Accessory Activities 3: Novel Flavoprotein Oxidoreductase from bacterial sources

Background.

Lignocellulose is the main component of woody plants´ cell wall and has evolved to fulfil mechanical requirements as well as protect plants against herbivores and saprobionts (Janusz et al. 2017 and references therein). Studies on lignin degradation have focused on white-rot and brown-rot fungi so far (Bugg et al. 2011 and references therein). In recent years, a number of bacteria were observed to be able to degrade and metabolize lignin components and derivatives as well. It is not well understood if bacterial lignin depolymerization follows pathways comparable to those from white rot fungi. Genome sequencing projects have revealed the presence of genes hypothetically annotated as lignin modifying enzymes across a range of mostly α- and γ-Proteobacteria and Actinobacteria (Cragg et al. 2015; Janusz et al. 2017). Besides laccase genes, many genes annotated as peroxidases (putatively dye-decolorizing peroxidases, catalase-peroxidases or versatile peroxidases rather than lignin or manganese peroxidases) were identified in Actinobacteria genomes (Brown et al. 2012; Brown and Chang 2014).

This suggests that such bacteria should also contain auxiliary enzymes such as aryl alcohol oxidase, glucose dehydrogenase, pyranose oxidase or cellobiose dehydrogenase (Janusz et al., 2017). These enzymes are implicated to be involved in lignin degradation by providing hydrogen peroxide for activation of peroxidases and for reduction during redox cycling of quinones (Martinez et al. 2005; de Gonzalo et al. 2016). All belong to the GMC oxidoreductase family of flavoproteins. Pyranose oxidase catalyzes the oxidation of aldopyranoses to 2-ketoaldoses and hydrogen peroxide, but most pyranose oxidases also use benzoquinone as electron acceptor, often with a higher efficiency than oxygen (Leitner et al. 2001; Pisanelli et al. 2009; Salaheddin et al. 2010). Pyranose dehydrogenase oxidizes mono- and disaccharides and uses only (substituted) quinones or complexed metal ions as electron acceptors (Peterbauer and Volc 2010). Cellobiose dehydrogenases additionally contain a heme domain (Zamocky et al. 2014) and play a major role in oxidative cellulose degradation by transferring electrons to Lytic Polysaccharide Monoxygenases (LPMO; Kracher et al. 2016). Genome data reveal a number of genes putatively encoding such enzymes in bacteria, however, biochemical information is almost non-existent. There is a single report of a bacterial pyranose 2-oxidase, from Pseudarthrobacter siccitolerans. The recombinantly expressed enzyme is a 64-kDa monomer with an FAD cofactor that oxidizes D-glucose, oxygen and 1,4-benzoquinone act as an electron acceptor (Mendes et al. 2016). Another pyranose oxidase (from Kitasatospora aureofaciens) was recently expressed in E. coli and characterized (Herzog, Sützl and Peterbauer, manuscript in press, DOI: 10.1128/AEM.00390-19).

Aims and methods.

Many Actinomycetes contain genes putatively encoding peroxidases, laccases and a peroxide-providing system like pyranose oxidases (Brown and Chang 2014). We propose to select a limited number of Actinomycete model organisms with sequenced genomes containing genes encoding peroxidases, laccases and carbohydrate oxidoreductases and that are capable of degradation of lignin or lignin derivatives, namely from the genera Streptomyces, Amycolatopsis and Kitasatospora, and characterize their auxiliary enzyme system (Auxiliary Activities Family 3). A key point will be the subcellular location of the enzyme(s).
Our investigation will include interactions with other enzymes to be tested in vitro, namely the activation of peroxidases through hydrogen peroxide (or another mechanism). In case enzymes are encountered that fulfil the structural requirements for activation of LPMOs (like cellobiose dehydrogenases, which have not been shown to be present in bacteria), the interaction between these enzymes will be investigated as well.

We further propose to study the inactivation of encoding genes and a phenotypical assessment for degradation of lignin in at least one model strain, to establish whether carbohydrate-oxidizing flavoproteins and similar auxiliary activities are involved in lignin modification and degradation. Unlike fungi, bacteria are less likely to contain genes or gene families encoding enzymes with identical, complementary or overlapping activities. This will make investigations into the biological role of individual enzymes more straight-forward, as a phenotypic effect can be expected upon inactivation of a single gene.

