Solubilization of crystalline chitin with a single LPMO: generating chito-oligosaccharides with unprecedented bioactivity

Abstract

Chitin is the second most abundant biopolymer in nature after cellulose and is very recalcitrant. The most valuable chitin-derived products are soluble polymeric chitosan and oligomeric chito-oligosaccharides, the production of which involves environmentally unfriendly processing steps with concentrated acid and alkali. Enzymatic valorization of chitin has not been explored industrially, and enzymatic routes from chitin to soluble oligomeric or polymeric products are lacking. Lytic polysaccharide monooxygenases (LPMOs) contribute to chitin turnover in nature because they promote the activity of chitinases. Here, we have explored whether such LPMOs can be used as stand-alone enzymes to solubilize chitin, yielding, in a single enzymatic step, bioactive oxidized chito-oligosaccharides (oxCHOS). We show that, indeed, chitin can be solubilized using only an LPMO, reaching up to 34% solubilization with a single enzyme treatment and 60% after four consecutive treatments. Importantly, the resulting soluble oxCHOS, with a degree of polymerization of 4–10, showed much stronger immunostimulatory activity on murine macrophages compared to non-oxidized CHOS. We also show that further diversification of the resulting oxCHOS may be achieved by subsequent treatment with chitin deacetylases. These results are poised to reshape current perceptions regarding the valorization of chitin biomass, opening a new green processing route, directly from chitin to products with high bioactivity.

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Green foundation

1. Chitin is abundant and can be valorized in several ways; however, today's valorization methods involve harsh chemical processing steps. This study presents a purely enzymatic method for converting chitin into bioactive chito-oligomers.

2. The novel method, based on innovative enzyme technology, provides a biological route for chitin valorization. Moreover, this method allows the generation of novel types of chitin-derived products that are inaccessible with existing methods and that have promising bioactivity.

3. Direct conversion of chitin to even longer oxidized chitin oligomers and a reduction in enzyme consumption should be pursued through process optimization and enzyme engineering. Additional comparative studies of the functionalities and application potential of the new products generated in this study and currently available, rather different, chitin-derived products should be conducted. As we show, the novel products could be diversified by using chitin deacetylases.

1. Introduction

Chitin is a recalcitrant linear polysaccharide consisting of β-1,4 linked N-acetyl-D-glucosamine (GlcNAc, A) that serves as the carbohydrate backbone of crustacean and insect exoskeletons and also as a structural component in the cell walls of fungi and yeasts.¹ The annual production of chitin in the biosphere is estimated to be approximately 10¹¹ tons, making it the second most abundant biopolymer in nature after cellulose.² Chitin is insoluble in water, acid or alkaline aqueous solutions and conventional organic solvents.³ Due to its crystalline nature, chitin is hard to degrade with enzymes, such as exo- and endo-acting chitinases,⁴ and additional chemical and/or physical pretreatments (concentrated acid, KOH/urea, ionic liquid, high pressure homogenization, and steam explosion) are needed to make this process efficient.⁵ The most common application of chitin relies on its partial chemical deacetylation, yielding chitosan, which is composed of β-1,4 linked GlcNAc and D-glucosamine (GlcN, D).⁶ Polymeric chitosan is soluble in dilute aqueous acid and may be further degraded by chitinases or chitosanases into oligomeric chito-oligosaccharides (CHOS). Both chitosan and CHOS have various applications in the food, medical, personal care and agricultural sectors.^7,8

Importantly, the conversion of chitin to chitosan requires large amounts of concentrated sodium hydroxide, which is environmentally unfriendly. Enzymatic conversion of chitin to (still polymeric) chitosan using chitin deacetylases has been proposed many times, but has not been demonstrated and seems impossible, as discussed previously.⁹ Truly “green” valorization of recalcitrant chitin, such as direct enzymatic conversion into bioactive longer CHOS with interesting properties, has not been achieved.

A major breakthrough was made with the discovery of lytic polysaccharide monooxygenases (LPMOs) by Vaaje-Kolstad et al. in 2010.¹⁰ LPMOs are secreted by aerobic microorganisms and these mono-copper enzymes catalyze oxidative cleavage of glycosidic bonds in crystalline regions of polysaccharides such as chitin and cellulose. By doing so, LPMOs make these recalcitrant polysaccharides more accessible for hydrolytic enzymes by creating new chain ends and amorphous regions.^11–13 LPMOs are classified as auxiliary activities (AA) in the carbohydrate-active enzyme (CAZy) database and to date comprise eight families (AA9–11 and AA13–17).¹⁴ Cellulose-active LPMOs may perform hydroxylation of C1 or C4 of the scissile glycosidic bond, whereas chitin-active LPMOs only act on C1. Such hydroxylation destabilizes the glycosidic bond, which is broken, generating one non-oxidized and one oxidized chain end.¹⁵ Since the same polymer chain may be cleaved twice by an LPMO, these enzymes release soluble products, which, for chitin-active LPMOs, are C1-oxidized CHOS carrying a terminal 2-(acetamido)-2-deoxy-D-gluconic acid (GlcNAc1A).¹⁶ Of note, this oxidation increases the solubility of CHOS, relative to non-oxidized fragments. Furthermore, soluble LPMO-generated products are not cleaved any further by the enzyme, whereas in reactions with chitinases, the enzymes degrade soluble fragments further to yield a product mixture largely consisting of mono- and di-saccharides. Thus, LPMOs have a unique potential to convert chitin into “longer” CHOS.

Originally, LPMOs were thought to catalyze a reductant-dependent monooxygenase reaction (R–H + O₂ + 2e⁻ + 2H⁺ → R–OH + H₂O),¹⁰ which entails that each catalytic cycle requires two externally delivered electrons and one molecule of O₂. An alternative catalytic scenario was proposed by Bissaro et al. who provided evidence for a H₂O₂-driven peroxygenase reaction (R–H + H₂O₂ → R–OH + H₂O) that is much faster than the monooxygenase reaction.¹⁷ It has been demonstrated that apparent monooxygenase reactions are limited by the in situ generation of H₂O₂, resulting from abiotic oxidation of the reductant and the oxidase side activity of the LPMO.^18,19

Both in vivo studies, e.g. using knock-out mutants, and in vitro studies with enzyme cocktails have demonstrated that LPMOs are important for efficient enzymatic solubilization of chitin by chitinases.^20,21 Importantly, so far, the oxidative degradation mediated by LPMOs has been predominantly regarded as ancillary within the complex enzyme systems that are required to solubilize chitin.^4,10 This prevailing perception partly stems from conventional experimental paradigms where only small quantities of soluble oxidized chito-oligosaccharides (oxCHOS) are detectable in chitin degradation reactions with enzyme cocktails. LPMO reactions are difficult to control and auto-catalytic enzyme inactivation is a problem both in reductant-driven reactions and in reactions fueled with externally added H₂O₂. LPMOs tend to be abundant in the secretomes of biomass degrading microorganisms, suggesting their importance. Remarkably, the capacity of LPMOs to directly solubilize solid chitin substrates is unknown.

