the average maximum wavelength (λmax) was plotted for each sample.

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Surface Charge Analysis of CT Initiator Complex

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The initial structure of CT-Initiator complex was built with Maestro built toolkit (Schrodinger)

5

using the crystal structure of CT from the Protein Data Bank (PDB ID 1YPH) as the starting

6

structure. To remove the bias and constraints of the starting point, the structure was subjected to

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a Simulated Annealing (SA) protocol using Desmond.48 This annealing protocol consisted of

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three stages with 100, 300, and 600 ps durations and temperature intervals from 300-400 K, 450-

9

300 K, and 300 K, respectively. The simulation system was prepared using Desmond’s system

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builder with the OPLS-2005 force field and SPC was chosen as a solvent model. An

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orthorhombic shape was chosen for the simulation box and its volume minimized with Desmond

12

tool with no ions added to neutralize the system. NVT ensemble and the Berendsen thermostat

13

method were used for temperature coupling with a relaxation time of 1 ps. A cutoff of 9 Å for

14

van der Waals interactions was applied, and the particle mesh Ewald algorithm was used for

15

Coulomb interactions with a switching distance of 9 Å. The total simulation time was 1 ns with

16

recording interval energy 1.2 ps and recording trajectory of 5 ps. The final structure obtained

17

after SA was then subjected molecular dynamics simulation (MD). Finally, a 10 ns MD

18

simulation was performed using Desmond at 300 K with a time-step bonded of 2 fs. Trajectory

19

energy values were recorded every 1.2 ps and structure energy was recorded every 4.8 ps. NPT

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ensemble, the ‘Nose-Hoover chain’ thermostat, and ‘Martyna-Tobia-Klein’ Barostat methods

21

were used with 2 ps relaxation time and isotropic coupling. The default relaxation model, a

22

cutoff of 9 Å for van der Waals interactions, and 200 force constant restrain one atom from the

9


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backbone were applied. The particle mesh Ewald algorithm was used for Coulomb interactions

2

with a switching distance of 9 Å and no ions were added to the solution.

3

Predicted ionization states of chymotrypsin-initiator complex at neutral pH and pH 1 were

4

determined using PROPKA 2.0.49 Surface charge analysis and molecular graphics for CT-Br

5

were obtained using electrostatic potential coulombic surface coloring in UCSF Chimera

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package.50

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RESULTS AND DISCUSSION

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CT Conjugate Synthesis and Polymer Characterization

10

To determine the relationship between polymer physicochemical properties and enzyme

11

activity, stability and mucin binding, we designed and synthesized four chymotrypsin-polymer

12

biohybrid conjugates. pCBAm (+/-) was zwitterionic with a net neutral charge, pOEGMA was

13

uncharged and neutral, pQA(+) was positively charged, and pSMA(-) was negatively charged.

14

The polymers were grown directly from the surface of chymotrypsin using atom-transfer radical

15

polymerization (ATRP) as described above. An idealistic representation of the final protein-

16

polymer conjugates is shown in Figure 1. The specific conditions for the synthesis of the

17

enzyme-polymer conjugates, as listed in Table 1, were selected in order to optimize the

18

polymerization with each of the monomers used.

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Figure 1 Schematic representations of CT-pOEGMA, CT-pCBAm (+/-), CT-pSMA

3

(-), and CT-pQA (+). The charge state of each polymer is shown at pH 7; below pH 4.5 the

4

carboxylic acid in pCBAm was protonated and pCBAm had an overall positive charge. The

5

charge states of the other polymers have no pH-dependence from pH 1-8.

6 7

Polymers were grown from 12 ATRP initiator sites (as calculated by MALDI-TOF-MS)

8

covalently attached to surface accessible lysine residues using NHS-ester/amine chemistry.

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Successful polymerization from chymotrypsin was confirmed using dynamic light scattering

10

(DLS), and each chymotrypsin-polymer conjugate had a similar increase in hydrodynamic

11

diameter (Dh) compared to native chymotrypsin (5.7 ± 2 nm) (Figure 2). To characterize

12

polymers grown from chymotrypsin, the polymers were cleaved from the surface of

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chymotrypsin using acid hydrolysis in 6 N HCl. Polymer molar mass was calculated using size

14

exclusion GPC, and polymer molar mass values correlated well with hydrodynamic diameters

15

measured by DLS. (

16

Table 1)

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Figure 2 The dependence of chymotrypsin-polymer hydrodynamic diameter on

3

charge state of the polymer. CT-pCBAm (+/-) (26.3 ± 3.2 nm), CT-pOEGMA (20.1 ± 2.0 nm),

4

CT-pQA(+) (34.5 ± 2.9 nm), and CT-pSMA (-) (17.2 ± 2.2 nm) hydrodynamic diameter values

5

were measured by dynamic light scattering (DLS) in 50 mM sodium phosphate (pH 7.0, 25 °C).

6

Native chymotrypsin hydrodynamic diameter is considerably smaller than that of the conjugates

7

(5.7 ± 2 nm).

