Abstract
Lytic polysaccharide monooxygenases (LPMOs) are a class of copper-dependent enzymes that can act on crystalline polysaccharides directly, which plays a critical role in cellulose degradation. In addition to reports on its structure and mechanism of action, it is important to study the auxiliary activity 9 (AA9) characteristics from different resources to support the mechanism research. The gene encoded ToLPMO9A was cloned from Trichoderma orientalis EU7-22 and first heterologously expressed in Pichia pastoris GS115. Both metal ions and reducing agent concentrations showed an important effect on ToLPMO9A. The ToLPMO9A exhibited maximal activity at 60 °C and a 6.0 pH. In addition, ToLPMO9A showed substrate specificity. The matrix-assisted laser desorption ionization-time of flight-mass spectrometry (MALDI-TOF-MS) analysis showed that ToLPMO9A cleaved the glycosidic bonds at C1 and C4/C6 position via oxidation. Concerning the synergistic effects on enzymic activity, ToLPMO9A exhibited promotion with endo-glucanase or exo-glucanase, but inhibition with β-glucosidases. In conclusion, ToLPMO9A could be a good choice for enzyme cocktails and provide theoretical support for subsequent action mechanisms and broader applications.
Download PDF
Full Article
Recombinant Expression of Lytic Polysaccharide Monooxygenase and its Functional Characterization
Jin Huang, Yong Xue, Jiang Han, Jian Liu,* Lihui Gan,* and Minnan Long
Lytic polysaccharide monooxygenases (LPMOs) are a class of copper-dependent enzymes that can act on crystalline polysaccharides directly, which plays a critical role in cellulose degradation. In addition to reports on its structure and mechanism of action, it is important to study the auxiliary activity 9 (AA9) characteristics from different resources to support the mechanism research. The gene encoded ToLPMO9A was cloned from Trichoderma orientalis EU7-22 and first heterologously expressed in Pichia pastoris GS115. Both metal ions and reducing agent concentrations showed an important effect on ToLPMO9A. The ToLPMO9A exhibited maximal activity at 60 °C and a 6.0 pH. In addition, ToLPMO9A showed substrate specificity. The matrix-assisted laser desorption ionization-time of flight-mass spectrometry (MALDI-TOF-MS) analysis showed that ToLPMO9A cleaved the glycosidic bonds at C1 and C4/C6 position via oxidation. Concerning the synergistic effects on enzymic activity, ToLPMO9A exhibited promotion with endo-glucanase or exo-glucanase, but inhibition with β-glucosidases. In conclusion, ToLPMO9A could be a good choice for enzyme cocktails and provide theoretical support for subsequent action mechanisms and broader applications.
Keywords: Lytic polysaccharide monooxygenases; Trichoderma orientalis EU7-22; Substrate specificity; Regioselectivity; Synergistic cooperation
Contact information: College of Energy, Xiamen University, Xiamen 361005.P.R. China;
* Corresponding authors: jianliu@xmu.edu.cn (Jian Liu), ganlihui@xmu.edu.cn (Lihui Gan)
INTRODUCTION
Lignocellulose is the richest renewable resource and widely distributed form of crop and forestry resources on the planet (Jönsson et al. 2013). It can provide renewable resources for the production of biofuels, chemicals, and polymers. However, due to the lack of effective technologies, burning is still the traditional method of utilization, which not only causes resource waste but aggravates the environmental pollution problem as well. Therefore, it is important to find ways to utilize it (Rodionova et al. 2017). Biochemical conversion of lignocellulose preserves the original carbohydrate structure in the form of monomeric sugars (as opposed to thermochemical conversion, which destroys carbohydrates). Enzymatic techniques are usually considered to be the key point of saccharification technology (Horn et al. 2012). Despite a lot of work that has been done over the past decade, the enzymatic hydrolysis efficiency of lignocellulosic biomass remains a critical limiting step in many biorefining methods (Klein-Marcuschamer et al. 2012). The limiting factor is the heterogeneity of plant cell walls (mainly cellulose, hemicellulose, and lignin) (Chundawat et al. 2011), and the inaccessibility and recalcitrance of its components. Thus, the enzymatic hydrolysis process is inefficient and costly (Lynd et al. 2008).
The classic scheme for cellulose degradation involves three enzymes, endo-β-1,4-glucanase, exo-β-1,4-glucanase, and β-glucosidases (Horn et al. 2006; Payne et al. 2011). These enzymes work synergistically, as the endoglucanase generates new reduced and non-reducing chain ends for exoglucanase and releases cellobiose, which is converted to glucose by β-glucosidase (Kostylev and Wilson 2012; Wood and McCrae 1979). Unlike the traditional way of cellulase mentioned above, LPMOs directly cleave glycosidic bonds in an oxidative manner to generate non-reducing chain ends and reducing chain ends, which provides more accessible sites for cellulase. This synergistic reaction can greatly improve the hydrolysis efficiency (Dimarogona et al. 2013; Gouvêa et al. 2019; Zhang et al. 2019). It has completely changed the enzymatic processing of polysaccharides (Chylenski et al. 2019).
