Fungal Lytic Polysaccharide Monooxygenases (LPMOs): Functional Adaptation and Biotechnological Perspectives
Abstract
1. Introduction
2. Classification of Fungal LPMO Families
3. Phylogenetic Distribution of Fungal LPMOs
4. Relationship Between LPMOs and Fungal Lifestyle
5. Catalytic Pathways and Specific Modulations of Fungal LPMOs
5.1. Catalytic Mechanism Overview
- Oxygen-Driven (O2) Mechanism: Traditionally, it was believed that LPMOs functioned as monooxygenases, employing molecular oxygen (O2) as the oxygen source [144]. In this pathway, the copper in the resting Cu(II) state must first be reduced to Cu(I) via an external electron donor (e.g., cellobiose dehydrogenase—CDH or phenolic compounds). The reduced Cu(I) center reacts with molecular oxygen to form a reactive oxygen species, which may be a Cu(II)-superoxide or Cu(II)-peroxide intermediate. This intermediate performs hydrogen atom abstraction (HAA) from the C–H bond of the polysaccharide substrate. The subsequent rebound of the hydroxyl group to the carbon leads to cleavage of the glycosidic bond, typically at the C1 or C4 position, depending on the enzyme’s regioselectivity. Although well-characterized in vitro, this O2-driven pathway is kinetically slow, as it requires two sequential electron transfers per catalytic event, one for reducing Cu(II) to Cu(I) and another to complete the O2 reduction [31,116,145].
- Peroxide-Driven (H2O2) Mechanism: In this route, the Cu(I) center reacts directly with hydrogen peroxide (H2O2), forming a highly reactive Cu(II)-oxyl species or a related high-valent intermediate. This pathway is significantly faster than the O2 route because only one electron is required to reduce Cu(II) to Cu(I) for catalytic turnover, and the reaction with H2O2 produces a potent oxidant capable of immediate hydrogen abstraction and glycosidic bond cleavage. It is important to highlight that in the H2O2-driven mechanism, LPMO stays in the reduced state (Cu(I)) at the end of the catalytical cycle, then multiple cycles are possible without additional reductant being necessary. However, this route carries inherent risks. Excessive or uncontrolled H2O2 levels can lead to enzyme self-inactivation through oxidative damage to the histidine brace or other nearby residues. Consequently, fungi have evolved intricate systems to regulate H2O2 production, including tight coupling with oxidoreductases (like CDH, glucose oxidase, and aryl-alcohol oxidases) and enzymes that scavenge excess H2O2 (such as catalases and peroxidases) [116,142,146,147,148,149].
5.2. Cofactor Requirements and Electron Donors
5.3. Substrate Specificity
- AA9: the most studied in fungi, primarily targets cellulose, with some members also acting on hemicelluloses like xyloglucan, glucomannan, and mixed-linkage β-glucans [25]. Oxidation patterns vary between C1, C4, and mixed C1/C4, depending on the enzyme [162]. This broad substrate range is largely attributed to the flexible architecture of AA9 surface loops (especially L2, L3, LC, and LS), which adapt the conserved LPMO β-sandwich fold to accommodate both crystalline and amorphous substrates [163,164].
- AA10 (rare in fungi): Fungal AA10s exhibit activity on both crystalline cellulose and chitin. They are capable of C1, C4, or mixed oxidation, similar to AA9 [25]. This dual specificity stems from a structurally open substrate-binding surface and conserved residues in the flat catalytic plane of the typical LPMO fold, allowing alignment with both β-1,4-glucans and β-1,4-N-acetylglucosamine chains [165,166].
- AA11: specializes in chitin, particularly in the modification of crystalline chitin [95], with oxidative cleavage at the C1 position being most common [31]. This specificity is associated with a narrower substrate-binding groove and distinct electrostatic surface potentials, tailored to interact with the acetylated chitin chains rather than the more hydrophobic cellulose [31,95].
