The impact of the carbohydrate-binding module on how a lytic polysaccharide monooxygenase modifies cellulose fibers.
AA9 LPMOs
CBM
Carbonyl detection
Cellulose
Enzymatic fiber engineering
Fluorescence
Functional variation
Oxidation
Size exclusion chromatography
Journal
Biotechnology for biofuels and bioproducts
ISSN: 2731-3654
Titre abrégé: Biotechnol Biofuels Bioprod
Pays: England
ID NLM: 9918300888906676
Informations de publication
Date de publication:
24 Aug 2024
24 Aug 2024
Historique:
received:
22
04
2024
accepted:
09
08
2024
medline:
26
8
2024
pubmed:
26
8
2024
entrez:
24
8
2024
Statut:
epublish
Résumé
In recent years, lytic polysaccharide monooxygenases (LPMOs) that oxidatively cleave cellulose have gained increasing attention in cellulose fiber modification. LPMOs are relatively small copper-dependent redox enzymes that occur as single domain proteins but may also contain an appended carbohydrate-binding module (CBM). Previous studies have indicated that the CBM "immobilizes" the LPMO on the substrate and thus leads to more localized oxidation of the fiber surface. Still, our understanding of how LPMOs and their CBMs modify cellulose fibers remains limited. Here, we studied the impact of the CBM on the fiber-modifying properties of NcAA9C, a two-domain family AA9 LPMO from Neurospora crassa, using both biochemical methods as well as newly developed multistep fiber dissolution methods that allow mapping LPMO action across the fiber, from the fiber surface to the fiber core. The presence of the CBM in NcAA9C improved binding towards amorphous (PASC), natural (Cell I), and alkali-treated (Cell II) cellulose, and the CBM was essential for significant binding of the non-reduced LPMO to Cell I and Cell II. Substrate binding of the catalytic domain was promoted by reduction, allowing the truncated CBM-free NcAA9C to degrade Cell I and Cell II, albeit less efficiently and with more autocatalytic enzyme degradation compared to the full-length enzyme. The sequential dissolution analyses showed that cuts by the CBM-free enzyme are more evenly spread through the fiber compared to the CBM-containing full-length enzyme and showed that the truncated enzyme can penetrate deeper into the fiber, thus giving relatively more oxidation and cleavage in the fiber core. These results demonstrate the capability of LPMOs to modify cellulose fibers from surface to core and reveal how variation in enzyme modularity can be used to generate varying cellulose-based materials. While the implications of these findings for LPMO-based cellulose fiber engineering remain to be explored, it is clear that the presence of a CBM is an important determinant of the three-dimensional distribution of oxidation sites in the fiber.
Sections du résumé
BACKGROUND
BACKGROUND
In recent years, lytic polysaccharide monooxygenases (LPMOs) that oxidatively cleave cellulose have gained increasing attention in cellulose fiber modification. LPMOs are relatively small copper-dependent redox enzymes that occur as single domain proteins but may also contain an appended carbohydrate-binding module (CBM). Previous studies have indicated that the CBM "immobilizes" the LPMO on the substrate and thus leads to more localized oxidation of the fiber surface. Still, our understanding of how LPMOs and their CBMs modify cellulose fibers remains limited.
RESULTS
RESULTS
Here, we studied the impact of the CBM on the fiber-modifying properties of NcAA9C, a two-domain family AA9 LPMO from Neurospora crassa, using both biochemical methods as well as newly developed multistep fiber dissolution methods that allow mapping LPMO action across the fiber, from the fiber surface to the fiber core. The presence of the CBM in NcAA9C improved binding towards amorphous (PASC), natural (Cell I), and alkali-treated (Cell II) cellulose, and the CBM was essential for significant binding of the non-reduced LPMO to Cell I and Cell II. Substrate binding of the catalytic domain was promoted by reduction, allowing the truncated CBM-free NcAA9C to degrade Cell I and Cell II, albeit less efficiently and with more autocatalytic enzyme degradation compared to the full-length enzyme. The sequential dissolution analyses showed that cuts by the CBM-free enzyme are more evenly spread through the fiber compared to the CBM-containing full-length enzyme and showed that the truncated enzyme can penetrate deeper into the fiber, thus giving relatively more oxidation and cleavage in the fiber core.
