Microplate-Based Detection of Lytic Polysaccharide Monooxygenase

Jan 26, 2017 - Most existing methods for screening the activity of lytic polysaccharide mono-oxygenases (LPMOs) on polysaccharides are based on the ...
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Microplate-Based Detection of Lytic Polysaccharide Monooxygenase Activity by Fluorescence-Labeling of Insoluble Oxidized Products Thu V. Vuong,† Bing Liu,‡ Mats Sandgren,‡ and Emma R. Master*,† †

Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto, Ontario M5S 3E5, Canada ‡ Department of Molecular Sciences, Swedish University of Agricultural Sciences, 750 07 Uppsala, Sweden S Supporting Information *

ABSTRACT: Most existing methods for screening the activity of lytic polysaccharide mono-oxygenases (LPMOs) on polysaccharides are based on the detection of soluble oxidized sugars. This approach might underestimate the total performance of LPMOs since oxidation events that do not lead to oligosaccharide release are not detected. Using PcLPMO9D as a model enzyme, a microplate-based method has been developed to detect C1oxidizing LPMO activity by covalently linking a water-soluble fluorophore to oxidized positions within the cellulose fiber. This fluorescence method was validated using X-ray photoelectron spectroscopy and then combined with high-performance anionexchange chromatography to track total PcLPMO9D activity.

1. INTRODUCTION The discovery of lytic polysaccharide monooxygenases (LPMOs)1 and their roles in breaking down biomass including cellulose,2 hemicelluloses,3 and starch4 have significantly advanced working models and methods for lignocellulose/ starch bioconversion.5 Briefly, LPMO action on glycosidic linkages of targeted polysaccharides mainly leads to the oxidation of either the free C1 position or C4 position, producing a lactone or 4-ketoaldose (eventually, aldonic acid and geminal diol in the presence of water), respectively. Since 2010, four new Auxiliary Activities (AA) families, including families 9, 10, 11, and 13, have been created for this enzyme class (www.cazy.org/). Whereas the numbers of protein sequences deposited into these four families are expanding, comparatively few enzymes have been functionally characterized in part due to challenges associated with their production as well as the lack of sensitive and high-throughput screening methods to detect LPMO activity. At present, the most common method to measure LPMO activity is to analyze oxidized, soluble mono- or oligosaccharides by high-performance chromatography, particularly highperformance anion-exchange chromatography (HPAEC). While sensitive, this chromatographic approach typically requires approximately 30 min per run. Synthesis of reference compounds, such as aldonic acid oligosaccharides, can also be costly; synthesis of 4-keto aldose sugars may even be impossible due to tautomerization.6 As LPMOs can also release nonoxidized sugars and 4-keto aldose sugars, methods for determining the reducing ends of sugars, such as 3,5dinitrosalicylic acid or p-hydroxybenzoic acid hydrazide methods might be used; these methods are fast, but they lack © XXXX American Chemical Society

sensitivity. In an attempt to develop a high-throughput method for measuring polysaccharide monooxygenase and glycoside hydrolase activities, polysaccharide hydrogels were embedded with chlorotriazine dyes, where the release of soluble dye resulting from enzyme activity can be measured in 96-well filter plates.7 This method can be optimized for high-throughput screening; however, enzyme accessibility to the targeted polysaccharides may be hindered by the presence of chlorotriazine groups. Another strategy is to increase product solubility by using chitin deacetylase,1 allowing larger oligosaccharides from chitin degradation to be detected. By detecting both soluble neutral and oxidized sugars, the methods above have clearly advanced the characterization of LPMOs; however, since these methods do not detect insoluble products generated by LPMOs, oxidation events that do not lead to solubilization are missed. Oxidation by LPMOs introduces an oxygen atom into aldoses, which can be detected by X-ray photoelectron spectroscopy (XPS).8 The consumption of oxygen during LPMO oxidation, if sufficiently large, can also be measured using an oxygraph system.6 While these methods account for oxidation events that do not lead to soluble products, they are not practical for screening LPMO activity given the need for specialized equipment or low throughput of the technique. Alternatively, the carboxylic acid group introduced by the LPMO could be conjugated with an amine-based fluorophore, thus allowing the detection of insoluble products using a Received: December 2, 2016 Revised: January 10, 2017

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DOI: 10.1021/acs.biomac.6b01790 Biomacromolecules XXXX, XXX, XXX−XXX

