The Podospora anserina lytic polysaccharide monooxygenase PaLPMO9H catalyzes oxidative cleavage of diverse plant cell wall matrix glycans
© The Author(s) 2017
Received: 2 November 2016
Accepted: 2 March 2017
Published: 11 March 2017
The enzymatic conversion of plant biomass has been recently revolutionized by the discovery of lytic polysaccharide monooxygenases (LPMO) that catalyze oxidative cleavage of polysaccharides. These powerful enzymes are secreted by a large number of fungal saprotrophs and are important components of commercial enzyme cocktails used for industrial biomass conversion. Among the 33 AA9 LPMOs encoded by the genome of Podospora anserina, the PaLPMO9H enzyme catalyzes mixed C1/C4 oxidative cleavage of cellulose and cello-oligosaccharides. Activity of PaLPMO9H on several hemicelluloses has been suggested, but the regioselectivity of the cleavage remained to be determined.
In this study, we investigated the activity of PaLPMO9H on mixed-linkage glucans, xyloglucan and glucomannan using tandem mass spectrometry and ion mobility–mass spectrometry. Structural analysis of the released products revealed that PaLPMO9H catalyzes C4 oxidative cleavage of mixed-linkage glucans and mixed C1/C4 oxidative cleavage of glucomannan and xyloglucan. Gem-diols and ketones were produced at the non-reducing end, while aldonic acids were produced at the reducing extremity of the products.
The ability of PaLPMO9H to target polysaccharides, differing from cellulose by their linkages, glycosidic composition and/or presence of sidechains, could be advantageous for this coprophilous fungus when catabolizing highly variable polysaccharides and for the development of optimized enzyme cocktails in biorefineries.
KeywordsAA9 LPMO Lignocellulose Biomass Polysaccharides Mass spectrometry Biorefinery
The use of plant biomass represents an attractive alternative to fossil-based technologies for the production of high-value chemicals . In nature, filamentous fungi produce lignocellulose-degrading enzymes to acquire carbon from plant biomass. Different types of mechanisms for the deconstruction of plant cell walls have been described in saprotrophic fungi , but the involvement of oxidative enzymes was largely underestimated. The recent discovery of a new class of oxidative enzymes, namely lytic polysaccharide monooxygenases (LPMOs), has dramatically broadened the concept of the enzymatic deconstruction of plant cell wall polysaccharides [3–5]. LPMOs represent key cellulolytic enzymes that act at the surface of fibers where they mediate oxidative cleavage of polysaccharide chains. In industry, addition of LPMOs to cellulolytic cocktails leads to the reduction of the enzyme loading required for efficient saccharification of cellulosic biomass [6, 7].
Known LPMOs are grouped into the families AA9, AA10, AA11 and AA13 in the CAZy classification . LPMOs all feature a similar histidine brace coordinating the copper ion responsible for the oxidative cleavage of the substrate . Members of the AA9 family are mainly active on cellulose. The few characterized members from AA11 and AA13 families are active on chitin and starch, respectively [10, 11]. In contrast, the AA10s are mostly found in bacteria and exhibit activity on cellulose or chitin . Beside cellulose, AA9 LPMOs are also known to act on xyloglucan and glucomannan  as well as soluble cellodextrins [14–16]. Activity on xylan was detected only when xylans were in complex with cellulose chains .
Lytic polysaccharide monooxygenases from the AA9 family are exclusively found in fungi with large expansion of genes in white-rot fungi and some ascomycetes. The coprophilous fungus Podospora anserina displays an impressive array of CAZymes [18, 19] with 189 glycoside hydrolases and 33 genes encoding AA9 LPMOs (PaLPMO9), of which seven have been characterized biochemically [15, 20]. These LMPOs are able to oxidatively cleave cellulose with different regioselectivity. For instance, PaLPMO9E was shown to produce exclusively C1-oxidized products, while PaLPMO9H was reported to release both C1- and C4-oxidized products from cellulose [15, 20]. In addition, the PaLPMO9H displayed relatively broad substrate specificity demonstrated by enzyme activity assays using cellulose and cello-oligosaccharides. Activity of PaLPMO9H on different kinds of hemicelluloses was suggested based on hydrogen peroxide assay results. Indeed, the repression of hydrogen peroxide production in the presence of different hemicellulose substrates was significant and concentration-dependent for barley β-glucan, glucomannan, lichenan, and xyloglucan while curdlan, pectin, and xylan had no effect on hydrogen peroxide production . The degradation products and regioselectivity of PaLPMO9H acting on these substrates remained undetermined.