CONCLUSIONS
CONCLUSIONS
These results demonstrate the capability of LPMOs to modify cellulose fibers from surface to core and reveal how variation in enzyme modularity can be used to generate varying cellulose-based materials. While the implications of these findings for LPMO-based cellulose fiber engineering remain to be explored, it is clear that the presence of a CBM is an important determinant of the three-dimensional distribution of oxidation sites in the fiber.
Identifiants
pubmed: 39182111
doi: 10.1186/s13068-024-02564-8
pii: 10.1186/s13068-024-02564-8
doi:
Types de publication
Journal Article
Langues
eng
Pagination
118Subventions
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Horizon 2020
ID : 773324
Organisme : Norges Forskningsråd
ID : 297907
Organisme : Norges Forskningsråd
ID : 297907
Organisme : Norges Forskningsråd
ID : 297907
Organisme : Bundesministerium für Land- und Forstwirtschaft, Umwelt und Wasserwirtschaft
ID : 101379
Organisme : Bundesministerium für Land- und Forstwirtschaft, Umwelt und Wasserwirtschaft
ID : 101379
Organisme : Bundesministerium für Land- und Forstwirtschaft, Umwelt und Wasserwirtschaft
ID : 101379
Organisme : Academy of Finland
ID : 326359
Organisme : Academy of Finland
ID : 326359
Organisme : Academy of Finland
ID : 326359
Organisme : Novo Nordisk Fonden
ID : 0061165
Informations de copyright
© 2024. The Author(s).
Références
Vaaje-Kolstad G, Westereng B, Horn SJ, Liu Z, Zhai H, Sørlie M, Eijsink VGH. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science. 2010;330(6001):219–22.
pubmed: 20929773
doi: 10.1126/science.1192231
Suzuki K, Suzuki M, Taiyoji M, Nikaidou N, Watanabe T. Chitin binding protein (CBP21) in the culture supernatant of Serratia marcescens 2170. Biosci Biotechnol Biochem. 1998;62(1):128–35.
pubmed: 9501524
doi: 10.1271/bbb.62.128
Vaaje-Kolstad G, Horn SJ, van Aalten DMF, Synstad B, Eijsink VGH. The non-catalytic chitin-binding protein CBP21 from Serratia marcescens is essential for chitin degradation. J Biol Chem. 2005;280(31):28492–7.
pubmed: 15929981
doi: 10.1074/jbc.M504468200
Merino ST, Cherry J. Progress and challenges in enzyme development for biomass utilization. Adv Biochem Eng Biotechnol. 2007;108:95–120.
pubmed: 17594064
Loose JSM, Forsberg Z, Fraaije MW, Eijsink VGH, Vaaje-Kolstad G. A rapid quantitative activity assay shows that the Vibrio cholerae colonization factor GbpA is an active lytic polysaccharide monooxygenase. FEBS Lett. 2014;588(18):3435–40.
pubmed: 25109775
doi: 10.1016/j.febslet.2014.07.036
Sabbadin F, Urresti S, Henrissat B, Avrova AO, Welsh LRJ, Lindley PJ, Csukai M, Squires JN, Walton PH, Davies GJ, et al. Secreted pectin monooxygenases drive plant infection by pathogenic oomycetes. Science. 2021;373(6556):774–9.
pubmed: 34385392
doi: 10.1126/science.abj1342
Sabbadin F, Hemsworth GR, Ciano L, Henrissat B, Dupree P, Tryfona T, Marques RDS, Sweeney ST, Besser K, Elias L, et al. An ancient family of lytic polysaccharide monooxygenases with roles in arthropod development and biomass digestion. Nat Commun. 2018;9(1):756.
pubmed: 29472725
pmcid: 5823890
doi: 10.1038/s41467-018-03142-x
Vu VV, Hangasky JA, Detomasi TC, Henry SJW, Ngo ST, Span EA, Marletta MA. Substrate selectivity in starch polysaccharide monooxygenases. J Biol Chem. 2019;294(32):12157–66.
pubmed: 31235519
pmcid: 6690692
doi: 10.1074/jbc.RA119.009509
Zhou X, Zhu H. Current understanding of substrate specificity and regioselectivity of LPMOs. Bioresour Bioprocess. 2020;7(1):11.