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Biomacromolecules

Table 1 Molecular weight and hydrodynamic diameter of chymotrypsin conjugates Cleaved Polymer

CT-pCBAm (+/-) CT-pOEGMA CT-pQA (+) CT-pSMA(-)

Cu/Ligand Pair

Mn (kDa)

PDI (Mw/Mn)

CuCl:Me6TREN CuCl:CuCl2:bpy CuBr:HMTETA CuCl:CuCl2:bpy

30.7 11.6 19.1 9.6

1.90 1.46 2.10 1.43

Conjugate Molar Mass (kDa) 393 165 254 140

Size (Dh) [nm] 26.3 ± 3.2 20.1 ± 2.0 34.5 ± 2.9 17.2 ± 2.2

2 3 4

In vitro mucin-binding of ATRP-synthesized free polymers

5

Exogenous enzymes modified with mucin-binding molecules could exhibit increased residence

6

time in the GI tract. In order to examine the in vitro pH-dependence of mucin-binding properties

7

of each of the polymers that were grown from chymotrypsin, we measured the turbidities of

8

mucin protein solutions (at 37 °C) with increasing free polymer content at pH 1 (167 mM HCl),

9

pH 4.5 (50 mM ammonium acetate buffer), and pH 8.0 (50 mM sodium phosphate buffer). In

10

this assay, an increase in the turbidity of mucin colloidal suspensions is driven by mucoadhesive

11

polymer-mediated crosslinking of mucin.51 The positively charged pQA (+) polymer exhibited

12

significant mucin binding across the range of pH values tested. The degree of mucin binding of

13

the zwitterionic polymer, pCBAm, was pH-dependent whereas the neutral polymer, pOEGMA,

14

was not mucoadhesive at any pH. The negatively charged polymer, pSMA (-), was also non-

15

mucin binding across the range of pH tested (Figure 3a-c).

13


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Figure 3 pH-Dependence of mucin-particle crosslinking by ATRP-synthesized free

3

polymers. (a) pH 1.0 (167 mM HCl), (b) pH 4.5 (50 mM ammonium acetate buffer), (c) pH 8

4

(50 mM sodium phosphate buffer), (d) 167 mM HCl with 10% ethanol or 0.2 M NaCl, (e) 50

5

mM ammonium acetate with ethanol or NaCl, and (f) 50 mM sodium phosphate with 10%

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ethanol, 0.2 M NaCl, or 0.5 M NaCl. Normalized absorbance at 400 nm (turbidity) at 37 °C was

2

used as a marker for mucin-particle crosslinking by free polymer.

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Mucoadhesion in vivo results from a balance of electrostatic interactions, hydrogen bonding,

5

and hydrophobic interactions with mucin.24 Sialic acid, a major component in mucin, is a

6

polysaccharide with carboxylic acid functionality giving mucin a net negative charge at neutral

7

pH.52 Since the positively charged polymer, pQA (+), increased the turbidity of mucin

8

suspensions from pH 1-8, we hypothesized that electrostatic interactions were the main driving

9

force for observed pQA (+) mucoadhesion. To test this hypothesis further we increased the ionic

10

strength of the mucin suspension with the addition of sodium chloride (NaCl). Separately, to rule

11

out hydrophobic interactions as the mucoadhesive driving force, we increased the hydrophobicity

12

of the solution with the addition of ethanol. We expected that an electrostatic attraction-mediated

13

mucin binding would be diminished by the salt-mediated increase in ionic strength, whereas a

14

hydrophobic interaction-mediated binding would be diminished by ethanol. Both pCBAm and

15

pQA (+) mucin suspension turbidities were unaffected by the addition of ethanol, but dependent

16

on ionic strength (Figure 3d-f). The addition of NaCl affected polymer mucin binding for both

17

pQA (+) and pCBAm by either decreasing the absolute turbidity or the shifting of turbidity

18

plateau to higher polymer:mucin ratios. From these results, it was clear that the mucoadhesion of

19

pQA (+) (at every pH) and pCBAm (at low pH) was due to electrostatic attraction of the

20

positively charged polymers with the negatively charged mucin. In order to further confirm this

21

hypothesis, the zeta potentials of the free polymers and mucin were measured at each pH (Table

22

2). The zeta potentials of each of the polymers correlated well with electrostatic interactions

23

being responsible for the behavior seen in the in vitro mucoadhesion experiments. The uncharged 15


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Page 16 of 36

1

free polymer, pOEGMA, did not show mucoadhesive properties at any of the pH values tested

2

which was not surprising given the existing literature.51 Predictably, pSMA (-) was also not

3

mucoadhesive, likely due to electrostatic repulsion of the negative charge in the polymer and

4

negatively charged mucin.

