In situ H2O2 Generation by Choline Oxidase and Its Application in Amino Polysaccharide Degradation by Coupling to Lytic Polysaccharide Monooxygenase

Chitin, the most abundant amino polysaccharide in Nature, has many applications in different fields. However, processing of this recalcitrant biopolymer in an environmentally friendly manner remains a major challenge. In this context, lytic polysaccharide monooxygenases (LPMOs) are of interest, as they can act on the most recalcitrant parts of chitin and related insoluble biopolymers such as cellulose. Efficient LPMO catalysis can be achieved by feeding reactions with H2O2, but careful control of H2O2 is required to avoid autocatalytic enzyme inactivation. Herein, we present a coupled enzyme system in which a choline oxidase from Arthrobacter globiformis is employed for controlled in situ generation of H2O2 that fuels LPMO‐catalyzed oxidative degradation of chitin. We show that the rate, stability and extent of the LPMO reaction can be manipulated by varying the amount of choline oxidase and/or its substrate, choline chloride, and that efficient peroxygenase reactions may be achieved using sub‐μM concentrations of the H2O2‐generating enzyme. This coupled system requires only sub‐stoichiometric amounts of the reductant that is needed to keep the LPMO in its active, reduced state. It is conceivable that this enzyme system may be used for bioprocessing of chitin in choline‐based natural deep eutectic solvents.


Introduction
Chitin is a natural amino polysaccharide of β(1!4) coupled Nacetylglucosamine units conferring mechanical stability to fungal cell walls and the exoskeletons of crustaceans and insects. Annually 6-8 million tons of chitin-rich waste shells from crab, shrimp and lobster are produced globally. [1] Chitin, and more so, its partially deacetylated and more soluble derivative chitosan, may find applications in the pharmaceutical, cosmetic, food, biomedical, chemical, and textile industries. [2] Biotechnological valorization of chitin through enzymatic conversion and depolymerization provides an attractive path towards a more sustainable economy, [3] but depends on the economic feasibility of available technologies. Lytic polysaccharide monooxygenases (LPMOs, EC 1.14.99.53-56) are mono-copper enzymes that catalyze oxidative cleavage of glycosidic bonds in recalcitrant polysaccharides such as chitin and cellulose. [4] Since the discovery of the catalytic abilities of these enzymes in 2010, [4a] LPMOs have received considerable attention due to their intriguing catalytic mechanism, [5] their importance for industrial bio-refineries [6] and their possible roles in microbial virulence. [7] LPMOs require an oxygen co-substrate, O 2 or H 2 O 2, and an external electron donor for their catalytic activity. [4a,8] When supplied with H 2 O 2 the enzyme-catalyzed peroxygenase reaction is orders of magnitude faster than reactions happening under "monooxygenase conditions" (i. e., reductant-driven, with O 2 as co-substrate). [8c,9] While high concentrations of H 2 O 2 may lead to autocatalytic enzyme inactivation, [8a,10] several experiments have shown that continuous supply of appropriate lower amounts of H 2 O 2 leads to efficient and stable LPMO reactions without notable enzyme inactivation. [8a,9e,10a,11] Steady supply of H 2 O 2 may be achieved by cascading the LPMO reaction with an (H 2 O 2 -generating) oxidase reaction that can be controlled through the concentration of the oxidase and/or its substrate. Indeed, when the peroxygenase activity of LPMOs was first described, [8a] it was shown that an Aspergillus niger glucose oxidase (AnGOx) can fuel the LPMO reaction in the presence of small amounts of a reductant (e. g., ascorbic acid) that prime the LPMO [LPMOÀ Cu(II)] to yield the catalytically active species LPMOÀ Cu(I). Furthermore, it has been shown for a few other H 2 O 2 -producing fungal flavoenzymes, such as wild-type and engineered cellobiose dehydrogenase, [9e,12] cellooligosaccharide dehydrogenase, [13] and arylÀ alcohol oxidase, [14] that they can drive LPMO reactions.
