Combining Xylose Reductase from Spathaspora arborariae with Xylitol Dehydrogenase from Spathaspora passalidarum to Promote Xylose Consumption and Fermentation into Xylitol by Saccharomyces cerevisiae

In recent years, many novel xylose-fermenting yeasts belonging to the new genus Spathaspora have been isolated from the gut of wood-feeding insects and/or wood-decaying substrates. We have cloned and expressed, in Saccharomyces cerevisiae, a Spathaspora arborariae xylose reductase gene (SaXYL1) that accepts both NADH and NADPH as co-substrates, as well as a Spathaspora passalidarum NADPH-dependent xylose reductase (SpXYL1.1 gene) and the SpXYL2.2 gene encoding for a NAD+-dependent xylitol dehydrogenase. These enzymes were co-expressed in a S. cerevisiae strain over-expressing the native XKS1 gene encoding xylulokinase, as well as being deleted in the alkaline phosphatase encoded by the PHO13 gene. The S. cerevisiae strains expressing the Spathaspora enzymes consumed xylose, and xylitol was the major fermentation product. Higher specific growth rates, xylose consumption and xylitol volumetric productivities were obtained by the co-expression of the SaXYL1 and SpXYL2.2 genes, when compared with the co-expression of the NADPH-dependent SpXYL1.1 xylose reductase. During glucose-xylose co-fermentation by the strain with co-expression of the SaXYL1 and SpXYL2.2 genes, both ethanol and xylitol were produced efficiently. Our results open up the possibility of using the advantageous Saccharomyces yeasts for xylitol production, a commodity with wide commercial applications in pharmaceuticals, nutraceuticals, food and beverage industries.


Introduction
Industrial biotechnology will play an increasing role in creating a more sustainable global economy. Lignocellulosic biomass, an abundant and renewable feedstock, is an attractive raw material for fuels and chemicals production, since it does not compete with food and feed production [1,2]. The major

Strains, Media and Growth Conditions
The yeast strains used in this study are listed in Table 1. Escherichia coli strain DH5α was used for cloning, and was grown in Luria broth (1% tryptone, 0.5% yeast extract, 0.5% sodium chloride) supplemented with ampicillin (100 mg/L). Yeasts were grown on rich YP medium (1% yeast extract, 2% Bacto peptone) or synthetic complete (YNB) medium (0.67% yeast nitrogen base without amino acids, supplemented with adequate auxotrophic requirements), containing 2% glucose or xylose. The pH of the medium was adjusted to pH 5.0 with HCl. When required, 2% Bacto agar, 200 mg/L geneticin sulfate (G-418, Sigma-Aldrich Brasil Ltda., São Paulo, SP, Brazil) or 0.5 g/L zeocin (Invivogen, San Diego, CA, USA) were added to the medium. Cells were grown aerobically at 28 • C with shaking (160 rpm) in cotton plugged Erlenmeyer flasks filled to 1/5 of the volume with medium. Cellular growth was followed by turbidity measurements at 600 nm (OD 600 nm ), and culture samples were harvested regularly, centrifuged (5000× g, 1 min), and their supernatants used for the determination of substrates and fermentation products, as described below. The maximum specific growth rate (µ max , h −1 ) was determined by the slope of a straight line between ln OD 600nm and time (h) during the initial (~48 h) exponential phase of growth on xylose. Table 1. Yeast strains, plasmids and primers used in this study.
For batch fermentations, cells were pre-grown in synthetic complete YNB medium containing 2% glucose for 20 h at 28 • C, the cells were collected by centrifugation at 6000× g for 5 min at 4 • C and washed twice with sterile water, and inoculated at a high cell density (10.0 ± 0.5 g of dry yeast/L), into 25 mL of synthetic YNB medium containing 2% xylose. Batch fermentations were performed at 30 • C in closed 50 mL bottles with a magnetic stir bar, to allow mild agitation (100 rpm). Samples were collected regularly and processed, as described above.