In the present work, a comprehensive oligosaccharide structural analysis was performed by tandem mass spectrometry (MS) and ion mobility–mass spectrometry on the released products of PaLPMO9H following incubation with four hemicellulosic substrates: lichenan, barley mixed-linkage glucan (MLG), konjac glucomannan (GM) and tamarind xyloglucan (XyG). These polysaccharides are widely represented in the cell walls of different monocotyledonous or dicotyledonous plants, and they exhibit different structural features . Similar to cellulose, all contain some (1 → 4)-linked β-d-glucose residues. Lichenan and MLG are polymers of β-d-glucosyl residues linked through (1 → 4) and (1 → 3) linkages. GM is composed of β-d-glucose and β-d-mannose residues, linked through (1 → 4) bonds, while XyG is a branched polysaccharide, with a backbone of (1 → 4)-linked β-d-glucose residues, most of which are substituted with (1 → 6)-linked α-d-xylose sidechains which are sometimes further extended with (1 → 2)-linked β-d-galactose residues. Our objective was to determine the specificity and selectivity of the PaLPMO9H enzyme for these structures, differing from cellulose by their linkages, glycosidic composition and/or the presence of sidechains.
Chemicals, substrates and standards
HPLC-grade methanol (MeOH) was purchased from Carlo-Erba (Peypin, France). Ultrapure Water was obtained from a milli-Q apparatus (Merck Millipore, Saint Quentin en Yvelines, France). All other chemicals were purchased from Sigma-Aldrich.
Polysaccharides including lichenan from Icelandic moss, barley MLG, tamarind XyG, konjac GM and β(1,3)/β(1,4)-oligosaccharides including G4A (Glc-β(1,3)-Glc-β(1,4)-Glc-β(1,4)-Glc), G4B (Glc-β(1,4)-Glc-β(1,4)-Glc-β(1,3)-Glc) and G4C (Glc-β(1,4)-Glc-β(1,3)-Glc-β(1,4)-Glc) were purchased from Megazyme (Bray, Ireland). XXXGXXXG (XGO14) was prepared and analyzed by HPAEC-PAD as described in McGregor et al. . Phosphoric acid cellulose was prepared as described in Bennati-Granier et al. .
PaLPMO9H from Podospora anserina (Genbank ID CAP61476) was expressed in Pichia pastoris, produced in a bioreactor and purified as described previously .
All the cleavage assays (300 µL liquid volume) contained 4.4 µM of PaLPMO9H, 1 mM of ascorbate and 0.05% (w/v) of hemicellulose or 0.2 mM XGO14 in milli-Q water. No buffer was used to avoid the formation of multiple salt adducts during the mass spectrometry analyses causing loss of sensitivity or misinterpretations. The enzyme reactions were performed in 2-mL tubes and incubated in a thermomixer (Eppendorf) at 40 °C and 500 rpm. After 24 h of incubation, the enzymatic reaction was stopped by filtration though a 10-kDa cutoff polyether sulfone membrane and the soluble products were analyzed by mass spectrometry.
Experiments were performed on a Synapt G2Si high-definition mass spectrometer (Waters Corp., Manchester, UK). Two types of mass measurements were performed on the samples. First, a mass profile was collected for a mass range of 350–2000 m/z. Second, ion mobility (IM) was activated to collect ion mobility–mass spectra separating possible isomers and/or reducing interference from sample impurities. IM wa performed in a traveling-wave ion mobility (TWIM) cell. The gas flows were held at 180 mL min−1 He in the helium cell and at 90 mL min−1 N2 in the mobility cell. The IM traveling-wave height was set to 40 V and its wave velocity was set to 450 m s−1 for positive ionization mode and 550 m s−1 for negative ionization mode. Ions of interest were fragmented after the gas-phase separation by collision-induced dissociation in the transfer cell of the instrument, with an adjustment of the collision energies such that a significant fragmentation was obtained while keeping an observable proportion of the intact precursor. Samples were diluted tenfold in MeOH/H2O (1:1, v/v) and infused at a flow rate of 5 μL min−1 in the instrument. The instrument was operated in positive or negative polarity and in “sensitivity” mode. Data acquisition was carried out using MassLynx software (V4.1).