Fueling LPMO reactions with oxidases acting on glucose or cellobiose may not always be favorable since the use of these enzymes will increase production of oxidized, rather than native, sugars. Furthermore, oxidase substrate levels will vary as saccharification reactions proceed, complicating process control. Thus, other enzymatic systems for driving LPMO reactions are of interest. Choline oxidase (E.C. 1.1.3.17) catalyzes the fourelectron, two-step oxidation of choline to glycine betaine via the intermediate betaine aldehyde, while simultaneously reducing two O 2 molecules to two H 2 O 2 molecules. [15] The enzyme belongs to the glucoseÀ methanolÀ choline (GMC) oxidoreductase superfamily, which also comprises enzymes like glucose oxidase and cellobiose dehydrogenase. Unlike the majority of flavoenzymes in this superfamily, in choline oxidase the flavin adenine dinucleotide (FAD) cofactor is covalently linked to the polypeptide chain. Interestingly, recent studies have shown the feasibility of using a choline oxidase from Arthrobacter nicotianae to drive H 2 O 2 -dependent enzyme-catalyzed reactions in the presence of choline-based natural deep eutectic solvents. [16] Of note, such solvents are also of interest in bioprocessing of chitin and cellulose. [17] In this study, we have assessed the ability of a choline oxidase from the soil bacterium Arthrobacter globiformis (AgChOx) [18] to fuel LPMO-catalyzed polysaccharide degradation, using chitin degradation by an LPMO from Serratia marcescens (SmAA10A, also known as CBP21) as a model system. To the best of our knowledge, no bacterial H 2 O 2 -producing enzyme has been tested or shown to fuel LPMO reactions. Next to a proof-of-concept demonstration that AgChOx indeed can fuel LPMO reactions, we provide an analysis of the interplay between the two enzyme reactions, including dose-response studies for AgChOx, its choline chloride (ChCl) substrate, and the priming reductant (Scheme 1). The results show how LPMO reactions can be fine-tuned and optimized using AgChOx, and provide general insight into features of the LPMO peroxygenase reaction and its potential applications.

Choline oxidase production and catalytic properties
The AgChOx enzyme was recombinantly expressed in Escherichia coli and purified by immobilized-metal ion affinity chromatography. Approximately 50 mg of purified enzyme was obtained from 1 L culture broth ( Figure S1). UV-Vis absorbance spectra of recombinant AgChOx showed maximum peaks at 373 nm and 452 nm, in accordance with previous observations. [19] The FAD cofactor is known to covalently bind to the enzyme during protein production, [20] and the FADloaded enzyme was thus obtained without addition of external cofactor.
The impact of pH and temperature on the efficiency of AgChOx as a H 2 O 2 generation catalyst is depicted in Figures S2  and S3. These data were obtained using the ABTS/HRP (2,2'azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)/horseradish peroxidase) method, because of the favorable high stability of ABTS in a wide range of conditions. [21] The enzyme was most active under neutral to slightly alkaline conditions (pH 7-10; Figure S2) and at ambient temperature (25-30°C; Figure S3). It is known that the first oxidation step catalyzed by choline oxidase involves the transfer of a hydride ion from the alcohol substrate to the enzyme-bound flavin, [19] which is more favorable at alkaline pH. [22] H 2 O 2 production by AgChOx at pH 7.0 was determined using the Amplex Red/HRP assay. [23] Figure 1 shows a linear response between the rate of H 2 O 2 production and the enzyme concentration within the tested range of 0-800 nM AgChOx. The enzyme steadily produced H 2 O 2 and product formation was linear over time, allowing determination of initial rates. From Figure 1, one can derive that, at the substrate concentration used, 1 mM ChCl, the rate of H 2 O 2 formation was 0.72 s À 1 . Deviations from linearity were observed when using longer measurement times or higher AgChOx concentrations. Of note,  excess levels of H 2 O 2 may lead to further oxidation of resorufin to yield less fluorescent compounds such as resazurin. [24] One would expect enzyme dose-dependent production of H 2 O 2 to be accompanied by dose-dependent depletion of O 2 , which was indeed observed ( Figure S4A). The longer incubation times used in measurements with the oxygen sensor showed that inactivation of AgChOx can occur, as one would expect when H 2 O 2 accumulates in the reaction. Importantly, this will not, or to a much lesser extent, happen if the reaction also contains an