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Polymer Conjugation to Enhance Cellulase Activity and Preserve Thermal and Functional Stability Thaiesha A. Wright, Melissa Lucius Dougherty, Benjamin Schmitz, Kevin M Burridge, Katherine Makaroff, Jamie M. Stewart, Henry D Fischesser, Jerry T Shepherd, Jason A. Berberich, Dominik Konkolewicz, and Richard C. Page Bioconjugate Chem., Just Accepted Manuscript • DOI: 10.1021/acs.bioconjchem.7b00518 • Publication Date (Web): 21 Sep 2017 Downloaded from http://pubs.acs.org on September 23, 2017

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Bioconjugate Chemistry

Polymer Conjugation to Enhance Cellulase Activity and Preserve Thermal and Functional Stability Thaiesha A. Wright, † Melissa Lucius Dougherty,† Benjamin Schmitz,† Kevin M. Burridge, † Katherine Makaroff, † Jamie M. Stewart, † Henry D. Fischesser, † Jerry T. Shepherd, † Jason A. Berberich, § Dominik Konkolewicz,*, † and Richard C. Page*,† †

Department of Chemistry and Biochemistry, 651 E. High Street, Miami University, Oxford, OH

45056 USA §

Department of Chemical, Paper, and Biomedical Engineering, 650 E. High Street, Miami

University, Oxford, OH 45056 USA

*Correspondence: [email protected] (R.C.P.), [email protected] (D.K.)

ACS Paragon Plus Environment

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ABSTRACT

A thermophilic cellulase, FnCel5a, from Fervidobacterium nodosum was conjugated with various functional polymers including cationic, anionic, and strongly and weakly hydrogen bonding polymers. The activity of FnCel5a toward a high molecular weight carboxymethyl cellulose substrate was enhanced by polymer conjugation. Activity enhancements of 50% or greater

observed

for

acrylamide

and

mixed

N,N-dimethyl

acrylamide/2-(N,N-

dimethylamino)ethyl methacrylate polymers suggesting that the greatest enhancements were caused by polymers capable of non-covalent interactions with the substrate. The conjugates were found to have nearly identical thermodynamic stability to the native enzyme as assessed by ∆G, ∆H, and T∆S parameters extracted from differential scanning fluorimetry. Polymers tended to confer comparable tolerance to high concentrations of dimethylformamide, with longer polymers typically enabling higher activity relative to shorter polymers. The new FnCel5a conjugates represent an advance in the production of cellulases which maintain activity at high temperatures or in the presence of denaturing organic solvents.


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INTRODUCTION With the increasing demand for energy, there are several promising options that complement fossil fuels, including biomass conversion.1,2 Biomass conversion is important since it creates liquid fuels, which are crucial for many energy end users.3 Currently, the majority of biofuels are biodiesel from soy, rapeseed and palm oil, or bioethanol fermented from corn-derived starch or sugarcane.3 Although the production of ethanol by fermentation of sugars derived from grain is a mature process,4 the process pulls grains away from other vital needs and exhibits intrinsic inefficiencies in the production process.3 In recognition of this, the Renewable Fuels Standard (RFS) Program caps production of bioethanol from grain-derived starches at approximately 16 billion gallons per year.5 RFS also sets a goal of 16 billion gallons of cellulosic ethanol production per year by 2022, thereby generating significant interest in cellulose as a feedstock for ethanol production to meet energy needs.6 However, a significant challenge in using cellulose derives from the β-1,4-glycosidic linkages in the cellulose backbone, which cannot be hydrolyzed by yeast and many other fermenting organisms.7 Therefore, it is imperative to develop stable and efficient cellulase enzymes that can degrade β-1,4-glycosidic linkages to produce sugars that are fermentable by yeast and other microbes. An additional requirement is the need for the cellulase to tolerate harsh conditions such as high temperatures.8 Herein we describe the production of bioconjugates between polymers and a thermophilic cellulase, FnCel5a, to create an active cellulase with resistance to heat and chemical denaturation. FnCel5a is a thermophilic endo-β-1,4-glucanase from glycosyl hydrolase family 5 with a 36 kDa catalytic domain.9 The cellulase adopts a canonical (β/α)8-barrel fold featuring an α-helix enclosed within a central 8-stranded β-barrel surrounded by eight α-helices.9 Native FnCel5a exhibits activity against carboxymethyl cellulose, regenerated amorphous cellulose, galactomannan, and barley β-


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D-glucan.9,10 FnCel5a activities toward carboxymethyl cellulose and regenerated amorphous cellulose are similar, and herein we utilize carboxymethyl cellulose as the substrate for activity assays with FnCel5a-polymer bioconjugates. The literature has highlighted that appropriate choice of polymer attached to a protein can enhance biological or enzymatic activity, as well as enhance the resistance to an array of challenges such as proteases, chemical denaturants, and extreme temperature.11-21 In the literature there has been significant effort to direct the synthesis of protein-polymer bioconjugates.22-27 Reversible deactivation radical polymerization (RDRP) methods have been used to attach functional polymers to enzymes as a tool to tune the activity and stability of the protein.11,14,15,17,20,28 Reversible addition-fragmentation chain transfer polymerization (RAFT)29 and atom transfer radical polymerization (ATRP)30 are RDRP methods that create well-defined polymers of complex architecture containing a wide variety of functional groups under mild conditions.31-33 This makes RAFT and ATRP well-suited for bioconjugation. There are two strategies for bioconjugation. One approach is to attach an initiating group to the protein surface and to grow the polymer off the protein surface, using a “grafting-from” (GF) approach.32 Alternatively a polymer can be synthesized in solution and subsequently coupled to the protein surface using a “grafting to” (GT) approach.32 RAFT and ATRP are well-suited for the synthesis of the next generation of biocatalysts using both the GF and GT approaches.32 Here, the thermophilic cellulase FnCel5a10 is modified with synthetic polymers using a GT approach to influence and enhance its enzymatic activity and potential for use in biofuels. In the literature, limited studies have been performed on cellulase polymer conjugates,34,35 and to the best of our knowledge, no systematic study of how polymer functionality affects cellulase


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activity and stability has been performed. The work herein creates a library of polymers of comparable molecular weight and varied functionality synthesized by RAFT and grafted to FnCel5a. Interestingly, this work indicates that enzymatic activity can be enhanced by polymer conjugation and increases in stability occur for certain conjugates. The bioconjugates created in this study are summarized in Scheme 1.