- AA13: targets starch (α-1,4-glucans), specifically disrupting the crystalline regions of amylose and amylopectin [167]. Its substrate preference reflects adaptations in the catalytic interface that favor the helical conformation of starch polysaccharides, diverging structurally from other LPMOs by modifications to loops around the active site cleft [58,168].
- AA14: exhibits activity on xylan but uniquely requires the presence of cellulose for activity, suggesting that it acts on xylan tightly bound to cellulose fibrils within the plant cell wall. The specificity of AA14 is hypothesized to rely on the positioning of the substrate-binding loops and electrostatic complementarity that favor xylan–cellulose complexes, consistent with the LPMO β-sandwich fold but with added surface constraints [62].
- AA15 (rare in fungi): demonstrates broad activity on both chitin and cellulose, with evidence of oxidation on both C1 and C4 carbons [25]. This functional versatility may reflect its evolutionary origin from arthropods and a more flexible binding surface, superimposed onto the typical LPMO fold, that tolerates multiple polysaccharide geometries [67,98].
- AA16: acts predominantly on cellulose, with oxidation typically at the C1 position. It may be complementary to AA9 enzymes, especially in fungal species specialized in degrading highly recalcitrant plant biomass [25]. The substrate preference of AA16 is linked to specific loop configurations and redox partner preferences that distinguish it from AA9, despite sharing the core β-sandwich fold and catalytic mechanism [70,71].
5.4. Oxidation Pattern Modulation (C1, C4, Mixed)
5.5. Synergy with Other Enzymes
5.6. Biological Modulation and Regulation
6. Fungal LPMOs with Industrial Applications
7. Future Perspectives
8. Conclusions
Author Contributions
Funding
Data Availability Statement
Conflicts of Interest
References
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AA Family | Taxonomic Distribution | Substrate Specificity | Oxidation Pattern | Structural Features | Biochemical Features | Ecological and Functional Roles | Fungi Examples | References |
---|---|---|---|---|---|---|---|---|
AA9 | Fungi | Primarily cellulose; also acts on hemicellulose (e.g., xylan, xyloglucan, galactoglucomannan) | C1, C4, or both (C1/C4) | β-sandwich fold with conserved histidine brace; variable loops (L2, L3, L8, LC) modulate substrate interaction; many members have carbohydrate-binding modules (CBMs) | High catalytic diversity; some thermostable; accept multiple electron donors (CDHs, lignin, H2O2); work synergistically with cellulases | Leading role in cellulose deconstruction; key in saprotrophic fungi for lignocellulose breakdown; industrial relevance in biofuels and biorefineries | Aspergillus fumigatus, Aspergillus nidulans, Malbranchea cinnamomea, Neurospora crassa, Pycnoporus sanguineus, Thermothelomyces fergusii Thermothelomyces thermophilus, Thermothielavioides terrestris, Thielavia australiensis, Trichoderma reesei | [23,25,26,27,28,49,50,51,52,53,54] |
AA10 | Mainly bacteria, some viruses and archaeans, minor reports in fungi | Cellulose, chitin | C1, C4, or both (C1/C4) | Similar β-sandwich fold to AA9; monocopper site; some contain CBMs | Oxidizes chitin and cellulose (C1/C4); requires O2 or H2O2; electron donors such as ascorbate or CDH | Originally bacterial; contributes to degradation of recalcitrant polysaccharides; cross-kingdom relevance, including minor fungal representatives | Ustilago maydis | [8,25,55] |
AA11 | Fungi, bacteria | Chitin (especially crystalline chitin from crustaceans, fungal cell walls) | Predominantly C1 | β-sandwich fold; distinct loop regions compared to AA9; conserved histidine brace with unique surrounding residues | Lower catalytic rates; electron donor systems less characterized; potential interaction with small molecule donors or fungal redox partners | Chitin degradation for nutrient acquisition; possible roles in fungal cell wall remodelling and defense against other fungi or arthropods | Aspergillus fumigatus, Trichoderma guizhouense | [25,56,57] |