H 2 O 2 -consuming enzyme system. The formation of H 2 O 2 is potentially limited by the oxygen transfer rate, i. e., the rate for transfer of oxygen from the reaction head space to the reaction mixture, especially in the higher solid matter conditions that apply to the reactions with insoluble chitin described below. Assessment of oxygen transfer to a reaction solution containing chitin in an open stirred vial ( Figure S4B) yielded an initial rate of 8.2 � 0.5 μM O 2 min À 1 . In the experiments described below, the closed reaction tubes had about 1.5 mL of headspace (equivalent to 14 μmol O 2 ) in 2 mL Eppendorf tubes to ensure an excess of oxygen and efficient transfer (note that 14 μmol corresponds to a total of 28 mM of O 2 when transferred to the 0.5 mL liquid reaction volume). The highest chitin oxidation rates observed in the reactions described below amount to some 5 μM min À 1 , when taking into account only soluble products, which represent the majority but not all [10b] of the oxidized products. Thus, it cannot be excluded that oxygen transfer may to some extent have been rate-limiting in the initial phases of the fastest reactions, although the heavy shaking during the reactions described below will have led to oxygen transfer rates that are higher than the 8.2 � 0.5 μM O 2 min À 1 derived from Figure S4B. Figure 2A shows that addition of AgChOx/ChCl drastically increased oxidative chitin degradation by SmAA10A in an AgChOx-dose dependent manner, for concentrations up to 100 nM. Without AgChOx (0 nM in Figure 2A), the reaction was slow, being driven by H 2 O 2 derived from the slow oxidase reaction of SmAA10A and the slow abiotic oxidation of ascorbic acid. With increasing amounts of AgChOx, leading to generation of increasing amounts of H 2 O 2 (Figure 1), the LPMO reactions became faster. Product formation levelled off over time, especially at the higher ChOx concentrations. This may be due to autocatalytic LPMO inactivation, which is expected to increase at higher H 2 O 2 concentrations [8a] and/or depletion of ChCl and/or inactivation of AgChOx, as discussed below.

Oxidative degradation of chitin driven by the AgChOx/ChCl system
At the highest AgChOx concentrations (200 nM, 400 nM and 800 nM), maximum product levels were reached at the first measuring point (30 min). This is typically observed for LPMO reactions with overfeeding of H 2 O 2 , which yield rapid initial product formation accompanied by rapid enzyme inactivation. While considerable product levels were still reached in the reactions with 200 nM and 400 nM the reaction with 800 nM AgChOx barely yielded detectable amounts of product, indicative of severe and almost immediate enzyme inactivation. Although H 2 O 2 production levels depicted in Figure 1 will not be equal to the levels of in situ generated H 2 O 2 in the reactions depicted in Figure 2A, due to variation in the presence of various redox active compounds such as ascorbic acid, the data depicted in Figure 1 do provide some insight into why enzyme inactivation occurs. Extrapolating from Figure 1, a reaction with 800 nM AgChOx would generate 34 μM of H 2 O 2 per minute, which is an excessively high feeding rate for reaction setups like this, with relatively low substrate concentrations. [10a] At these high H 2 O 2 concentrations, non-substrate bound SmAA10A, which will transiently emerge in between catalytic steps, will be highly sensitive for inactivation. In this respect it is worth noting that in the absence of substrate, SmAA10A becomes completely and irreversibly inactivated upon incubation with 20 μM H 2 O 2 for 10 min. [9a] Control reactions, depicted in Figure 2B showed that product formation by AgChOx/ChCl-driven SmAA10A, i. e., product formation beyond what is achieved in reactions with only 0.1 mM ascorbic acid, depends on the presence of ChCl, and ascorbic acid, and the LPMO. These control reactions confirm that a peroxygenase reaction is taking place and that the H 2 O 2 driving this reaction is generated as a result of AgChOx-catalyzed oxidation of ChCl. A control reaction with a 10-fold increased concentration (1 mM) of ascorbic acid and lacking AgChOx, i. e. a typical standard set-up for reductantdriven LPMO reactions, showed slow but steady formation of LPMO products. This reaction reached higher final product concentrations, compared to the reactions with the AgChOx/ ChCl system, which is likely due to a combination of low LPMO inactivation (since in situ generation of H 2 O 2 is slow) and the absence of depletion effects that occur in the AgChOx/ChCl driven reaction. In the latter