A)

C)

B)

Scheme 1. A) Synthesis of poly(DMAm) by RAFT polymerization and B) attachment of poly(DMAm) to the cellulase enzyme. C) Structure of all monomers and co-monomers used in this study. RESULTS AND DISCUSSION A series of hydrophilic polymers were synthesized for conjugation to FnCel5a. The polymers each targeted a degree of polymerization of 30 units, and differ in their ability to engage in hydrogen bonding, i.e. acrylamide (Am) vs. N,N-dimethyl acrylamide (DMAm), with Am being able to engage more effectively with the protein through hydrogen bonds. Additionally, cationizable polymers containing 2-(N,N-dimethylamino)ethyl methacrylate (DMAEMA) or anionizable polymers containing acrylic acid (AA) co-monomers were synthesized to investigate the impact of charge on the properties of the conjugate. The ionizable groups were incorporated


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into polymers of DMAm to introduce potential ionic interactions between the polymer and the protein into the polymer with a smaller propensity to form non-covalent hydrogen bonds between polymer and protein, and thereby more directly probe the impact of the polymer’s ionic groups on the bioconjugate’s performance. Homopolymers targeting 30 units of Am and DMAm were created, as well as copolymers targeting 30 total units with ratios of 4:1 DMAm:DMAEMA and 4:1 DMAm:AA were synthesized. These well-defined polymers were synthesized using RAFT polymerization, as demonstrated for the synthesis of p(DMAm) in Scheme 1 with similar approaches used for the other polymers, adapting strategies outlined in the literature.18,19 Polymer characterization data are in Table 1 and Figure S1. Table 1. Monomer ratios and polymerization results. Monomer Monomer Targeted Measured Sample 1 [M1]:[M2] [M1]:[M2]a 2

Mn-Th

Mn a

Mw/Mn

pAm

Am

-

30:0

33:0

2342

2600

1.03b

pDMAm

DMAm

-

30:0

35:0

3183

3700

1.16c,d

pDMAm/

DMAm

AA

24:6

27:7

3021

3400

1.39c

DMAm

DMAEMA

24:6

34:7

3532

4700

1.31c

AA pDMAm/ DMAEMA a

As determined by NMR.

b

c

As determined from MALDI-TOF mass spectrometry.

As determined by GPC. dMw/Mn determined by MALDI-TOF = 1.09.

Each polymer was conjugated to FnCel5a using an in situ EDC/NHS coupling.19,36 FnCel5a has 26 lysine groups plus one N-terminal amine.10 This is demonstrated in Scheme 1 for the poly(DMAm) system, and the same approach is used for each polymer. The ratios of


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amine:polymer for conjugation are denoted as lower grafting density (L) for the 1:14 amine:polymer system, and higher grafting density (H) for the 1:20 amine:polymer system. For both H and L systems, similar conjugation efficiencies were observed for samples that were allowed to react for 2 h vs 6 h. Polyacrylamide gel electrophoresis (PAGE) of all conjugates studied in these experiments (Figure 1) indicates that all samples contain minimal native protein, less than 5%, assuming that the staining of native enzyme and bioconjugate are similar. This low fraction of unmodified enzyme is likely to have minimal impact on the interpretation of activity and stability measurements. Details of the conjugation efficiency are estimated in Table S1 and Figure S2. The L series has one to three attachments while the H series has two to five attachments. The efficacy of the centrifugation based purification protocol is seen in comparing Figure S2C (after purification) to Figure S2D (before purification), with the UV-Vis spectra of samples before purification dominated by polymer end-group (~310 nm), while after purification the UV-Vis spectra showed a main protein peak (~280 nm) with a small shoulder due to the grafted polymers (~310 nm). The surface area of FnCel5a was estimated to be 133.7 nm2 using the GETAREA method,37 implying that the L series has on average between 0.007 and 0.02 polymers/nm2 of protein, while the H series has on average between 0.02 and 0.04 polymers/nm2. As indicated in Figure S3, there is negligible evidence of aggregation of the bioconjugates as determined by dynamic light scattering (DLS). This shows that the native enzyme and bioconjugates are stable, and the size is consistent with FnCel5a bioconjugate monomers.


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Figure 1. Poly(acrylamide) gel electrophoresis (PAGE) of native FnCel5a and conjugates with Am, DMAm, DMAm/AA and DMAm/DMAEMA polymers. Lanes: 1. Ladder; 2. Native Cellulase; 3. Am-L; 4-DMAm-L; 5-DMAm/AA-L; 6. DMAm/DMAEMA-L;7. Am-H; 8. DMAm-H; 9. DMAm/AA-H; 10. DMAm/DMAEMA-H. The library of FnCel5a-polymer bioconjugates were assessed for enzymatic activity. Carboxymethyl cellulose (CMC), of average molecular weight 200,000, was used as a soluble cellulosic substrate for the enzyme assays, Figure 2A. Activity assays were performed across a range of temperatures and pH values, indicating that peak activity occurs at 80 °C and at pH = 5 for native FnCel5a (Figure S4). The activity assays were conducted using 150 mM CMC, a concentration well in excess of the 0.1 µM Km value of FnCel5a for CMC.9 Literature studies of


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FnCel5a,10 confirm that peak enzymatic activity occurs at 80 °C and at pH = 5, therefore these conditions were chosen for subsequent assays of FnCel5a-polymer conjugates. It is important to note that this temperature and pH is attained by external control of the pH and temperature after synthesis and purification of the bioconjugate. The colorimetric assay detects production of glucose reducing end groups through reaction with 3,5-dinitrosalicyclic acid (DNS).10 Figure 2B shows the relative enzymatic activity of the bioconjugates compared to the native (unmodified) FnCel5a protein. All samples had at least the same activity as the native enzyme, suggesting that conjugation with synthetic polymers did not lead to major steric hindrance between the enzyme’s active site and the 200,000 molecular weight CMC. Remarkably, the bioconjugate typically had enhanced activity compared to the native enzyme. Figure 2B also shows that 6 bioconjugates had statistically significant activity compared to the native at the 5% level, with 4 of these conjugates having statistically significant activity compared to the native at the 1% level, as determined using a two-way Student’s t-test of means. In the uncharged Am and DMAm conjugates, lower grafting density (L) lead to higher activity, while higher grafting density (H) reduces activity. Presumably these effects are due to lower steric repulsions between the conjugate and the substrate for Am-L and DMAm-L conjugates.