Results and discussion
Different sources of hemicelluloses were incubated with PaLPMO9H under ascorbate conditions and their reaction products were characterized by mass spectrometry. Four substrates were considered: lichenan, MLG, XyG and GM. Cellulose was also included in the study as a reference substrate. The activity of PaLPMO9H products on cellulose has been described in detail and was shown to generate singly and doubly oxidized products at the C1 and/or C4 positions. This results in the formation of gem-diols/ketones at the newly formed “non-reducing” end or aldonic acids at the newly formed “reducing” end .
PaLPMO9H demonstrates broad specificity toward β-(1 → 4)-linked glucans
These results unambiguously confirmed that PaLPMO9H enzyme exhibits a broad specificity toward hemicelluloses compared to other AA9 LPMOs. Subtle differences were observed with respect to the action on cellulose, among which the apparent absence of double-oxidation. In-depth characterization of the reaction products was further carried out by tandem MS to explore the regioselectivity of the oxidative cleavage.
PaLPMO9H oxidatively cleaves β-(1 → 4)- and β-(1 → 4; 1 → 3)-linked glucosidic substrates
The results thus showed that DP4 species released from lichenan and MLG by PaLPMO9H had two main types of structures: one displaying three consecutive β-(1 → 4) linkages, the other encompassing at least one β-(1 → 3) linkage. MLG and lichenan contain a significant proportion of β-(1 → 3) linkages. On average, the ratio of (1 → 3) to (1 → 4) β-linkages is of 1:3 to 1:2 in these two polymers. In barley MLG, cellodextrin units of two (DP3) or three (DP4) adjacent β-(1 → 4) linkages are predominant, while longer cellodextrin sequences contributed to less than 10% of the total polysaccharide chain in the starchy endosperm . If one assumes that DP4 structures of different shapes have comparable ionization efficiency in MS, the peak area in Fig. 2 gives an estimate of the proportion of the two populations of DP4 differentiated by their ion mobility. The DP4 made of three consecutive β-(1 → 4) bonds was released in high proportion following action of PaLPMO9H on MLG and lichenan substrates. DP5 species were also released from MLG and lichenan (Additional file 1: Figure S3). The most abundant peak (dt = 92 bins) falls at the same drift time as the one measured from cellulose. Without excluding that those species arise in part from long cellodextrin stretches in the polymer, their abundance suggests that β-(1 → 3) bonds linking consecutive cellotetraosyl units were cleaved by PaLPMO9H. Thus, in addition to β-(1 → 4) linkages, PaLPMO9H appears able to catalyze the oxidative cleavage of β-(1 → 3) linkages in β-(1 → 4; 1 → 3)-linked glucosidic substrates (MLG and lichenan).
PaLPMO9H is active toward β-(1 → 4)-linked glucosidic substrates with branched sidechains
To examine the mode of action of PaLPMO9H on xyloglucan, the tetradecasaccharide XXXGXXXG was used as a model substrate. Named according to the standard linear nomenclature, in which “G” represents an unbranched β-(1 → 4)-linked glucosyl residue and “X” represents β-(1 → 4)-linked Glc bearing an α-(1 → 6)-linked xylosyl branch , XXXGXXXG presents a highly decorated backbone consisting of eight glucosyl residues. The MS spectrum of the products of cleavage of the XXXGXXXG preparation by PaLPMO9H (Fig. 1e) showed the presence of minor peaks, whose masses indicated the presence of sidechain galactosylation. They likely arose from a DP15 XyG, which was present as minor impurity (roughly 5%) of the XXXGXXXG preparation as confirmed by both MS and HPAEC-PAD chromatogram of the substrate before incubation with PaLPMO9H (Additional file 1: Figure S4). In the latter structure (likely XXLGXXXG and/or XXXGXXLG), one of the laterally branched xylose bears a β-(1 → 2)-linked galactosyl (represented as “L” ).