Chitin is a well-known pathogen-associated molecular pattern (PAMP)^22,23 and chitin fragments are known signalling molecules in various biological systems.^24,25 Plant or animal hosts do not contain chitin and recognize chitin fragments, derived from, for example, pathogenic fungi, as stress signals, eliciting intracellular immune signaling via mechanisms such as the mitogen-associated protein kinase (MAPK) pathways, ultimately preventing pathogen infections.^26–28 Interestingly, chitin-active LPMOs have been associated with microbial virulence and fungal cell wall remodelling^29,30 and, while causal relationships remain unclear, the fact is that oxCHOS are likely to be formed in nature, for example during the interaction between a fungal pathogen and its host. The signalling potential of such oxCHOS has so far not been explored.

In this study, we have explored the feasibility of directly depolymerizing chitin using only a chitin-active LPMO, thus producing oxCHOS. We show that, by tuning the processing conditions, considerable depolymerization of chitin can be achieved. Furthermore, the biological role of the LPMO-generated oxCHOS was investigated, revealing strong immunostimulatory activity. Finally, we show how chitin deacetylases (CDA) can be used to tune the acetylation pattern of the oxCHOS, which would diversify the resulting oxCHOS and may affect their bioactivity. These results are poised to reshape current perceptions regarding the enzymatic conversion of chitin to useful products, while simultaneously expanding our understanding of the biological roles of chitin-derived molecules, in particular oxCHOS. These advancements provide a foundation for developing innovative strategies in green biomass processing and high-value utilization of chitin-rich bioresources.

2. Experimental section

2.1 Materials

Chitin was ball-milled in-house using a PM 200 planetary ball mill (Retsch, Haan, Germany) equipped with a stainless-steel container and zirconium dioxide beads and fractionated to different particle sizes of 75–200 μm, 200–500 μm or 500–800 μm. Chitin with a particle size of 75–200 μm was used for all LPMO reactions except for the comparative experiments with different particle sizes. β-Chitin originating from squid pen (batch: 20140101) was purchased from France Chitin (Orange, France). Colloidal chitin was obtained from β-chitin as previously reported.³¹ Basically, 175 mL of 10 M HCl containing 10 g of chitin was kept at 4 °C for 16 h. After incubation, 1 L of ethanol was added to the mixture to precipitate colloidal chitin, and the suspension was kept at 4 °C for an additional 16 h. The precipitated colloidal chitin was collected by filtering and washed with distilled water. Alkaline- and acid-pretreated shrimp shell α-chitin originating from Pandalus Borealis was purchased from Chitinor AS (Norway). Ascorbic acid was prepared in ddH₂O as 100 mM stock solutions and stored in aliquots for single use, at −20 °C.

Fetal bovine serum (FBS) was purchased from Gibco (Carlsbad, California, California CA, USA). DMEM/high glucose was purchased from Cytiva (Hangzhou, China). Lipopolysaccharides (LPS) and 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2-H-tetrazolium bromide (MTT) were purchased from Solarbio Science & Technology Co. (Beijing, China). Mouse TNF-α ELISA and mouse IL-6 ELISA kits were purchased from Solarbio Science & Technology Co. (Beijing, China). A mixture of native chito-oligosaccharides (CHOS) ((GlcNAc)_1–5) was purchased from Tokyo Chemical Industry Co., Ltd (Tokyo, Japan) and the monomer was removed by precipitating GlcNAc_2–5 with ethanol, followed by lyophilization. Oxidized CHOS (oxCHOS) with a low degree of polymerization (DP) were prepared with chito-oligosaccharide oxidase³² using these precipitated native CHOS as the starting material. Briefly, 5 μM chitooligosaccharide-oxidase (ChitO) was mixed with a CHOS mixture at a final concentration of 20 mg mL⁻¹ in 20 mM Tris-HCl, pH 8.0, and the reaction was incubated at 20 °C for 24 h. Individual oxidized CHOS were obtained in a similar manner by treating CHOS (Megazyme, Ireland) with chito-oligosaccharide oxidase.

All other reagents used in this study were obtained from Sigma-Aldrich.

2.2 Enzyme expression and purification

The chitin-active LPMO SmAA10A from Serratia marcescens (also known as CBP21) was produced as described previously.³³ The enzyme was purified using a column containing chitin resin (New England Biolabs, Inc.) as an affinity matrix with a low-pressure chromatography system. For the purpose of better experimental repeatability, the purified enzyme was incubated with a 3-fold molar excess of CuSO₄ in 20 mM Tris-HCl, pH 8.0, for 30 min at room temperature to produce a copper-saturated version. Subsequently, excess copper was removed by several steps of concentration and dilution in 20 mM Tris-HCl, pH 8.0, using an Amicon Ultra-15 centrifugal filter with 10 kDa MWCO (Merck, Darmstadt, Germany), after which the enzyme was stored in the same buffer, at 4 °C.

A chitin deacetylase from a marine Arthrobacter species (ArCE4A) was produced as described previously.³⁴E. coli strain BL21 Star (DE3) harboring the expression plasmid was used to inoculate 0.5 L TB-medium supplemented with kanamycin (50 μg ml⁻¹) and containing 0.011% Antifoam 204 (Sigma, Steinheim, Germany), followed by incubation at 37 °C in a Harbinger system (LEX-48 Bioreactor, Harbinger Biotech, Markham, Canada). At an OD₆₀₀ of approximately 0.8, isopropyl-D-thiogalactopyranoside (IPTG) was added at a final concentration of 0.2 mM to induce recombinant gene expression, followed by incubation at 28 °C overnight. After harvesting the cells by centrifugation (7000g, 20 min), the cell pellet was resuspended and sonicated (5 s on/10 s off cycles, 10 min, 25% amplitude, on ice). Cell debris was removed by ultracentrifugation (40 000g, 30 min) and the protein was purified by subjecting the clarified lysate to immobilized metal affinity chromatography (IMAC) using a HisTrap FF 5 mL column (Cytiva, Uppsala, Sweden) pre-equilibrated with binding buffer (20 mM MOPS, 20 mM imidazole, 500 mM NaCl, 5% (v/v) glycerol, pH 7.4). After washing, the bound protein was eluted with elution buffer (20 mM MOPS, 500 mM imidazole, 500 mM NaCl, 5% glycerol, pH 7.4). Further purification of ArCE4A was achieved through size exclusion chromatography (SEC) using a HiLoad 16/600 Superdex 75 pg column connected to an ÄKTA Pure 25 system (Cytiva, Uppsala, Sweden). The SEC process used a running buffer of 25 mM Tris-HCl, 150 mM NaCl, at pH 8.0. The solution with purified enzyme was concentrated, with concomitant buffer exchange to 20 mM Tris-HCl, pH 8.0, 100 mM NaCl, using Amicon Ultra-15 centrifugal filters with 10 kDa MWCO (Merck Millipore, Cork, Ireland).