OR

A

O

HO RO

O

HO

OR O OR RO j

RO

OR O

OH OR j

OH OR

k

pH = 5, 80 oC

+ OR O

HO RO

OH OR

k R = H or CH 2COOH or CH 2COO-


9


B

200

L-Series H-Series

*** Relative Activity (%)

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150

*

*** ***

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***

*

100

50

0

Figure 2. A) Schematic of FnCel5a hydrolysis of CMC. B) Activity of non-ionic Am and DMAm, and of ionic DMAm/AA and DMAm/DMAEMA based conjugates compared to the native enzyme. (* denotes statistically significant difference in activity between conjugate and native enzyme at the 5% level, and *** denotes statistically significant difference in activity between conjugate and native enzyme at the 1% level) Figure 2B also investigates the enzymatic activity of conjugates of FnCel5a with charged polymers. Figure 2B suggests that polymer charge could be used modulate enzymatic activity. In particular, across the H series the Am-H has an activity of 110±10% of the native, DMAm-H sample has an activity of 127±3% of the native, the DMAm/AA-H has an activity of 112±6% of the native, while DMAm/DMAEMA-H has an activity of 160±10% of the native. The DMAm-


10

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H, DMAm/AA-H, and DMAm/DMAEMA-H polymers are essentially the same except for charge present along the backbone. This is an interesting result, especially given that the DMAm/DMAEMA-L conjugate, with fewer polymer chains, had lower activity than the DMAm/DMAEMA-H conjugate with more conjugated polymer chains, in contrast to the DMAm and Am based materials. This trend in the H materials and the similarity of the Am-L, DMAm-L and DMAm/DMAEMA-L activities suggests that these effects are pronounced at higher grafting densities. A potential explanation for the trend seen in the DMAm-H, DMAm/AA-H, and DMAm/DMAEMA-H materials could be electrostatic effects, particularly at the higher grafting density. At pH=5 many of the AA units should be deprotonated, leading to a net negative charge on the DMAm/AA based conjugates, while the vast majority of the DMAEMA units should be protonated leading to a net positive charge on the DMAm/DMAEMA based conjugates. Since the substrate for the FnCel5a assay is anionic CMC, we hypothesized that the anionic AA containing conjugate would be repelled from CMC, decreasing enzymatic turnover, compared to purely DMAm based conjugates. Conversely, we hypothesized that the anionic CMC would likely be attracted to the conjugates containing cationic DMAEMA residues, causing an increase in turnover seen in the comparison of DMAmH and DMAm/DMAEMA-H conjugates. Evidence of the electrostatic nature of the interactions between enzyme and substrate as well as its bioconjugates is given in Figure S5, which shows activity at different NaCl concentrations. At high salt concentrations, such as 100 mM NaCl, the activity of all conjugates is reduced, presumably due to electrostatic screening due to the high salt concentration. Control experiments in Figure S6 indicate that the polymers alone lead to negligible hydrolytic activity. Further, Figure S7 indicates that mixtures of native enzyme and free


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polymers generally lead to no increase in enzymatic hydrolysis, with the exception of samples containing equivalents of 1:5 enzyme:free DMAm/DMAEMA polymer, which lead to an increase of 27% over native enzyme alone, and the 1:5 enzyme:free DMAm polymer that lead to a 15% increase in activity over the native enzyme alone. These differences in activity of the native enzyme and the native plus 5 equivalents of DMAm or DMAm/DMAEMA polymer were statistically significant at the 5% level, but not the 1% level. This is still much smaller than the H-DMAm/DMAEMA bioconjugate that had an increase in activity of 64% over the native enzyme, and 27% increase for the H-DMAm bioconjugate compared to native. Thus, while an increase in enzyme activity in the presence of free DMAm/DMAEMA polymers was observed, higher increases in activity are gained when the same polymer is conjugated to the FnCel5a. This suggests that the DMAm/DMAEMA polymer may mediate interactions between the substrate and enzyme, however these interactions are enhanced when the polymer is conjugate to FnCel5a. While the extent and nature of these interactions warrant further study, the tools needed to specifically characterize the extent of protein-polymer interactions have not been developed. Therefore, in this study we have focused on characterizing the effect of polymers upon the activity of FnCel5a. In addition to enzymatic activity, the stability of the protein must be assessed. Earlier work in our group has shown that protein-polymer conjugate stability is a complex phenomenon including, but not limited to, thermal stability and chemical stability. Thermal protein stability was assessed through differential scanning fluorimetry (DSF). This method uses SYPRO Orange, a solvatofluorochromic dye with strongly quenched fluorescence in aqueous media. Upon protein unfolding, hydrophobic residues in the protein are exposed, and the dye partitions to the exposed hydrophobic residues resulting in increased fluorescence. Raw DSF fluorescence