These results clearly indicated that PaLPMO9H was able to oxidatively cleave β-(1 → 4)-linked glucose chains even in the presence of extensive lateral branching.
Activity of PaLPMO9H on β-(1 → 4)-linked hetero-polysaccharides
Glucomannans are mainly straight-chain polymers of β-(1 → 4)-linked d-mannose and d-glucose. The product released at 705.18 m/z corresponding to a +16 Da mass shift from the non-modified DP4 was fragmented in tandem MS. The observed fragment ions showed the coexistence of two oxidized species with (i) the loss of 46 Da from the parent ion indicating a C1 oxidation (aldonic acid) at the reducing end, and (ii) a double loss of water from the precursor ion indicating a gem-diol at the non-reducing end (Additional file 1: Figure S5). On the other hand, fragmentation of the −2 Da species showed unambiguously a single species, corresponding to a ketone form at the non-reducing end. It can thus be concluded that PaLPMO9H oxidatively cleaved GM by forming oxidized species both at the reducing end and the non-reducing end. Since the main chain of GM consists of β-(1 → 4)-linked residues, it can be presumed that the oxidized positions are at C1 and C4, respectively.
auxiliary activity enzyme
Podospora anserina AA9 LPMO
lytic polysaccharide monooxygenase
barley mixed-linkage β-glucan
degree of polymerization
HR and JGB designed the research. MF, NM and SG performed the research experiments. MF, SG, DR, NM, HB, HR and JGB analyzed the data. MF, HR and JGB drafted the manuscript. All authors read and approved the final manuscript.
The authors would like to thank M. Haon and S. Grisel for technical assistance. N.M. thanks the Natural Sciences and Engineering Research Council of Canada (NSERC) for an Alexander Graham Bell Canada Graduate Doctoral Scholarship.
The authors declare that they have no competing interests.
Availability of supporting data
All data generated or analyzed during this study are included in this published article and its Additional files.
Consent for publication
All authors approved the final version of the manuscript.
This study was funded by the AMIDEX foundation (Funcopper project, Grant Number A*M-AAP-EI-13-13-130115-15.37 and MicrobioE project, Grant Number ANR-11-IDEX-0001-02). Work in Vancouver was additionally supported by NSERC (Discovery Grant), the Canada Foundation for Innovation and the British Columbia Knowledge Development Fund.
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- Gupta VK, et al. Fungal enzymes for bio-products from sustainable and waste biomass. Trends Biochem Sci. 2016;41(7):633–45.View ArticleGoogle Scholar
- Sigoillot J-C, et al. Fungal strategies for lignin degradation. In: Jouann L, Lapierre C, editors. Lignins: biosynthesis, biodegradation and bioengineering. Cambridge: Academic Press; 2012. p. 263–308.View ArticleGoogle Scholar
- Vaaje-Kolstad G, et al. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science. 2010;330(6001):219–22.View ArticleGoogle Scholar
- Beeson WT, et al. Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases. J Am Chem Soc. 2012;134(2):890–2.View ArticleGoogle Scholar
- Quinlan RJ, et al. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc Natl Acad Sci USA. 2011;108(37):15079–84.View ArticleGoogle Scholar
- Harris PV, et al. Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: structure and function of a large, enigmatic family. Biochemistry. 2010;49(15):3305–16.View ArticleGoogle Scholar
- Johansen KS. Discovery and industrial applications of lytic polysaccharide mono-oxygenases. Biochem Soc Trans. 2016;44:143–9.View ArticleGoogle Scholar
- Levasseur A, et al. Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes. Biotechnol Biofuels. 2016;6:14.Google Scholar