5 6

Table 2 Zeta potential (ζ) measurements of free polymer in mucoadhesive relevant solutions Zeta potential (ζ) [mV] Mucin

pCBAm

pOEGMA

pQA (+)

pSMA (-)

50 mM Citric acid (pH 2.3)

0.7 ± 0.4

15 ± 6

1.9 ± 0.6

34 ± 10

-22 ± 3.6

+

-3.6 ± 0.5 -7.1 ± 0.7

0.3 ± 0.4 -2.0 ± 1.8

2.9 ± 2.3 1.2 ± 4.5

29 ± 5.9 7.8 ± 4.0

-25 ± 2.6 -22 ± 5.1

50 mM (NH4 ) acetate (pH 4.5) 50 mM NaPhos (pH 8.0) 7 8

While it was clear that pQA (+) and pCBAm did indeed have mucoadhesive properties, several

9

unexpected and interesting trends resulted for these polymers. The pH responsive behavior of

10

pCBAm was likely due to the ionization state of the carboxylic acid in the polymer at each of the

11

test pH values. In highly acidic conditions (pH 1), protonation occurred in pCBAm, resulting in a

12

net positive charge. However, at pH 4.5 and pH 8, no mucoadhesion was observed for pCBAm

13

due to deprotonation and a net neutral charge. While pQA (+) was mucoadhesive at each pH

14

tested, the normalized absorbance values after incubation for pQA (+) polymers were much

15

higher at pH 8 compared to both pH 1 and pH 4.5. This result was likely due to the reduced

16

number of negatively charged crosslinking sites in mucin at pH 1. As described earlier,

17

carboxylic acid functionality is responsible for the negative charge in mucin, so it was not

18

surprising to see less of a crosslinking effect at low pH values. Interestingly, at pH 4.5, pQA (+)

19

normalized turbidity initially increased before returning to baseline levels at 0.3 w/w ratios. At 16


Page 17 of 36

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Biomacromolecules

1

this pH, it was likely that, at higher ratios of pQA (+), the polymer fully encapsulated mucin

2

particles rather than crosslink between particles, resulting in solubilization and lower turbidity.

3

These results led us to hypothesize that the CT-pQA (+) conjugate would also be mucin binding.

4

As we hypothesized, CT-pQA (+) conjugates were mucin binding at pH 1.0, pH 4.5, and pH 8

5

(Figure 4). At pH 8, neither CT-pCBAM (+/-), CT-pOEGMA, nor CT-pSMA (-) showed mucin

6

binding behavior. As in the case of the free polymer, CT-pCBAm was mucin-binding at pH 1.0

7

and pH 4.5. We expect this was due to the protonation of the carboxylic acid at low pH and a

8

resulting net positive charge. Mucin-binding properties of CT-pOEGMA and CT-pSMA (-)

9

conjugates were not determined at low pH due to protein structural unfolding induced by the

10

polymers.

11

17


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Page 18 of 36

1 2

Figure 4 pH dependence of mucin particle crosslinking by chymotrypsin-polymer

3

conjugates. (a) pH 1.0 (167 mM HCl), (b) pH 4.5 (50 mM ammonium acetate buffer), (c) pH 8

4

(50 mM sodium phosphate buffer). Chymotrypsin polymer conjugates exhibited mucin binding

5

properties consistent with free polymers. At pH 1.0 and pH 4.5, only enzyme conjugates

6

structurally stable to those conditions (CT-pQA, CT-pCBAm) were tested to eliminate the effect

7

of unfolded protein. Native protein showed no mucin-binding properties at equivalent

8

concentrations.

9 18


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Biomacromolecules

1

Impact of polymer charge state on the activity of chymotrypsin-polymer conjugates

2

Kinetic rate (kcat) and substrate binding (KM) constants for the ATRP-synthesized CT-polymer

3

conjugates were determined using a short peptide substrate, N-succinyl-Ala-Ala-Pro-Phe-p-

4

nitroanilide, in 100 mM sodium phosphate buffer (pH 6-8) at 37 °C. Conjugate activity was

5

dependent on the properties of the covalently attached polymer in the protein-polymer conjugate

6

(Figure 5). Relative turnover number (kcat) values for each conjugate were independent of pH,

7

and all decreased compared to native chymotrypsin. CT-pSMA (-) and CT-pOEGMA activity

8

values were both less than half that of native chymotrypsin, while CT-pQA (+) and CT-pCBAm

9

maintained approximately 70% of native chymotrypsin activity after modification. A reduction

10

in kcat has often been observed for enzyme-polymer conjugates with the prevailing hypothesis

11

being that the polymer causes a structural stiffening of the enzyme, though definitive

12

mechanisms have never been determined.53 Our group has previously observed a decrease in

13

relative kcat values for CT-pSBAm-b-pNIPAM conjugates.43 The more significant decrease in

14

activity for CT-pSMA (-) and CT-pOEGMA could have been due to interactions between those

15

specific polymers and chymotrypsin. A large increase in substrate affinity was observed for CT-

16

pQA (+) conjugates, as evidenced by the decrease in KM values. CT-pOEGMA had decreased

17

substrate affinity relative to the native unmodified enzyme, whereas the CT-pCBAm had similar

18

substrate affinity. The KM values for both conjugates were not pH-dependent from pH 6-8.

19

Interestingly, the CT-pSMA (-) conjugate relative KM values were pH-dependent and substrate

20

affinity was decreased at pH 6. The effect of polymer on KM of the CT-pSMA (-) and CT-pQA

21

(+) conjugates for the substrate were most likely due to electrostatic repulsion and attraction,

22

respectively, between the polymer coat around the enzyme and the negatively charged substrate.