A chitin deacetylase from Aspergillus nidulans FGSC A4 (AnCDA) was produced as reported previously.³⁵ The expression vector was transformed into E. coli TOP 10 cells and the transformant was cultured at 37 °C in 2 × TY medium containing 100 mg of ampicillin per liter until the OD₆₀₀ reached 0.6, after which gene expression was induced by adding 0.02% (w/v; final concentration) arabinose. After overnight incubation at 28 °C, cells were harvested by centrifugation and lysed by sonication, after which the protein from the lysate was purified through IMAC and SEC, following the protocol for ArCE4A purification. Protein concentrations for all enzymes were measured by recording the absorbance at 280 nm and the proteins’ molar extinction coefficients were calculated using the ExPASy-ProtParam tool.³⁶

2.3 Degradation of chitin with CBP21

Chitin degradation was performed in reductant-driven reactions containing 1.0 μM CBP21 and 10 g L⁻¹ chitin, in 20 mM Tris-HCl, pH 8.0, which were started by the addition of ascorbic acid (AscA) to a final concentration of 1 mM.¹⁸ The reaction mixtures were incubated in an Eppendorf ThermoMixer Comfort (Eppendorf, Hamburg, Germany) set to 37 °C with 850 rpm shaking. For the AscA supplementation experiments, AscA was added to a final concentration of 1 mM at intervals of 24 hours. Samples were taken at various time points. Solubilized products were separated from the insoluble fraction by filtration using a MultiScreen™ 96-well filter plate (Merck, Darmstadt, Germany) and the filtrates were stored at −20 °C prior to analysis by HPLC and MALDI-TOF MS (see below). The remaining insoluble chitin was resuspended in ddH₂O and collected by filtration. This washing process was repeated five times and the resulting chitin was dried at 60 °C in a drying oven prior to further experiments.

For quantification of the total amount of soluble oxidized products, 30 μL of the filtrate was treated with 1.0 μM chitobiase from S. marcescens, SmCHB,³⁷ at 37 °C for 16 h, yielding a mixture of the oxidized dimer (GlcNAcGlcNAc1A) and the native monomer (GlcNAc), which were quantified by HPLC as described below, to obtain the concentration of soluble oxidized chito-oligosaccharides produced in the LPMO reactions. The degree of chitin conversion was calculated by summing up the molar concentration of GlcNAc (C_GlcNAc) and twice the molar concentration of GlcNAcGlcNAc1A (C_{GlcNAcGlcNAc1A}), obtaining the total amount of solubilized GlcNAc, which was compared to the total molar amount of GlcNAc added to the reaction in the form of chitin, as summarized in the following equation:

where m_chitin is the mass concentration of the original chitin and 203 is the molar mass of GlcNAc. Of note, this method will also capture soluble non-oxidized chito-oligosaccharides in the reaction solution. From earlier work (e.g. Vaaje-Kolstad et al. in 2010¹⁰) it is known that CBP21 hardly produces non-oxidized soluble products, as also shown by the MALDI-TOF MS spectra of product mixtures included in this study.

The XRD patterns of pre- and post-reaction β-chitin were acquired using a Bruker® AXS D8 Advance diffractometer with copper radiation operating at 40 kV and 40 mA. The scanning measurements were performed applying the radiation λ_Kα = 1.5406 Å with the light scattering angle ranging from 5° to 50° at a 1.5° min⁻¹ scan rate.

2.4 Analysis of reaction products

Soluble reaction products were analyzed by hydrophilic interaction liquid chromatography (HILIC) using a Dionex UltiMate 3000 RSLC setup with a diode array detector (DAD). Chromatography was performed using an Acquity UPLC BEH amide column (2.1 mm × 150 mm) and a BEH amide VanGuard pre-column (2.1 mm × 5 mm), both having a particle size of 1.7 μm (Waters Corp., USA), with a column temperature of 30 °C. The binary mobile phase consisted of 15 mM Tris-HCl buffer, pH 8.0 (A), and pure acetonitrile (B), applied at a flow rate of 0.4 mL min⁻¹. The elution gradient started at 26% eluent A and 74% eluent B, held for 5 min, followed by a 2 min linear gradient to 62% B, held for 1 min. Column reconditioning was performed with a 2 min gradient back to starting conditions, which were then maintained for 2 min. All chromatograms were recorded using Chromeleon 7.0 software. In all figures, A_nox refers to (GlcNAc)_n−1GlcNAc1A, i.e., the C1-oxidized form (in equilibrium between lactone and aldonic acid) of chito-oligosaccharides composed of n glycosyl units.

Chromatographic analysis of the oxidized dimer (GlcNAcGlcNAc1A) and the monomer (GlcNAc) in chitobiase-treated samples was performed using a 100 × 7.8 mm Rezex RFQ-Fast Acid H + (8%) (Phenomenex, Torrance, CA, USA) column operated at 85 °C in an RSLC system (Dionex, Sunnyvale, CA, USA). Isocratic elution was achieved using 5 mM sulfuric acid with a flow rate of 1 mL min⁻¹, and analytes were monitored by measuring absorbance at 195 nm. Quantification was performed by creating a standard curve (25–1600 μM) for the oxidized dimer (GlcNAcGlcNAc1A) and a standard curve (25–1600 μM) for the native monomer (GlcNAc). The oxidized standards were created in-house by incubating N-acetyl-chitobiose (Megazyme, Bray, Ireland, 95% purity) with a chito-oligosaccharide oxidase from Fusarium graminearum, as described previously.^32,37

Mass spectrometry (MS) analysis of soluble reaction products was performed using an Ultraflex™ TOF/TOF mass spectrometer (Bruker Daltonics GmbH, Bremen, Germany) as described previously.¹⁰ In short, 1 μL of LPMO reaction filtrates was mixed with 2 μL of matrix solution (15 mg mL⁻¹ 2,5-dihydroxybenzoic acid) and spotted on a 384-Spot MALDI Plate, followed by drying under a stream of air. MALDI-TOF MS spectra were obtained from m/z 150 to 2500 with an acceleration voltage of 25 kV, a reflector voltage of 26, and pulsed ion extraction of 40 ns in the positive ion mode. Peak lists were generated from the MS spectra using Bruker FlexAnalysis software.