Molecular Biology Techniques
Standard methods for bacterial transformation, DNA manipulation and analysis were employed [32]. Yeast transformation was performed by the lithium acetate method [33]. To over-express the XKS1 gene encoding xylulokinase in S. cerevisiae CEN.PK2-1C, the promoter region of this gene was modified according to the polymerase chain reaction (PCR)-based gene replacement procedure, as described previously [29]. Briefly, the kanMX-P ADH1 module from plasmid pFA6a-kanMX6-PADH1 (Table 1) was amplified with primers XKS1-Kanr-F and XKS1-PADH1-R (Table 1), and the resulting PCR product of 2394 bp (flanked by~40 bp of homology to the promoter and start regions of the XKS1 gene) containing the truncated and constitutive promoter of the ADH1 gene was used to transform competent yeast cells. After 2 h cultivation on YP-2% glucose, the transformed cells were plated on the same medium containing G-418 and incubated at 28 • C. G-418-resistant isolates were tested for proper genomic integration of the kanMX-P ADH1 cassette at the XKS1 locus by diagnostic colony PCR using 3 primers (V-XKS1-F, V-XKS1-R and V-kan r -F; Table 1). This set of 3 primers amplified a 1557-bp fragment (primers V-XKS1-F and V-XKS1-R) from a normal XKS1 locus, or yielded a 3871-bp fragment (primers V-XKS1-F and V-XKS1-R) and a 2889-bp fragment (primers V-kan r -F and V-XKS1-R) if the kanMX-P ADH1 module was correctly integrated at the promoter region of the XKS1 gene, producing strain ASY-1 (KanMX-P ADH1 ::XKS1, Table 1).
We further improved the capacity of xylose utilization in this strain by deleting the PHO13 gene of S. cerevisiae, a gene encoding for an alkaline phosphatase known to suppress xylose utilization by recombinant yeast strains [34][35][36]. Briefly, the LoxP-Ble R -LoxP knockout cassette from plasmid pUG66 ( Table 1) was amplified with primers DE-PHO13-F and DE-PHO13-R (Table 1), and the resulting PCR product of 1265-bp (flanked by~40 bp of homology to the upstream and downstream regions of the PHO13 locus) containing the Ble R gene was used to transform ASY-1 competent yeast cells. After 2-h cultivation on YP-2% glucose, the transformed cells were plated on the same medium containing zeocin and incubated at 28 • C. Zeocin-resistant isolates were tested for proper genomic integration of the LoxP-Ble R -LoxP cassette at the PHO13 locus by diagnostic colony PCR using 3 primers (V-PHO13-F, V-PHO13-R and V-Ble r -F; Table 1). This set of 3 primers amplified a 1654-bp fragment (primers V-PHO13-F and V-PHO13-R) from a normal PHO13 locus, or yielded a 1900-bp fragment (primers V-PHO13-F and V-PHO13-R) and a 965-bp fragment (primers V-Ble r -F and V-PHO13-R) if the LoxP-Ble R -LoxP module replaced and deleted the PHO13 gene, producing strain ASY-2 (KanMX-P ADH1 ::XKS1 and pho13∆::LoxP-Ble R -LoxP, Table 1).
Based on the genome sequence of Sp. arborariae [37] and Sp. passalidarum [21], primers were designed (Table 1) to amplify the xylose reductase encoded by the SaXYL1 or SpXYL1.1 genes, and the xylitol dehydrogenase encoded by the SpXYL2.2 gene, introducing restriction sites for cloning into multicopy shuttle vectors containing strong and constitutive promoters and terminators (pPGK, p423-TEF and p423-GPD, Table 1), as well as the URA3 or HIS3 genes used as selective markers. The genomic DNA from the Sp. passalidarum and Sp. arborariae strains was purified using a YeaStar Genomic DNA Kit TM (Zymo Research, Irvine, CA, USA). The amplified DNA fragments, originally cloned into the pPGK plasmid, had their 5 and 3 ends sequenced (ACTGene Analíses Moleculares Ltda., Alvorada, RS, Brazil) using primers Prom_PGK_54_F and Ter_PGK_65_R (Table 1) to confirm the identity of the cloned genes.