doi: 10.1186/s40643-020-0300-6
Yu X, Zhao Y, Yu J, Wang L. Recent advances in the efficient degradation of lignocellulosic metabolic networks by lytic polysaccharide monooxygenase. Acta Biochim Biophys Sin. 2023;55(4):529–39.
pubmed: 37036250
pmcid: 10195143
doi: 10.3724/abbs.2023059
Forsberg Z, Courtade G. On the impact of carbohydrate-binding modules (CBMs) in lytic polysaccharide monooxygenases (LPMOs). Essays Biochem. 2023;67(3):561–74.
pubmed: 36504118
doi: 10.1042/EBC20220162
Boraston AB, Bolam DN, Gilbert HJ, Davies GJ. Carbohydrate-binding modules: fine-tuning polysaccharide recognition. Biochem J. 2004;382(3):769–81.
pubmed: 15214846
pmcid: 1133952
doi: 10.1042/BJ20040892
Levasseur A, Drula E, Lombard V, Coutinho PM, Henrissat B. Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes. Biotechnol Biofuels. 2013;6(1):41.
pubmed: 23514094
pmcid: 3620520
doi: 10.1186/1754-6834-6-41
Bissaro B, Røhr ÅK, Müller G, Chylenski P, Skaugen M, Forsberg Z, Horn SJ, Vaaje-Kolstad G, Eijsink VGH. Oxidative cleavage of polysaccharides by monocopper enzymes depends on H
pubmed: 28846668
doi: 10.1038/nchembio.2470
Jones SM, Transue WJ, Meier KK, Kelemen B, Solomon EI. Kinetic analysis of amino acid radicals formed in H
pubmed: 32414932
pmcid: 7275769
doi: 10.1073/pnas.1922499117
Kont R, Bissaro B, Eijsink VGH, Väljamäe P. Kinetic insights into the peroxygenase activity of cellulose-active lytic polysaccharide monooxygenases (LPMOs). Nat Commun. 2020;11(1):5786.
pubmed: 33188177
pmcid: 7666214
doi: 10.1038/s41467-020-19561-8
Chang H, Gacias Amengual N, Botz A, Schwaiger L, Kracher D, Scheiblbrandner S, Csarman F, Ludwig R. Investigating lytic polysaccharide monooxygenase-assisted wood cell wall degradation with microsensors. Nat Commun. 2022;13(1):6258.
pubmed: 36271009
pmcid: 9586961
doi: 10.1038/s41467-022-33963-w
Bissaro B, Eijsink VGH. Lytic polysaccharide monooxygenases: enzymes for controlled and site-specific Fenton-like chemistry. Essays Biochem. 2023;67(3):575–84.
pubmed: 36734231
doi: 10.1042/EBC20220250
Lim H, Brueggemeyer MT, Transue WJ, Meier KK, Jones SM, Kroll T, Sokaras D, Kelemen B, Hedman B, Hodgson KO, et al. Kβ X-ray emission spectroscopy of Cu(I)-lytic polysaccharide monooxygenase: Direct observation of the frontier molecular orbital for H
pubmed: 37441786
pmcid: 10557184
doi: 10.1021/jacs.3c04048
Golten O, Ayuso-Fernández I, Hall KR, Stepnov AA, Sørlie M, Røhr ÅK, Eijsink VGH. Reductants fuel lytic polysaccharide monooxygenase activity in a pH-dependent manner. FEBS Lett. 2023;597(10):1363–74.
pubmed: 37081294
doi: 10.1002/1873-3468.14629
Kittl R, Kracher D, Burgstaller D, Haltrich D, Ludwig R. Production of four Neurospora crassa lytic polysaccharide monooxygenases in Pichia pastoris monitored by a fluorimetric assay. Biotechnol Biofuels. 2012;5(1):79.
pubmed: 23102010
pmcid: 3500269
doi: 10.1186/1754-6834-5-79
Phillips CM, Beeson WT, Cate JH, Marletta MA. Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa. ACS Chem Biol. 2011;6(12):1399–406.
pubmed: 22004347
doi: 10.1021/cb200351y
Eijsink VGH, Petrović D, Forsberg Z, Mekasha S, Røhr ÅK, Várnai A, Bissaro B, Vaaje-Kolstad G. On the functional characterization of lytic polysaccharide monooxygenases (LPMOs). Biotechnol Biofuels. 2019;12(1):58.
