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Figure S1. Partial protein alignment of the closest neighbors of PEL12 and separation of the group A and B for further reverse conservation analysis. C. rosea paralogs BN869_T0006915 (PEL12) and BN869_T00007653 (PEL13) align to the group B, whereas the protein BN869_T0008859 (PEL2), Fusaria and some members of Sordariales are separated by several hallmarks forming the group A. Figure S2. SDS-PAGE gel showing the putative expression of the PEL12 protein (arrows) at 37 kDa in the fermentation broths of two independent T. reesei PEL12 overexpression strains pel12OE-1 and pel12OE-2. The strains were cultivated for 48 h in Mandels-Anderotti medium supplemented with 1% glucose. Figure S3. Carbon source utilization of T. reesei WT and pel12OE mutants on 95 different carbon sources based on BIOLOG Phenotypic assay for filamentous fungi. Mycelial density is shown for 96 h of incubation at 28 °C in darkness. Black, gray and white circles represent mycelial density of T. reesei QM9414 (wt) and the mutants pel12OE-1 and − 2, respectively. Figure S4. Validation of pel12 mutant strains using PCR and RT-PCR. A: PCR verification of Δpel12 using primers located within the hygB cassette (Hyg F /Hyg R) in combination with primers located upstream and downstream regions from the deletion cassette (6915 ko F / 6915 ko R). A PCR product of ~ 2.5 kb using primers 6915 ko F / Hyg R and 6915 ko R / Hyg F were expected from a correct gene replacement. M, gene ruler DNA ladder mix; 1–3, independent Δpel12 mutants; 4, WT strain. B: RT-PCR analysis of pel12 gene expression in WT and deletion strains using pel12 specific primers. A PCR product of 92 bp was expected from the WT. M, gene ruler DNA ladder mix; 1, WT; 2–4, independent deletion strains. Primer combinations used for PCR and RT-PCR are given above the images. Figure S5. Carbon source utilization of C. rosea WT and ∆pel12 mutants (∆pel12 KOA, KOB, KOC) on 95 different carbon sources based on BIOLOG Phenotypic assay for filamentous fungi. Mycelial density is shown for 144 h of incubation at 28 °C in darkness. Figure S6. Measurement of B. cinerea necrotic lesions area on detached leaves of Arabidopsis thaliana. The leaves were inoculated with C. rosea strains (A) or T. reesei (B) strains spores 30 min before application of B. cinerea spores. Necrotic lesion area was measured under the microscope using a DeltaPix camera and software 56 h post inoculations. Only B. cinerea inoculated leaves were used as control. Error bars represent standard deviation based on three biological replicates. Different letters indicate statistically significant differences (P ≤ 0.05) based on the Fisher test. (PDF 633 kb)

Figure S1. Alternative phylogenetic placements of Clonostachys rosea in Hypocreales. The alternative topologies were retrieved from the literature, while branch lengths were determined based on a four-gene alignment including actin, glyceraldehyde 3-phosphate dehydrogenase, DNA-directed RNA polymerase II subunit B and translation elongation factor 1 alpha, using MEGA ver. 6. The species phylogeny was calibrated to the fossil record by setting the split between H. minnesotensis and H. sinensis to 29 million years. (A) Clonostachys rosea (Bionectriaceae) as sister taxon with Fusaria (Nectriaceae) and (B) C. rosea (Bionectriaceae) as basal lineage in Hypocreales. The included species represents different families within the order Hypocreales: H. minnesotensis, H. thompsonii and H. sinensis (Ophiocordycipitaceae), M. robertsii (Clavicipitaceae), T. atroviride, T. virens and T. reesei (Hypocreaceae), F. graminearum and F. solani (Nectriaceae), C. rosea (Bionectriaceae) and the outgroup species N. crassa (Sordariales, Sordariaceae). E = entomopathogenic, M = mycoparasitic, N = nematode parasitic, P = plant pathogenic, S = saprotrophic. Figure S2. Growth of Clonostachys rosea on milk powder plates to assess extracellular protease activity. Clonostachys rosea was inoculated to plates that contained (A) water agar, (B) potato dextrose agar, (C) malt extract agar and (D) Czapek dox agar, and the clearing zones (indicative of extracellular protease activity) were assessed both from above and below the agar plates three and five days post inoculation. (DOCX 2101 kb)

Presently, plant biomass is considered as one of the major future renewable sources for the production of second-generation biofuels. While the first generation biofuels essentially are based on starch and sucrose rich feed stocks and which production may compete with food production, the second-generation biofuels may be based on lignocellulose as feedstock, which is less problematic from an ethical point of view. The degradation of carbohydrates in plant biomass to fermentable sugars requires the concerted action of several diverse classes of carbohydrate active enzymes (CAZymes) for a total and efficient conversion of the plant biomass. Through a carefully balanced synergism mechanistically different CAZymes are able to degrade the stable and recalcitrant pol- ymers in the plant cell walls, such as cellulose, to soluble and fermentable monosaccharides. It is crucial to study the properties and function of these enzymes if we want to strive for a sustainable production of chemicals and biofuels, as they serve as a reservoir of environmentally friendly molecular tools. The main focus of the research work presented in this thesis is biochemical and structure-function characterizations of two classes of CAZymes: fungal glycoside hydrolase family 3 (GH3) β-glucosidases, and bacterial lytic polysaccharide monooxygenases, often referred to as LPMOs. GH3 β- glucosidases catalyse the conversion of disaccharides, produced by other CAZymes e.g. cellulases, to glucose. H. jecorina Cel3A, R. emersonii Cel3A and N. crassa NcGH3-3 are three industrially relevant fungal GH3 β-glucosidases for which the structures have been determined using X-ray crystallographic methods. The H. jecorina Cel3A, R. em- ersonii Cel3A enzymes has also been characterized biochemically. The LPMOs act in the very initial stage of plant cell wall degradation and cleave glycosidic bonds in crys- talline polysaccharides via an oxidative mechanism, which facilitates access to new chain ends for other CAZymes. To elucidate the structural and biochemical properties of LPMOs with bacterial origin, the structure of an AA10 LPMO the LPMO10A from Enterococcus faecalis was determined using X-ray crystallography. Furthermore, structural changes of the active site metal configuration by so-called X-ray induced photoreduction, were determined. During this reduction process, which mimics the active enzyme, the bound active site copper atom is reduced from Cu(I) to Cu(II), which causes changes in the ligation configuration.