reaction, substrate depletion effects did occur, as demonstrated by experiments described below. Considering the low temperature optimum of AgChOx (Figure S3), gradual depletion of the oxidase may also have occurred, which would reduce the efficiency of the AgChOx/ ChCl reactions at later time points. Figure 2C shows a clear dependency of chitin oxidation by the AgChOx/ChClÀ SmAA10A system on the ChCl concentration in the tested 0.1 mM to 10 mM range. This is not surprising considering that (1) the K m of AgChOx for ChCl is in the order of 0.5 mM, [18b] and (2) the obtained levels of oxidized products show that considerable amounts of ChCl are consumed as the reaction proceeds. Thus, the system is not saturated with AgChOx substrate and ChCl concentrations will matter. Maximum production of chitobionic acid was obtained in the reactions with 2.5 mM and 5 mM ChCl, which yielded 900-1000 μM soluble product after 25 hours of incubation. Of note, these product levels are similar to those obtained in a standard reaction with 1 mM ascorbic acid ( Figure 2B), but the initial rates of the two reaction types are very different. The reactions with AgChOx, only 100 μM ascorbic acid and 2.5 mM or 5 mM ChCl ( Figure 2C) showed clearly higher production rates for the first 2 h with approximately 600 μM product compared to about 200 μM during standard reaction conditions. The fact that increasing the ChCl concentration from 2.5 to 5 mM did not increase the final production yield may be due to an offset between increased LPMO activity and increased LPMO inactivation in the reaction with 5 mM ChCl. Alternatively, it is possible that under these reaction conditions the maximum attainable levels of oxidation of β-chitin amount to generation of about 1 mM chitobionic acid. Of note, this product level shows that some 2 % of the sugars in the β -chitin have been oxidized; higher levels of oxidation, up to 7 %, have been observed in earlier work, using another type of β-chitin. [4a] Product formation in the reaction with the highest tested ChCl concentration (10 mM), which was hard to assess accurately due to interference of ChCl with chromatographic resolution, showed a pattern typical of LPMO reactions with high levels of H 2 O 2 : a high initial rate combined with rapid enzyme inactivation.
Reduction of the amount of ascorbic acid in reactions with 100 nM AgChOx and 1 mM ChCl from 100 μM to 12.5 μM had a minimal impact on the yield of oxidized products ( Figure 2D). This supports a peroxygenase mechanism, since such a mechanism only requires priming amounts of reductant to drive the reaction. It is worth noting that the concentration of chitobionic acid generated in the experiment with 12.5 μM ascorbic acid was about 300 μM, which means that at least 24 oxidative cleavages were catalyzed per consumed ascorbic acid molecule. This number corresponds well with previously reported turnovers per consumed reductant in reactions with externally supplied H 2 O 2 , which include 18 per ascorbic acid for SmAA10A, [8d] 15 per ascorbic acid for the LPMOs in the commercial cellulase cocktail Cellic CTec2, [8a] and 20 per electron delivered by an enzymatic redox partner, for NcAA9C. [8c] Of note, these numbers will depend on reaction parameters that affect the frequency of LPMO re-oxidation, such as the concentrations of substrate and H 2 O 2 . Figure 2D also shows that the reductant became rate-and yield-limiting at concentrations below 12.5 μM.
Due to the use of largely different reaction conditions, it is difficult to compare the present results with the results of studies in which LPMO catalysis is driven by other enzyme systems. Figure 2C shows that when using 2.5 mM ChCl, the combination of 100 nM AgChOx and 1 μM SmAA10A generate about 1 mM of oxidized products in 25 h. For comparison it has been shown that, at pH 6.0 (and not 7.0 as in this study), one would need about 1 μM of wild-type cellobiose dehydrogenase (CDH) to reach the same product formation by 1 μM of SmAA10A. [12] When using an engineered CDH with drastically enhanced oxidase activity, only about 30 nM of enzyme was needed to reach this product level. [12] Thus it would seem that the AgChOx/ChCl system described herein is much more effective than wild-type CDH and performs similarly to the engineered CDH. Of note, the E. coli expressed AgChOx was used as is, which, based on previous reports on recombinant expression of ChOx, [22] implies that a significant fraction of the enzyme lacked the FAD co-factor. It would thus seem that there is room for further improvement of the AgChOx/ChCl system.