AA13 | Fungi | Retrograded (gelatinized) starch; inactive on crystalline cellulose | Mainly C1 | β-sandwich fold; shallow binding groove adapted for α-glucans rather than flat crystalline surfaces | High efficiency on retrograded starch; activity enhanced by small molecule donors; natural redox partners still unclear | Enables fungi to utilize recalcitrant starch in natural or anthropogenic environments; adaptation to transient starch-rich substrates | Aspergillus nidulans, Aspergillus oryzae | [25,58,59,60] |
AA14 | Fungi | Xylan, but only when associated with cellulose microfibrils | C1 | β-sandwich fold with surface adapted to interact with xylan helices embedded in cellulose | Requires xylan to be in the context of the cellulose-xylan network; facilitates loosening of plant cell wall matrices | Plays a role in disrupting hemicellulose-cellulose interactions; facilitates access for other glycoside hydrolases; niche role in complex biomass degradation | Phlebia radiata, Sordaria brevicollis Talaromyces rugulosus. Trametes coccinea (syn. Pycnoporus coccineus) | [25,61,62,63,64,65] |
AA15 | Fungi, viruses, archaeans, animals, plants | Cellulose, chitin | C1 and C4 (dual oxidation possible) | β-sandwich fold; higher structural plasticity; maintains histidine brace; adaptable binding surfaces | Poorly characterized in fungi; functional versatility seen in arthropods suggests potential for broad substrate recognition | Ecological role in fungi remains unclear; may be involved in specific symbiotic or pathogenic interactions, or degradation of insect-derived biomass | No robust biochemical characterizations have been published yet confirming the expression, enzymatic activity, and specific substrate of AA15 in fungi. In most databases, such as CAZy and JGI MycoCosm, AA15 are often annotated as “putative” or “hypothetical” enzymes in fungi | [25,66,67] |
AA16 | Fungi, viruses, oomycetes | Cellulose, chitin | C1 | β-sandwich fold; monocopper active site | Acts primarily on cellulose; C1 oxidation predominant; works synergistically with cellulases | Contributes to cellulose breakdown; improves saccharification yields in industrial enzyme blends | Peniophora sp., Pleurotus eryngii, Thermothelomyces thermophilus | [26,68,69,70,71] |
Aspect | O2-Driven Pathway | H2O2-Driven Pathway |
---|---|---|
Speed (Turnover Rate) | Slow | Fast (10–100× faster) |
Electron Requirement | Two electrons per cycle | One electron per cycle |
Control Complexity | Lower (less prone to ROS damage) | Higher (requires strict H2O2 control) |
Risk of Inactivation | Minimal | High if H2O2 is in excess |
Physiological Prevalence | Secondary, backup under low H2O2 conditions | Dominant under most natural biomass degradation |
Evolutionary Implication | Possibly ancestral or for oxygen-rich conditions | Adaptation to lignocellulose-rich microenvironments |
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Contato, A.G.; Conte-Junior, C.A. Fungal Lytic Polysaccharide Monooxygenases (LPMOs): Functional Adaptation and Biotechnological Perspectives. Eng 2025, 6, 177. https://doi.org/10.3390/eng6080177
Contato AG, Conte-Junior CA. Fungal Lytic Polysaccharide Monooxygenases (LPMOs): Functional Adaptation and Biotechnological Perspectives. Eng. 2025; 6(8):177. https://doi.org/10.3390/eng6080177
Chicago/Turabian StyleContato, Alex Graça, and Carlos Adam Conte-Junior. 2025. "Fungal Lytic Polysaccharide Monooxygenases (LPMOs): Functional Adaptation and Biotechnological Perspectives" Eng 6, no. 8: 177. https://doi.org/10.3390/eng6080177
APA StyleContato, A. G., & Conte-Junior, C. A. (2025). Fungal Lytic Polysaccharide Monooxygenases (LPMOs): Functional Adaptation and Biotechnological Perspectives. Eng, 6(8), 177. https://doi.org/10.3390/eng6080177