- Walton PH, Davies GJ. On the catalytic mechanisms of lytic polysaccharide monooxygenases. Curr Opin Chem Biol. 2016;31:195–207.View ArticleGoogle Scholar
- Hemsworth GR, et al. Discovery and characterization of a new family of lytic polysaccharide rnonooxygenases. Nat Chem Biol. 2014;10(2):122–6.View ArticleGoogle Scholar
- Lo Leggio L, et al. Structure and boosting activity of a starch-degrading lytic polysaccharide monooxygenase. Nat Commun. 2015;6:9.View ArticleGoogle Scholar
- Forsberg Z, et al. Comparative study of two chitin-active and two cellulose-active AA10-type lytic polysaccharide monooxygenases. Biochemistry. 2014;53(10):1647–56.View ArticleGoogle Scholar
- Agger JW, et al. Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc Natl Acad Sci USA. 2014;111(17):6287–92.View ArticleGoogle Scholar
- Isaksen T, et al. A C4-oxidizing lytic polysaccharide monooxygenase cleaving both cellulose and cello-oligosaccharides. J Biol Chem. 2014;289(5):2632–42.View ArticleGoogle Scholar
- Bennati-Granier C, et al. Substrate specificity and regioselectivity of fungal AA9 lytic polysaccharide monooxygenases secreted by Podospora anserina. Biotechnol Biofuels. 2015;8:14.View ArticleGoogle Scholar
- Frandsen KEH, et al. The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases. Nat Chem Biol. 2016;12(4):298.View ArticleGoogle Scholar
- Frommhagen M, et al. Discovery of the combined oxidative cleavage of plant xylan and cellulose by a new fungal polysaccharide monooxygenase. Biotechnol Biofuels. 2015;8:12.View ArticleGoogle Scholar
- Espagne E, et al. The genome sequence of the model ascomycete fungus Podospora anserina. Genome Biol. 2008;9(5):22.View ArticleGoogle Scholar
- Couturier M, et al. Plant biomass degrading ability of the coprophilic ascomycete fungus Podospora anserina. Biotechnol Adv. 2016;34(5):976–83.View ArticleGoogle Scholar
- Bey M, et al. Cello-oligosaccharide oxidation reveals differences between two lytic polysaccharide monooxygenases (Family GH61) from Podospora anserina. Appl Environ Microbiol. 2013;79(2):488–96.View ArticleGoogle Scholar
- Scheller HV, Ulvskov P. Hemicelluloses. In: Merchant S, Briggs WR, Ort D, editors. Annual review of plant biology. Santa Clara: Palo Alto; 2010. p. 263–89.Google Scholar
- McGregor N, et al. Structure-function analysis of a mixed-linkage beta-glucanase/xyloglucanase from the key ruminal bacteroidetes Prevotella bryantii B(1)4. J Biol Chem. 2016;291(3):1175–97.View ArticleGoogle Scholar
- Burton RA, Fincher GB. Current challenges in cell wall biology in the cereals and grasses. Front Plant Sci. 2012;3:6.View ArticleGoogle Scholar
- Tuomivaara ST, et al. Generation and structural validation of a library of diverse xyloglucan-derived oligosaccharides, including an update on xyloglucan nomenclature. Carbohydr Res. 2015;402:56–66.View ArticleGoogle Scholar
- Jagadeeswaran G, et al. A family of AA9 lytic polysaccharide monooxygenases in Aspergillus nidulans is differentially regulated by multiple substrates and at least one is active on cellulose and xyloglucan. Appl Microbiol Biotechnol. 2016;100(10):4535–47.View ArticleGoogle Scholar
- Nekiunaite L, et al. FgLPMO9A from Fusarium graminearum cleaves xyloglucan independently of the backbone substitution pattern. FEBS Lett. 2016;590(19):3346–56.View ArticleGoogle Scholar
- Kojima Y, et al. A lytic polysaccharide monooxygenase with broad xyloglucan specificity from the brown-rot fungus Gloeophyllum trabeum and its action on cellulose-xyloglucan complexes. Appl Environ Microbiol. 2016;82(22):6557–72.View ArticleGoogle Scholar
- Varki A, et al. Symbol nomenclature for glycan representation. Proteomics. 2009;9(24):5398–9.View ArticleGoogle Scholar