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data is given in Figures S8-S11. As seen in Figure S9, free polymers alone showed essentially flat DSF signal, similar to pure buffer. Figure S10 indicates that the DSF trace of protein plus free polymer is essentially the same as the DSF trace of the bioconjugate (Figure S8), indicating that the polymer is not interfering with the DSF signal or providing a polymer specific signal. As outlined in the supporting information and adapting the method outlined in the literature,38 the DSF data was transformed into fraction of unfolded (Punfolded) and folded (Pfolded) protein. This analysis invoked a simple two state model, which was deemed appropriate since the DSF data showed a smooth transition from minimal to maximal fluorescence with no evidence of intermediate folding states. Figure 3 shows the proportion of unfolded protein as a function of temperature, as determined from the DSF data. Activity assays were performed at 80 °C, and at that temperature the fraction of unfolded enzyme is between 9 and 24%, depending on the conjugate, as determined from Figure 3. This slight loss of folded enzyme is offset by the enhanced enzymatic hydrolysis rate at higher temperature. The key features of the transition between folded and unfolded protein are that the native and bioconjugates all have essentially the same Tm values, where Punfolded = Pfolded = 0.5. The Tm of each conjugate varied by less than 1.5 °C from the native enzyme which has a Tm = 84.8 °C. Details are given in Table S2. Each conjugate transitions from an essentially fully folded state at 75 °C to an essentially fully unfolded state at 90 °C. In addition to the proportion of folded and unfolded enzyme, the DSF data enables extrapolation of thermodynamic parameters, as shown in Figure 4, following the method outlined in the literature,38 and highlighted in Supporting Information. In general, a more stable protein will have a standard larger Gibbs free energy of unfolding, ∆uG°. Earlier work,20 showed that when the polymer interacts with the protein surface through non-covalent interactions that a increase in the standard enthalpy of unfolding, ∆uH°, occurs due to the new


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non-covalent bonds that must be broken. Also, the polymer must point away from the protein surface, which could lead to a slight decrease in configurations in the folded state, leading to a slight increase in the standard entropy of unfolding, ∆uS°. In particular, the standard Gibbs free energy of unfolding, ∆uG°, is an important measure of protein stability, since it gives a thermodynamic measure to the protein preference for the folded state. ∆uG° is determined by extrapolation to 298 K. The ∆uG° values for each conjugate are provided in Table 2, ∆uH° and ∆uS° values are provided in Table S2. In general, the thermodynamic parameters of each bioconjugate, ∆uH° and ∆uS°, show less than 5% variation from the parameters of the native. The thermodynamic parameters indicate that conjugation of the studied polymers has negligible impact on the thermodynamic stability of FnCel5a. The key conclusions from the thermal denaturation data are that the FnCel5a-polymer conjugates exhibited similar stability and resistance to unfolding, as evidenced by a similar ∆uG° values (Table 2). Although there are statistically significant differences in activity as determined by the Student’s t-test of means, (Am-H, DMAm-H and DMAm/DMAEMA-L at the 5% level in ∆uG°), these differences in ∆uG° are less than 6% of the native ∆uG°. This suggests that all bioconjugates have practically the same thermal stability as the native.


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Figure 3. Fraction of unfolded enzyme (Punfolded) as a function of temperature, as determined from DSF data.

Table 2. Summary of thermal and functional chemical stability studies.

Sample

Tm (°C)a

∆Tm (°C)

∆uG° (kJ/mol) b

% Relative ∆% Relative Activityc Activity

Native

84.8±0.3

0

67.8±0.7

32 ± 7

0

Am-L

85.0±0.1

0.2

66.2±0.5

38 ± 7

6

Am-H

84.0±0.1

0.2

65.1±0.9

44 ± 7

12

DMAm-L

84.5±0.1

-0.3

68.0±0.7

40 ± 10

8

DMAm-H

84.0±0.1

-0.8

64±0.3

24 ± 2

-8

DMAm/AA-L

84.0±0.1

-0.8

67±1

25 ± 4

-7


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DMAm/AA-H

83.5±0.1

-1.3

68.6±0.7

30 ± 10

-2

DMAm/DMAEMA-L

85.0±0.1

0.2

70.4±0.1

28 ± 5

-4

DMAm/DMAEMA-H 84.8±0.3 0 68.3±0.3 a Melting temperatures (Tm) determined from DSF.

38 ± 4

6

b

Gibbs free energy of unfolding (∆uG°) extracted from differential scanning fluorimetry data. Additional thermodynamic parameters available in Table S2. c

Percent relative activities for native FnCel5a and all conjugates compared to the native before chemical denaturation by DMF.

Figure 4. Plot of experimental ∆uG as a function of temperature calculated from DSF data for all conjugates and native protein. A linear fit was included used to determine the thermodynamic parameters. Inset extrapolated thermodynamic parameters to calculate the standard Gibbs freeenergy of unfolding (∆uG).


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Stability against chemical denaturants, such as polar aprotic solvents, is particularly important for cellulase enzymes since mixtures including the solvents dimethylsulfoxide (DMSO) or dimethylformamide (DMF) are often used to solubilize cellulose to aid its degradation.39 Thus, we also assessed protein stability as resistance to permanent inactivation by chemical denaturation. A functional chemical stability assay was used to determine the enzyme’s and the bioconjugates’ ability to withstand chemical challenges, using DMF as the chemical denaturant. In this functional chemical stability assay, native FnCel5a or its polymer conjugates were incubated at ambient temperature in a medium that consisted of 76 vol% DMF and 24 vol% pH 5 buffer for 2h. Subsequently a DNS assay was run at pH 5 and 80 °C to quantify the extent of enzyme inactivation caused by the chemical denaturant. The data were plotted as enzymatic activity of the native FnCel5a or FnCel5a-polymer conjugate denatured by DMF compared to the activity of the native FnCel5a prior to denaturation. As indicated in Figure 5 there is a significant loss of enzymatic activity caused by DMF denaturant for each system. The key conclusion from Figure 5 is that polymer modification does not lead to substantial differences in the enzyme’s ability to withstand chemical challenges, with all samples displaying statistically insignificant differences in enzymatic activity after exposure to the chemical denaturant, DMF. Additionally, Figure S11 shows the residual activity of each conjugate after exposure to DMF compared to its activity prior to DMF exposure. Despite the statistically insignificant differences in residual activity, the results are encouraging as they show that polymer conjugation does not lead to a loss of stability against a chemical challenge, despite the gains in enzymatic activity.


