Overcoming inefficient cellobiose fermentation by cellobiose phosphorylase in the presence of xylose
© Chomvong et al.; licensee BioMed Central Ltd. 2014
Received: 19 February 2014
Accepted: 21 May 2014
Published: 7 June 2014
Cellobiose and xylose co-fermentation holds promise for efficiently producing biofuels from plant biomass. Cellobiose phosphorylase (CBP), an intracellular enzyme generally found in anaerobic bacteria, cleaves cellobiose to glucose and glucose-1-phosphate, providing energetic advantages under the anaerobic conditions required for large-scale biofuel production. However, the efficiency of CBP to cleave cellobiose in the presence of xylose is unknown. This study investigated the effect of xylose on anaerobic CBP-mediated cellobiose fermentation by Saccharomyces cerevisiae.
Yeast capable of fermenting cellobiose by the CBP pathway consumed cellobiose and produced ethanol at rates 61% and 42% slower, respectively, in the presence of xylose than in its absence. The system generated significant amounts of the byproduct 4-O-β-d-glucopyranosyl-d-xylose (GX), produced by CBP from glucose-1-phosphate and xylose. In vitro competition assays identified xylose as a mixed-inhibitor for cellobiose phosphorylase activity. The negative effects of xylose were effectively relieved by efficient cellobiose and xylose co-utilization. GX was also shown to be a substrate for cleavage by an intracellular β-glucosidase.
Xylose exerted negative impacts on CBP-mediated cellobiose fermentation by acting as a substrate for GX byproduct formation and a mixed-inhibitor for cellobiose phosphorylase activity. Future efforts will require efficient xylose utilization, GX cleavage by a β-glucosidase, and/or a CBP with improved substrate specificity to overcome the negative impacts of xylose on CBP in cellobiose and xylose co-fermentation.
KeywordsCellobiose Cellobiose phosphorylase Glucopyranosyl-xylose Inhibition Xylose
Cellulosic biofuels could make significant contributions to meet the ever-rising demand for energy. To economically produce fuels from cellulosic biomass, sugars derived from cellulose as well as hemicellulose must be utilized completely[1–3]. The co-fermentation of cellobiose derived from cellulose and xylose derived from hemicellulose allows these sugars to be consumed simultaneously, and may enable continuous biofuel production. In this system, cellobiose and xylose are transported into engineered S. cerevisiae using a cellodextrin transporter (that is, CDT-1 from Neurospora crassa) and endogenous hexose transporters, respectively[4, 6]. Intracellular cellobiose is then hydrolyzed into two molecules of glucose by an intracellular β-glucosidase (NCU00130; GH1-1). At the same time, xylose is consumed by an oxidoreductive pathway, comprising xylose reductase and xylitol dehydrogenase, that converts xylose to xylulose[7, 8]. Alternatively, xylose can be utilized by xylose isomerase, which converts xylose directly to xylulose. Glucose and xylulose are then metabolized using glycolysis and the pentose phosphate pathway, respectively, resulting in ethanol production.
A pathway potentially better suited to the anaerobic environment of large-scale biofuels production substitutes cellobiose phosphorolysis for the hydrolytic reaction of β-glucosidase[10, 11]. This pathway comprises cellobiose phosphorylase (CBP), which cleaves intracellular cellobiose into glucose and glucose-1-phosphate (G1P). The phosphorolytic pathway requires one less ATP for each molecule of cellobiose to be metabolized by glycolysis. This is because glucose generated by hydrolysis of cellobiose requires two ATP molecules for hexokinase generation of glucose-6-phosphate (G6P), whereas CBP uses inorganic phosphate in place of one of the ATP molecules to produce G1P. G1P can then be converted to G6P without the need for ATP by the enzyme phosphoglucomutase. Under anaerobic conditions, in which glycolysis generates only two ATP molecules per glucose, increased ATP can result in increased biomass at the expense of ethanol product yield. However, the phosphorolytic pathway can be engineered to perform better than the hydrolytic pathway in terms of product yield in stressful conditions like those expected in lignocellulosic hydrolysates.
Although the cellobiose phosphorolytic pathway has potential advantages, its efficiency, hereafter defined as incomplete or low rate of consumption, in the context of cellobiose and xylose co-fermentation is not known. Previously, cellobiose phosphorolytic pathways were combined with xylose isomerase pathways to construct anaerobic cellobiose and xylose co-fermentation systems in Saccharomyces cerevisiae and in Escherichia coli[16, 17]. However, they are inefficient in terms of sugar consumption and ethanol production rates, or the systems remain to be fully optimized. CBP from Ruminococcus albus NE1 (RaCBP) uses xylose as a substrate for the reverse of the phosphorolytic reaction. We therefore hypothesized that the inefficiency in cellobiose and xylose co-fermentation previously observed was due to xylose interference with cellobiose consumption via CBP. The presence of xylose is unavoidable because it is a major component of hemicellulose, which has to be utilized for economical biofuel production[1–3]. We therefore tested the effect of xylose on CBP cellobiose fermentation, as well as two potential approaches to alleviate inefficient CBP-mediated cellobiose fermentation in the presence of xylose.