23

As a negatively charged substrate molecule, the affinity of the substrate for chymotrypsin has 19


Biomacromolecules

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Page 20 of 36

1

previously been shown to be affected by polymer conjugation.54 The significant KM effects drove

2

increased CT-pQA (+) productivity (kcat/KM) values relative to the native unmodified

3

chymotrypsin. Both CT-pSMA (-) and CT-pOEGMA productivities were lower than native

4

chymotrypsin at each pH, and CT-pCBAm conjugates showed similar productivity to native

5

chymotrypsin at each pH. Importantly, activity for each of the chymotrypsin-polymer conjugates

6

was measured against a four peptide long substrate and activity of these conjugates could be

7

different if activity was tested with a larger protein substrate.55 In vivo, chymotrypsin-polymer

8

conjugates would work most effectively in the intestines, because neutral pH is optimum and

9

foodstuff proteins would already be broken down to peptides by pepsin and acid in the stomach.

10 11 12 13 14 15

20


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Biomacromolecules

1 2

Figure 5 pH-Dependence of kinetics for chymotrypsin- and chymotrypsin-polymer

3

conjugate-catalyzed hydrolysis of a negatively charged substrate. Kinetic constants (a) kcat,

4

(b) KM, and (c) kcat/KM were measured for native chymotrypsin (black open upside down triangle)

5

from pH 6-8 at 37 °C in 100 mM sodium phosphate buffer. Relative kinetic constants (d) kcat, (e)

6

KM, and (f) kcat/KM were calculated for CT-pSMA (purple diamond), CT-pOEGMA (green

7

triangle), CT-pQA (blue circle), and CT-pCBAm (red square) in the same conditions and plotted

8

relative to native chymotrypsin. 21


Biomacromolecules

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1 2

Page 22 of 36

Impact of polymer charge state on the stability of chymotrypsin-polymer conjugates at pH 1.0

3

The rate of irreversible inactivation of the CT conjugates at low pH was determined by

4

incubating conjugates in 167 mM HCl at 37 °C and measuring residual activity at specified time

5

points. (Figure 6a) The stability of CT conjugates in acid was dependent on the polymer

6

attached to chymotrypsin. Both CT-pCBAm (+, at low pH) and CT-pQA (+) conjugates were

7

more stable than native chymotrypsin, with stability profiles similar to what we have observed

8

previously for CT-pSBAm43 and CT-pQA conjugates.43 CT-pOEGMA and CT-pSMA (-)

9

conjugates both lost activity more rapidly than native chymotrypsin, showing that both

10

pOEGMA and pSMA (-) had a destabilizing effect on chymotrypsin. The addition of free pQA

11

(+) or pCBAm (+) polymers to native unmodified chymotrypsin did not stabilize the enzyme

12

from acid-mediated irreversible inactivation, indicating that the covalent attachment of polymers

13

was required for increased stability. (Figure 6b)

14 15 16 17

22


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Biomacromolecules

1 2

Figure 6 Rate of acid-mediated irreversible inactivation of chymotrypsin-polymer

3

conjugates. (a) Native chymotrypsin (black upside down triangle), CT-pCBAm (red square),

4

CT-pOEGMA (green triangle), CT-pQA (+) (blue circle), CT-pSMA (-) (purple diamond) were

5

incubated in 167 mM HCl at 37 °C. (b) Native CT (black upside down triangle) was incubated

6

with pOEGMA (green triangle), pCBAm (red square), pSMA (-) (purple diamond), and pQA (+)

7

(blue circle) free polymers. Activity assays were completed using 288 µM substrate (NS-AAPF-

8

pNA) in 100 mM sodium phosphate (pH 8.0) at 37 °C.

9

The addition of pSMA (-) free polymers to unmodified chymotrypsin actually decreased

10

activity compared to native chymotrypsin, confirming the destabilization effect of pSMA (-)

11

towards chymotrypsin. Conversely, while CT-pOEGMA conjugates showed similar low stability

12

to CT-pSMA (-) conjugates when incubated in 167 mM HCl, native chymotrypsin incubated

13

with free pOEGMA had a similar stability profile to native chymotrypsin, chymotrypsin with

14

pQA (+), and chymotrypsin with pCBAm (+). These data help contribute to our understanding of

15

a proposed mechanism by which the covalently attached polymers either stabilize or destabilize

16

enzymes. In order to develop that mechanism further, we examined the effect of ATRP-grown

23


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Page 24 of 36

1

polymers on the structural integrity of the enzyme by following intrinsic tryptophan fluorescence

2

for each of the conjugates after exposure to low pH. (Figure 7)

3 4

Figure 7 Acid-mediated changes in chymotrypsin-polymer conjugate tertiary

5

structure. Tryptophan intrinsic fluorescence wavelength of maximum emission intensity values

6

(λmax) after incubation in 167 mM HCl (pH 1) at 37 °C for native chymotrypsin (black upside

7

down triangle) and chymotrypsin conjugates; CT-pCBAm (red square), CT-pOEGMA (green

8

triangle), CT-pQA (+) (blue circle), CT-pSMA (-) (purple diamond). An increase in λmax

9

indicates protein unfolding. Time = 0 minutes indicates tryptophan intrinsic fluorescence at pH

10

8, 37 ⁰C.