Due to its glycoside hydrolase activity, the AA9 family was defined as the GH61 (glycoside hydrolase) family. However, compared to other endocellulases derived from the same genus, the hydrolytic activity of the GH61 family is notably lower than other enzymes (Karlsson et al. 2001). Also, the crystal structure of GH61 family enzymes is different from the glycoside hydrolases structure. The GH family structure does not contain the clustering of conserved catalytic acidic side chains, which constitutes the classic catalytic mechanism of this class of proteins (Karkehabadi et al. 2008; Harris et al. 2010). In 2010, the GH61 family was first demonstrated as a cellulose-boosting enzyme to meaningfully improve hydrolysis efficiency (Harris et al. 2010). Since then, additional studies have demonstrated that there are both oxidized and non-oxidized products in the hydrolysate. Thus, the GH61 family is considered as a polysaccharide monooxygenase (Quinlan et al. 2011; Westereng et al. 2011; Beeson et al. 2012). In 2012, the GH61 family was reclassified as lytic polysaccharide monooxygenases (LPMO). In 2013, LPMO and lignin-degrading enzymes were classified into one group and constitute new carbohydrate active enzymes named “Auxiliary Activities”. In addition, the LPMOs are distributed in seven Auxiliary Activity families in CAZy database (www.cazy.org), with various origins, including eukaryota, bacteria, virus and archaea. LPMO-encoding genes are highly abundant in nature, and LPMOs are predicted to play a significant role in global carbon flux (Frandsen and Lo Leggio 2016). Therefore, there are several reviews on LPMO (Frommhagen et al. 2018b; Zhang 2020; Zhou and Zhu 2020), such as the substrate specificity, regioselectivity, electron-donating systems and the application. Besides, the GH61 family was reclassified into the AA9 family (Levasseur et al. 2013).
Currently, there is limited public data that addresses the possible relationship between the structural characteristics of LPMOs and the regioselectivity on C1/C4/C6 (Frandsen and Lo Leggio 2016; Vaaje-Kolstad et al. 2017). Therefore, more LPMO-substrate interactions must be found to improve the understanding of the structure-function relationship of LPMOs. A large number of biochemical, structural, functional, and regiospecific data show that the research of LPMO in fungi mainly focuses on the AA9 family (Payne et al. 2011; Phillips et al. 2011; Aachmann et al. 2012; Beeson et al. 2012; Kittl et al. 2012; Bey et al. 2013; Wu et al. 2013; Vu et al. 2014).
In this study, the AA9 LPMO, the gene encoding ToLPMO9A, was first cloned from Trichoderma orientalis EU7-22 and was heterologous expressed in Pichia pastoris GS115. The enzymatic properties, substrate specificity, and regioselectivity of ToLPMO9A were investigated. Meanwhile, the synergistic actions with three cellulases were studied.
EXPERIMENTAL
Methods
Strains and enzymes
The T. orientalis EU7-22 was used as the source of the ToLPMO9A gene. Escherichia coli DH5α (Invitrogen, Carlsbad, CA, USA) was used to construct and amplify the recombinant plasmids, and P. pastoris GS115 (Invitrogen) was used for the heterologous expression of the recombinant ToLPMO9A. Commercial cellulase was purchased from Novozymes (Nanjing, China). The recombinant enzymes EGII (endo-β-1,4-glucanase), CBHII (exo-β-1,4-glucanase), and BGLI (β-glucosidases) from T. orientalis EU7-22 were expressed in P. pastoris and preserved in the laboratory. The culture environment of the plate was 30 °C, and the liquid medium was incubated in the rotary shaker at 30 °C and 180 rpm.
Cloning of ToLPMO9A gene
The frozen spore powder of T. orientalis EU7-22 was stored at -70 °C and inoculated into the potato dextrose agar (PDA) plate and incubated for 4 days. The strain colony was then injected into the PDA liquid medium and incubated for 24 h. The mycelia suspensions above were collected and incubated in the inducing medium (Xue et al. 2020). After induction culture for 48 h, the total RNA was isolated using an RNA prep pure plant kit (DP432, Tiangen, Beijing, China), and then it was reverse-transcribed into cDNA by polymerase chain reaction (PCR). The ToLPMO9A gene was cloned from T. orientalis EU7-22 cDNA with a pair of primers by PCR. The forward primer was 5′-CCGGAATTCCGGACACATCAACAACATTG-3′, and the reverse primer was 5’-AAGGAAAAAAGCGGCCGCCTAGCTAAGGCACTGGGCATAG-3′ (the restriction endonuclease sites EcoRI and NotI were underlined).
The PCR product of the ToLPMO9A gene and the expression vector pPIC9K (Invitrogen) were both digested by the EcoRI and NotI. The gene was then ligated to the pPIC9K to generate pPIC9K-ToLPMO9A plasmids. The recombinant plasmid was transformed into E. coli DH5α, and the correct transformants were identified by PCR and DNA sequencing.
Expression of ToLPMO9A in P. pastoris and enzyme purification
The pPIC9K-ToLPMO9A recombinant plasmid was extracted by the TIANprep midi plasmid kit (Tiangen, Beijing, China). The plasmid pPIC9K-ToLPMO9A was linerized by Bgl I and then electrotransformed into P. pastoris GS115. After 3 days of culture on minimal dextrose (MD) medium (1.34% yeast nitrogen base without amino acids, 2% glucose) plates, the transformants were verified by PCR with a sequencing primer, 5’AOX1 (5′-GACTGGTTCCAATTGACAAGC-3′), and 3’AOX1 (5′- GGCAAATGGCATTCTGACAT-3′). The positive recombinants were inoculated into the BMGY medium until the culture reached an optical density of 2.0 to 6.0 at 600 nm (OD600). The yeast cells were collected by centrifugation (4 °C, 6000 × g, 10 min) from the culture medium. The cells were resuspended in the BMMY medium, and methanol was added daily, while keeping the concentration at 1%. The BMMY medium was centrifuged. The fermentation broth was collected by centrifugation (12000 × g, 5 min) to collect the fermentation broth, which was stored at -20 °C for later use.