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Figure 1. Labeling scheme of PcLPMO9D products. PcLPMO9D cleaves cellulose (1) creating insoluble fragments comprising a carboxyl group at the C1 position (2), which forms an unstable O-acylisourea intermediate (4) in the presence of EDAC (3), allowing the fluorophore ANDA (5) to cross-link to form a fluorescence-labeled product (6).

fluorescence microplate reader (Figure 1). For example, carbodiimides, particularly, water-soluble 1-ethyl-3-[3(dimethylamino)propyl]carbodiimide (EDAC) has been used to couple carboxyl groups of acidic sugars, including gluconic acid, to primary amines at room temperature.9 A fluorescence dye that contain a primary amine, 7-amino-1,3-naphthalenedisulfonic acid (ANDA), is suitable for a screening assay as it is relatively cheap and water-soluble. The predicted pKa of ANDA is 2.7 (as by ACD/I-Lab Web service, https://ilab.acdlabs.com/ ), which even permits a one-step reaction with EDAC. The genome of the wood-degrading white-rot fungus Phanerochaete chrysosporium has at least 11 genes encoding LPMO9 single-module proteins or LPMO9s fused to a carbohydrate-binding module.10 C1-oxidizing LPMO9D from P. chrysosporium strain K-3 (PcLPMO9D, also known as PcGH61D) was overexpressed in Pichia pastoris KM71H.10 When acting on cellulose, PcLPMO9D released soluble, reduced and oxidized cello-oligosaccharides.10 However, the production of insoluble, oxidized products by PcLPMO9D had not been previously investigated. In this study, we developed a fluorescence, microplate-based approach for screening the activity of C1-oxidizing LPMOs on insoluble substrates. Enzymatic reactions of PcLPMO9D on cellulose were conducted in filter-microplates, and the insoluble, oxidized products were labeled with the fluorophore ANDA via EDAC activation. The entire process was incorporated into a semiautomated robotic screening system with the potential for full automation. In combination with detection of solubilized products, this high-throughput approach can provide a complete picture of PcLPMO9D action on polysaccharides.

ethanesulfonic acid (MES) were purchased from Sigma-Aldrich. EDAC was stored as crystal aliquots at −20 °C. Bacterial microcrystalline cellulose (BMCC) was a gift from Prof. David Wilson (Cornell University): BMCC was washed extensively five times on a glass filter with Milli-Q water, resuspended completely by stir-bar mixing for 24 h, and stored in Milli-Q water at 5 mg/mL at 4 °C. Nanocrystalline cellulose was a gift from Prof. Yaman Boluk (University of Alberta). 2.2. Gene Expression and Protein Purification. The cDNA of PcLPMO9D was inserted between BsrXI and NotI restriction sites of the pGAPZαA vector (Invitrogen, USA). Pichia pastoris KM71H (Invitrogen, USA) was transformed with linearized DNA of the constructed expression plasmid by electroporation. Transformants were selected by growing on YPD plates containing 100 μg/mL Zeocin at 30 °C for 2 days. PcLPMO9D was produced using an 8 L bioreactor (Belach Bioteknik AB, Sweden) with 4 L basal salts medium containing100 g/L glucose and 1.2% (v/v) PTM1 trace salts as described in the manufacturer guidelines (Invitrogen, USA). A preculture was prepared by incubating a 100 mL culture in a 500 mL shaking flask at 30 °C, overnight. Batch production of the protein was carried out by incubating at 28 °C for 3 days with a stirring rate of 180−200 rpm to keep the level of dissolved oxygen above 20%. After harvesting the culture broth, Pichia cells were removed by centrifugation at 6000 × g for 30 min, at 25 °C, and the culture supernatant was filtered sequentially through 1, 0.45, and 0.2 μm poly(ether sulfone) membranes (Millipore, USA). Culture supernatant adjusted to 1 M ammonium sulfate was loaded onto a 250 mL PhenylSepharose 6 Fast Flow column (GE Healthcare Biosciences AB, Sweden). After washing the column with 20 mM Tris buffer pH7.5, bound proteins were eluted with a reverse gradient from 1 to 0 M ammonium sulfate in 20 mM Tris buffer pH 7.5. Fractions containing PcLPMO9D were pooled and the buffer was exchanged to 20 mM BisTris buffer pH 6.0 using a 500 mL BioGel P-6DG column (Biorad Laboratories, USA). Deglycosylation was performed by a 2-day incubation at 25 °C after the addition of Endo H at a ratio of 1:200 (w/w, EndoH: PcLPMO9D). The EndoH (endo-β-N acetyl glucosaminidase, EC 3.2.1.96), from Streptomyces plicatus, was a kind gift from Genencor (Palo Alto, USA). The deglycosylated protein was purified by ion-exchange chromatography using a 10 mL Source 30Q column (GE Healthcare Biosciences AB, Sweden) with gradient