3.1. Expression and characterization of bacterial pyranose oxidoreductases

DNA sequences encoding putative pyranose oxidases and related enzymes from Actinomycetes will be synthesized by commercial service providers. We will use established platforms like pET-21 for E. coli, as has been published for fungal enzymes (Pisanelli et al. 2009). We will, however, consider using gram-positive secretory expression systems based on Bacillus spp. or Streptomyces spp. (Anne et al. 2014), if sequence analysis suggests that the target genes encode secretory proteins, as this will more closely resemble the natural situation as well as facilitate downstream processing. Recombinant proteins will be expressed containing a His6 tag for easy purification based on metal affinity chromatography (IMAC). The enzymes will be biochemically characterized with respect to electron donor (carbohydrate substrate), electron acceptor and basic kinetic characteristics (e.g., Pisanelli et al. 2009; Salaheddin et al. 2010; Brugger et al. 2014).

3.2. Transcription and location of GMC oxidoreductases
Pyranose oxidases often do not contain recognizable signal peptides and their subcellular localization is unclear, even though activation of extracellular peroxidases through hydrogen peroxide suggests an extracellular location. We will cultivate selected species on glucose, cellulose (derivatives), lignocellulose and lignin model substrates and determine the expression profiles of peroxidase and oxidoreductase genes, including LPMO genes, in order to assess the biological role of individual enzymes. Previously obtained biochemical data will allow us to establish the localization of the enzymes in the original organisms, as enzymatic assays can be performed on the supernatant, the cell pellet fraction and cell free extracts after homogenization.
Streptomycetes are amenable to genetic manipulation (Anne et al. 2014). We will use established protocols to express the respective genes in the original organism as a cell fusion with a fluorescent marker, e.g., the Green Fluorescent Protein, and detect the localization during growth on lignin-containing substrates in vivo.

3.3. Interaction with other enzymes
Peroxidase activation, which happens indirectly through hydrogen peroxide, will be studied in vitro using purified enzymes with appropriate controls (addition of catalase) as was shown for lignin and manganese peroxidases. LPMOs were shown to be activated through interdomain electron transfer from cellobiose dehydrogenases (Courtade et al. 2016, Kracher and Scheiblbrandner et al. 2017). Available information to date does not suggest that such enzymes are present in bacteria. However, activation of LPMO has been shown through unspecific reduction by plant phenolic compounds and lignin fractions, as well as by plant-derived diphenols or quinones acting as redox mediators (Kracher and Scheiblbrandner et al. 2017). This could be a link between lignin and cellulose degradation, and we will investigate bacterial oxidoreductases if they can fulfil that role. Lastly there is the hypothesis that LPMO activity depends on hydrogen peroxide as a required co-substrate (Bissaro et al. 2017), which could point to yet another function for carbohydrate active oxidoreductases in the context of biomacromolecule degradation.

3.4. Functional genomic approach
In order to complement the biochemical data generated in the first work packages we will utilize the bacterial propensity for small and parsimonious genomes and their – relative – lack of multigenicity. The highly versatile CRISPR/Cas9 system for genome editing has been established in Streptomyces spp. (Cobb et al. 2015). This will allow us to selectively inactivate/knock out genes encoding GMC-oxidoreductases singly and in combination in much shorter time and with less effort than previously using integrative knock-out-cassettes, and study the phenotypic consequences particularly with respect to growth on lignocellulosic substrates, production of characteristic lignin degradation products and depolymerization of crystalline cellulose: depending on the hypothesis, knock-out of a (all) hydrogen peroxide-producing oxidoreductases should result in a lack of peroxidase activation and lignin degradation, perhaps also in a lowered LPMO reduction for cellulose attack and reduced capacity to degrade crystalline cellulose fractions. This approach will shed light on the particularly enigmatic role of GMC-oxidoreductases during lignocellulose depolymerization.