The fermentation supernatant was concentrated in a 10 kDa Millipore ultrafiltration centrifuge tube and loaded on a dextran G-50 gel filtration column. Afterwards, the culture supernatant was eluted with a phosphate buffer at a flow rate of 0.6 mL/min. The recombinant protein expression was verified with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and the separation gel concentration and concentrated gel were 13% and 5%, respectively.
Substrate specificity
Five kinds of substrate (microcrystalline cellulose, CMC, corn cob, filter paper, beechwood xylan) were used for the experiments for substrate specificity of ToLPMO9A.
The components in the reaction system were composed of 0.5% (w/v) substrate, 10 mmol/L NH4Ac-HAc (pH5.0), 1 mmol/L ascorbic acid, 1 mmol/L CuSO4, and 5 µmol/L ToLPMO9A enzyme. The reaction system was incubated at 50 °C and 200 rpm for 48 h, and the reaction was terminated by a boiling water bath.
The supernatant of all incubations was obtained by centrifugation (4 °C, 6000 × g, 10 min). The enzyme activity was further determined by measuring the reducing sugar concentration of the reaction solution using the reported method (Miller 1959). All reactions were performed in triplicate.
Effect of reducing agent and metal ions concentration
To study the effect of reducing agents on ToLPMO9A, different ascorbic acid concentrations were added to the enzymatic reaction system from 1 mmol/L to 10 mmol/L.
To study the effect of metal ions on ToLPMO9A, five metal ions (Cu2+, Mg2+, Co2+, Mn2+, Ca2+, and Fe3+) were added to the enzymatic reaction system at the terminal concentration of 1 mmol/L and 5 mmol/L.
The components in the reaction system were the same as above, and the substrate used in the reaction was microcrystalline cellulose.
Effects of temperature and pH on the stability of ToLPMO9A
To study the effect of temperature on ToLPMO9A, the enzyme activity was measured at 30 °C, 40 °C, 50 °C, 60 °C, 70 °C, 80 °C, and 90 °C, and the highest enzyme activity was 100%.
To study the effect of pH on ToLPMO9A, the reaction was incubated with a buffer with a range of 3.0 to 9.0, and the highest enzyme activity was 100%.
The components in the reaction system were the same as above, and the substrate used in the reaction was microcrystalline cellulose.
Mass spectrometry
The oxidation product type was identified by matrix-assisted laser desorption ionization-time of flight-mass spectrometry (MALDI-TOF-MS). The substrate used in mass spectrometry was phosphoric acid swollen cellulose (PASC), which was prepared from Avicel PH-101 (Sigma, St. Louis, MO, USA) following instructions from Wood (1988). The reaction solution for enzymatic hydrolysis and the test conditions followed previous reported Vaaje-Kolstad et al. (2010).
The analysis was completed on a MALDI-TOF 5800 (AB SCIEX, Singapore). For the MALDI-TOF-MS analysis, the instrument operated in a linear mode and acquired spectra in the range of 0 to 7000 m/z.
Enzyme synergistic hydrolysis
To study the synergistic effect of ToLPMO9A enzymes, three former studied cellulases were introduced in this research: EGII, CBHII, and BGLI. They were cloned from EU7-22 and heterologously expressed in P. pastoris GS115.
The supernatant of all incubations was obtained by centrifugation (4 °C, 6000 × g, 10 min). The experiment of synergistic hydrolysis followed the method from Zhu et al. (2018). The synergistic system was incubated at 60 °C, pH 6.0 and 200 rpm for 48 h, and the reaction was terminated by a boiling water bath.
To further understand the synergistic mechanism, three formerly investigated enzymes in the cellulase degradation system were selected in this study. As a parameter, DS (degree of synergism) was employed to describe the effect of synergism, as shown in Eq. 1,
(1)
where DS means the degree of synergism, Cab is the reducing sugar concentration obtained from the combined system with both ToLPMO9A and cellulase, Ca is that from the simple system with ToLPMO9A only, and Cb is that from the simple system only containing cellulase.
RESULTS AND DISCUSSION
Cloning of ToLPMO9A Gene
Using T. orientalis EU7-22 as a template, the genes were amplified by PCR with the primers given above. The results on agarose gel electrophores is shown in Fig. 1.
Fig. 1. Cloning of the gene encoding ToLPMO9A from T. orientalis EU7-22; Lane M: DNA marker; Lane 1: cDNA (1044 bp)
Figure 1 indicates that the full length of the ToLPMO9A gene cDNA sequence including the removed signal peptide sequence was 981 bp. The obtained sequence was compared with the existing ToLPMO9A in the entire genome sequence of EU7-22 by DNAMAN, and the similarity was 100%, which proved that the cDNA sequence of ToLPMO9A in EU7-22 was successfully cloned.
Expression of ToLPMO9A in P. pastoris
The ToLPMO9A expressed in P. pastoris was analyzed by SDS-PAGE. The theoretical molecular weight of ToLPMO9A was 33 kDa, while the molecular weight was approximately 68 kDa (Fig. 2) due to the over glycosylation of ToLPMO9A in P. pastoris.
The result of function domain analysis shows that ToLPMO9A have the carbohydrate-binding module (CBM) domain, which have been reported for both LPMO9s and LPMO10s (Forsberg et al. 2016; Courtade et al. 2018; Chalak et al. 2019; Laurent et al. 2019).
Fig. 2. SDS-PAGE of ToLPMO9A; Lane M: protein marker; Lane 1: unpurifiedToLPMO9A; and Lane 2: purified ToLPMO9A
Substrate Specificity
There are multiple excurrent loops in LPMOs that could shape the binding surface of the substrate to recognize and bind polysaccharides. The substrate specificity of AA9 was determined by the loops, which contained various hydrophilic and aromatic residues related to the substrate-binding of LPMO (Aachmann et al. 2012; Wu et al. 2013; Borisova et al. 2015).