2. MATERIALS AND METHODS 2.1. Materials. 1-Ethyl-3-[3-(dimethylamino)propyl]carbodiimide hydrochloride (EDAC), 7-amino-1,3-naphthalene-disulfonic acid monopotassium salt monohydrate (ANDA), and 2-(N-morpholino)B

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Figure 2. Flowchart for analyzing PcLPMO9D-oxidized soluble and insoluble products. Soluble fractions were analyzed by HPAEC, while insoluble fractions were analyzed by XPS and fluorescent labeling with ANDA.

Figure 3. HPAEC analysis of BMCC oxidation by PcLPMO9D. BMCC (black solid line) or incubated with PcLPMO9D, either 100 nM (dash line) or 200 nM (dotted dash line), in the presence of 10 mM ascorbic acid and 50 nM CuSO4 at 40 °C for 16 h were filtered using 10 kDa molecular mass cutoff membranes, and the flow-through was analyzed by HPAEC. The standard (gray solid line) includes glucose and cello-oligosaccharides (degrees of polymerization (DP) of 2−6) as well as their counterparts that were oxidized by GOOX (indicated by subscripted ox). elution from 0 to 0.1 M NaCl in 180 mL of 20 mM BisTis buffer, pH 6.0. Size-exclusion chromatography was applied for final polishing using a 120 mL Superdex 75 column (GE Healthcare Biosciences AB, Sweden). The protein concentration was determined by gel densitometry using bovine serum albumin as the reference. 2.3. Microplate-Based Fluorescence Labeling. BMCC (0.2%) in 96-well filter plates (0.22 μm, PVDF) (Millipore, USA) was vacuum-washed (500 mbar, 30 s) with 50 mM Tris-HCl buffer pH 7.5 using a Tecan liquid handler (Tecan Trading AG, Switzerland) (Figure

S1). PcLPMO9D (100 and 200 nM) was then added, together with 10 mM ascorbic acid and 50 nM CuSO4. The 250-μL reaction was sealed with adhesive films and incubated at 40 °C with 500 rpm vertical shaking in an Eppendorf Thermo mixer for 16 h. The reaction was then vacuum-filtered (500 mbar, 30 s) using a Tecan liquid handler to separate the flow-through and the insoluble fractions. The flow-through was collected into another 96-well microplate for high-performance anion-exchange chromatography, and a sample set of the insoluble fraction was kept for XPS analyses. C