Anne J, Vrancken K, van Mellaert L, van Impe J, Bernaerts K (2014) Protein secretion biotechnology in Gram-positive bacteria with special emphasis on Streptomyces lividans. Biochim Biophys Acta 1843(8): 1750-1761.
Bissaro B, Røhr ÅK, Mülle G, Chylenski P, Skaugen M, Forsberg Z, Horn SJ, Vaaje-Kolstad G, Eijsink VGH (2017) Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2. Nat Chem Biol 13: 1123-1127
Brown ME, Chang MC (2014) Exploring bacterial lignin degradation. Curr Opin Chem Biol 19: 1-7.
Brown ME, Barros T, Chang MC (2012) Identification and Characterization of a Multifunctional Dye Peroxidase from a Lignin-Reactive Bacterium. ACS Chem Biol 7: 2074-2081
Brugger D, Krondorfer I, Zahma K, Stoisser T, Bolivar JM, Nidetzky B, Peterbauer CK, Haltrich D (2014) Convenient microtiter plate-based, oxygen-independent activity assays for Flavin-dependent oxidoreductases based on different redox dyes. Biotechnol J 9:474-482
Bugg TDH, Ahmad M, Hardiman EM, Singh R (2011) The emerging role for bacteria in lignin degradation and bio-product formation. Curr Op Biotechnol 22: 394-400
Cobb RE, Wang Y, Zhao H (2015) High-Efficiency Multiplex Genome Editing of Streptomyces Species Using an Engineered CRISPR/Cas System. ACS Synth Biol 4:723-728
Courtade G, Wimmer R, Røhr ÅK, Preims M, Felice AKG, Dimarogona M, Vaaje-Kolstad G, Sørlie M, Sandgren M, Ludwig R, Eijsink VGH, Aachmann FL (2016)
Interactions of a fungal lytic polysaccharide monooxygenase with β-glucan substrates and cellobiose dehydrogenase. Proc Natl Acad Sci USA 113: 5922-5927
Cragg SM, Beckham GT, Bruce NC, Bugg TDH, Distel DL, Dupree P, Green Etxabe A, Goodell BS, Jellison J, McGeehan JE, McQueen-Mason SJ, Schnorr K, Walton PH, Watts JEM, Zimmer M (2015) Lignocellulose degradation mechanisms across the Tree of Life. Curr Op Chem Biol 29: 108-119
de Gonzalo G, Colpa DI, Habibb MHM, Fraaije MW (2016) Bacterial enzymes involved in lignin degradation. J Biotech 236: 110–119
Janusz G, Pawlik A, Sulej J, Świderska-Burek U, Jarosz-Wilkolazka A, Paszczyński A (2017) Lignin degradation: microorganisms, enzymes involved, genomes analysis and evolution. FEMS Microbiol Rev 41: 941-962
Kracher D, Scheiblbrandner S, Felice AKG, Breslmayr E, Preims M, Ludwicka K, Haltrich D, Eijsink VGH, Ludwig R (2016) Extracellular electron transfer systems fuel cellulose oxidative degradation. Science 352: 1098-1101
Leitner C, Volc J, Haltrich D (2001) Purification and Characterization of Pyranose Oxidase from the White Rot Fungus Trametes multicolor. Appl Environ Microbiol 67(8): 3636–3644
Martinez AT, Speranza M, Ruiz-Duenas FJ, Ferreira P, Camarero S, Guillen F, Martinez MJ, Gutierrez A, del Rio JC (2005) Biodegradation of lignocellulosics: microbial, chemical, and enzymatic aspects of the fungal attack of lignin. Int Microbiol 8(3): 195-204
Mendes S, Banha C, Madeira J, Santos D, Miranda V, Manzanera M, Ventura MR, van Berkel WJH, Martins LO (2016) Characterization of a bacterial pyranose 2-oxidase from Arthrobacter siccitolerans. J Mol Catal B: Enzymatic. S1381-1177(16): 30218-1
Peterbauer CK, Volc J (2010) Pyranose dehydrogenases: biochemical features and perspectives of technological applications. Appl Microbiol Biotechnol 85: 837-848
Pisanelli I, Kujawa M, Spadiut O, Kittl R, Halada P, Volc J, Mozuch MD, Kersten P, Haltrich D, Peterbauer C (2009) Pyranose 2-oxidase from Phanerochaete chrysosporium--expression in E. coli and biochemical characterization. J Biotechnol 142(2): 97-106
Salaheddin, C., Takakura, Y., Tsunashima, M., Stranzinger, B., Spadiut, O., Yamabhai, M., Peterbauer, C.K., Haltrich, D. (2010) Characterisation of recombinant pyranose oxidase from the cultivated mycorrhizal basidiomycete Lyophyllum shimeji (hon-shimeji). Microb Cell Fact 9:57
Zámocky M, Hallberg M, Ludwig R, Divne C, Haltrich D (2004) Ancestral gene fusion in cellobiose dehydrogenase reflects a specific evolution of GMC oxidoreductases in fungi. Gene 338: 1-14