11 12

Intrinsic tryptophan fluorescence is a recognized sentinel for changes in protein tertiary

13

structure. Protein unfolding leads to increases in the maximum wavelength of fluorescence

14

emission (λmax). Each CT conjugate was incubated at pH 1.0 in 167 mM HCl and fluorescence

15

emission spectrum was measured from 300-400 nm after excitation at 270 nm. Native 24


Page 25 of 36

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Biomacromolecules

1

chymotrypsin λmax increased from 320 nm to 334 nm over 60 minutes. This result was expected

2

as native chymotrypsin has been previously shown to irreversibly unfold at low pH.56,57 Not

3

surprisingly, both CT-pSMA (-) and CT-pOEGMA also had increased lambda max values

4

compared to native chymotrypsin starting at t=0 min. Conversely, CT-pQA (+) had increased

5

λmax values over the course of the experiment, but the increase was not as large in magnitude as

6

native chymotrypsin, indicating significantly less extensive irreversible unfolding for CT-pQA

7

(+) at low pH. CT-pCBAm (+ at pH 1.0) conjugates showed the least amount of unfolding

8

during this experiment as the λmax remained almost unchanged during the course of the

9

experiment. From this experiment, it was clear that the loss in activity for both CT-pOEGMA

10

and CT-pSMA (-) conjugates was due to unfolding and not enzyme autolysis. The results of the

11

intrinsic fluorescence experiments correlated well with residual activity measurements, where

12

both CT-pQA (+) and CT-pCBAm (+ low pH) were more stable than native chymotrypsin. CT-

13

pCBAm (+ at low pH) had higher activity during the course of the experiment. In addition, both

14

CT-pSMA (-) and CT-pOEGMA lost activity quickly during residual activity experiments, and

15

this reduced activity coincided with a large increase in λmax values during intrinsic fluorescence

16

experiments.

17

Mechanism of stabilization of enzymes by conjugated “grown-from” polymers

18

The data generated for the impact of polymer charge on both activity and tertiary structure

19

align well with data generated previously when determining the impact of anionic nanoparticles

20

on the activity and unfolding of chymotrypsin.58 In that work, it was hypothesized that the

21

anionic nanoparticles selectively associated with a cationic core of amino acid residues around

22

the chymotrypsin active site. In addition, the authors hypothesized the hydrophobic nature of the

23

nanoparticles also led to detrimental effects on chymotrypsin stability and activity due to a 25


Biomacromolecules

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Page 26 of 36

1

region of hydrophobic residues also near the active site. Of the destabilizing polymers used in

2

this study, one was negatively charged (pSMA) (-) and the other was amphiphilic (pOEGMA).

3

While the inhibition and destabilizing properties of negatively charged molecules seemed to be

4

conserved in this study, the surface charge of initiator modified chymotrypsin (CT-Br) must be

5

considered rather than native chymotrypsin. Indeed, a large amount of positively charged surface

6

area was lost when coupling the ATRP initiator onto surface lysine residues, which bear a

7

positive charge in native chymotrypsin at neutral pH. We therefore performed a 10 ns molecular

8

dynamics simulation of CT-Br in water to obtain a representative structure of the initiator

9

complex. Ionization states of CT-Br at pH 1 and 7 were predicted using PROPKA 2.0 after

10

which surface charge analysis was possible by calculating electrostatic potentials according to

11

Coulomb’s Law. At pH 7 molecular dynamic simulations showed that the region of positive

12

charge near the active site remains in the initiated enzyme and, thus, this region could exhibit

13

specific interactions with charged conjugated polymers. (Figure 8a) In addition, at pH 1, where

14

the stability experiments were conducted for this study, CT-Br bore a global positive surface

15

charge. (Figure 8b)

16 17 18 19

26


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Biomacromolecules

1 2

Figure 8 Electrostatic potential coulombic surface coloring for CT-Br. CT-Br

3

structures were obtained after a 10 ns molecular dynamics simulations in water. Molecular

4

graphics and surface charge analyses were performed with the UCSF Chimera package at (a)

5

neutral pH 7.0 and (b) pH 1.0. The PROPKA method was used for the prediction of the

6

ionization states in the initiator complex at both pH values.

7 8

Other studies have observed that denaturing osmolytes (urea, guanidine hydrochloride)

9

preferentially accumulated at the protein surface, whereas stabilizing osmolytes (TMAO,

10

betaine) were preferentially excluded from the surface.59 This preferential accumulation or

11

exclusion of osmolytes was due to either specific interactions of the osmolytes with the protein

12

or a global alteration of water structure.60 In any case, stabilizing osmolytes resulted in a stronger

13

hydration layer which strengthened protein structural stability, and denaturing osmolytes

14

displaced water molecules in the hydration layer causing lower stability.61 Destabilizing

15

osmolytes interact with the protein by reducing the thermodynamic penalty for exposing 27


Biomacromolecules

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Page 28 of 36

1

hydrophobic residues usually confined to the protein core. Conversely, stabilizing osmolytes

2

increase the thermodynamic barrier for proteins to transfer to the unfolded from the folded state.

3

Specifically for protein-polymer conjugates, other reports indicated that proteins were

4

stabilized after polymer conjugation due to favorable interactions between the protein and

5

polymer.55,62 In these examples, the authors hypothesized that the polymer non-covalently bound

6

with the protein and stabilized the protein through a mechanism similar to cross-linking.