2.5 Immunostimulatory activity of oxidized chito-oligosaccharides

The mouse macrophage RAW 264.7 cell line was purchased from the Type Culture Collection of the Chinese Academy of Sciences (Shanghai, China). The cells were cultured in DMEM/high glucose supplemented with 10% fetal bovine serum and maintained under a humidified atmosphere with 5% CO₂ at 37 °C.

The viability of RAW264.7 cells exposed to oxCHOS was measured using the MTT assay as described previously.³⁸ Briefly, cells in the logarithmic growth phase were adjusted to a density of 1 × 10⁶ cells per mL and seeded into 96-well plates (100 μL per well), cultured overnight, and then exposed to different concentrations of oxCHOS for 24 h. Afterwards, the medium was removed and 100 μL MTT solution (0.5 mg mL⁻¹ in medium) was added to each well, followed by incubation for another 4 h at 37 °C. After removing the MTT solution, the formazan crystals (reflecting the number of viable cells that are capable of reducing MTT to its insoluble formazan) were dissolved in 150 μL DMSO. The absorbance of the solutions in each well was recorded at 490 nm using a microplate reader. The experiment included three well replicates (independent biological repeats) for each dosage of oxCHOS.

The experimental design for testing immunostimulatory activity was as follows: CK group (negative control, treated with only DMEM/high glucose), LPS group (positive control, treated with 1 μg ml⁻¹ LPS in DMEM/high glucose), and experimental group (treated with oxCHOS at different concentrations in DMEM/high glucose). Three reactions were set up for each group (independent biological repeats). The cells were inoculated to a density of 1 × 10⁶ cells per mL and seeded into 96-well plates (100 μL per well), followed by culturing for 12 h. Then, the medium was removed and 100 μL oxCHOS solution, fresh DMEM/high glucose, or LPS solution was added to each well according to the set groups, followed by culturing for another 24 h. After this incubation, the medium was collected for the determination of NO and inflammatory cytokines.

For determination of the NO content, the cell culture supernatant was mixed 1 : 1 with the Griess reagent.³⁹ The Griess reagent was prepared as follows: Griess A (100 mg sulfanilamide dissolved in 10 mL of 5% phosphoric acid solution) and Griess B (10 mg N-(1-naphthyl) ethylenediamine dihydrochloride dissolved in 10 mL of ultrapure water). The working reagent was freshly prepared by mixing Griess A and B solutions at a 1 : 1 ratio immediately before use. Cell culture supernatants were mixed with the prepared Griess reagent in a volume ratio of 1 : 1, followed by incubation in the dark for 10 min at room temperature, after which the absorbance at 540 nm was measured with a microplate reader, and the NO content was calculated according to an NaNO₂ standard curve. Enzyme-linked immunosorbent assays (ELISA) were used to detect the inflammatory cytokines TNF-α and IL-6 using mouse-specific ELISA kits, according to the manufacturer's instructions.

The experimental data were analyzed using SPSS Statistics (version 27.0, IBM Corp., Armonk, NY, USA) and are presented as mean ± standard deviation. Statistical comparisons were performed through one-way analysis of variance (ANOVA), followed by Duncan's multiple range test. Asterisks show statistically significant differences between the experimental groups (*p < 0.05, ** p < 0.01, *** p < 0.001) and “ns” indicates “not significant”.

2.6 Enzymatic deacetylation of oxidized chito-oligosaccharides

Reaction mixtures for deacetylation contained 1.0 mg ml⁻¹ substrate, 10 μM CoCl₂ and 1 μM enzyme in 50 mM Tris-HCl, pH 8.0. The reaction mixtures were incubated at 37 °C, in a shaker at 225 rpm for 15 min, 30 min, 1 h, 2 h, 4 h, 8 h, and 24 h. Reactions were quenched by heating for 10 min at 95 °C, followed by filtration using a MultiScreen™ 96-well filter plate. The resulting solutions were used for the determination of released acetate. Samples (1.2 mL per sample) to be used for NMR characterization were lyophilized. Alternatively, as indicated in the text, reactions for which products were only analyzed with MALDI-TOF MS were either not quenched at all or quenched by adding acetonitrile to reach 50% (v/v).

Quantification of released acetate was done by ion chromatography using an RSLC system (Dionex, Sunnyvale, CA, USA) equipped with a Dionex IonPac AS11 organic acid column, using the following gradient: 0–8 min, 1 mM KOH; 8–9 min, from 1 to 60 mM KOH; 9–16 min, 60 mM KOH; 16–16.1 min, from 60 to 1 mM KOH; 16.1–22 min, 1 mM KOH. The flow rate was 0.375 ml min⁻¹. The amount of released acetate was quantified using acetic acid (glacial, anhydrous (Merck, Darmstadt, Germany)) as the standard. Operation of the chromatographic system and processing of chromatograms were performed using the Chromeleon 7 software (Dionex Corp.).

2.7 Characterization of oxidized chito-oligosaccharides after deacetylation

Lyophilized oxCHOS before and after enzymatic deacetylation (1.2 mL reaction mixture was lyophilized) were dissolved in 600 μL D₂O. All NMR experiments were performed with a Bruker Avance 400 MHz spectrometer equipped with a 5 mm BBI probe. NMR spectra were recorded using a zg30 pulse sequence at 25 °C and analyzed using TopSpin 3.2 software. One-dimensional ¹H NMR spectra were recorded using a 90° pulse of 8.55 μs with solvent suppression provided by excitation sculpting.⁴⁰ Spectra were recorded using a spectral window of 8012 Hz and a relaxation delay of 2.0 s, with 256 scans.

MALDI-TOF MS analysis of the products of deacetylation reactions was performed as described above.