When ToLPMO9A acted on different substrates, the reducing sugar yield of the reaction system was also different. As shown in Fig. 3, ToLPMO9A can lead to the catalytic degradation of four substrates: microcrystalline cellulose, CMC, corn cob, and xylan. Furthermore, the reducing sugar yield was the highest when microcrystalline cellulose was used as the substrate, followed by the corn cob. In addition, ToLPMO9A was active with xylan, which was the same as previous research (Jagadeeswaran et al. 2016; Ladevèze et al. 2017; Simmons et al. 2017).
Fig. 3. Reducing sugar yields from cellulosic specimens treated with ToLPMO9A
Effect of Reducing Agent and Metal Ions Concentration
In 2010, a study that revealed the oxidative mechanism behind LPMO activity showed that LPMO requires electrons (Vaaje-Kolsatand et al. 2010). To catalyze the oxidation of polysaccharides, LPMO relies on reducing agent, which provides electrons to reduce copper in the active site and activate molecular oxygen (Frandsen and Lo Leggio 2016). The effect of the reducing agent concentration on ToLPMO9A was investigated. There was an increase in enzymatic activity when the ascorbic acid concentration increased from 1 to 10 mmol/L (Fig. 4a). Moreover, when the ascorbic acid concentration reached 10 mmol/L, the conversion amount of reducing sugar increased by 6.46 times.
Fig. 4. Effect of reducing agent and metal ions on enzyme activity: (a) effect of reducing agent on ToLPMO9A activity; (b) effect of metal ions on ToLPMO9A activity
The effect of the metal ion concentrations on ToLPMO9A was investigated. Except for the reaction with Fe3 +, the enzyme activity increased in all reactions with various metals (Fig. 4b). In particular, the metal ions Co2+ and Cu2+ notably promoted substrate degradation at low concentrations. At a concentration of 5 mmol/L, the increasing amount of reducing sugar was 1.65 and 1.97, respectively, indicating that ToLPMO9A is a metal-dependent enzyme. The results in this research are consistent with the previous report (Naghshehechi 2014).
Effects of Temperature and pH on the Stability of ToLPMO9A
The relative enzyme activity of ToLPMO9A at different temperatures is shown in Fig. 5a. The results show that the optimal reaction temperature of ToLPMO9A was 60 °C. In the range of 30 °C to 60 °C, the relative enzyme activity of ToLPMO9A gradually increased with an increase of reaction temperature. At a temperature higher than 60 °C, the relative enzyme activity of ToLPMO9A decreased rapidly, while at 90 °C, ToLPMO9A was almost completely inactivated.
There have been few studies on the effect of pH on LPMO. According to previous studies, the pH preference of different sources of LPMO is different. For example, LPMO from Pestalotiopsis sp. has the highest activity at pH 5.5 (Patel et al. 2016), and the LPMO from Gloeophyllum trabeum is the most active at pH 7.0 (Hegnar et al. 2018). The optimum pH was 6.0 for ToLPMO9A, which is consistent with the preference pH of T. orientalis EU7-22 (Fig. 5b).
Regioselectivity Identification by Mass Spectrometry
The oxidization acting sites by AA9 LPMOs are located on either C1 (Frommhagen et al. 2016, 2017; Liu et al. 2018), C 4 (Frandsen et al. 2016; Liu et al. 2017) or both the C1 and C4 position (Vu et al. 2014; Song et al. 2018), and occasionally the C6 position (Chen et al. 2018).
Fig. 5. Effect of temperature and pH on enzyme activity: (a) effect of temperature on ToLPMO9A activity; (b) effect of pH on ToLPMO9A activity
Figure 6 shows the results of the MALDI-TOF-MS analysis on the PASC degradation products by ToLPMO9A. According to the mass spectrometry analysis, the product contained sodium adducts of oligosaccharides with the degree of polymerization (DPs) ranging from 3 to 6, and the m/z ratios of 527, 689, 851, and 1013. In addition to the various non-oxidized oligosaccharides, the mass spectrometry results also contained oxidized oligosaccharides. The oxidation peaks at the m/z values of 543, 705, 867, and 1029 correspond to the sodium adducts of C1 oxidized oligosaccharides (m/z + 16). The peaks at m/z of 525, 687, 849, and 1011 correspond to the sodium adducts of C4 oxidized oligosaccharides (C4-ketoaldose, C6-hexodialdose, m/z-2). Accordingly, ToLPMO9A cleaved the polysaccharide chain of cellulose at both the C1 and C4/C6 positions. More detailed information on its cleavage pattern is reported in the references below (Frommhagen et al. 2018a).
For LPMO application in the renewable biomass industry, more enzymatic and structural characterizations of LPMO from different sources are needed to clarify the substrate specificity and oxidative regioselectivity.
Fig. 6. MALDI-TOF-MS analysis of the degradation products by ToLPMO9A
Synergistic Hydrolysis by ToLPMO9A with Three Cellulases
The LPMOs have received widespread attention since their discovery due to their unique oxidation mechanism and their effective contribution to the cellulase activity (Vaaje-Kolstad et al. 2010; Aachmann et al. 2012).
The degradation ability of three enzymes were increased to varying degrees after the addition of ToLPMO9A (Fig. 7). The initial reducing sugar concentration was 0.950 mg/mL and 0.096 mg/mL for the EGII and CBHII enzymes, respectively. The reducing sugar concentration increased to 2.56 and 0.32 mg/mL for the system with ToLPMO9A and the DS were 2.27 and 1.16, respectively. For BGLI enzymes, the initial reducing sugar concentrations were 0.47 mg/mL, but with the addition of ToLPMO9A, the reducing sugar concentrations were 0.55 mg/mL and the DS was 0.9. Therefore, for both EGII and CBHII, ToLPMO9A exhibited positive synergistic effects, but the negative effect was shown in the synergism of ToLPMO9A and BGLI.