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Biomacromolecules A second sample set of the insoluble fraction was vacuum-washed five times (500 mbar, 120 s) with 100 mM MES pH 4.75, and then 2 mM EDAC (the activating reagent) and 10 mM ANDA (the fluorescence dye), both prepared fresh, were added. The 250-μL reaction was then transferred into an Infinite 200 plate reader (Tecan Trading AG, Switzerland) and incubated in the dark at 25 °C for 4 h with shaking (amplitude of 4 mm and orbital mode). Bound fluorescence fractions were separated by vacuum-filtering (500 mbar, 30 s) using the liquid handler and washed five times with Milli-Q water (500 mbar, 120 s) until no fluorescence background was observed in the flow-through. Bound fractions were suspended in 250 μL Milli-Q water, and fluorescence intensity was measured in black microplates with the λex of 310 nm and the λem of 450 nm using the microplate reader (Tecan Trading AG, Switzerland). In addition to BMCC, using the same procedure, fluorescence labeling was tested with other cellulose preparations, including 0.2% (v/v) Avicel (Avicel pH-101) and nanocrystalline cellulose, 0.4% (v/v) regenerated amorphous cellulose (produced from Sigmacell cellulose Type 2011), and 1.36% (w/v) filter paper (Whatman qualitative filter paper, grade 1). 2.4. High-Performance Anion-Exchange Chromatography. The flow-through fractions collected through the vacuum filtration described above were filtered with 10 kDa molecular mass cutoff membranes, and analyzed by high-performance anion-exchange chromatography (HPAEC), following a previously published method.12,13 Briefly, soluble oxidized sugars were detected using an ICS5000 HPAEC-PAD system (Dionex, USA) with a CarboPac PA1 (2 × 250 mm) analytical column (Dionex, USA). The HPAEC-PAD samples were eluted at 0.25 mL/min using NaOAc gradient (0−1 M) in 0.1 M NaOH. Chromatograms were viewed and analyzed using Chromeleon 7.2 (Dionex, USA). The aldonic acid standard was produced by oxidizing glucose and its oligomers, from cellobiose to cellohexaose, using Sarocladium strictum gluco-oliogosaccharie oxidase (GOOX, family AA7).14,15 2.5. X-ray Photoelectron Spectroscopy. Following PcLPMO9D treatment, the insoluble BMCC fraction was vacuumwashed five times with Milli-Q water using the Tecan liquid handler (Tecan Trading AG, Switzerland). The washed BMCC in the 96-well, filtered microplate was then dried at 37 °C for 16 h; the resulting cellulose discs were then recovered for analysis by XPS using a Thermo Scientific Theta Probe instrument (East Grinstead, UK). Survey spectra were collected with 200 eV pass energy, and highresolution scans were collected with 50 eV pass energy. The C 1s peak at 285 eV was used to calibrate the binding energy scale. Gaussian peak profiles were used for spectral deconvolution of C 1s and O 1s spectra.

Figure 4. Fluorescence detection of PcLPMO9D-oxidized BMCC. After incubation with PcLPMO9D (100 or 200 nM), BMCC was mixed with 2 mM EDAC and 10 mM ANDA, and the fluorescence intensity of derivatized BMCC was measured at λex = 310 nm and λem = 450 nm.

The carboxyl group concentration in a 250-μL reaction of 0.2% chemically pretreated pulp was estimated to be lower than 40 μM.18 Therefore, the amounts of EDAC and ANDA were added at millimolar levels, in great excess, in order to label as many carboxyl groups as possible in minimal time, which is affordable due to the low cost of ANDA. Furthermore, as both chemicals are highly water-soluble, the excess amount can be easily washed away from insoluble cellulose samples. Previous studies have shown that similar efficiencies are achieved through fluorescence-labeling of either insoluble or solventdissolved pulps.19 Therefore, herein the fluorescence dye was added directly to a cellulose suspension, and the labeling reaction was mixed for 4 h, an incubation time much longer than the half-life (a few seconds)17 of the reaction between EDAC and carboxylic acids. 3.3. Cellulose Choice for Fluorescence Labeling. Plantderived cellulose samples, including Avicel, nanocrystalline cellulose, regenerated amorphous cellulose and filter paper, as well as BMCC were first directly labeled with the dye (ANDA) to measure background fluorescence. The presence of both ANDA and EDAC gave higher fluorescence intensity than when ANDA was added alone, indicating that the increase in fluorescence is due to chemical labeling rather than physical binding of the dye to cellulose. All plant-derived cellulose, particularly filter paper, displayed fluorescence intensity at least 3 times higher than that of BMCC (Figure S3). Carboxyl groups in plant-derived cellulose might be introduced at different carbon positions by natural aging and oxidative stress in plants, or during pulping and bleaching processes, or by the presence of hemicellulosic materials (uronic acid moieties). Whatman filter paper no. 1, used in this study, was reported to contain a carboxyl (mainly restricted to uronic acids) concentration of 8 μmol/g.20 Using the same quantification method, the carboxyl content in pulps was reported to be even higher, as it was 18.9 μmol/g in hardwood sulfite pulp and 9.2 μmol/g in prehydrolysis hardwood kraft pulp.18 Due to its low fluorescence background, BMCC was, therefore, used for further experiments. 3.4. BMCC Oxidation by PcLPMO9D. After the incubation of PcLPMO9D with BMCC, the soluble fractions were analyzed by HPAEC, while the insoluble fractions were either labeled with ANDA for fluorescence detection or analyzed by XPS (Figure 2).