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Figure 5. Residual activity of native FnCel5a, non-ionic Am and DMAm conjugates, and ionic DMAm/AA and DMAm/DMAEMA conjugates after incubation in 76 vol% DMF, compared to native FnCel5a prior to DMF exposure. CONCLUSIONS In summary, a series of cellulase bioconjugates were synthesized by attaching well-defined functional polymers to FnCel5a, a thermophilic cellulase. The resulting bioconjugates displayed enhanced activity against a carboxymethyl cellulose substrate with the greatest activity occurring when the polymer can attract the substrate through non-covalent interactions. All conjugates had similar thermal stability to the native, and retained activity even after being exposed to chemical denaturants, such as N,N-dimethylformamide. These results suggest that using polymers with functional groups complementary to the enzyme substrate can tune the activity of an enzymepolymer conjugate. Furthermore, this study suggests that increased activity can be achieved without coming at the cost of stability in the face of thermal and chemical challenges.


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EXPERIMENTAL METHODS Materials Dimethylaminoethoxy methacrylate (DMAEMA) was purchased from TCI. Dimethylacrylamide (DMAm), acrylic acid (AA), dinitrosalicylic acid (DNS), and d-DMSO were obtained from Acros Organics. Acrylamide (Am), N-hydroxysuccinimide (NHS), and Dglucose were from Thermo-Fisher. Azobisisobutyronitrile (AIBN), 2-bromopropionic acid, carbon disulfide, and ethanethiol were purchased from Sigma Aldrich. 1-ethyl-3-(3dimethylaminopropyl)

carbodiimide

(EDC)

was

obtained

from

Carbosynth,

Carboxymethylcellulose (CMC, average molecular weight 200,000) was from Millipore Corp and SYPRO Orange was purchased from Life Technologies. All materials were used without further purification unless otherwise specified. SDS-poly(acrylamide) gel electrophoresis was done using Bio-Rad Mini PROTEAN TGX 4-20% gradient gels. The plasmid for FnCel5a was purchased from GenScript. GelCode Blue protein stain was obtained from Thermo Scientific. Cloning, Expression, and Purification of FnCel5a. FnCel5a was cloned into a pET-15b vector to produce a plasmid encoding for a N-terminal His6 tag, a TEV protease cleavage site, followed by residues 11-320 of FnCel5a with a stop codon inserted immediately after residue 320. FnCel5a was expressed in BL21(DE3) E. coli cells grown in Terrific Broth. Cells were harvested and resupsended in 20 mM HEPES, pH 7.5, 150 mM NaCl, 1 mg/mL lysozyme (Sigma), 20 µg/mL DNaseI (MP Biomedicals), and 100 µg/ml 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride (Gold Biotechnology). Resuspended cells were frozen in liquid nitrogen, then thawed overnight at 4°C with rotation. Cell lysate was centrifuged at 17,000 × g for 45 min and the supernatant was filtered through a 0.45 µM filter (Fisher). Filtered lysate was heated for 1 hour at 60 °C to precipitate non-thermostable proteins and centrifuged at 17,000 × g for 15


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min. Following centrifugation, heat-treated lysate was loaded onto a HiTrap nickel affinity column (GE Healthcare) and FnCel5a was eluted in 20 mM HEPES, pH 7.5, 50 mM NaCl, 500 mM imidazole. Eluted FnCel5a was exchanged into 20 mM HEPES, pH 7.6, 50mM NaCl prior to use for polymer conjugation. Purity of FnCel5a was confirmed by polyacrylamide gel electrophoresis. Synthesis of PAETC. The chain transfer agent 2-(((ethylthio)-carbonothioyl)thio)propanoic acid (PAETC) was used in the synthesis of polymers by RAFT polymerization.2 To produce PAETC, potassium hydroxide (14.6 g, 0.26 mol) was dissolved in 15.0 mL of distilled water. This solution was added dropwise to a stirring mixture of ethanethiol (18.6 mL, 0.26 mol) in acetone (150 mL) on an ice bath. Next, carbon disulfide (16.1 mL, 0.27 mol) was added to the mixture and stirred to react for 30 minutes. The reaction mixture was removed from the ice bath and 2-bromopropionic acid (23.0 mL, 0.26 mol) was added dropwise with continued stirring. The reaction was stirred at room temperature overnight after which the solvent was removed by rotary evaporation. The resulting yellow residue was dissolved in 200 mL of ether. The ether layer was washed with seven equivalents (200 mL) of water, followed by one wash with brine (200 mL). Solvent was separated from the product on a rotary evaporator. The resulting product, PAETC (32.7 g, 0.155 mol, 59%) is a viscous yellow liquid which can be frozen to solidify. 1HNMR (300 MHz, CDCl3) δ ppm 4.87 (1H, q, J = 7.4 Hz, CH3CH(S)COOH), 3.38 (2H, q, J = 7.4 Hz, CH3CH2S), 1.63 (3H, d, J = 7.4 Hz, CH3CH(S)COOH), 1.36 (3H, t, J =7.4 Hz, CH3CH2S). RAFT Polymerization of water-soluble polymers. Water-soluble polymers were synthesized by RAFT polymerization with targeted chain lengths of 30 repeat units. Polymerizations proceeded as shown in Scheme 1A. Dimethylacrylamide (1.4903 g, 15.03 mmol), PAETC (0.1082 g, 0.51 mmol), and AIBN (0.0172 g, 0.10 mmol) were dissolved in 2.1 mL of ethanol in