It is speculated that this synergy is due to the oxidative cleavage of polysaccharide crystalline regions by LPMOs, which can bring more accessible sites for glycoside hydrolases (Vermaas et al. 2015) as EGII and CBHII enzymes. Concurrently, the EGII and CBHII enzymes degraded the polysaccharide chains and unlocked new binding sites to contribute to the LPMO reaction (Bissaro et al. 2017). In the following experiments, it is possible to adjust the ratio of enzymes and substrates so as to obtain more suitable operating conditions for industrial application. Therefore, AA9 can meaningfully increase the EGII and CBHII enzyme activity, and it can improve the synergy between AA9 and EGII/CBHII. However, as noted previously, the oxidization at the C1 position by AA9 produces gluconic acid that inhibits glucosidase activity (Hemsworth et al. 2015). Thus, the main interactions between BGL and ToLPMO9A finally resulted in inhibition.
Fig. 7. Synergistic hydrolysis of PASC by ToLPMO9A with three cellulases
CONCLUSIONS
- An LPMO gene (ToLPMO9A) was cloned from T. orientalis EU7-22 and it was heterologously expressed in Pichia pastoris GS115 successfully. ToLPMO9A exhibited maximal activity at 60 °C and pH 6.0.
- Both metal ions (Co2+, Cu2+) and reducing agent (ascorbic acid) concentrations showed a noteworthy effect on ToLPMO9A. The ToLPMO9A could degrade microcrystalline cellulose, corn cob, CMC and xylan, and the enzymatic activity on them were different. The MALDI-TOF-MS analysis showed that the ToLPMO9A cleaved the glycosidic bonds at C1 and C4/C6 positions oxidatively.
- The ToLPMO9A exhibited positive synergistic effects with EGII or CBHII, and DS were 2.27 and 1.16, while ToLPMO9A exhibited inhibitory effects with BGLI.
ACKNOWLEDGMENTS
The authors gratefully acknowledge the financial support from National Natural Science Foundation of China (No. 21978249) and Xiamen Science and Technology Plan Project (No. 3502Z20193022).
REFERENCES CITED
Aachmann, F. L., Sørlie, M., Skjåk-Bræk, G., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2012). “NMR structure of a lytic polysaccharide monooxygenase provides insight into copper binding, protein dynamics, and substrate interactions,” Proceedings of the National Academy of Sciences 109(46), 18779-18784. DOI: 10.1073/pnas.1208822109
Beeson, W. T., Phillips, C. M., Cate, J. H. D., and Marletta, M. A. (2012). “Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases,” Journal of the American Chemical Society 134(2), 890-892. DOI: 10.1021/ja210657t
Bey, M., Zhou, S., Poidevin, L., Henrissat, B., Coutinho, P. M., Berrin, J.-G., and Sigoillot, J.-C. (2013). “Cello-oligosaccharide oxidation reveals differences between two lytic polysaccharide monooxygenases (family GH61) from Podospora anserina,” Applied and Environmental Microbiology 79(2), 488-496. DOI: 10.1128/AEM.02942-12
Bissaro, B., Røhr, Å. K., Müller, G., Chylenski, P., Skaugen, M., Forsberg, Z., Horn, S. J., Vaaje-Kolstad, G., and Eijsink, V. G. H. (2017). “Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2,” Nature Chemical Biology 13(10), 1123. DOI: 10.1038/nchembio.2470
Borisova, A. S., Isaksen, T., Dimarogona, M., Kognole, A. A., Mathiesen, G., Várnai, A., Røhr, Å. K., Payne, C. M., Sørlie, M., Sandgren, M., and Eijsink, V. G. H. (2015). “Structural and functional characterization of a lytic polysaccharide monooxygenase with broad substrate specificity,” The Journal of Biological Chemistry 290(38), 22955-22969. DOI: 10.1074/jbc.M115.660183
Chalak, A., Villares, A., Moreau, C., Haon, M., Grisel, S., d’Orlando, A., Herpoël-Gimbert, I., Labourel, A., Cathala, B., and Berrin, J.-G. (2019). “Influence of the carbohydrate-binding module on the activity of a fungal AA9 lytic polysaccharide monooxygenase on cellulosic substrates,” Biotechnology for Biofuels, 12(1), 206. DOI: 10.1186/s13068-019-1548-y
Chen, C., Chen, J., Geng, Z., Wang, M., Liu, N., and Li, D. (2018). “Regioselectivity of oxidation by a polysaccharide monooxygenase from Chaetomium thermophilum,” Biotechnology for Biofuels 11(1), 155. DOI: 10.1186/s13068-018-1156-2
Chundawat, S. P. S., Beckham, G. T., Himmel, M. E., and Dale, B. E. (2011). “Deconstruction of lignocellulosic biomass to fuels and chemicals,” Annual Review of Chemical and Biomolecular Engineering 2(1), 121-145. DOI: 10.1146/annurev-chembioeng-061010-114205