In industrial processes, α-chitin is a more common substrate and therefore activity towards α-chitin was also investigated (Figure 3, with MALDI-TOF MS product identification shown in Figure S5). The results show that the system also works for αchitin and underpin that this substrate is less suitable for SmAA10A compared to β-chitin. [25] With α-chitin, product formation reached a plateau after 120 min, and only 160 μM product from was obtained, compared with 300 μM product generated from β-chitin.

Conclusions
Herein, we present an enzyme cascade system that allows for fine-tuning of in situ generation of H 2 O 2 , using AgChOx acting on choline chloride. As discussed above, system efficiency, i. e., reaction kinetics, LPMO stability, and total yield of oxidized products, can be manipulated by varying various reaction parameters. For applications in biomass processing, further, application-specific optimization will be needed, since LPMO action and optimal supply rates of H 2 O 2 depend on the nature of the substrate, the substrate concentration, the presence of other redox-active compounds, and the interplay with other degradative enzymes (e. g., chitinases or cellulases). [26] Interestingly, solvent engineering plays an important role in current attempts to develop greener routes for biomass processing. [17b,27] For example, deep eutectic solvents (DES) have been used for extracting and partially deacetylating chitin. [17a,28] Choline chloride is the most commonly used salt in DES, and processing scenarios entailing initial chitin extraction with DES followed by dilution in water and AgChOx-fueled LPMOcatalyzed modification or degradation are conceivable. While ChCl concentrations may be high in such scenarios, in situ production of H 2 O 2 may be controlled by varying other parameters, such as the concentration of AgChOx. In this respect, recent studies by Ma et al., showing that the reaction of an unspecific peroxygenase in DES can de driven by choline oxidase activity, are encouraging. [16b] The combined use of the AgChOx/LPMO system and ChCl-containing DES may offer novel opportunities in chitin bioprocessing and valorization.

Experimental Section
Production and purification of choline oxidase (AgChOx). The synthetic codA gene (Accession No. AY304485) from Arthrobacter globiformis [29] was codon-optimized for bacterial expression with an N-terminal His 6 -tag sequence and inserted into a pET22b(+) plasmid, which was electro-transformed into E.coli BL21 DE3. The transformant strain was grown in Luria-Bertani (LB) liquid medium with shaking at 150 rpm or on LB agar plates at 37°C. Media were supplemented with 100 μg·mL À 1 ampicillin. AgChOx expression was induced by addition of isopropyl β-d-1-thiogalactopyranoside (IPTG, final concentration 0.05 mM) at an OD 600 of 0.6, and cultivation was continued for 16 h at 18°C. Cells were harvested by centrifugation (4000 × g, 10 min, 4°C) and stored at À 20°C until further use. The cell pellet was thawed on ice and resuspended in buffer A (20 mM sodium phosphate buffer, 0.5 M sodium chloride, pH 7.5). Then cells were disrupted in a French press at 1.350 bar (TS 0.75, Constant Systems, UK). After centrifugation for 30 min at 30 000 × g, the soluble fraction containing AgChOx was passed through a 0.2 μm sterile filter, before loading on a 1 mL HisTrap FF crude column (GE Life Sciences, USA) installed in an ÄKTA purifier system (Amersham Bioscience, USA) at a flow rate of 3 mL·min À 1 . After washing out unbound protein, using 5 column volumes (CV) of buffer A, the retained protein was eluted by applying a 25 CV linear gradient towards 60 % buffer B (20 mM sodium phosphate buffer, 0.5 M NaCl, 0.5 M imidazole, pH 7.5) at a flow rate of 2 mL·min À 1 . The fractions containing AgChOx were pooled and desalted using a HiPrep 26/10 desalting column (GE Life Sciences, USA) to 20 mM sodium phosphate buffer pH 7.0. The protein concentration was determined by UV-Vis spectroscopy (A 280 ) using the theoretical extinction coefficient calculated using the ExPASy-ProtParam tool. [30] The absorption spectrum of the FAD cofactor was recorded using an Evolution 201 UV-Vis spectrophotometer and a Hellma Suprasil quartz cuvette (Thermo Fisher Scientific, Waltham, MA, USA). A wavelength scan from 300-700 nm of a solution containing purified AgChOx in 20 mM sodium phosphate buffer pH 7.0 was performed at 25°C. The enzyme was then freezedried and stored at À 20°C for later studies.