pubmed: 30923566
pmcid: 6423801
doi: 10.1186/s13068-019-1392-0
Vu VV, Beeson WT, Phillips CM, Cate JHD, Marletta MA. Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases. J Am Chem Soc. 2014;136(2):562–5.
pubmed: 24350607
doi: 10.1021/ja409384b
Borisova AS, Isaksen T, Dimarogona M, Kognole AA, Mathiesen G, Várnai A, Røhr AK, Payne CM, Sørlie M, Sandgren M, et al. Structural and functional characterization of a lytic polysaccharide monooxygenase with broad substrate specificity. J Biol Chem. 2015;290(38):22955–69.
pubmed: 26178376
pmcid: 4645601
doi: 10.1074/jbc.M115.660183
Hansson H, Karkehabadi S, Mikkelsen N, Douglas NR, Kim S, Lam A, Kaper T, Kelemen B, Meier KK, Jones SM. High-resolution structure of a lytic polysaccharide monooxygenase from Hypocrea jecorina reveals a predicted linker as an integral part of the catalytic domain. J Biol Chem. 2017;292(46):19099–109.
pubmed: 28900033
pmcid: 5704490
doi: 10.1074/jbc.M117.799767
Courtade G, Forsberg Z, Heggset EB, Eijsink VGH, Aachmann FL. The carbohydrate-binding module and linker of a modular lytic polysaccharide monooxygenase promote localized cellulose oxidation. J Biol Chem. 2018;293(34):13006–15.
pubmed: 29967065
pmcid: 6109919
doi: 10.1074/jbc.RA118.004269
Chalak A, Villares A, Moreau C, Haon M, Grisel S, d’Orlando A, Herpoël-Gimbert I, Labourel A, Cathala B, Berrin JG. Influence of the carbohydrate-binding module on the activity of a fungal AA9 lytic polysaccharide monooxygenase on cellulosic substrates. Biotechnol Biofuels. 2019;12:206.
pubmed: 31508147
pmcid: 6721207
doi: 10.1186/s13068-019-1548-y
Danneels B, Tanghe M, Desmet T. Structural features on the substrate-binding surface of fungal lytic polysaccharide monooxygenases determine their oxidative regioselectivity. Biotechnol J. 2019;14(3): e1800211.
pubmed: 30238672
doi: 10.1002/biot.201800211
Laurent CVFP, Sun P, Scheiblbrandner S, Csarman F, Cannazza P, Frommhagen M, van Berkel WJH, Oostenbrink C, Kabel MA, Ludwig R. Influence of lytic polysaccharide monooxygenase active site segments on activity and affinity. Int J Mol Sci. 2019;20(24):6219.
pubmed: 31835532
pmcid: 6940765
doi: 10.3390/ijms20246219
Sun P, Valenzuela SV, Chunkrua P, Javier Pastor FI, Laurent CVFP, Ludwig R, van Berkel WJH, Kabel MA. Oxidized product profiles of AA9 lytic polysaccharide monooxygenases depend on the type of cellulose. ACS Sustain Chem Eng. 2021;9(42):14124–33.
pubmed: 34722005
pmcid: 8549066
doi: 10.1021/acssuschemeng.1c04100
Srivastava A, Nagar P, Rathore S, Adlakha N. The linker region promotes activity and binding efficiency of modular LPMO towards polymeric substrate. Microbiol Spectr. 2022;10(1): e0269721.
pubmed: 35080440
doi: 10.1128/spectrum.02697-21
Várnai A, Siika-aho M, Viikari L. Carbohydrate-binding modules (CBMs) revisited: Reduced amount of water counterbalances the need for CBMs. Biotechnol Biofuels. 2013;6(1):30.
pubmed: 23442543
pmcid: 3599012
doi: 10.1186/1754-6834-6-30
Forsberg Z, Stepnov AA, Tesei G, Wang Y, Buchinger E, Kristiansen SK, Aachmann FL, Arleth L, Eijsink VGH, Lindorff-Larsen K, et al. The effect of linker conformation on performance and stability of a two-domain lytic polysaccharide monooxygenase. J Biol Chem. 2023;299(11): 105262.
pubmed: 37734553
pmcid: 10598543
doi: 10.1016/j.jbc.2023.105262
Kuusk S, Eijsink VGH, Väljamäe P. The “life-span” of lytic polysaccharide monooxygenases (LPMOs) correlates to the number of turnovers in the reductant peroxidase reaction. J Biol Chem. 2023;299(9): 105094.