Chylenski, P., Bissaro, B., Sørlie, M., Røhr, Å. K., Várnai, A., Horn, S. J., and Eijsink, V. G. H. (2019). “Lytic polysaccharide monooxygenases in enzymatic processing of lignocellulosic biomass,” ACS Catalysis 9(6), 4970-4991. DOI: 10.1021/acscatal.9b00246
Courtade, G., Forsberg, Z., Heggset, E. B., Eijsink, V. G. H., and Aachmann, F. L. (2018). “The carbohydrate-binding module and linker of a modular lytic polysaccharide monooxygenase promote localized cellulose oxidation,” Journal of Biological Chemistry, American Society for Biochemistry and Molecular Biology, 293(34), 13006-13015. DOI: 10.1074/jbc.RA118.004269
Dimarogona, M., Topakas, E., and Christakopoulos, P. (2013). “Recalcitrant polysaccharide degradation by novel oxidative biocatalysts,” Applied Microbiology and Biotechnology, 97(19), 8455-8465. DOI: 10.1007/s00253-013-5197-y
Forsberg, Z., Nelson, C. E., Dalhus, B., Mekasha, S., Loose, J. S. M., Crouch, L. I., Røhr, Å. K., Gardner, J. G., Eijsink, V. G. H., and Vaaje-Kolstad, G. (2016). “Structural and functional analysis of a lytic polysaccharide monooxygenase important for efficient utilization of chitin in Cellvibrio japonicus,” Journal of Biological Chemistry, American Society for Biochemistry and Molecular Biology, 291(14), 7300-7312. DOI: 10.1074/jbc.M115.700161
Frandsen, K. E. H., and Lo Leggio, L. (2016). “Lytic polysaccharide monooxygenases: A crystallographer’s view on a new class of biomass-degrading enzymes,” International Union of Crystallography 3(6), 448-467. DOI: 10.1107/S2052252516014147
Frandsen, K. E. H., Simmons, T. J., Dupree, P., Poulsen, J. N., Hemsworth, G. R., Ciano, L., Johnston, E. M., Tovborg, M., Johansen, K. S., Freiesleben, P., et al. (2016). “The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases,” Nature Chemical Biology 12(4), 298-303. DOI: 10.1038/nchembio.2029
Frommhagen, M., Koetsier, M. J., Westphal, A. H., Visser, J., Hinz, S. W. A., Vincken, J., van Berkel, W. J. H., Kabel, M. A., and Gruppen, H. (2016). “Lytic polysaccharide monooxygenases from Myceliophthora thermophila C1 differ in substrate preference and reducing agent specificity,” Biotechnology for Biofuels 9, 186. DOI: 10.1186/s13068-016-0594-y
Frommhagen, M., Mutte, S. K., Westphal, A. H., Koetsier, M. J., Hinz, S. W. A., Visser, J., Vincken, J.-P., Weijers, D., van Berkel, W. J. H., Gruppen, H., and Kabel, M. A. (2017). “Boosting LPMO-driven lignocellulose degradation by polyphenol oxidase-activated lignin building blocks,” Biotechnology for Biofuels 10, 121. DOI: 10.1186/s13068-017-0810-4
Frommhagen, M., Westphal, A. H., Hilgers, R., Koetsier, M. J., Hinz, S. W. A., Visser, J., Gruppen, H., van Berkel, W. J. H., and Kabel, M. A. (2018a). “Quantification of the catalytic performance of C1-cellulose-specific lytic polysaccharide monooxygenases,” Applied Microbiology and Biotechnology, 102(3), 1281-1295. DOI: 10.1007/s00253-017-8541-9
Frommhagen, M., Westphal, A. H., van Berkel, W. J. H., and Kabel, M. A. (2018b). “Distinct substrate specificities and electron-donating systems of fungal lytic polysaccharide monooxygenases,” Frontiers in Microbiology 9, 1080. DOI: 10.3389/fmicb.2018.01080
Gouvêa, P. F. de, Gerolamo, L. E., Bernardi, A. V., Pereira, L. M. S., and Dinamarco*, S. A. U. and T. M. (2019). “Lytic polysaccharide monooxygenase from Aspergillus fumigatus can improve enzymatic cocktail activity during sugarcane bagasse hydrolysis,” Protein & Peptide Letters, 26(5), 377-389. DOI: 10.2174/0929866526666190228163629
Harris, P. V., Welner, D., McFarland, K. C., Re, E., Navarro Poulsen, J., Brown, K., Salbo, R., Ding, H., Vlasenko, E., et al. (2010). “Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: Structure and function of a large, enigmatic family,” Biochemistry 49(15), 3305-3316. DOI: 10.1021/bi100009p
Hegnar, O. A., Petrovic, D. M., Bissaro, B., Alfredsen, G., Várnai, A., and Eijsink, V. G. H. (2018). “pH-dependent relationship between catalytic activity and hydrogen peroxide production shown via characterization of a lytic polysaccharide monooxygenase from Gloeophyllum trabeum,” Applied and Environmental Microbiology 85(5), e02612-18. DOI: 10.1128/AEM.02612-18