3. Results and discussion

3.1 One-enzyme solubilization of crystalline chitin

To evaluate the one-enzyme conversion of chitin, we first conducted standard reductant-driven LPMO reactions using CBP21 and three chitin forms. Reactions with α-chitin (Fig. 1A, Fig. S1) and β-chitin (Fig. 1C, Fig. S1) showed a predominant production of soluble oxidized products with a degree of polymerization (DP) of DP4 to DP8, consistent with previous reports.⁴¹ Notably, β-chitin demonstrated superior degradability compared to α-chitin under identical conditions (Fig. 1G and Fig. S1), although both exhibited relatively low conversion (1.4% for α-chitin vs. 7.3% for β-chitin at 48 h). This substrate preference aligns with reported structural differences between the two chitin allomorphs: α-chitin has a more compact structure than β-chitin⁴² and is generally observed to be less susceptible to enzymatic degradation. When testing colloidal chitin, a more amorphous substrate commonly used in chitinase reactions, we observed products of similar DP distribution, but with a relative increase in odd-numbered oligosaccharides (Fig. 1E and Fig. S1), as one would expect as the substrate becomes more amorphous.¹⁰ The conversion after 48 h reached 12.5% (Fig. 1G), representing a 1.7-fold improvement over crystalline β-chitin.

Fig. 1 Product formation and polymer solubilization in LPMO reactions with different chitin materials and reaction conditions. Panels A, C and E show standard reactions containing 1 μM CBP21, 10 g L⁻¹ of α-chitin, β-chitin or colloidal chitin, respectively, and 1 mM ascorbic acid (AscA) in 20 mM Tris-HCl, pH 8.0. Panels B, D and F show the same reactions, supplemented with 1 mM AscA at 24 h, as indicated by the arrow. All reactions were incubated at 37 °C, with shaking at 850 rpm. The levels of individually oxidized products are quantified as peak areas (from HPLC chromatograms). In panels A–F, products are named Anox, where A represents a GlcNAc unit. Anox refers to (GlcNAc)_n−1GlcNAc1A, i.e., the C1-oxidized form of chito-oligosaccharides composed of n glycosyl units (GlcNAc1A stands for GlcNAc oxidized at C1). Panel G shows the total conversion (solubilization) of chitin for each of the six reactions after 48 h, which was determined after conversion of all products into a mixture of GlcNAc and the oxidized dimer, which were quantified. Panel H shows conversion of β-chitin with repeated additions of 1 mM AscA, as indicated. The values shown are the average of three independent experiments with standard deviations shown as error bars.

The nonlinear progression of LPMO reactions suggests potential enzyme inactivation through oxidative damage or reactant depletion (e.g., reductant consumption). Supplementation of the reactions with another 1 mM of ascorbic acid at 24 h led to increased product formation (Fig. 1B, D and F), showing that, under these reaction conditions, the enzyme was not fully inactivated after 24 h and that reductant depletion limited the reaction. This approach enhanced chitin solubilization across all substrates, by 5.6-fold, 2.7-fold and 1.6-fold for α-chitin (7.7% conversion), β-chitin (19.8% conversion) and colloidal β-chitin (18.6% conversion), respectively (Fig. 1G). For two of the substrates, the increase in chitin solubilization was well above a factor of two. This likely relates to an initial delay in the release of soluble products, since such release requires multiple cleavages that are close to each other and in the same polysaccharide chain. It is also possible that this phenomenon reflects heterogeneity in the substrate, with the outer layers being less easily solubilized than the inner layers. It is also worth noting that while β-chitin was still more efficiently solubilized than α-chitin, β-chitin and colloidal β-chitin were solubilized with similar efficiency in the experiments with additional ascorbic acid supplementation. This latter observation may also result from variation in the outer and inner layers of the substrates. It is conceivable that the colloidal particles have a more accessible outer surface, hence the more rapid conversion during the early phase of the reaction, whereas the accessibility of the underlying polysaccharide chains is similar in both β-chitin forms.

Extended time-course experiments with β-chitin and multiple 24-hour AscA additions (Fig. 1H) demonstrated sustained LPMO activity up to 72 h, achieving nearly 30% total conversion. However, supplementation of AscA beyond 72 h did not further increase product yields, suggesting that the enzyme had become inactive.

To better understand the factors limiting the reaction, we then conducted a series of experiments with substrate concentrations varying from 0.5 to 50 g L⁻¹. These reactions were run for 72 h with the addition of 1 mM AscA at 0, 24 and 48 hours (Fig. 2) or without the addition of extra AscA (Fig. S2). Fig. 2A and S2A show that neither the substrate concentration nor the supplementation with AscA had major effects on the product profile, showing that the LPMO interacts with the substrate in the same manner, in all reactions.

Fig. 2 Generation of soluble products in LPMO reactions with different chitin concentrations. The reactions contained 1 μM CBP21, 1 mM AscA, and different concentrations (0.5, 1, 2.5, 5, 10, 20, 30, 40, and 50 g L⁻¹) of β-chitin in 20 mM Tris-HCl, pH 8.0, and were incubated at 37 °C, with shaking at 850 rpm. Fresh AscA to a final concentration of 1 mM was added at 24 h and 48 h (see Fig. S2 for a control experiment without extra additions of AscA). Panel A shows analysis of the formation of each individually oxidized chito-oligosaccharide at 72 h. Anox stands for (GlcNAc)_n−1GlcNAc1A, i.e., the C1-oxidized form of chito-oligosaccharides composed of n glycosyl units. Panels B and C show a quantitative analysis of the formation of soluble products, which was conducted after treating these products with SmCHB. The values shown are the average of three independent experiments with standard deviations shown as error bars. Panel D was generated using PyMOL 3.1 and ChemDraw 22.0.0 and illustrates the various productive and non-productive reactions that are at play and their substrate dependency. The active site, which is visualized in the lower half of the panel, is located near the enzyme surface, permitting interaction with insoluble chitin.

In the control experiment without repetitive addition of AscA, close to maximum product levels were reached after 24 h for all substrate concentrations, and these levels did not increase at higher substrate concentrations (Fig. S2). This indicates that the reactions were limited by depletion of AscA, which may be due to the productive use of AscA, implying that the H₂O₂ resulting from oxidation of AscA leads to chitin cleavage, but may also reflect inactivation of the LPMO. Inactivation of the LPMO, which results from non-productive reactions with H₂O₂ and which is promoted at low substrate concentrations, leads to the release of copper, which will drastically speed up the abiotic oxidation of AscA, leading to even more H₂O₂ and even faster enzyme inactivation.^19,43,44 Which process dominates depends on the substrate concentration, as discussed further below (Fig. 2D).

The substrate concentration had large effects on the level and kinetics of product formation in reactions with supplementation of AscA. Up to substrate concentrations of about 5 g L⁻¹, the reaction was more or less finished at 24 h, meaning that adding extra ascorbic acid did not lead to additional product formation (Fig. 2B and C). In these reactions, the final product levels increased with substrate concentration. Clearly, these reactions are limited by substrate availability, which affects both product levels and enzyme stability (Fig. 2D). The lower the substrate concentration, the more prone the LPMO will be to oxidative damage and the faster the substrate will be depleted (which again will lead to more LPMO damage). The 5 g L⁻¹ experiment shows that enzyme inactivation plays a major role: the product level in this reaction is 3.5 times (i.e. more than two times) lower than the highest product level in the 10 g L⁻¹ experiment, which proves that the substrate was not depleted and that enzyme inactivation is the reason why adding additional ascorbic acid did not lead to additional product formation.