7

Separately, Price et al. has extensively examined the effect of PEGylation on protein

8

stabilization.63 They found that PEGylation can stabilize proteins both by PEG extension into the

9

solution or PEG interaction with the protein surface.64 Importantly, they determined that

10

PEGylation can be stabilizing or destabilizing in the WW domain of human protein pin 1

11

depending on the location of attachment,65 and that conjugation strategy66 and length of PEG67

12

both heavily influence conformational stability. We hypothesize that grown from pQA (+) and

13

pCBAM (+) stabilized chymotrypsin to low pH structural unfolding by preferential exclusion of

14

the polymer from chymotrypsin surface. (Figure 9) Since polymer interactions with

15

chymotrypsin were thermodynamically unfavorable, the water hydration layer was strengthened,

16

and thereby increased structural stability. We also hypothesized that pSMA (-) and pOEGMA

17

destabilized chymotrypsin through a preferential interaction between the polymer and the protein

18

surface. The differing stabilization profiles for native chymotrypsin incubated with free pSMA (-

19

) and pOEGMA polymer indicated that the mechanism of destabilization may be different for

20

these two polymers. We believe that growing polymers from the surface of a protein will be

21

stabilizing if the polymer and protein surface are designed to not interact strongly with each

22

other (and vice versa). In addition, to see a stabilizing effect, the solvent environment around the

23

protein must not reduce the penalty of exposed hydrophobic residues. Conversely, destabilizing 28


Page 29 of 36

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Biomacromolecules

1

polymers manipulate the solvent environment to reduce the penalty of exposed hydrophobic

2

residues. Naturally, electrostatic forces, hydrogen bonding, and hydrophobic interactions all

3

drive protein surface-“grown from” polymer interactions. Our data with CT-pOEGMA

4

demonstrate that minimizing hydrophobic interactions between the polymer and the protein

5

surface was important, but pSMA (-) destabilization indicates more than just hydrophobic

6

interactions are important for conformational stability. One of the most exciting aspects of

7

growing polymers from the surface of proteins is our ability to target and tune the properties of

8

the polymer. As we continue to learn about the mechanism through which “grown from”

9

polymers stabilize or destabilize enzymes we will be well positioned to take full advantage of the

10

potential of polymer-based protein engineering.

29


Biomacromolecules

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Page 30 of 36

1 2

Figure 9 Hypothesized effect of polymer conjugation on hydration shell of

3

chymotrypsin. For CT-pSMA(-) and CT-pOEGMA, the polymers interacted with chymotrypsin,

4

displacing water molecules via preferential binding which resulted in a decrease in stability.

5

Conversely, CT-pQA (+) and CT-pCBAm(+) were excluded from chymotrypsin due to

6

unfavorable interactions between polymer and protein, resulting in preferential hydration which

7

increased stability to strongly acidic conditions.

8 9

CONCLUSION

10

In this study, four different chymotrypsin-polymer conjugates were synthesized using surface

11

initiated ATRP polymer-based protein engineering. The four conjugates, CT-pCBAm (+/-), CT30


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Biomacromolecules

1

pOEGMA, CT-pQA (+), and CT-pSMA (-), each had different mucoadhesive, bioactivity, and

2

stability profiles. CT-pQA (+) and CT-pCBAm (+/-) conjugates were mucoadhesive and

3

maintained bioactivity at all pH values tested, whereas CT-pOEGMA and CT-pSMA (-) were

4

not mucoadhesive and had reduced activity. Most importantly, CT-pQA (+) and CT-pCBAm (+/)

5

conjugates stabilized chymotrypsin, whereas CT-pSMA (-) and CT-pOEGMA destabilized the

6

enzyme to the low pH structural denaturation. We hypothesized that the different stabilization

7

properties were due to preferential accumulation of the destabilizing polymers and preferential

8

exclusion of the stabilizing polymers at the enzyme surface. This accumulation and exclusion

9

likely influenced the integrity of the surface hydration layer which led to structural

10

destabilization and stabilization, respectively. We are now in a position to determine whether

11

other enzymes can be stabilized to a low pH environment in a similar manner. Due to their

12

increased stability and maintained activity, CT-pCBAm (+/-) and CT-pQA (+) would be better

13

candidates than CT-pOEGMA or CT-pSMA (-) as an exogenous chymotrypsin enzyme

14

replacement therapy.

15 16 17 18 19 20

31


Biomacromolecules

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Page 32 of 36

1

AUTHOR INFORMATION

2

Corresponding Author

3

*Alan Russell, [email protected]

4

Author Contributions

5

The manuscript was written through contributions of all authors. All authors have given

6

approval to the final version of the manuscript.

7

Funding Sources

8

The authors thank Carnegie Mellon University and the Center for Polymer-based Protein

9

Engineering for funding this work.