Hemsworth, G. R., Johnston, E. M., Davies, G. J., and Walton, P. H. (2015). “Lytic polysaccharidemonooxygenases in biomass conversion,” Trends in Biotechnology 33(12), 747–761. DOI: 10.1016/j.tibtech.2015.09.006
Henrissat, B. (1991). “A classification of glycosyl hydrolases based on amino acid sequence similarities,” Biochemical Journal 280(2), 309-316. DOI: 10.1042/bj2800309
Horn, S. J., Vaaje-Kolstad, G., Westereng, B., and Eijsink, V. (2012). “Novel enzymes for the degradation of cellulose,” Biotechnology for Biofuels 5, 45. DOI: 10.1186/1754-6834-5-45
Jagadeeswaran, G., Gainey, L., Prade, R., and Mort, A. J. (2016). “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,” Applied Microbiology and Biotechnology 100(10), 4535-4547. DOI: 10.1007/s00253-016-7505-9
Jönsson, L. J., Alriksson, B., and Nilvebrant, N. (2013). “Bioconversion of lignocellulose: Inhibitors and detoxification,” Biotechnology for Biofuels 6, 16. DOI: 10.1186/1754-6834-6-16
Karkehabadi, S., Hansson, H., Kim, S., Piens, K., Mitchinson, C., and Sandgren, M. (2008). “The first structure of a glycoside hydrolase family 61 member, Cel61B from Hypocrea jecorina, at 1.6 A resolution,” Journal of Molecular Biology 383(1), 144-154. DOI: 10.1016/j.jmb.2008.08.016
Karlsson, J., Saloheimo, M., Siika‐aho, M., Tenkanen, M., Penttilä, M., and Tjerneld, F. (2001). “Homologous expression and characterization of Cel61A (EG IV) of Trichoderma reesei,” European Journal of Biochemistry 268(24), 6498-6507. DOI: 10.1046/j.0014-2956.2001.02605.x
Kittl, R., Kracher, D., Burgstaller, D., Haltrich, D., and Ludwig, R. (2012). “Production of four Neurospora crassa lytic polysaccharide monooxygenases in Pichia pastoris monitored by a fluorimetric assay,” Biotechnology for Biofuels 5(1), 79. DOI: 10.1186/1754-6834-5-79
Klein-Marcuschamer, D., Oleskowicz-Popiel, P., Simmons, B. A., and Blanch, H. W. (2012). “The challenge of enzyme cost in the production of lignocellulosic biofuels,” Biotechnology and Bioengineering 109(4), 1083-1087. DOI: 10.1002/bit.24370
Kostylev, M., and Wilson, D. (2012). “Synergistic interactions in cellulose hydrolysis,” Biofuels 3(1), 61-70. DOI: 10.4155/bfs.11.150
Ladevèze, S., Haon, M., Villares, A., Cathala, B., Grisel, S., Herpoël-Gimbert, I., Henrissat, B., and Berrin, J. (2017). “The yeast Geotrichum candidum encodes functional lytic polysaccharide monooxygenases,” Biotechnology for Biofuels 10, 215. DOI: 10.1186/s13068-017-0903-0
Laurent, C. V. F. P., Sun, P., Scheiblbrandner, S., Csarman, F., Cannazza, P., Frommhagen, M., van Berkel, W. J. H., Oostenbrink, C., Kabel, M. A., and Ludwig, R. (2019). “Influence of lytic polysaccharide monooxygenase active site segments on activity and affinity,” International Journal of Molecular Sciences, Multidisciplinary Digital Publishing Institute, 20(24), 6219. DOI: 10.3390/ijms20246219
Levasseur, A., Drula, E., Lombard, V., Coutinho, P. M., and Henrissat, B. (2013). “Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes,” Biotechnology for Biofuels 6, 41. DOI: 10.1186/1754-6834-6-41
Liu, B., Kognole, A. A., Wu, M., Westereng, B., Crowley, M. F., Kim, S., Dimarogona, M., Payne, C. M., and Sandgren, M. (2018). “Structural and molecular dynamics studies of a C1-oxidizing lytic polysaccharide monooxygenase from Heterobasidion irregulare reveal amino acids important for substrate recognition,” The FEBS Journal 285(12), 2225-2242. DOI: 10.1111/febs.14472
Liu, B., Olson, Å., Wu, M., Broberg, A., and Sandgren, M. (2017). “Biochemical studies of two lytic polysaccharide monooxygenases from the white-rot fungus Heterobasidion irregulare and their roles in lignocellulose degradation,” PLOS ONE 12(12), e0189479. DOI: 10.1371/journal.pone.0189479
Lynd, L. R., Laser, M. S., Bransby, D., Dale, B. E., Davison, B., Hamilton, R., Himmel, M., Keller, M., McMillan, J. D., Sheehan, J., et al. (2008). “How biotech can transform biofuels,” Nature Biotechnology 26(2), 169-172. DOI: 10.1038/nbt0208-169
Miller, G. L. (1959). “Use of dinitrosalicylic acid reagent for determination of reducing sugar,” Analytical Chemistry 31(3), 426-428. DOI: 10.1021/ac60147a030
Naghshehechi, K. (2014). The Influence of Divalent Metal Ions on a Lytic Polysaccharide Monooxygenase, Master’s Thesis, Norwegian University of Life Sciences, Ås, Norway.