pubmed: 37507015
pmcid: 10458328
doi: 10.1016/j.jbc.2023.105094
Gao W, Li T, Zhou H, Ju J, Yin H. Carbohydrate-binding modules enhance H
pubmed: 38122901
doi: 10.1016/j.jbc.2023.105573
Stepnov AA, Eijsink VGH, Forsberg Z. Enhanced in situ H
pubmed: 35414104
pmcid: 9005612
doi: 10.1038/s41598-022-10096-0
Eibinger M, Ganner T, Bubner P, Rosker S, Kracher D, Haltrich D, Ludwig R, Plank H, Nidetzky B. Cellulose surface degradation by a lytic polysaccharide monooxygenase and its effect on cellulase hydrolytic efficiency. J Biol Chem. 2014;289(52):35929–38.
pubmed: 25361767
pmcid: 4276861
doi: 10.1074/jbc.M114.602227
Vuong TV, Liu B, Sandgren M, Master ER. Microplate-based detection of lytic polysaccharide monooxygenase activity by fluorescence-labeling of insoluble oxidized products. Biomacromol. 2017;18(2):610–6.
doi: 10.1021/acs.biomac.6b01790
Solhi L, Li J, Li J, Heyns NMI, Brumer H. Oxidative enzyme activation of cellulose substrates for surface modification. Green Chem. 2022;24(10):4026–40.
doi: 10.1039/D2GC00393G
Mathieu Y, Raji O, Bellemare A, Di Falco M, Nguyen TTM, Viborg AH, Tsang A, Master E, Brumer H. Functional characterization of fungal lytic polysaccharide monooxygenases for cellulose surface oxidation. Biotechnol Biofuels Bioprod. 2023;16(1):132.
pubmed: 37679837
pmcid: 10486138
doi: 10.1186/s13068-023-02383-3
Sulaeva I, Støpamo FG, Melikhov I, Budischowsky D, Rahikainen J, Borisova A, Marjamaa K, Kruus K, Eijsink VGH, Várnai A, et al. Beyond the surface: a methodological exploration of enzyme impact along the cellulose fiber cross-section. Biomacromol. 2024;25(5):3076–86.
doi: 10.1021/acs.biomac.4c00152
Støpamo FG, Sulaeva I, Budischowsky D, Rahikainen J, Marjamaa K, Potthast A, Kruus K, Eijsink VGH, Várnai A. Oxidation of cellulose fibers using LPMOs with varying allomorphic substrate preferences, oxidative regioselectivities, and domain structures. Carbohydr Polym. 2024;330: 121816.
pubmed: 38368098
doi: 10.1016/j.carbpol.2024.121816
Rahikainen J, Ceccherini S, Molinier M, Holopainen-Mantila U, Reza M, Väisänen S, Puranen T, Kruus K, Vuorinen T, Maloney T, et al. Effect of cellulase family and structure on modification of wood fibres at high consistency. Cellulose. 2019;26(8):5085–103.
doi: 10.1007/s10570-019-02424-x
Swerin A, Odberg L, Lindström T. Deswelling of hardwood kraft pulp fibers by cationic polymers. Effect Wet Press Sheet Propert. 1990;5(4):188–96.
Zhang YHP, Cui J, Lynd LR, Kuang LR. A transition from cellulose swelling to cellulose dissolution by o-phosphoric acid: evidence from enzymatic hydrolysis and supramolecular structure. Biomacromol. 2006;7(2):644–8.
doi: 10.1021/bm050799c
Stepnov AA, Forsberg Z, Sørlie M, Nguyen GS, Wentzel A, Røhr ÅK, Eijsink VGH. Unraveling the roles of the reductant and free copper ions in LPMO kinetics. Biotechnol Biofuels. 2021;14(1):28.
pubmed: 33478537
pmcid: 7818938
doi: 10.1186/s13068-021-01879-0
Tuveng TR, Jensen MS, Fredriksen L, Vaaje-Kolstad G, Eijsink VGH, Forsberg Z. A thermostable bacterial lytic polysaccharide monooxygenase with high operational stability in a wide temperature range. Biotechnol Biofuels. 2020;13(1):194.