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a glass vial. The contents were transferred to a round bottom flask and deoxygenated using nitrogen gas. The mixture was stirred in a 65 °C oil bath for 21-22 hours. Conversion of monomer was determined using 1H NMR. To precipitate, the polymer solution was added dropwise to hexanes on ice. Polymer was separated from solvent using centrifugation and the resulting product was left in a vacuum oven until dry. Polymers were characterized by 1H NMR on a 500 MHz Bruker instrument. All remaining polymers were synthesized using the same general protocol. Matrix-Assisted Laser Desorption/Ionization Mass Specrtometry (MALDI-MS). Polymers were characterized using MALDI-TOF MS. Target spots were prepared directly on the plate by mixing 0.5 µL of polymer (~100 µM) and 0.5 µL of 2,5-dihydroxybenzoic acid (DHB) matrix and allowing to air dry. Samples were assessed in positive ion linear mode. Poly(Am) and Poly(DMAm) peaks were fitted with a Gaussian distribution to estimate dispersity following an earlier procedure.18 Gel Permeation Chromatography (GPC). Polymers were characterized further using GPC. pDMAm, pDMAm/AA, and pDMAm/DMAEMA samples were each dissolved in approximately 2 mL of DMF with 2 drops of toluene. Polymer solutions were filtered through a 0.2 µm PTFE filter prior to analysis. An Agilent 1260 gel permeation chromatography system equipped an isocratic pump, a degasser, an autosampler, a guard and 2 x Polargel-M columns, and a refractive index detector was used for size exclusion chromatography (SEC) analysis. N,Ndimethylformamide with 0.1 wt% LiBr was used as the eluent with a flow rate of 1mL/min at 50 °C. This SEC system was calibrated using poly(methyl methacrylate) standards in the range of 617,000 to 1,010.


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Conjugation of Polymers to Cellulase. Using a N-hydroxysuccinimide/1-ethyl-3-(3dimethylaminopropyl) carbodiimide (NHS/EDC) coupling reaction, polymers with a reactive carboxylic acid end group from PAETC were attached to amine groups on FnCel5a as shown in Scheme 1C. Conjugation conditions were optimized by varying the ratio of amine:polymer-CTA from 1:3, 1:8, 1:14, and 1:20. Coupling reactions were completed with 1 mg/mL enzyme and 5 mM NHS final concentrations. In a representative reaction, 1:3 polyacrylamide (pAm), pAmCTA (6.2 mg, 2.43 µmol), EDC (0.1 mL of 4.3 mg/mL stock, 2.24 µmol), and NHS (0.1 mL of 5.8 mg/mL stock, 5.04 µmol, 5.04 mM final concentration) were mixed in FnCel5a (0.52 mL of 1.92 mg/mL stock) and 0.28 mL of 0.1 M potassium phosphate buffer, pH 9 for a total volume of 1 mL. This reaction mixture was stirred at room temperature. Samples were taken at 2- and 6hour time points to determine optimal reaction time. Each sample was quenched with 0.2 M glycine buffer (2.5 µL) for a final concentration of 1 mM glycine and stirred to mix. Samples were diluted to 5 mL with buffer (25 mM phosphate, pH 8) in 5 mL 10,000 MWCO PES Centricon tubes and centrifuged in a 4 °C rotor at 6000 rpm for approximately 10 minutes. Filtration was repeated until at least 15 mL of buffer was dialyzed through each sample, and a stable UV-Vis spectrum was obtained. An SDS PAGE gel was used to confirm extent of polymer attachment. UV-Visible Spectroscopy. All UV-Vis analysis was performed using a BioTek Synergy H1 microplate reader. Maximum absorbance values of native cellulase (280nm) and a poly(Am) chain capped with a trithiocarbonate group (306nm) were determined using UV-Visible Spectroscopy. A calibration was created with increasing ratios of protein to polymer (1:1, 1:3, 1:6, 1:9, 1:12, 1:15). However, a shift in maximum absorbance was observed with increasing


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ratios of polymer. This shift was not observed at 270nm, and a calibration was set up involving the ratio of the absorbance at 306 nm to the absorbance at 270 nm. Dynamic Light Scattering (DLS). DLS experiments were performed on a Malvern Instruments Zetasizer nano series at 25 oC in disposable 40 µL cuvettes. Cellulase Activity Assay. Cellulase activity was determined using a colorimetric assay measuring degradation of carboxymethylcellulose as previously described.40 Carboxymethylcellulose (CMC), glucose standards, and conjugate samples were prepared in acetate buffer (100mM sodium acetate, 20 mM calcium chloride, 10% Tween 20, pH 5). Glucose standards (6 µL of 0, 1, 5, 10, 20, 30, 40, 50 µM solutions), enzyme samples (6 µL of 32 nM solutions), and 150 mM CMC substrate were pre-incubated for 5 minutes at 81 °C in a Life Technologies Proflex 3x32-Well PCR System. After incubation, 60 µL of CMC substrate was added to the glucose standards and enzyme samples which were allowed to continue incubating. At each 2.5-minute time point, 30 µL of DNS stop solution (0.4 M 3,5-dinitrosalicylic acid, 0.4 M sodium hydroxide, 0.4 M potassium hydroxide, 1 M potassium sodium tartrate tetrahydrate) was added to the corresponding sample and incubated for 10 minutes at 90 °C. Following the final incubation, samples were cooled on ice for 5 minutes then centrifuged to mix for 10 s. Samples were plated in 40 µL aliquots on a 384-well plate. Absorbance measurements were taken at 530 nm on a BioTek Synergy H1 microplate reader. Thermal Denaturation. The fluorescence of SYPRO Orange dye was measured as a function of temperature to determine the thermal stability of cellulase conjugates. The 5000x SYPRO Orange (Thermo Fisher Scientific, Inc.) stock was diluted to 11.44x in 5 mL of HEPES buffer (20 mM HEPES, 150 mM NaCl, pH 7.8). This solution was used to dilute enzyme samples to 5