Patel, I., Kracher, D., Ma, S., Garajova, S., Haon, M., Faulds, C. B., Berrin, J., Ludwig, R., and Record, E. (2016). “Salt-responsive lytic polysaccharide monooxygenases from the mangrove fungus Pestalotiopsis sp. NCi6,” Biotechnology for Biofuels 9,108. DOI: 10.1186/s13068-016-0520-3
Payne, C. M., Bomble, Y. J., Taylor, C. B., McCabe, C., Himmel, M. E., Crowley, M. F., and Beckham, G. T. (2011). “Multiple functions of aromatic-carbohydrate interactions in a processive cellulase examined with molecular simulation,” Journal of Biological Chemistry 286(47), 41028-41035. DOI: 10.1074/jbc.M111.297713
Phillips, C. M., Beeson, W. T., Cate, J. H., and Marletta, M. A. (2011). “Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa,” ACS Chemical Biology 6(12), 1399-1406. DOI: 10.1021/cb200351y
Quinlan, R. J., Sweeney, M. D., Lo Leggio, L., Otten, H., Poulsen, J. N., Johansen, K. S., Krogh, K. B. R. M., Jorgensen, C. I., Tovborg, M., Anthonsen, A., et al. (2011). “Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components,” Proceedings of the National Academy of Sciences 108(37), 15079-15084. DOI: 10.1073/pnas.1105776108
Rodionova, M. V., Poudyal, R. S., Tiwari, I., Voloshin, R. A., Zharmukhamedov, S. K., Nam, H. G., Zayadan, B. K., Bruce, B. D., Hou, H. J. M., and Allakhverdiev, S. I. (2017). “Biofuel production: Challenges and opportunities,” International Journal of Hydrogen Energy 42(12), 8450-8461. DOI: 10.1016/j.ijhydene.2016.11.125
Simmons, T. J., Frandsen, K. E. H., Ciano, L., Tryfona, T., Lenfant, N., Poulsen, J. C., Wilson, L. F. L., Tandrup, T., Tovborg, M., Schnorr, K., et al. (2017). “Structural and electronic determinants of lytic polysaccharide monooxygenase reactivity on polysaccharide substrates,” Nature Communications 8, 1064. DOI: 10.1038/s41467-017-01247-3
Song, B., Li, B., Wang, X., Shen, W., Park, S., Collings, C., Feng, A., Smith, S. J., Walton, J. D., and Ding, S. (2018). “Real-time imaging reveals that lytic polysaccharide monooxygenase promotes cellulase activity by increasing cellulose accessibility,” Biotechnology for Biofuels 11, 41. DOI: 10.1186/s13068-018-1023-1
Vaaje-Kolstad, G., Forsberg, Z., Loose, J. S., Bissaro, B., and Eijsink, V. G. (2017). “Structural diversity of lytic polysaccharide monooxygenases,” Current Opinion in Structural Biology 44, 67-76. DOI: 10.1016/j.sbi.2016.12.012
Vaaje-Kolstad, G., Westereng, B., Horn, S. J., Liu, Z., Zhai, H., Sørlie, M., and Eijsink, V. G. H. (2010). “An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides,” Science 330(6001), 219-222. DOI: 10.1126/science.1192231
Vermaas, J. V., Crowley, M. F., Beckham, G. T., and Payne, C. M. (2015). “Effects of lytic polysaccharide monooxygenase oxidation on cellulose structure and binding of oxidized cellulose oligomers to cellulases,” The Journal of Physical Chemistry B 119(20), 6129-6143. DOI: 10.1021/acs.jpcb.5b00778
Vu, V. V., Beeson, W. T., Phillips, C. M., Cate, J. H. D., and Marletta, M. A. (2014). “Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases,” Journal of the American Chemical Society 136(2), 562-565. DOI: 10.1021/ja409384b
Westereng, B., Ishida, T., Vaaje-Kolstad, G., Wu, M., Eijsink, V. G. H., Igarashi, K., Samejima, M., Ståhlberg, J., Horn, S. J., and Sandgren, M. (2011). “The putative endoglucanase PcGH61D from Phanerochaete chrysosporium is a metal-dependent oxidative enzyme that cleaves cellulose,” PLOS ONE 6(11), e27807. DOI: 10.1371/journal.pone.0027807
Wu, M., Beckham, G. T., Larsson, A. M., Ishida, T., Kim, S., Payne, C. M., Himmel, M. E., Crowley, M. F., Horn, S. J., Westereng, B., Igarashi, K., Samejima, M., Ståhlberg, J., Eijsink, V. G. H., and Sandgren, M. (2013). “Crystal structure and computational characterization of the lytic polysaccharide monooxygenase GH61D from the Basidiomycota Fungus Phanerochaete chrysosporium,” Journal of Biological Chemistry 288(18), 12828-12839. DOI: 10.1074/jbc.M113.459396
Wood, T. (1988). “Preparation of crystalline, amorphous, and dyed cellulase substrates,” Methods in Enzymology 160, 19-25.
Wood, T. M., and McCrae, S. I. (1979). “Synergism between enzymes involved in the solubilization of native cellulose,” in: Hydrolysis of Cellulose: Mechanisms of Enzymatic and Acid Catalysis, American Chemistry Society, Washington, DC., pp. 181-209. DOI: 10.1021/ba-1979-0181.ch010
Xue, Y., Han, J., Li, Y., Liu, J., Gan, L., and Long, M. (2020). “Promoting cellulase and hemicellulase production from Trichoderma orientalis EU7-22 by overexpression of transcription factors Xyr1 and Ace3,” Bioresource Technology 296. DOI: 10.1016/j.biortech.2019.122355
Zhang, R., Liu, Y., Zhang, Y., Feng, D., Hou, S., Guo, W., Niu, K., Jiang, Y., Han, L., Sindhu, L., and Fang, X. (2019). “Identification of a thermostable fungal lytic polysaccharide monooxygenase and evaluation of its effect on lignocellulosic degradation,” Applied Microbiology and Biotechnology, 103(14), 5739-5750. DOI: 10.1007/s00253-019-09928-3
Zhang, R. (2020). “Functional characterization of cellulose-degrading AA9 lytic polysaccharide monooxygenases and their potential exploitation,” Applied Microbiology and Biotechnology. DOI: 10.1007/s00253-020-10467-5
Zhou, X., and Zhu, H. (2020). “Current understanding of substrate specificity and regioselectivity of LPMOs,” Bioresources and Bioprocessing, 7(1), 11. DOI: 10.1186/s40643-020-0300-6
Zhu, M., Chen, J., Liu, N., Guo, X., and Li, D. (2018). “Oxidation mode of polysaccharide monooxygenases from Scytalidium thermophilum,” Mycosystema 38(03), 362-371. DOI: 10.13346/j.mycosystema.180247
Article submitted: March 26, 2020; Peer review completed: June 29, 2020; Revised version received and accepted: July 28, 2020; Published: July 31, 2020.
DOI: 10.15376/biores.15.3.7143-7158