Yeasts Have Evolved Divergent Enzyme Strategies To Deconstruct and Metabolize Xylan

Abstract

Together with bacteria and filamentous fungi, yeasts actively take part in the global carbon cycle. Over 100 yeast species have been shown to grow on the major plant polysaccharide xylan, which requires an arsenal of carbohydrate active enzymes. However, which enzymatic strategies yeasts use to deconstruct xylan and what specific biological roles they play in its conversion remain unclear. In fact, genome analyses reveal that many xylan-metabolizing yeasts lack expected xylanolytic enzymes. Guided by bioinformatics, we have here selected three xylan-metabolizing ascomycetous yeasts for in-depth characterization of growth behavior and xylanolytic enzymes. The savanna soil yeast Blastobotrys mokoenaii displays superior growth on xylan thanks to an efficient secreted glycoside hydrolase family 11 (GH11) xylanase; solving its crystal structure revealed a high similarity to xylanases from filamentous fungi. The termite gut-associated Scheffersomyces lignosus, in contrast grows more slowly, and its xylanase activity was found to be mainly cell surface-associated. The wood-isolated Wickerhamomyces canadensis, surprisingly, could not utilize xylan as the sole carbon source without the addition of xylooligosaccharides or exogenous xylanases or even co-culturing with B. mokoenaii, suggesting that W. canadensis relies on initial xylan hydrolysis by neighboring cells. Furthermore, our characterization of a novel W. canadensis GH5 subfamily 49 (GH5_49) xylanase represents the first demonstrated activity in this subfamily. Our collective results provide new information on the variable xylanolytic systems evolved by yeasts and their potential roles in natural carbohydrate conversion. IMPORTANCE Microbes that take part in the degradation of the polysaccharide xylan, the major hemicellulose component in plant biomass, are equipped with specialized enzyme machineries to hydrolyze the polymer into monosaccharides for further metabolism. However, despite being found in virtually every habitat, little is known of how yeasts break down and metabolize xylan and what biological role they may play in its turnover in nature. Here, we have explored the enzymatic xylan deconstruction strategies of three underexplored yeasts from diverse environments, Blastobotrys mokoenaii from soil, Scheffersomyces lignosus from insect guts, and Wickerhamomyces canadensis from trees, and we show that each species has a distinct behavior regarding xylan conversion. These findings may be of high relevance for future design and development of microbial cell factories and biorefineries utilizing renewable plant biomass.

Keywords: CAZymes; microbial co-cultures; xylan; xylanase; yeast.

FIG 1

Overview of predicted endo-xylanases in known xylanolytic yeasts. Putative xylanases are marked next to known xylanolytic yeasts (highlighted in bold and by numbers). Asterisks (*) indicate the three yeasts selected for characterization. GH, glycoside hydrolase.

FIG 2

Yeast growth in beechwood glucuronoxylan and xylanolytic activity localization. (A to C) Yeasts were grown in 10 mL Delft minimal medium with 2% beechwood glucuronoxylan or 2% xylose as the sole carbon source in biological triplicates. XOs, xylooligosaccharides. (D to F) Bright-field microscopy showing yeast morphology from 96-h xylan cultures. (G) Yeast subcellular xylanase activity originating from secretome (Sec), cells (Cell), and intracellular (Intra) from lysed cell fractions were compared using DNS reducing sugar assays in at least triplicate experiments. Values are means ± standard deviations as error bars. Asterisks indicate statistical significance in subcellular activity levels between W. canadensis and W. canadensis supplemented with XOs. P values of ≤0.0001 (***) were considered significant (n = 12) and evaluated using one-way analysis of variance (ANOVA) with Tukey’s test. (H) β-Xylosidase activity was quantified using p-nitrophenyl-β-d-xylopyranoside; (I) acetyl esterase activity using p-nitrophenyl-acetate; (J) α-l-arabinofuranosidase activity using p-nitrophenyl-α-l-arabinofuranoside; and (K) α-d-glucuronidase activity using p-nitrophenyl-α-d-glucuronide.