At substrate concentrations above 5 g L⁻¹, supplementation of AscA at 24 h and 48 h markedly increased the accumulation of soluble oligosaccharide products (Fig. 2B and C), showing that at these substrate concentrations, a sufficient number of LPMO molecules remained active to productively use in situ generated H₂O₂ for cleavage of chitin. At these substrate concentrations (>10 g L⁻¹), product levels did not increase and even slightly decreased with substrate concentrations. This effect of the substrate concentration likely reflects a combination of multiple effects, including substrate saturation effects and the known fact that the fraction of soluble products generated by LPMOs may go down at higher substrate concentrations (see legend to Fig. S2 for a detailed explanation). In addition, mass transfer and oxygen transfer limitations may play a role. Finally, at these higher substrate concentrations, the LPMO reaction may become limited by available H₂O₂.

In terms of substrate conversion, the reactions at lower substrate concentrations (0.5–5 g L⁻¹) showed similar results (25–30%; Fig. 2C & S2C). Since product levels hardly changed at higher substrate concentrations, substrate conversion decreased with increasing substrate concentration above 10 g L⁻¹ (Fig. 2C). In terms of conversion, the best results were obtained at 10 g L⁻¹, achieving 34% conversion after 72 h, which, most importantly, is comparable to conversions obtained with chitinases acting on chitins that first had to be subjected to environmentally detrimental chemical pretreatment.^45,46

3.2 Potential of LPMO-mediated biocatalysis for chitin solubilization

XRD analysis of pre- and post-reaction β-chitin revealed a minimal reduction in crystallinity (70.9% → 68.5%; Fig. 3A), suggesting that the substrate had not changed much and could be amenable to further degradation with CBP21. To determine the maximum solubilization potential, the remaining insoluble chitin from the degradation reaction with 10 g L⁻¹ chitin (Fig. 2) was collected after being washed with ddH₂O to remove possible soluble products and then subjected to another treatment (at 10 g L⁻¹ substrate concentration) with CBP21. This was repeated twice, so that the total number of treatments was four (Fig. 3B). In contrast to the first reaction (Fig. 2C), these subsequent three reactions all showed enzyme inactivation at 24 h, which is shown by the fact that additions of AscA at 24 h and 48 h hardly increased product formation. This inactivation shows that the number of enzyme-accessible binding sites per gram of chitin has been reduced as a result of the first reaction. This is likely due to the presence of oxidized sites in the LPMO treated chitin that reduce the efficiency of LPMO binding. Interestingly, however, each of the three steps solubilized approximately 15% of the remaining chitin, implying that after four steps, the total conversion of the original chitin reached 60% (Fig. 3B). Thus, most importantly, the major part of the chitin could indeed be solubilized with one single enzyme. Clearly, there is a hitherto overlooked potential for solubilization of chitin by LPMOs alone.

Fig. 3 Solubilization potential of crystalline chitin with a single LPMO (CBP21). Reactions contained 1 μM CBP21, 1 mM AscA and 10 g L⁻¹ β-chitin in 20 mM Tris-HCl, pH 8.0, and were incubated at 37 °C, with shaking at 850 rpm. Fresh AscA was added to obtain a final concentration of 1 mM at 24 h and 48 h. (A) XRD analysis of β-chitin before (blue) and after (red) treatment with CBP21. The CBP21-treated material was derived from a 72 hour reaction as presented in Fig. 2. The crystallinity index (CrI, %) was calculated using the equation CrI = [(I₁₁₀ − I_am)/I₁₁₀] × 100, where I₁₁₀ is the maximum intensity at around 20° (2θ) and I_am is the intensity for the amorphous region at 16.0° (2θ).⁴⁷ The CrI values for non-treated and treated chitin were determined to be 70.9% and 68.5%, respectively. (B and C) After the first LPMO reaction (as shown in Fig. 2), the remaining insoluble chitin was collected by filtration and a new reaction was set up using the same conditions (10 g L⁻¹ substrate concentration) as in the first reaction (2^nd); this was repeated two times (3^rd and 4^th). Quantitative analysis of the soluble products was performed after treating these products with SmCHB. This quantitative analysis was used to calculate the mass of the solubilized chitin and of the remaining insoluble chitin. Cumulative conversion was calculated by summing the amounts of chitin solubilized in each step. The values shown are the average of three independent experiments with standard deviations shown as error bars.

Comparative analysis of three β-chitin fractions with distinct particle sizes in a standard 72 h reaction with repeated AscA supplementation (Fig. S3A) revealed minor size-dependent effects on the degree and kinetics of degradation. While the small- and medium-particle substrates gave very similar results, the reaction with the largest particles exhibited an accelerated reaction rate during the initial 48 h, while the reaction then slowed down, leading to lower final conversion, compared to the other two substrates. MALDI-TOF MS analysis of the products after 72 h (Fig. S3B) showed similar product profiles for all reactions.

To facilitate scaled production of oxidized chito-oligosaccharides, we then carried out LPMO-mediated chitin degradation in sealed glass bottles (Fig. S4). Experiments at 10 mL and 50 mL scales showed reduced enzymatic operational stability, with product formation limited to 48 h, and reduced final conversion (20.5% and 17.8%, respectively) compared to the reactions in microtubes that are described above. Although not very visible in the literature, years of LPMO research have shown that the outcome of reductant-driven LPMO reactions depends on incubation conditions such as the headspace and the shaking regime. Further process optimization is clearly possible.^48,49 One option is the gradual dosing of only small amounts of reductant, to keep the LPMO reduced, combined with gradual, controlled feeding of H₂O₂ to be used as the co-substrate. Gradual addition of fresh enzyme and/or fresh substrate could also be considered. Even without such further optimization, the 50 mL reaction enabled efficient preparation of >100 mg oxidized oligosaccharides, establishing a practical and truly green platform for subsequent bioactivity assays and other further studies.