3. RESULTS AND DISCUSSION 3.1. PcLPMO9D Purity and Reactivity. The purity and concentration of PcLPMO9D were evaluated by SDS-PAGE, which were loaded with both known concentrations of bovine serum albumin and unknown concentrations of PcLPMO9D (Figure S2). Single bands were seen on SDS-PAGE gels, and migrated close to the position for the expected molecular mass of PcLPMO9D (23 kDa). 3.2. Choice of Fluorescence-Labeling Condition. EDAC was chosen for this study as the activation mechanism is well understood;16 furthermore, the optimization conditions to apply EDAC and ANDA for labeling acidic monosaccharides (including gluconic acid9) as well as carboxymethyl cellulose17 were previously studied. For instance, the amidation of carboxymethyl cellulose with ANDA using EDAC was found most effective at pH 4.75 and when using a high ratio of EDAC/ANDA.17 Accordingly, these conditions were applied for labeling cellulose samples. MES was the buffer choice, as it does not contain either carboxylic groups or primary amine groups. D

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Biomacromolecules Table 1. Elemental Composition and High-Resolution C 1s Peaks of BMCC Samples elemental composition (%) BMCC BMCC + 100 nM PcLPMO9D BMCC + 200 nM PcLPMO9D a

C 1s peak analysis (%)

O

C

O/C ratio

C−C/C−H

C−OH

CO

C−OO

39.5 ± 1.8 39.9 ± 1.8 40.7 ± 1.6

60.5 ± 1.8 60.1 ± 1.8 59.3 ± 1.6

0.65 0.66 0.69

14.8 ± 3.6 13.2 ± 4.8 11.3 ± 3.6

68.0 ± 1.5 69.3 ± 4.1 71.3 ± 1.7

15.2 ± 2.4 15.0 ± 0.7 14.7 ± 1.6

2.0 ± 0.2 2.50 ± 0.03a 2.7 ± 0.1a

One-way ANOVA analyses showed p < 0.05 for these samples, compared with untreated BMCC; errors represent standard deviations; n = 3.

has previously been shown to release soluble sugars from phosphoric acid swollen cellulose, Avicel and cellulose nanofibrils10 along with steam-exploded spruce.21 3.5. Fluorescence-Labeling of PcLPMO9D-Oxidized Sugars. HPAEC indicated that PcLPMO9D oxidized BMCC, releasing soluble sugars. To then account for the production of insoluble oxidized products, the insoluble BMCC fraction was fluorescence-labeled with ANDA. In these experiments, ANDA in the presence of BMCC was used to generate the standard curve for fluorescent measurements. Notably, a linear relation between fluorescence and ANDA concentration was observed in the nanomolar range (Figure S4), facilitating the detection of low carboxyl contents. BMCC and PcLPMO9D-treated BMCC displayed no significant difference in binding ANDA in the absence of EDAC. Labeled samples and free dye exhibited the same fluorescence emission maxima, and the influence of BMCC on fluorescence intensity readings was negligible (Figure S5). Low levels of ANDA were retained by BMCC incubated with EDAC alone, probably due to cross-linking between ANDA and pre-existing carboxyl groups; however, PcLPMO9D treatment increased fluorescence intensity by more than five times (Figure 4). Since fluorescence labeling was restricted to insoluble cellulose products, this analysis directly confirms that PcLPMO9D action introduces carboxyl groups that are retained in the cellulose fiber. The concentration of carboxyl groups introduced to BMCC by PcLPMO9D was less than 0.2 μmol/g, which was an order of magnitude lower than the reported carboxyl concentration presenting in filter paper.20 It is expected, however, that most of existing carboxyl groups in commercial cellulose samples is at the C6 position of glucose instead of at the anomeric carbon.18 Increasing the enzyme dose from 100 nM to 200 nM impacted the amount of soluble sugars more so than the fluorescence intensity of the insoluble fraction. More specifically, doubling the amount of PcLPMO9D nearly doubled the production of soluble products (Figure 3), whereas the fluorescence intensity of the insoluble product increased by only 18% (Figure 4), suggesting that PcLPMO9D might preferentially act upon amorphous cellulose,10 resulting from the enzyme action, leading to more oligosaccharide products. 3.6. XPS Analysis of PcLPMO9D-Oxidized BMCC. XPS analysis of BMCC before and after PcLPMO9D treatment was performed to validate the fluorescent measurements. Signals corresponding to nitrogen were not detected in XPS spectra of PcLPMO9D-treated BMCC (Figure S6), confirming that the protein was removed prior to fluorescence labeling. Instead, signals corresponding to C and O were measured in all XPS spectra at 285 and 532 eV, respectively. The O/C ratio of samples ranged from 0.65 to 0.69 (Table 1), which is higher than those of bleached kraft pulp- derived nanocellulose (0.58)22 and plant-derived, acetone-extracted cellulosic fibers (0.52).23 The theoretical O/C ratio of pure cellulose was