10 11 12 13 14 15 16 17 18 19 20 21 22 23 32


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1

Biomacromolecules

Table of Contents Figure

2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 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

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REFERENCES (1) Hong, J.; Gong, P.-J.; Yu, J.-H.; Xu, D.-M.; Sun, H.-W.; Yao, S. J. Mol. Catal. B: Enzym. 2006, 42, 99. (2) Falatach, R.; Li, S.; Sloane, S.; McGlone, C.; Berberich, J. A.; Page, R. C.; Averick, S.; Konkolewicz, D. Polymer 2015, 72, 382. (3) Sandanaraj, B. S.; Vutukuri, D. R.; Simard, J. M.; Klaikherd, A.; Hong, R.; Rotello, V. M.; Thayumanavan, S. J. Am. Chem. Soc. 2005, 127, 10693. (4) Blow, D. M. Biochem. J. 1969, 112, 261. (5) Scheidig, A. J.; Hynes, T. R.; Pelletier, L. A.; Wells, J. A.; Kossiakoff, A. A. Protein Sci. 1997, 6, 1806. (6) Günther, R.; Thust, S.; Hofmann, H.-J.; Bordusa, F. Eur. J. Biochem. 2000, 267, 3496. (7) Wysocka, M.; Lesner, A.; Legowska, A.; Jaskiewicz, A.; Miecznikowska, H.; Rolka, K. Protein Pept. Lett. 2008, 15, 260. (8) Asgeirsson, B.; Bjarnason, J. B. Comp. Biochem. Physiol., Part B: Biochem. Mol. Biol. 1991, 99, 327. (9) Simon, L. M.; Kotormán, M.; Garab, G.; Laczkó, I. Biochem. Biophys. Res. Commun. 2001, 280, 1367. (10) Klibanov, A. M. Nature 2001, 409, 241. (11) Graham , D. Y. N. Engl. J. Med. 1977, 296, 1314. (12) Geokas, M. C.; Conteas, C. N.; Majumdar, A. P. Clin. Geriatr. Med. 1985, 1, 177. (13) Webb, S. Nat. Biotechnol. 2010, 28, 772. (14) Trapnell, B.; Jerkins, T.; Haupt, M.; Chen, S.; Bodhani, A.; Seo, B.; Kapoor, M. Pediatr. Pulm. 2014, 49, 406. (15) Somaraju, U. R.; Solis-Moya, A. Cochrane Db. Syst. Rev. 2014. (16) Lisowska, A.; Kaminska, B.; Grzymislawski, M.; Herzig, K.-H.; Walkowiak, J. J. Cystic Fibrosis 2006, 5, 253. (17) Fieker, A.; Philpott, J.; Armand, M. Clin. Exp. Gastroenterol. 2011, 4, 55. (18) DiPalma, J. A.; Collins, M. S. J. Clin. Gastroenterol. 1989, 11, 290. (19) Montalto, M.; Curigliano, V.; Santoro, L.; Vastola, M.; Cammarota, G.; Manna, R.; Gasbarrini, A.; Gasbarrini, G. World J. Gastroenterol. 2006, 12, 187. (20) Heyman, M. B. Pediatrics 2006, 118, 1279. (21) Watson, L. Clin. Oncol. UK 2010, 22, 391. (22) Lowe, M. E.; Whitcomb, D. C. Gastroenterology 2015, 149, 1678. (23) Borowitz, D.; Goss, C. H.; Limauro, S.; Konstan, M. W.; Blake, K.; Casey, S.; Quittner, A. L.; Murray, F. T. J. Pediatr. 2006, 149, 658. (24) Smart, J. D. Adv. Drug Delivery Rev. 2005, 57, 1556. (25) Davidovich-Pinhas, M.; Bianco-Peled, H. Expert Opin. Drug Delivery 2010, 7, 259. (26) al-Ajlan, A.; Bailey, G. S. Arch. Biochem. Biophys. 1997, 348, 363. (27) Castillo-Yanez, F. J.; Pacheco-Aguilar, R.; Lugo-Sanchez, M. E.; GarciaSanchez, G.; Quintero-Reyes, I. E. Food Chem. 2009, 112, 634. (28) Kurinomaru, T.; Kameda, T.; Shiraki, K. J. Mol. Catal. B: Enzym. 2015, 115, 135. 34