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µM final concentration in PCR tubes. Each sample was pipetted in 50 µL aliquots onto a 96-well plate and analyzed in a BioRad CFX96 RT-PCR. DSF curves were collected by measuring the fluorescence of each sample at 570 nm while temperature increased in 0.5 °C increments from 25 to 95 °C with a 5 second equilibrium at each temperature. Functional Chemical Stability. Following chemical denaturation by dimethylformamide (DMF), the colorimetric activity assay previously described was used to determine the functional chemical stability of cellulase-polymer conjugates. Conjugate samples (5 µL of 20 µM stock) were combined with acetate buffer (100 mM sodium acetate, 20 mM calcium chloride, 10% Tween 20, pH 5) and 76 % DMF (v/v%) in PCR tubes. Final volumes were 100 µL per tube with final enzyme concentrations of 1 µM. Solutions were well-mixed then left to sit at room temperature for one hour to incubate. After one hour, samples were diluted to 32 nM and were assayed following the protocol for the colorimetric carboxymethylcellulose assay previously described. Samples for this experiment were analyzed at 0, 6, and 12-minute time points following the same process as the original activity assay. ASSOCIATED CONTENT The Supporting Information is available free of charge on the ACS Publications website at DOI: Details for extracting thermodynamic parameters and polymer characterization data, FnCel5apolymer conjugate SDS-PAGE and UV-Vis analyses, activity assays for polymer-only and nonconjugated polymer control samples, relative activities of FnCel5a-polymer conjugates after DMF exposure, differential scanning calorimetry (PDF)


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AUTHOR INFORMATION Corresponding Author *Email: [email protected] *Email: [email protected] ORCID Richard C. Page: 0000-0002-3006-3171 Dominik Konkolewicz: 0000-0002-3828-5481 Author Contributions T.A.W., M.L.D., J.A.B., D.K., and R.C.P. designed the research. T.A.W., M.L.D., B.S., K.M.B., K.M, J.M.S., H.D.F., and J.T.S. performed the research. M.L.D, T.A.W, J.A.B, D.K, and R.C.P. wrote the manuscript. Notes The authors declare no competing financial interest. ACKNOWLEDGMENT We acknowledge support from Miami University through startup funding (DK) and the Robert H. and Nancy J. Blayney Professorship (RCP). RCP acknowledges support from the US National Science Foundation (Award No. MCB 1552113). We are grateful to Dr. Theresa Ramelot for experimental assistance. ABBREVIATIONS Am, acrylamide; ATRP, atom transfer radical polymerization; CMC, carboxymethyl cellulose; DMAm, N,N-dimethyl acrylamide; DMAEMA, 2-(N,N-dimethylamino)ethyl methacrylate;


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DMF, dimethylformamide; DSF, differential scanning fluorimetry; GF, grafting from; GT, grafting to; RAFT, reversible addition-fragmentation chain transfer polymerization; RDRP, reversible deactivation radical polymerization. REFERENCES (1) de Vries, B. J. M., van Vuuren, D. P., and Hoogwijk, M. M. (2007) Renewable energy sources: Their global potential for the first-half of the 21st century at a global level: An integrated approach. Energy Policy 35, 2590–2610. (2) Panwar, N. L., Kaushik, S. C., and Kothari, S. (2011) Role of renewable energy sources in environmental protection: A review. Renewable and Sustainable Energy Reviews 15, 1513– 1524. (3) Field, C. B., Campbell, J. E., and Lobell, D. B. (2008) Biomass energy: the scale of the potential resource. Trends Ecol. Evol. (Amst.) 23, 65–72. (4) Lin, Y., and Tanaka, S. (2006) Ethanol fermentation from biomass resources: current state and prospects. Appl. Microbiol. Biotechnol. 69, 627–642. (5) Environmental Protection Agency. (2010) Regulation of Fuels and Fuel Additives: Changes to Renewable Fuel Standard Program; Final Rule. Federal Register 75, 14669–14904. (6) Wyman, C. E. (2007) What is (and is not) vital to advancing cellulosic ethanol. Trends Biotechnol. 25, 153–157. (7) Kricka, W., Fitzpatrick, J., and Bond, U. (2014) Metabolic engineering of yeasts by heterologous enzyme production for degradation of cellulose and hemicellulose from biomass: a perspective. Front Microbiol 5, 174. (8) Rastogi, G., Bhalla, A., Adhikari, A., Bischoff, K. M., Hughes, S. R., Christopher, L. P., and Sani, R. K. (2010) Characterization of thermostable cellulases produced by Bacillus and Geobacillus strains. Bioresource Technology 101, 8798–8806. (9) Zheng, B., Yang, W., Zhao, X., Wang, Y., Lou, Z., Rao, Z., and Feng, Y. (2012) Crystal structure of hyperthermophilic endo-β-1,4-glucanase: implications for catalytic mechanism and thermostability. J. Biol. Chem. 287, 8336–8346. (10) Wang, Y., Wang, X., Tang, R., Yu, S., Zheng, B., and Feng, Y. (2010) A novel thermostable cellulase from Fervidobacterium nodosum. J. Mol. Catal. B: Enzym. 66, 294–301. (11) Mancini, R. J., Lee, J., and Maynard, H. D. (2012) Trehalose glycopolymers for stabilization of protein conjugates to environmental stressors. J. Am. Chem. Soc. 134, 8474– 8479. (12) Grover, G. N., and Maynard, H. D. (2010) Protein-polymer conjugates: synthetic approaches by controlled radical polymerizations and interesting applications. Curr Opin Chem Biol 14, 818–827. (13) Pelegri-O'Day, E. M., Lin, E.-W., and Maynard, H. D. (2014) Therapeutic Protein-Polymer Conjugates: Advancing Beyond PEGylation. J. Am. Chem. Soc. 136, 140930092104000–14332. (14) Cummings, C., Murata, H., Koepsel, R., and Russell, A. J. (2013) Tailoring enzyme activity and stability using polymer-based protein engineering. Biomaterials 34, 7437–7443. (15) Cummings, C., Murata, H., Koepsel, R., and Russell, A. J. (2014) Dramatically increased pH and temperature stability of chymotrypsin using dual block polymer-based protein engineering. Biomacromolecules 15, 763–771.