While enzymes capable of depolymerizing chitin exist widely in nature, the efficiency of these enzymes in hydrolyzing natural crystalline chitin is low, particularly when acting alone. Process efficiency may be improved by reducing crystallinity using different chemical or physical methods. Chemical dissolution–precipitation steps, using chemicals such as phosphoric acid, hydrochloric acid, sodium or potassium hydroxide, urea and/or methanol increase enzymatic solubilization yields, but the yields obtained with single enzymes remain low, typically below 15%.^50,51 Colloidal chitin is relatively easy to degrade, but its preparation is not straightforward and requires concentrated hydrochloric acid.⁵² Physical pretreatment methods, such as ultrasonication, steam explosion and high-pressure homogenization, reduce crystallinity and increase enzymatic solubilization, but also when using such methods, solubilization yields tend to be on the order of only 20%.^45,53 These physical methods do not result in pollution and seem promising in breaking the crystalline structure of chitin, but they require relatively expensive process infrastructure. Importantly, the studies referred to above, with solubilization yields not exceeding 20%, all yielded simple mixtures of mono- and di-sugars with limited value. The LPMO approach described above gives higher solubilization yields, without any pretreatment or use of harsh chemicals, and yields more interesting products.

3.3 Immunostimulatory activity of oxidized chito-oligosaccharides

Macrophages play a pivotal role in both innate and adaptive immunity through their direct and indirect involvement in immune responses.^54,55 It is well known that chitin fragments (i.e. non-oxidized CHOS) may stimulate macrophages eliciting immune activation.^28,56,57 So far, the bioactivity of oxCHOS has not been studied, whereas data on other oligosaccharides suggest that oxidation may modulate bioactivity.^58,59 We therefore investigated the immunomodulatory effects of LPMO-generated oxCHOS on macrophage activation using RAW264.7 cells. The effects were assessed by monitoring the secretion of nitric oxide (NO) and of the pro-inflammatory cytokines interleukin-6 (IL-6) and tumor necrosis factor-alpha (TNF-α), which are all well-established indicators of macrophage activation.

Fig. 4A shows that treatment with oxCHOS enhanced NO secretion in macrophages after 6 hours of exposure (Fig. 4A). The difference became significant at about 6 h and then grew drastically. Fig. 4B shows that the effect was oxCHOS concentration-dependent. Notably, when oxCHOS concentrations exceeded 15 μg mL⁻¹, their capacity to stimulate NO secretion was comparable to that of LPS, a well-known potent inducer of macrophage activation. ELISA assays demonstrated that treatment with oxCHOS also led to upregulation of the two key pro-inflammatory cytokines interleukin-6 (IL-6) and tumor necrosis factor-alpha (TNF-α), in a dose-dependent manner (Fig. 4C). Importantly, cell viability assays showed that oxCHOS have low cytotoxicity (Fig. S5).

Fig. 4 Immunostimulatory activity of oxCHOS assessed using RAW 264.7 macrophages. Panels A and B show NO release over time by RAW 264.7 macrophages treated with oxCHOS (A, 15 μg ml⁻¹ oxCHOS) and after 24 h at different oxCHOS concentrations (B). Panel C shows the release of IL-6 and TNF-α after 24 h by RAW 264.7 macrophages treated with different concentrations of oxCHOS. RAW264.7 cells were stimulated with LPS (1 μg mL⁻¹) or oxCHOS for 24 h. Culture medium of non-stimulated cells was used as a blank control (CK). Panels D, E and F show the comparison of the immunostimulatory effects of low DP CHOS and oxCHOS (DP2 to DP5). (D) NO release by RAW 264.7 macrophages treated for 24 h with different concentrations of CHOS or oxCHOS (DP2–5), or with 1 μg mL⁻¹ LPS, or not treated (CK). (E & F) The production of IL-6 and TNF-α by these same cells. The data are presented as means ± SD of three independent biological replicates. Asterisks show statistically significant differences between the experimental groups (*p < 0.05, ** p < 0.01, *** p < 0.001); “ns” indicates “not significant”.

When comparing CHOS and oxCHOS for their immunostimulatory activities, it is required to use oligomers with the same DP, since DP may affect bioactivity.⁶⁰ Considering the poor solubility of, in particular, native oligomers with DP > 6, we prepared a mixture of low DP oxCHOS containing dimers to pentamers by treating a mixture of DP2–5 chito-oligosaccharides with a chito-oligosaccharide oxidase³² (Fig. S6). Assays similar to those shown in Fig. 4 demonstrated that the immunostimulatory activity of the oxidized oligomers was one to two orders of magnitude higher compared to their non-oxidized counterparts (Fig. 4D–F). It is noteworthy that these shorter oxCHOS (DP2–5), tested in a concentration range of 50–800 μg mL⁻¹, required higher concentrations to reach LPS-like immunostimulation, compared to the longer oligomers produced through the newly developed LPMO-based chitin solubilization method described above (Fig. 4B and D). This shows that higher molecular weight oxCHOS possess superior immunostimulatory activity, likely playing a significant role in stimulating host defense mechanisms. These results underpin the value of the one enzyme solubilization process developed in this study. Of note, longer oxCHOS can only be produced using LPMOs, since longer non-oxidized oligomers, which in principle could be oxidized to yield longer oxCHOS, have low or no solubility.

3.4 Modification of oxCHOS by enzymatic deacetylation

Deacetylation represents one of the most critical modifications of chitin and CHOS, changing properties varying from crystallinity and solubility to enzymatic degradability and bioactivity.^61,62 Chitin deacetylases (CDAs) selectively remove one or multiple acetyl groups from CHOS to produce partially or fully deacetylated oligomers. Enzymatic deacetylation of the oxCHOS produced here would give access to a whole new range of potentially bioactive compounds and was therefore explored, using ArCE4A, a marine-derived CDA that can catalyze multi-site deacetylation of chito-oligosaccharides.³⁴ Of note, CDAs with varying deacetylation specificities exist⁶³ and could be explored to create an even wider variation of oligomers.