Enzyme Assays
Cell-free extracts for assays of xylose metabolizing enzymes were prepared with the yeast protein extraction reagent Y-PER (Pierce, Rockford, IL, USA) after cultivation yeast cells on YP-2% xylose, or 2% glucose (for S. cerevisiae strains over-expressing the cloned genes). Protein concentrations in the cell-free extracts were determined with the Micro-BCA kit (Pierce, Thermo Fisher Scientific Inc., Sinapse Biotecnologia, São Paulo, SP, Brazil), using a Biowave II spectrophotometer (Biochrome WPA, Cambridge, UK). Xylose reductase activity was measured by monitoring the oxidation of NADH or NADPH at 340 nm [10,38] at 30 • C in 45.5 mM potassium phosphate buffer (pH 6.0), using 0.15 mM NADH or NADPH and 200 mM xylose as substrate. The kinetic parameters of xylose reductase for each substrate was determined using 1-400 µM NADH or NADPH, or 0.5-600 mM xylose, under the conditions described above. Xylitol dehydrogenase activity was measured by monitoring the reduction of NAD + or NADP + at 340 nm [10,38] at 35 • C in 50 mM Tris-HCl buffer (pH 9.0) containing 50 mM MgCl 2 , 300 mM xylitol, and 1 mM NAD + or NADP + . The kinetic parameters of xylitol dehydrogenase for each substrate was determined using 1-4000 µM NAD + , or 0.1-500 mM xylitol, under the same conditions. The xylulokinase activity was determined by a coupled assay to measure ADP production as previously described [39] in 0.2 M Tris-HCl buffer (pH 7.0) containing 2.3 mM MgCl 2 , 10 mM NaF, 2.5 mM ATP, 0.25 mM phosphoenolpyruvate, 3.5 mM reduced glutathione, 10 U of pyruvate kinase, 15 U of lactate dehydrogenase, 0.2 mM NADH, and 4.25 mM xylulose. One unit of enzyme activity was defined as the amount of enzyme that reduced or oxidized 1 µmol of NAD(P) + or NAD(P)H per minute. These enzymatic activities were determined using a Cary 60 UV-VIS spectrophotometer (Agilent Technologies, Santa Clara, CA, USA). The kinetic parameters (K m and V max ) of the cloned xylose reductases and xylitol dehydrogenase enzymes expressed in S. cerevisiae were determined by fitting the experimental data to the Michaelis-Menten equation, using SigmaPlot v. 11.0 (Systat Software Inc., San Jose, CA, USA).

Analytical Methods
Xylose, ethanol, xylitol, glycerol, and acetate were determined by high performance liquid chromatography (HPLC), equipped with a refractive index detector (RI-2031Plus; JASCO, Tokyo, Japan) using an Aminex HPX-87H column (Bio-Rad Laboratories, Hercules, CA, USA). The HPLC apparatus was operated at 40 • C using 5 mM H 2 SO 4 as the mobile phase at a flow rate of 0.1 mL/min and 0.01 mL injection volume. The following calculations were considered: products yield (Y p/s , g/g) were determined by correlating ∆P produced (xylitol, ethanol, glycerol or acetate) with ∆S consumed (xylose or glucose) at time of maximal substrate consumption. The xylitol volumetric productivity (Q p , g/L/h) was determined by the slope of a straight line between xylitol concentration (g/L) and time (h) during maximum xylitol production.

Results and Discussion
Our analysis for xylose reductase in Sp. arborariae UFMG-HM19.1A T showed activity with both co-substrates (NADH and NADPH); nevertheless, the NADH-dependent activity was~25% of that observed with NADPH. The NAD + -dependent xylitol dehydrogenase, however, was not active in the presence of NADP + (Table 2). Similarly, the Sp. passalidarum strain UFMG-CM-Y474 showed xylose reductase activity with both NADH and NADPH, and the highest xylitol dehydrogenase (NAD + -dependent) activity (~1.3 U/[mg protein], Table 2) among all the Spathaspora yeasts analyzed (data not shown). Based on the Sp. passalidarum and Sp. arborariae genome sequences [21,37], we designed primers to amplify the SaXYL1 and SpXYL1.1 genes (both open reading frames -ORFswith 957 nucleotides, encoding for proteins of 318 amino acids), and the SpXYL2.2 gene (an ORF with 1089 nucleotides, encoding for a protein of 362 amino acids), and cloned these three genes into the pPGK multicopy plasmid for expression in the S. cerevisiae CEN.PK2-1C yeast strain. As shown in Table 3, these genes were functional in S. cerevisiae, with a xylose reductase activity encoded by the SaXYL1 gene that accepts both co-substrates, with a NADH-dependent activity that is~30% the activity measured with NADPH, while the xylose reductase encoded by the SpXYL1.1 gene accepted only NADPH as co-substrate. The determination of the kinetic parameters of the SaXYL1 enzyme cloned in S. cerevisiae (Table 4) revealed an enzyme with high-affinity for both NADH and NADPH, as described also for Sp. passalidarum [18], but with a maximal capacity (V max ) with NADH, which is~35-40% the V max determined with NADPH. Indeed, this gene is closely related to other known yeast xylose reductases that accept both co-substrates (NADH and NADPH) from Candida tropicalis [40], C. parapsilosis [41], C. intermedia [42], and Scheffersomyces stipitis [43]. It is worth noting that the predicted (but not functionally characterized) xylose reductase from Sp. roraimanenses (a species with strictly NADPH-dependent xylose reductase activity, see [19]) is 98% identical to the enzyme (encoded by SaXYL1) cloned from Sp. arborariae, differing in only two amino acids: an aspartate (D 29 ) in Sp. roraimanenses xylose reductase is substituted by a glutamate (E 29 ) in SaXYL1, and the amino acid arginine (R 60 ) from the Sp. roraimanenses enzyme is replaced by a lysine (K 60 ) in SaXYL1. Further work would be required to verify if these two amino acid substitutions are indeed responsible for the NADH-dependent xylose reductase activity of SaXYL1. The cloned SpXYL1.1 xylose reductase, a gene with 93% identity with SaXYL1, is a NADPH-dependent enzyme with affinities for NADPH and xylose similar to the enzyme cloned from Sp. arborariae (Table 4), but with much higher maximal capacity (V max =~4.5 U/[mg protein]) than the cloned SaXYL1 enzyme. Thus, this enzyme is similar to other NADPH-dependent xylose reductases from Pachysolen tannophilus [44] and Debaryomyces nepalensis [45]. Regarding the SpXYL2.2 gene (Table 3), it also encodes for a functional and highly active (2.2 U/[mg protein]) xylitol dehydrogenase when expressed in S. cerevisiae, but showing relatively low affinity (Table 4) for both substrates (NAD + and xylitol), when compared with Sp. passalidarum cells [18]. Unfortunately, we do not have any kinetic parameters of the SpXYL2.1 enzyme cloned in S. cerevisiae by Mamoori and co-workers [23]. Nevertheless, the enzyme encoded by SpXYL2.2 is similar to xylitol dehydrogenases cloned and characterized from other yeasts [46,47].
We next tested the functionality of these enzymes by their co-expression in S. cerevisiae. For the first approach, we expressed the SaXYL1 or SpXYL1.1 genes from plasmid pPGK (with URA3 as the selectable marker) and the SpXYL2.2 gene in plasmid p423-TEF (with HIS3 as the selectable marker) in the strain CEN.PK2-1C. Although the activities were expressed as expected in the yeast strain (see Tables 3-5), these cells were unable to grow or consume xylose from the medium (data not shown). To allow growth on xylose, the xylulokinase gene (XKS1) gene from S. cerevisiae was over-expressed to increase the flux of carbon into the pentose-phosphate pathway. While the CEN.PK2-1C strain showed a xylulokinase activity of 0.04 ± 0.01 U/[mg protein], the engineered P ADH1 ::XKS1 strain ASY-1 (Table 1) showed a~4-fold increase in xylulokinase activity (0.15 ± 0.04 U/[mg protein]). As such, it was observed growth and xylose consumption when the S. cerevisiae ASY-1 strain was transformed with either the pPGK-SpXYL1.1 and pTEF-SpXYL2.2, or the pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids (Figure 1).    Table 6 shows that the ASY-1 strain over expressing the SaXYL1 xylose reductase that accepts both co-substrates (NADH and NADPH) consumes more xylose that the same strain over-expressing the NADPH-dependent SpXYL1.1 xylose reductase, although the differences are not statistically significant. For both strains the only product of xylose consumption (besides biomass) was ~2 g/L xylitol, and, while the final yields were similar for both strains, the volumetric xylitol productivity by the strain over-expressing the SaXYL1 xylose reductase was higher than the strain over-expressing the SpXYL1.1 enzyme (Table 6).   Figure 1 and Table 6 shows that the ASY-1 strain over expressing the SaXYL1 xylose reductase that accepts both co-substrates (NADH and NADPH) consumes more xylose that the same strain over-expressing the NADPH-dependent SpXYL1.1 xylose reductase, although the differences are not statistically significant. For both strains the only product of xylose consumption (besides biomass) was~2 g/L xylitol, and, while the final yields were similar for both strains, the volumetric xylitol productivity by the strain over-expressing the SaXYL1 xylose reductase was higher than the strain over-expressing the SpXYL1.1 enzyme (Table 6).