pubmed: 33292445
pmcid: 7708162
doi: 10.1186/s13068-020-01834-5
Ståhlberg J, Divne C, Koivula A, Piens K, Claeyssens M, Teeri TT, Jones TA. Activity studies and crystal structures of catalytically deficient mutants of cellobiohydrolase I from Trichoderma reesei. J Mol Biol. 1996;264(2):337–49.
pubmed: 8951380
doi: 10.1006/jmbi.1996.0644
Müller G, Várnai A, Johansen KS, Eijsink VGH, Horn SJ. Harnessing the potential of LPMO-containing cellulase cocktails poses new demands on processing conditions. Biotechnol Biofuels. 2015;8:187.
pubmed: 26609322
pmcid: 4659242
doi: 10.1186/s13068-015-0376-y
Röhrling J, Potthast A, Rosenau T, Lange T, Ebner G, Sixta H, Kosma P. A novel method for the determination of carbonyl groups in cellulosics by fluorescence labeling .1. Method development. Biomacromol. 2002;3(5):959–68.
doi: 10.1021/bm020029q
Sulaeva I, Budischowsky D, Rahikainen J, Marjamaa K, Støpamo FG, Khaliliyan H, Melikhov I, Rosenau T, Kruus K, Várnai A, et al. A novel approach to analyze the impact of lytic polysaccharide monooxygenases (LPMOs) on cellulosic fibres. Carbohydr Polym. 2024;328: 121696.
pubmed: 38220335
doi: 10.1016/j.carbpol.2023.121696
Ernst O, Zor T. Linearization of the Bradford protein assay. J Vis Exp. 2010;38: e1918.
Várnai A, Viikari L, Marjamaa K, Siika-aho M. Adsorption of monocomponent enzymes in enzyme mixture analyzed quantitatively during hydrolysis of lignocellulose substrates. Bioresour Technol. 2011;102(2):1220–7.
pubmed: 20736135
doi: 10.1016/j.biortech.2010.07.120
Stepnov AA, Eijsink VGH. Looking at LPMO reactions through the lens of the HRP/Amplex Red assay. Methods Enzymol. 2023;679:163–89.
pubmed: 36682861
doi: 10.1016/bs.mie.2022.08.049
Filandr F, Man P, Halada P, Chang H, Ludwig R, Kracher D. The H
pubmed: 32158501
pmcid: 7057652
doi: 10.1186/s13068-020-01673-4
Brander S, Tokin R, Ipsen JØ, Jensen PE, Hernández-Rollán C, Nørholm MHH, Lo Leggio L, Dupree P, Johansen KS. Scission of glucosidic bonds by a Lentinus similis lytic polysaccharide monooxygenases is strictly dependent on H
doi: 10.1021/acscatal.1c04248
Potthast A, Radosta S, Saake B, Lebioda S, Heinze T, Henniges U, Isogai A, Koschella A, Kosma P, Rosenau T, et al. Comparison testing of methods for gel permeation chromatography of cellulose: coming closer to a standard protocol. Cellulose. 2015;22(3):1591–613.
doi: 10.1007/s10570-015-0586-2
Rieder L, Stepnov AA, Sørlie M, Eijsink VGH. Fast and specific peroxygenase reactions catalyzed by fungal mono-copper enzymes. Biochemistry. 2021;60(47):3633–43.
pubmed: 34738811
doi: 10.1021/acs.biochem.1c00407
Müller G, Chylenski P, Bissaro B, Eijsink VGH, Horn SJ. The impact of hydrogen peroxide supply on LPMO activity and overall saccharification efficiency of a commercial cellulase cocktail. Biotechnol Biofuels. 2018;11:209.
pubmed: 30061931
pmcid: 6058378
doi: 10.1186/s13068-018-1199-4
Kracher D, Andlar M, Furtmüller PG, Ludwig R. Active-site copper reduction promotes substrate binding of fungal lytic polysaccharide monooxygenase and reduces stability. J Biol Chem. 2018;293(5):1676–87.
pubmed: 29259126
doi: 10.1074/jbc.RA117.000109
Raji O, Eijsink VGH, Master E, Forsberg Z. Modularity impacts cellulose surface oxidation by a lytic polysaccharide monooxygenase from Streptomyces coelicolor. Cellulose. 2023;30(17):10783–94.
doi: 10.1007/s10570-023-05551-8