Page 35 of 36

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

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

Biomacromolecules

(29) Xenos, K.; Kyroudis, S.; Anagnostidis, A.; Papastathopoulos, P. Eur. J. Drug Metab. Ph. 1998, 23, 350. (30) Rodriguez, E.; Wood, Z. A.; Karplus, P. A.; Lei, X. G. Arch. Biochem. Biophys. 2000, 382, 105. (31) Abian, O.; Grazú, V.; Hermoso, J.; González, R.; García, J. L.; FernándezLafuente, R.; Guisán, J. M. Appl. Environ. Microbiol. 2004, 70, 1249. (32) Qi, K. K.; Wu, J.; Deng, B.; Li, Y. M.; Xu, Z. W. Animal 2015, 9, 1481. (33) Turner, K. M.; Pasut, G.; Veronese, F. M.; Boyce, A.; Walsh, G. Biotechnol Lett 2011, 33, 617. (34) Zhang, Y.; Cremer, P. S. Curr. Opin. Chem. Biol. 2006, 10, 658. (35) Baldwin, R. L. Biophys. J. 1996, 71, 2056. (36) Yin, L.; Ding, J.; He, C.; Cui, L.; Tang, C.; Yin, C. Biomaterials 2009, 30, 5691. (37) Park, H.; Robinson, J. R. Pharm. Res. 1987, 4, 457. (38) Rodríguez-Martínez, J. A.; Rivera-Rivera, I.; Solá, R. J.; Griebenow, K. Biotechnol Lett 2009, 31, 883. (39) Werle, M.; Bernkop-Schnürch, A. Amino Acids 2006, 30, 351. (40) Wang, W. Int. J. Appl. Pharm. 2000, 203, 1. (41) Ciampolini, M.; Nardi, N. Inorg. Chem. 1966, 5, 41. (42) Kostina, N. Y.; Rodriguez-Emmenegger, C.; Houska, M.; Brynda, E.; Michálek, J. Biomacromolecules 2012, 13, 4164. (43) Cummings, C.; Murata, H.; Koepsel, R.; Russell, A. J. Biomacromolecules 2014, 15, 763. (44) Murata, H.; Cummings, C. S.; Koepsel, R. R.; Russell, A. J. Biomacromolecules 2013, 14, 1919. (45) Averick, S.; Simakova, A.; Park, S.; Konkolewicz, D.; Magenau, A. J. D.; Mehl, R. A.; Matyjaszewski, K. ACS Macro Lett. 2012, 1, 6. (46) Murata, H.; Cummings, C. S.; Koepsel, R. R.; Russell, A. J. Biomacromolecules 2014, 15, 2817. (47) Millard, P.-E.; Mougin, N. C.; Böker, A.; Müller, A. H. E. In Controlled/Living Radical Polymerization: Progress in ATRP; American Chemical Society: 2009; Vol. 1023, p 127. (48) Bowers, K. J.; Chow, D. E.; Xu, H.; Dror, R. O.; Eastwood, M. P.; Gregersen, B. A.; Klepeis, J. L.; Kolossvary, I.; Moraes, M. A.; Sacerdoti, F. D. In SC 2006 Conference, Proceedings of the ACM/IEEE; IEEE: 2006, p 43. (49) Bas, D. C.; Rogers, D. M.; Jensen, J. H. Proteins: Struct., Funct., Bioinf. 2008, 73, 765. (50) Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.; Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. J. Comput. Chem. 2004, 25, 1605. (51) Fuhrmann, G.; Grotzky, A.; Lukić, R.; Matoori, S.; Luciani, P.; Yu, H.; Zhang, B.; Walde, P.; Schlüter, A. D.; Gauthier, M. A.; Leroux, J.-C. Nat. Chem. 2013, 5, 582. (52) Leach, S. A. Nature 1963, 199, 486. (53) Rodríguez-Martínez, J. A.; Solá, R. J.; Castillo, B.; Cintrón-Colón, H. R.; RiveraRivera, I.; Barletta, G.; Griebenow, K. Biotechnol. Bioeng. 2008, 101, 1142. (54) Riccardi, C. M.; Cole, K. S.; Benson, K. R.; Ward, J. R.; Bassett, K. M.; Zhang, Y.; Zore, O. V.; Stromer, B.; Kasi, R. M.; Kumar, C. V. Bioconjugate Chem. 2014, 25, 1501. 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

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24

Page 36 of 36

(55) Lucius, M.; Falatach, R.; McGlone, C.; Makaroff, K.; Danielson, A.; Williams, C.; Nix, J. C.; Konkolewicz, D.; Page, R. C.; Berberich, J. A. Biomacromolecules 2016, 17, 1123. (56) Kijima, T.; Yamamoto, S.; Kise, H. Enzyme Microb. Technol. 1996, 18, 2. (57) Desie, G.; Boens, N.; De Schryver, F. C. Biochemistry 1986, 25, 8301. (58) Fischer, N. O.; McIntosh, C. M.; Simard, J. M.; Rotello, V. M. P. Natl. Acad. Sci. U.S.A. 2002, 99, 5018. (59) Street, T. O.; Bolen, D. W.; Rose, G. D. P. Natl. Acad. Sci. U.S.A. 2006, 103, 13997. (60) Bennion, B. J.; Daggett, V. P. Natl. Acad. Sci. U.S.A. 2004, 101, 6433. (61) Timasheff, S. N. P. Natl. Acad. Sci. U.S.A. 2002, 99, 9721. (62) Mancini, R. J.; Lee, J.; Maynard, H. D. J. Am. Chem. Soc. 2012, 134, 8474. (63) Lawrence, P. B.; Price, J. L. Curr. Opin. Chem. Biol. 2016, 34, 88. (64) Chao, S.-H.; Matthews, S. S.; Paxman, R.; Aksimentiev, A.; Gruebele, M.; Price, J. L. J. Phys. Chem. B 2014, 118, 8388. (65) Lawrence, P. B.; Gavrilov, Y.; Matthews, S. S.; Langlois, M. I.; Shental-Bechor, D.; Greenblatt, H. M.; Pandey, B. K.; Smith, M. S.; Paxman, R.; Torgerson, C. D. J. Am. Chem. Soc. 2014, 136, 17547. (66) Lawrence, P. B.; Billings, W. M.; Miller, M. B.; Pandey, B. K.; Stephens, A. R.; Langlois, M. I.; Price, J. L. ACS Chem. Biol. 2016. (67) 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. Bioconjugate Chem. 2013, 24, 796.

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Design of Stomach Acid-Stable and Mucin-Binding Enzyme Polymer

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