Figure 5. C 1s deconvolution of BMCC and PcLPMO9D-treated BMCC. (A) Untreated BMCC, (B) BMCC treated with 100 nM PcLPMO9D, and (C) BMCC treated with 200 nM PcLPMO9D.

Incubating PcLPMO9D with BMCC generated several peaks that eluted in the region of acidic oligosaccharides (Figure 3). This observation confirmed that PcLPMO9D was active on BMCC, oxidizing and breaking down microcrystalline cellulose to release both neutral and acidic soluble sugars. PcLPMO9D E

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calculated as 0.83;24 accordingly, the BMCC used in this study is relatively pure. The deconvolution of C 1s signals gave four peaks in all samples corresponding to C−C/C−H, C−OH, CO, and C− OO, respectively (Figure 5). Indicative of cellulose oxidation, the relative intensity of the signal corresponding to C−OO functional groups increased following PcLPMO9D treatment. Moreover, similar to the fluorescent labeling experiment, doubling the amount of enzyme did not double the C−O O signal, which increased by only 8% (Table 1). This lower increase, compared with the fluorescence approach, likely reflects the restriction of XPS measurements to sample surfaces.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.6b01790. Liquid handler incorporated with a vacuum system and a plate reader; gel densitometry for mass determination; fluorescence-labeling of different cellulose samples; fluorescence standard curve; fluorescence intensity of BMCC and ANDA; XPS survey spectrum of PcLPMO9D-treated BMCC (PDF)