In order to improve xylose consumption, we deleted in the ASY-1 strain the PHO13 gene of S. cerevisiae, a gene encoding for an alkaline phosphatase known to suppress xylose utilization by recombinant S. cerevisiae strains [34]. Indeed, it has been recently reported that PHO13 deletion leads to up-regulation of the pentose phosphate genes, including TAL1 (encoding transaldolase) and TKL1 (encoding transketolase), an up-regulation mediated by the transcription factor STB5, enhancing xylose consumption by recombinant S. cerevisiae cells [35,36]. As can be seen in Figure 1, the P ADH1 :XKS1 plus pho13∆ strain ASY-2 transformed with the same pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids, which showed improved xylose consumption and significantly higher growth rates on this carbon source, increased the production of xylitol to~5.5 g/L, and, consequently, significantly higher volumetric productivities (Table 6). A recent publication [48] also overexpressed the Sp. passalidarum xylose metabolizing enzymes, as well as a Piromyces sp. xylose isomerase, in Aureobasidium pullulans, in order to improve xylose utilization by this yeast-like fungus. The best xylose consumption, and product production (pullulan and melanin), were obtained when the SpXYL1.2 and SpXYL2.2 genes were introduced into A. pullulans, a direct consequence of the higher activities achieved with these xylose reductase and xylitol dehydrogenases enzymes from Sp. passalidarum [48]. Figure 2 shows the xylose fermentation kinetics during batch fermentations with high cell densities by the ASY-2 strain transformed with the pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids. Xylitol (~10 g/L) was the main product of xylose fermentation, but some ethanol (<2 g/L), glycerol (<0.5 g/L), and acetate (<0.3 g/L) were also produced ( Figure 2 and Table 6). Due to this acetate production, the pH of the medium dropped from an initial pH of 5.0 into pH~3.5. Considering that xylose was not completely consumed during the batch fermentation, in a further attempt to increase the consumption of xylose, we changed the plasmids/promoters of the two cloned genes to improve their activities. When the ASY-2 strain was transformed with the pGPD-SaXYL1 and pPGK-SpXYL2.2 plasmids, a 2-fold higher xylose reductase activity and a 6-fold higher xylitol dehydrogenase activity were obtained (Table 5). However, xylose consumption was not improved in this new recombinant strain. Although the recombinant strain produced no acetate (and consequently the pH of the medium dropped just to pH 4.5) and less xylitol and ethanol, the production of glycerol was only slightly increased ( Figure 2, Table 6), indicating that other factors may limit xylose utilization by our engineered yeast strains. For example, the bottleneck may be a consequence of the relatively low affinity of the cloned SpXYL2.2 xylitol dehydrogenase for both NAD + and xylitol (Table 4), or the intracellular pools of reduced or oxidized and NADH/NADPH ratio of co-substrates [49,50], or even the low affinity of yeast sugar permeases for xylose transport [5,14,51]. Nevertheless, the ASY-2 strain transformed with the pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids (Table 6) showed xylitol yields (Y p/s = 0.614 g xylitol/g xylose) and volumetric productivities (Q p = 0.513 g/L/h) as good as or superior to those reported by other naturally xylose fermenting yeasts [52][53][54][55] or even engineered S. cerevisiae strains [38,49,50]. It is important to note that the maximal expected theoretical yield for the biotransformation of xylose into xylitol is 0.905-0.917 g xylitol/g xylose consumed, depending on how the cells will regenerate the NADH/NADPH consumed in the reduction of xylose, and that no carbon is used for cell growth or production of other metabolites [56]. Finally, Figure 3 shows the fermentation kinetics during batch co-fermentations of 2% glucose plus 2% xylose with high cell densities by the ASY-2 strain transformed with the pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids. As expected for S. cerevisiae, glucose is rapidly consumed in the first 5 h of incubation, while xylose consumption was significantly slower and only 66.45 ± 3.19% of the pentose was consumed after 50 h of incubation. Ethanol (~9.7 g/L, Yp/s ethanol = 0.460 ± 0.001 g ethanol/g glucose), glycerol (~2 g/L, Yp/s glycerol = 0.094 ± 0.001 g glycerol/g glucose), and a small amount of acetate (<0.5 g/L, Yp/s acetate = 0.021 ± 0.001 g acetate/g glucose) were produced during glucose consumption, although we cannot