FIG 3

Co-cultures of yeast growth in xylan. (A) Co-cultures of B. mokoenaii and W. canadensis at different starting ratios in 2% beechwood glucuronoxylan in triplicates. Values are means ± standard deviations as error bars. Asterisks indicate statistical significance in OD levels at different time points between B. mokoenaii mono- and co-culture treatments with W. canadensis. P values of ≤0.05 (*) and ≤0.01 (**) were considered significant (n = 3) and evaluated using one-way ANOVA with Tukey’s test. (B) Distribution in fluorescence particle area (%) of co-cultures of B. mokoenaii (blue) and W. canadensis (yellow) at an initial ratio of 1:1. (C to E) Representative images from fluorescence microscopy of B. mokoenaii and W. canadensis (1:1 ratio) (C) after 24 h, (D) after 48 h, and (E) after 72 h. (F and G) Co-cultures on agar plates with (F) 0.4% beechwood glucuronoxylan and (G) 0.4% wheat arabinoxylan over time with (+Bm) or without (–Bm) the addition of B. mokoenaii. The clearing zones correlate with xylanase-mediated xylooligosaccharide release and enable W. canadensis growth (red arrows). Bm, Blastobotrys mokoenaii; Sl, Scheffersomyces lignosus; Wc, Wickerhamomyces canadensis.

FIG 4

Effect of xylanase treatment on beechwood glucuronoxylan and wheat arabinoxylan and boosting of yeast growth. (A and B) Hydrolysis of (A) beechwood glucuronoxylan and (B) wheat arabinoxylan by recombinant enzymes over 24 h using 0.1 μM enzyme in triplicates. (C) Xylooligosaccharide formation from beechwood glucuronoxylan and wheat arabinoxylan after 24 h in duplicates. X1, Xylose; X2, xylobiose; X3, xylotriose; X4, xylotetraose; X5, xylopentaose; X6, xylohexaose; XOs, xylooligosaccharide. (D) Boosting of W. canadensis growth in 2% beechwood glucuronoxylan by supplying the cultures with BmXyn11A and WcXyn5_49A in lower (50 μg/g xylan) and higher (250 μg/g xylan) concentrations in biological triplicates.

FIG 5

Crystal structure of BmXyn11A and comparison with other GH11 members. (A) The overall fold of BmXyn11A (chain A), with β-sheets in yellow, α-helices in red, loops in green, and catalytic Glu residues as cyan sticks. (B) Closeup of the active site cleft of BmXyn11A (green) superimposed with the bound oligosaccharide in yellow ball-and-stick representation derived from the Aspergillus niger GH11 xylanase (PDB: 2QZ2, inactivated E170A variant, crystallized with X5); (C) shown in the same orientation.

FIG 6

Predicted AlphaFold 2 structures for GH5 and GH10 xylanases and β-xylosidases characterized in this study. (A) SlXyn10A. (B) WcXyn5_49A. (C) WcXyn5_22A. (i) Overview of each enzyme. The helical bundle in WcXyn5_49A is shown as green sticks. The catalytic Glu residues are magenta sticks. In the GH5 structures, a superimposed oligonucleotide from the structure of XEG5A (PDB: 4W89) is shown as yellow sticks; in the GH10 structure the oligonucleotide is superimposed from CbXyn10C (PDB: 5OFK). (ii) Closeups of each active site. (iii) Surface representation, highlighting the different shapes of the binding clefts. In panel Ciii, a 90° rotated view reveals the deep cleft of WcXyn5_22A with limited surface accessibility. (iv) Reference structures of CbXyn10C and XEG5A. Figure S7 visualizes the pLDDT scores for each predicted model.

FIG 7

Schematic view of the xylanolytic strategies of the three investigated species. (A) Predicted xylanolytic CAZymes (10) of B. mokoenaii, S. lignosus, and W. canadensis represented by color (yellow, red, and blue, respectively). Arrows from individual enzyme families indicate enzyme activity in beechwood glucuronoxylan degradation. CE, carbohydrate esterase; GH, glycoside hydrolase. (B) Xylan. B. mokoenaii utilizes a secreted xylanase which depolymerizes xylan away from the cells and liberates oligosaccharides in the process. S. lignosus instead maintains the xylanase activity close to the cell surface, which might give a competitive advantage in its native environment. W. canadensis secretes xylanases similarly to B. mokoenaii, but only after sensing xylooligosaccharides released by xylanase action from neighboring cells. Putative enzymes, as indicated by activity on chromogenic model substrates, and therefore remaining speculative, are shown in gray together with assumed CAZy family memberships based on previous work (10). Acetyl groups are shown as black dots. XHT, xylose/glucose transporters.