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REFERENCES

(1) Vaaje-Kolstad, G.; Westereng, B.; Horn, S. J.; Liu, Z.; Zhai, H.; Sorlie, M.; Eijsink, V. G. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 2010, 330 (6001), 219−222. (2) Beeson, W. T.; Vu, V. V.; Span, E. A.; Phillips, C. M.; Marletta, M. A. Cellulose degradation by polysaccharide monooxygenases. Annu. Rev. Biochem. 2015, 84, 923−946. (3) Agger, J. W.; Isaksen, T.; Varnai, A.; Vidal-Melgosa, S.; Willats, W. G. T.; Ludwig, R.; Horn, S. J.; Eijsink, V. G. H.; Westereng, B. Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (17), 6287−6292. (4) Lo Leggio, L.; Simmons, T. J.; Poulsen, J. C.; Frandsen, K. E.; Hemsworth, G. R.; Stringer, M. A.; von Freiesleben, P.; Tovborg, M.; Johansen, K. S.; De Maria, L.; Harris, P. V.; Soong, C. L.; Dupree, P.; Tryfona, T.; Lenfant, N.; Henrissat, B.; Davies, G. J.; Walton, P. H. Structure and boosting activity of a starch-degrading lytic polysaccharide monooxygenase. Nat. Commun. 2015, 6, 5961. (5) Gupta, V. K.; Kubicek, C. P.; Berrin, J. G.; Wilson, D. W.; Couturier, M.; Berlin, A.; Filho, E. X.; Ezeji, T. Fungal enzymes for bio-products from sustainable and waste biomass. Trends Biochem. Sci. 2016, 41 (7), 633−645. (6) Cannella, D.; Mollers, K. B.; Frigaard, N. U.; Jensen, P. E.; Bjerrum, M. J.; Johansen, K. S.; Felby, C. Light-driven oxidation of polysaccharides by photosynthetic pigments and a metalloenzyme. Nat. Commun. 2016, 7, 11134. (7) Kracun, S. K.; Schuckel, J.; Westereng, B.; Thygesen, L. G.; Monrad, R. N.; Eijsink, V. G.; Willats, W. G. A new generation of versatile chromogenic substrates for high-throughput analysis of biomass-degrading enzymes. Biotechnol. Biofuels 2015, 8, 70. (8) Selig, M. J.; Vuong, T. V.; Gudmundsson, M.; Forsberg, Z.; Westereng, B.; Felby, C.; Master, E. R. Modified cellobiohydrolase− cellulose interactions following treatment with lytic polysaccharide monooxygenase CelS2 (ScLPMO10C) observed by QCM-D. Cellulose 2015, 22 (4), 2263−2270. (9) Mechref, Y.; El Rassi, Z. Capillary zone electrophoresis of derivatized acidic monosaccharides. Electrophoresis 1994, 15 (1), 627− 634. (10) Westereng, B.; Ishida, T.; Vaaje-Kolstad, G.; Wu, M.; Eijsink, V. G.; Igarashi, K.; Samejima, M.; Stahlberg, J.; Horn, S. J.; Sandgren, M. The putative endoglucanase PcGH61D from Phanerochaete chrysosporium is a metal-dependent oxidative enzyme that cleaves cellulose. PLoS One 2011, 6 (11), e27807. (11) Foumani, M.; Vuong, T. V.; MacCormick, B.; Master, E. R. Enhanced polysaccharide binding and activity on linear beta-glucans through addition of carbohydrate-binding modules to either terminus of a gluco-oligosaccharide oxidase. PLoS One 2015, 10 (5), e0125398. (12) Vuong, T. V.; Master, E. R. Fusion of a xylan-binding module to gluco-oligosaccharide oxidase increases activity and promotes stable immobilization. PLoS One 2014, 9 (4), e95170. (13) Forsberg, Z.; Vaaje-Kolstad, G.; Westereng, B.; Bunaes, A. C.; Stenstrom, Y.; MacKenzie, A.; Sorlie, M.; Horn, S. J.; Eijsink, V. G. Cleavage of cellulose by a CBM33 protein. Protein Sci. 2011, 20 (9), 1479−1483. (14) Foumani, M.; Vuong, T. V.; Master, E. R. Altered substrate specificity of the gluco-oligosaccharide oxidase from Acremonium strictum. Biotechnol. Bioeng. 2011, 108 (10), 2261−2269. (15) Vuong, T. V.; Vesterinen, A. H.; Foumani, M.; Juvonen, M.; Seppala, J.; Tenkanen, M.; Master, E. R. Xylo- and cellooligosaccharide oxidation by gluco-oligosaccharide oxidase from Sarocladium strictum and variants with reduced substrate inhibition. Biotechnol. Biofuels 2013, 6 (1), 148. (16) Nakajima, N.; Ikada, Y. Mechanism of amide formation by carbodiimide for bioconjugation in aqueous media. Bioconjugate Chem. 1995, 6 (1), 123−130. (17) Madison, S. A.; Carnali, J. O. pH optimization of amidation via carbodiimides. Ind. Eng. Chem. Res. 2013, 52 (38), 13547−13555.

4. CONCLUSION The microplate-based method developed in this study was used to detect the formation of LPMO-oxidized insoluble products, providing a more complete picture of product profile and substrate preference of PcLPMO9D. Using bacterial microcrystalline cellulose with low fluorescence background as the substrate, PcLPMO9D appeared to preferentially oxidize de novo amorphous regions. In addition to screening for new C1oxiding LPMO activities, this fluorescence assay would be useful when testing the roles of other enzymes, such as cellobiose dehydrogenases, in donating electrons to LPMOs, as it does not interfere with soluble oxidized sugars generated by these enzymatic reductants.

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AUTHOR INFORMATION

Corresponding Author

*Address: Department of Chemical Engineering and Applied Chemistry University of Toronto, 200 College Street, Toronto, ON, M3S 3E5. Phone: +1−416-946-7861; Fax: +1−416-9788605; E-mail: [email protected]. ORCID

Thu V. Vuong: 0000-0002-4753-6975 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We would like to thank Profs. David Wilson (Cornell University) and Yaman Boluk (University of Alberta) for sharing BMCC and nanocrystalline cellulose with us, respectively. This study was funded by the Government of Ontario for the project “Forest FAB: Applied Genomics for Functionalized Fibre and Biochemicals” (ORF-RE-05-005), and by the Natural Sciences and Engineering Research Council of Canada for the Strategic Network Grant “Industrial Biocatalysis Network”. F

DOI: 10.1021/acs.biomac.6b01790 Biomacromolecules XXXX, XXX, XXX−XXX

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DOI: 10.1021/acs.biomac.6b01790 Biomacromolecules XXXX, XXX, XXX−XXX