rule out the possibility that some of these products could come from the ~3.6 g/L of xylose assimilated during this period of glucose consumption. From xylose consumption, ~8.5 g/L of xylitol (Yp/s xylitol = 0.614 ± 0.030 g xylitol/g xylose) and ~0.8 g/L of acetate (Yp/s acetate = 0.061 ± 0.012 g acetate/g xylose) were produced ( Figure 3). While the xylitol yield was similar to the one obtained during batch fermentation of 2% xylose (see Table 6), the acetate produced during the cofermentation of glucose and xylose also promoted a drop of the pH of the medium to pH ~3.5. Furthermore, we also observed a drop in the volumetric xylitol productivity (Qp = 0.238 ± 0.025 g/L / h) during glucose-xylose co-fermentation, when compared with fermentation of just xylose by these cells (see Table 6). From a biorefinery perspective, higher xylose concentrations (>100 g/L) should be tested, in order to obtain higher xylitol titers (>60 g/L), as these fermented broths containing xylitol need to be clarify, concentrated (>750 g/L), and cooled in order to favor its crystallization, all this downstream processing contributing significantly to the overall costs of the product [57,58]. Thus, further research should improve the fermentative production of this interesting sugar alcohol with wide commercial applications in pharmaceuticals, nutraceuticals, food and beverage industries. Finally, Figure 3 shows the fermentation kinetics during batch co-fermentations of 2% glucose plus 2% xylose with high cell densities by the ASY-2 strain transformed with the pPGK-SaXYL1 and pTEF-SpXYL2.2 plasmids. As expected for S. cerevisiae, glucose is rapidly consumed in the first 5 h of incubation, while xylose consumption was significantly slower and only 66.45 ± 3.19% of the pentose was consumed after 50 h of incubation. Ethanol (~9.7 g/L, Y p/s ethanol = 0.460 ± 0.001 g ethanol/g glucose), glycerol (~2 g/L, Y p/s glycerol = 0.094 ± 0.001 g glycerol/g glucose), and a small amount of acetate (<0.5 g/L, Y p/s acetate = 0.021 ± 0.001 g acetate/g glucose) were produced during glucose consumption, although we cannot rule out the possibility that some of these products could come from the~3.6 g/L of xylose assimilated during this period of glucose consumption. From xylose consumption,~8.5 g/L of xylitol (Y p/s xylitol = 0.614 ± 0.030 g xylitol/g xylose) and~0.8 g/L of acetate (Y p/s acetate = 0.061 ± 0.012 g acetate/g xylose) were produced ( Figure 3). While the xylitol yield was similar to the one obtained during batch fermentation of 2% xylose (see Table 6), the acetate produced during the co-fermentation of glucose and xylose also promoted a drop of the pH of the medium to pH~3.5. Furthermore, we also observed a drop in the volumetric xylitol productivity (Q p = 0.238 ± 0.025 g/L/h) during glucose-xylose co-fermentation, when compared with fermentation of just xylose by these cells (see Table 6). From a biorefinery perspective, higher xylose concentrations (>100 g/L) should be tested, in order to obtain higher xylitol titers (>60 g/L), as these fermented broths containing xylitol need to be clarify, concentrated (>750 g/L), and cooled in order to favor its crystallization, all this downstream processing contributing significantly to the overall costs of the product [57,58]. Thus, further research should improve the fermentative production of this interesting sugar alcohol with wide commercial applications in pharmaceuticals, nutraceuticals, food and beverage industries.

Conclusions
In the present work, we cloned and expressed, in S. cerevisiae, the Sp. arborariae xylose reductase (SaXYL1) gene that accepts both NADH and NADPH as cofactors, as well as a Sp. passalidarum NADPH-dependent xylose reductase (SpXYL1.1 gene), and the SpXYL2.2 gene encoding a NAD + -dependent xylitol dehydrogenase. The co-expression of both the SaXYL1 and SpXYL2.2 enzymes in a PADH1:XKS1/pho13Δ S. cerevisiae strain allowed efficient growth and xylose consumption by the cells, producing xylitol as the major fermentation product.

Conclusions
In the present work, we cloned and expressed, in S. cerevisiae, the Sp. arborariae xylose reductase (SaXYL1) gene that accepts both NADH and NADPH as cofactors, as well as a Sp. passalidarum NADPH-dependent xylose reductase (SpXYL1.1 gene), and the SpXYL2.2 gene encoding a NAD + -dependent xylitol dehydrogenase. The co-expression of both the SaXYL1 and SpXYL2.2 enzymes in a P ADH1 :XKS1/ pho13∆ S. cerevisiae strain allowed efficient growth and xylose consumption by the cells, producing xylitol as the major fermentation product.