26

Page 27 of 29

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


(16) Murata, H., Cummings, C. S., Koepsel, R. R., and Russell, A. J. (2014) Rational tailoring of substrate and inhibitor affinity via ATRP polymer-based protein engineering. Biomacromolecules 15, 2817–2823. (17) Gauthier, M. A., and Klok, H.-A. (2010) Polymer – protein conjugates: an enzymatic activity perspective. Polym. Chem. 1, 1352–1373. (18) Falatach, R., Li, S., Sloane, S., McGlone, C., Berberich, J. A., Page, R. C., Averick, S., and Konkolewicz, D. (2015) Why synthesize protein–polymer conjugates? The stability and activity of chymotrypsin-polymer bioconjugates synthesized by RAFT. Polymer 72, 382–386. (19) Falatach, R., McGlone, C., Al-Abdul-Wahid, M. S., Averick, S., Page, R. C., Berberich, J. A., and Konkolewicz, D. (2015) The best of both worlds: active enzymes by grafting-to followed by grafting-from a protein. Chem. Commun. 51, 5343–5346. (20) Lucius, M., Falatach, R., McGlone, C., Makaroff, K., Danielson, A. P., Williams, C., Nix, J. C., Konkolewicz, D., Page, R. C., and Berberich, J. A. (2016) Investigating the Impact of Polymer Functional Groups on the Stability and Activity of Lysozyme-Polymer Conjugates. Biomacromolecules 17, 1123–1134. (21) De, P., Li, M., Gondi, S. R., and Sumerlin, B. S. (2008) Temperature-regulated activity of responsive polymer-protein conjugates prepared by grafting-from via RAFT polymerization. J. Am. Chem. Soc. 130, 11288–11289. (22) Boyer, C., Bulmus, V., Davis, T. P., Ladmiral, V., Liu, J., and Perrier, S. (2009) Bioapplications of RAFT polymerization. Chem. Rev. 109, 5402–5436. (23) Boyer, C., Bulmus, V., Liu, J., Davis, T. P., Stenzel, M. H., and Barner-Kowollik, C. (2007) Well-defined protein-polymer conjugates via in situ RAFT polymerization. J. Am. Chem. Soc. 129, 7145–7154. (24) Boyer, C., Stenzel, M. H., and Davis, T. P. (2011) Building nanostructures using RAFT polymerization. J. Polym. Sci., Part A: Polym. Chem. 49, 551–595. (25) Cobo, I., Li, M., Sumerlin, B. S., and Perrier, S. (2015) Smart hybrid materials by conjugation of responsive polymers to biomacromolecules. Nat Mater 14, 143–159. (26) Sumerlin, B. S. (2011) Proteins as Initiators of Controlled Radical Polymerization: Graftingfrom via ATRP and RAFT. ACS Macro Lett. 1, 141–145. (27) Averick, S., Simakova, A., Park, S., Konkolewicz, D., Magenau, A. J. D., Mehl, R. A., and Matyjaszewski, K. (2012) ATRP under Biologically Relevant Conditions: Grafting from a Protein. ACS Macro Lett. 1, 6–10. (28) Murata, H., Cummings, C. S., Koepsel, R. R., and Russell, A. J. (2013) Polymer-based protein engineering can rationally tune enzyme activity, pH-dependence, and stability. Biomacromolecules 14, 1919–1926. (29) Chiefari, J., Chong, Y. K. B., Ercole, F., Krstina, J., Jeffery, J., Le, T. P. T., Mayadunne, R. T. A., Meijs, G. F., Moad, C. L., Moad, G., Rizzardo, E., and Thang, S. H. (1998) Living FreeRadical Polymerization by Reversible Addition−Fragmentation Chain Transfer: The RAFT Process. Macromolecules 31, 5559–5562. (30) Wang, J.-S., and Matyjaszewski, K. (1995) Controlled/“living” radical polymerization. atom transfer radical polymerization in the presence of transition-metal complexes. J. Am. Chem. Soc. 117, 5614–5615. (31) Hill, M. R., Carmean, R. N., and Sumerlin, B. S. (2015) Expanding the Scope of RAFT Polymerization: Recent Advances and New Horizons. Macromolecules 48, 5459–5469. (32) Averick, S., Mehl, R. A., Das, S. R., and Matyjaszewski, K. (2015) Well-defined biohybrids using reversible-deactivation radical polymerization procedures. J Control Release 205, 45–57.


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(33) Matyjaszewski, K. (2012) Atom Transfer Radical Polymerization (ATRP): Current Status and Future Perspectives. Macromolecules 45, 4015–4039. (34) Mackenzie, K. J., and Francis, M. B. (2013) Recyclable thermoresponsive polymer-cellulase bioconjugates for biomass depolymerization. J. Am. Chem. Soc. 135, 293–300. (35) Mackenzie, K. J., and Francis, M. B. (2014) Effects of NIPAm polymer additives on the enzymatic hydrolysis of Avicel and pretreated Miscanthus. Biotechnol. Bioeng. 111, 1792–1800. (36) Fischer, M. J. E. (2010) Amine coupling through EDC/NHS: a practical approach. Methods Mol. Biol. 627, 55–73. (37) Fraczkiewicz, R. (1998) Exact and efficient analytical calculation of the accessible surface areas and their gradients for macromolecules. J Comput Chem. (38) Wright, T. A., Stewart, J. M., Page, R. C., and Konkolewicz, D. (2017) Extraction of Thermodynamic Parameters of Protein Unfolding Using Parallelized Differential Scanning Fluorimetry. J Phys Chem Lett 8, 553–558. (39) Sen, S., Martin, J. D., and Argyropoulos, D. S. (2013) Review of Cellulose NonDerivatizing Solvent Interactions with Emphasis on Activity in Inorganic Molten Salt Hydrates. ACS Sustainable Chem. Eng. 1, 858–870. (40) König, J., Grasser, R., Pikor, H., and Vogel, K. (2002) Determination of xylanase, betaglucanase, and cellulase activity. Anal Bioanal Chem 374, 80–87.


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TABLE OF CONTENTS FIGURE

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Polymer Conjugation to Enhance Cellulase Activity ... - ACS Publications

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