Characterization of choline oxidase. Choline oxidase activity was determined spectrophotometrically by recording the increase in absorbance at 420 nm resulting from the oxidation of 2,2'-azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) by (H 2 O 2 -fueled) horseradish peroxidase (HRP). [31] HRP was obtained from Sigma-Aldrich. For the determination of pH and temperature optima,  Apparent H 2 O 2 production by AgChOx. Experiments aimed at monitoring the production of H 2 O 2 were based on a previously described method. [23] 90 μL reaction mixtures containing AgChOx, HRP and Amplex Red (AR) in 50 mM sodium phosphate buffer, pH 7.0, were incubated at 30°C for 5 minutes in a 96 well plate. The reactions were initiated by adding 10 μL of a choline chloride (ChCl) solution. The production of H 2 O 2 was determined by monitoring the formation of resorufin at 563 nm using a Varioscan LUX plate reader (Thermo Fisher Scientific, Waltham, MA, USA) for 40 minutes. The H 2 O 2 standard curve was prepared in 50 mM sodium phosphate buffer, pH 7.0. The final concentrations of AgChOx, HRP, AR, and ChCl were 0-800 nM, 5 U·mL À 1 , 100 μM and 1 mM, respectively.
Measurement of oxygen levels. Oxygen consumption by AgChOx was recorded in a 2 mL open vial containing reaction mixtures with AgChOx (100, 200, 800, or 3200 nM), choline chloride (10 mM), and β-chitin (10 g·L À 1 ) in 1 mL 50 mM sodium phosphate buffer, pH 7.0, at 25°C, with magnetic stirring. β-chitin was added to mimic the conditions of reactions with SmAA10A. O 2 uptake was measured using a reaction mixture containing 1 mL degassed 50 mM sodium phosphate buffer, pH 7.0 containing 10 g·L À 1 chitin in a 2 mL open vial, at 25°C. The oxygen meter (Firesting GO2, PyroScience GmbH, Germany) was pre-calibrated via a two-point calibration assuming an oxygen solubility of 258 μM at 25°C for saturated conditions and of 0 μM in a sodium dithionite solution.
Aliquots were taken at various time points, and the reactions were quenched by removing the insoluble substrate using a 96-well filter plate (Millipore, Burlington, MA, USA) operated with a Millipore vacuum manifold. For quantification of soluble LPMO products, the filtrated reaction supernatants were transferred to new tubes and supplemented with 0.5 μM SmCHB, followed by static incubation at 37°C for 24 hours. Treatment with SmCHB converts longer native and oxidized chito-oligosaccharides to N-acetylglucosamine (GlcNAc) and oxidized chitobiose (GlcNAcGlcNAc1A), which simplifies quantification of soluble oxidized reaction products. [32] Quantification of oxidized products. Quantification of chitobionic acid (GlcNAcGlcNAc1 A) was done by chromatography using an RSLC system (Dionex, Sunnyvale, CA, USA) equipped with a 100 × 7.8 mm Rezex RFQ-Fast Acid H + (8 %) (Phenomenex, Torrance, CA, USA) column operated at 85°C. Samples of 8 μL were injected to the column, and sugars were eluted isocratically with 5 mM sulfuric acid as mobile phase and a flow rate of 1 mL·min À 1 . Standards of GlcNAcGlcNAc1A (10-500 μM) were used for quantification. GlcNAcGlcNAc1 A was generated in house by complete oxidation of N-acetyl-chitobiose (Megazyme, Bray, Ireland; 95 % purity) with a chitooligosaccharide oxidase from Fusarium graminearum, [33] as previously described. [32]