Treatment of oxCHOS with ArCE4A led to rapid initial release of acetate, reaching about 30% after 15 min, and this fast phase was followed by a phase of decelerating deacetylation, reaching a final degree of deacetylation (DDA) of 65% after 24 hours (Fig. 5A). Nuclear magnetic resonance (NMR) analysis corroborated the time-dependent deacetylation that was deducted from acetate measurements, but the obtained DDA values were generally somewhat lower, with the maximum DDA being 57% (Fig. 5B and C). This discrepancy arises from the absence of the reducing-end H1 proton in oxCHOS, which disrupts the theoretical 1 : 3 area ratio between the H1 and CH₃ signals of chitin oligomers (see the legend to Fig. 5 for a more detailed explanation). Deacetylation is easily visible when zooming in on the δ 4.30–4.60 ppm region of the NMR spectra (Fig. 5B inset): over time the GlcNAc H1 signal at δ 4.45–4.55 ppm decreases while new H1 signals emerge at δ 4.39–4.45 ppm, corresponding to deacetylated GlcN residues.⁶⁴ Interestingly, the H1 signals at δ ∼4.36 ppm remain relatively unmodified, indicating ArCE4A's inability to deacetylate specific GlcNAc residues. These are likely GlcNAc units near the reducing end that are not deacetylated, in accordance with the previously reported substrate specificity of ArCE4A.³⁴

Fig. 5 Deacetylation of oxCHOS with ArCE4A, analyzed by measuring acetate release and NMR. Reactions contained 1 μM ArCE4A, 10 μM CoCl₂, and 1 mg mL⁻¹ (A–C) or 2 mM (D and E) substrate in 50 mM Tris-HCl, pH 8.0, and were incubated at 37 °C with shaking at 225 rpm. In the reactions depicted in panels A–C, 100% deacetylation of the substrate would yield an acetate concentration of 4.9 mM. (A) Quantitative analysis of the acetic acid concentration over time. (B) ¹H NMR spectra for reaction samples recorded at different time points. The inset shows the H1 region. (C) DDA (%) was estimated based on acetic acid release, assuming one acetylation per sugar unit in the starting material, or based on NMR data for which the DDA (%) was calculated using the equation . A_CH3 and A_H1 are the peak areas of the CH₃ and H1 signals in the ¹H NMR spectra and thus represents the degree of acetylation. Note: the NMR method leads to a slight underestimation of the DDA due to the absence of the reducing-end H1 proton in oxCHOS [(GlcNAc)_n−1GlcNAc1A]. MALDI-TOF MS spectra of LPMO-generated oxidized chito-oligosaccharides are shown in Fig. S7. (D) Acetic acid release in reactions with individual native (CHOS, A_n) or oxidized chito-oligosaccharides (oxCHOS, A_n-ox) after 1 h. (E) Acetic acid release over time in reactions with A₆ and A₆-ox. The panels show experimental data points with means ± SD for three replicates. In panel D, asterisks show statistically significant differences between the experimental groups (*p < 0.05, *** p < 0.001).

Mass spectrometric analysis of the products revealed distinct substrate preferences of ArCE4A towards oxCHOS of varying chain lengths. Within the initial 30 min of reaction (Fig. S7), deacetylation products were almost exclusively observed for oligomers with DP ≥ 6, including DA₅ (hexamer), DA₆ (heptamer), DA₇, D₂A₆, and D₃A₅ (octamers). Notably, in this early phase of the reaction, the number of deacetylated glucosamine residues appeared positively correlated with substrate chain length. This suggests that, when acting on oxCHOS, ArCE4A prefers longer-chain substrates. Later in the reaction (Fig. S7), deacetylated forms of the shorter oligomers also appeared.

This chain-length dependency and the possible impact of oxidation on CDA performance were further analyzed through several experiments. Deacetylation reactions with individual native and oxidized oligomers showed that oxidation substantially reduces the deacetylation efficiency for DP2–5 and much less so for DP6 (Fig. 5D). This explains the faster deacetylation of the longer oxCHOS discussed above and shows that the aldonic acid moieties of oxCHOS impair enzyme–substrate interactions. Progress curves with hexameric substrates showed that deacetylation of A₆-ox is slower and less complete compared to the native hexamer (Fig. 5E). Thus, while ArCE4A retains catalytic competence toward oxidized oligosaccharide substrates, structural modifications at the reducing end impose significant constraints on both substrate recognition and catalytic turnover.

Overall, further diversification of LPMO-generated oxCHOS could be achieved by enzymatic deacetylation. As we show here, oxidation does affect the impact and efficiency of deacetylases, but deacetylases can still be used for the eco-friendly production of oxidized chitosan oligosaccharides with widely varying degrees of deacetylation. To substantiate this, we verified that, besides ArCE4A, oxidized chito-oligosaccharides can also be deacetylated by other chitin deacetylases, such as an AnCDA from Aspergillus nidulans FGSC A4 (Fig. S8). Importantly, highly specific deacetylases can be engineered,⁶⁵ which allows for targeted deacetylation of one or more residues in the oxCHOS. Deacetylation transforms the CHOS into binary oligomers and exponentially increases structural and chemical diversity. Moreover, the concurrent presence of free amine groups (from deacetylation) and carboxyl groups (from terminal oxidation) creates unique zwitterionic properties. Next to potentially providing a wide range of bioactivities, the diversity of oxCHOS that can be created may help unravel the glycan code governing chitin functions in nature.

4. Conclusion

We show the feasibility of producing oxidized chito-oligosaccharides directly from crystalline chitin using LPMOs and show that this process gives access to otherwise hard to produce oligomeric products with strong bioactivity. Here, LPMOs are used as stand-alone enzymes to solubilize non-pretreated recalcitrant chitin with decent efficiency, which has never been described and has for long been considered “impossible”. It is interesting to contemplate the role of LPMO-catalyzed solubilization of chitin in nature and the signaling functions of the resulting oxidized oligomers. Our data suggest a crucial role of the terminal aldonic acid in determining the properties and functionalities of chito-oligosaccharides. The present results provide a truly eco-friendly alternative for the catalytic valorization of chitin biomass and at the same time provide an avenue towards novel chitin-derived products. The extended oligosaccharide library made accessible through the work described above adds a new dimension to understanding the role of chitin in nature and to the valorization of chitin biomass.

Author contributions

K. L., P. E. K. and V. G. H. E. designed the study. K. L., P. E. K., S. Z. and A. L. performed the experiments. O. G. and T. R. T. contributed to the LPMO and CDA methodology. K. L., P. E. K., S. Z., A. L. and V. G. H. E. performed the analysis and interpreted the data. K. L. and V. G. H. E. wrote the initial manuscript.

Conflicts of interest

The authors declare no competing interests.

Data availability

All the data provided in this article are included within the figures and tables of this article and the supplementary information (SI). Supplementary information is available. See DOI: https://doi.org/10.1039/d6gc00449k.

Acknowledgements

This study was supported by the National Natural Science Foundation of China (42576119 and 42276097), the Project for Future Network from the Chinese Academy of Sciences (058GJHZ2023044FN), the Natural Science Foundation of Shandong Province (ZR2022YQ37), and the Earmarked Fund for Modern Agro-industry Technology Research System (CARS-49). In addition, this study was supported by the European Union's Horizon Europe research and innovation programme under grant agreement no 101059786 “Valorisation of fungal biomass using novel enzymatic technology” (VALUABLE) and by the ChitoVal project funded by the Research Council of Norway, with project number 353185.

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