Inefficient cellobiose fermentation in the presence of xylose
Strains and plasmids used in this study
MATalpha, leu2, his3, ura3, and can1
ald6Δ of the evolved strain D452-2 leu2::LEU2 P TDH3 -XYL1-T TDH3 ; ura3::URA3 P TDH3 -XYL1- T TDH3 P PGK1 -XYL2- T PGK1 P TDH3 -XYL3- T TDH3 ; his1::HIS1 P PGK1 -XYL2- T PGK1 P TDH3 -XYL3- T TDH3
Courtesy of Dr. Soo Rin Kim
pRS426- P PGK1 -cdt-1(F213L)-eGFP-t CYC1
pRS425- P PGK1 -SdCBP-t CYC1
pRS426-P PGK1 -cdt-1(F213L)-eGFP-t CYC1 -P PGK1 -SdCBP-t CYC1
Cellobiose consumption rate (g/L · h)
Ethanol production rate (g/L · h)
3.6 ± 0.05
1.5 ± 0.03
Cellobiose + xylose
1.4 ± 0.04
0.85 ± 0.02
3.7 ± 0.09
1.5 ± 0.04
Cellobiose + xylose
2.9 ± 0.08
1.6 ± 0.04
In vitro and in vivo production of glucopyranosyl-xylose
Using GX synthesized in vitro as a standard, GX was detected in the fermentation broth when cellobiose and xylose were supplied to yeast engineered with the CBP cellobiose consumption pathway (Figure 1E). Interestingly, the concentration of GX initially increased but started to decrease after 36 hours, with its highest concentration reaching approximately 30 g/L (Figure 1E). Extracellular concentrations of xylitol and GX combined accounted for 88% to 100% of the imported xylose (Additional file1: Figure S2). Thus, yeast utilizing cellobiose by the CBP consumption pathway formed GX from intracellular xylose and G1P, in addition to converting some of the imported xylose to xylitol.
Competition assays identified xylose as a mixed-inhibitor
Reduced glucopyranosyl-xylose formation in an efficient xylose utilizing strain, SR8-a
The cellobiose consumption rate of the SR8-a strain supplemented with xylose was not as rapid as that of the D452-2 strain with no xylose present, showing a 22% decrease (Figure 1A, Figure 4A, Table 2). However, the negative effect of xylose on cellobiose consumption was alleviated in comparison to the 61% decrease in cellobiose consumption rate with the D452-2 strain in the presence of xylose (Figure 1A).
Cleavage of glucopyranosyl-xylose by intracellular β-glucosidase and β-xylosidase
Kinetic parameters of β-glucosidase GH1-1 for glucopyranosyl-xylose and cellobiose
3.5 ± 1.4
0.49 ± 0.05
Vmax (μM · min-1 · nM-1)
0.48 ± 0.10
1.3 ± 0.04
In this study, we identified GX production as a competing off-pathway product of cellobiose and xylose co-fermentation when cellobiose is consumed using a CBP-mediated pathway. The production of GX results in a decreased efficiency of cellobiose fermentation (Figure 1), especially in the absence of a xylose-consumption pathway. The decrease in extracellular xylose concentration (Figure 1C) indicates that xylose is transported into the cell, likely via endogenous hexose transporters. Though lacking a xylose utilization pathway, the engineered yeast strain D452-2 expressing the pCS plasmid is expected to convert some xylose to xylitol by means of endogenous aldose reductase (Gre3) activity, consistent with the fact that xylitol was detected in the fermentation medium (Figure 1D). We further verified that much of the remaining xylose loss in the intracellular pool was due to its conversion to GX by the reverse phosphorolysis reaction catalyzed by CBP between G1P and xylose (Additional file1: Figure S2). Interestingly, we found that the extracellular concentration of GX eventually starts to decrease, and at the same time that of xylose begins to recover (Figure 1C,E).
The reported catalytic efficiency (kcat,app/KM,app) of RaCBP for xylose in the reverse phosphorolysis reaction is only 1% of that for glucose. However, with SdCBP, a substantial amount of GX formation is observed (Figure 1E). This may be explained by differences between the two CBPs, or by the high concentrations of intracellular xylose and low concentrations of intracellular glucose present in our experiments. When the xylose utilization pathway is absent, high intracellular xylose concentrations are expected. Although we did not measure the intracellular concentration of xylose directly, it is expected to be similar to the extracellular concentration due to the fact that xylose is imported by hexose transporters, which are facilitators[23, 24]. Thus, the imported xylose not accounted for by xylitol production (Figure 1C,D), would result in an intracellular concentration of xylose near or above the reported KM,app of RaCBP for xylose (25 mM). By contrast, the intracellular concentration of glucose is expected to be small because glucose can be efficiently converted to G6P and consumed by glycolysis. The maximal reported free intracellular glucose concentration is 2 to 3 mM, slightly above the KM,app of RaCBP on glucose (1.5 mM). Furthermore, the CBP reverse reaction is thermodynamically favorable, with a ΔG° = -3.6 kJ mol-1 for cellobiose formation[10, 12]. Thus, the amount of GX formation we observed was consistent with the known thermodynamic and kinetic properties of the enzymes used in the CBP-mediated cellobiose consumption pathway, especially due to the drive from a high intracellular xylose concentration.
By carrying out cellobiose and xylose co-fermentation using an efficient xylose-utilizing strain, SR8-a, we were able to increase the cellobiose consumption rate and decrease GX titer (Figure 4A,B), likely by keeping the steady-state intracellular xylose concentration low. The low concentrations of xylose present inside the cell would improve the apparent kinetic properties of cellobiose phosphorylation, resulting in a faster cellobiose consumption rate (Figure 3). Furthermore, with less intracellular xylose present, less substrate is available for GX formation. Thus, smaller amounts of GX are made, exported out of the cell and re-imported, reserving the capacity of the cellodextrin transporter and CBP for cellobiose phosphorolysis (Figure 6). Finally, less GX formation allows G1P to more efficiently enter into glycolysis by its conversion to G6P, thereby increasing the ethanol production rate. Thus, an efficient xylose utilization pathway can be used to alleviate cellobiose fermentation inefficiencies due to the use of CBP.
For bacteria with CBP, ORFs encoding xylose isomerase are found to co-exist, suggesting that cellobiose and xylose co-fermentation by anaerobic bacteria may be common, for example in S. degradans, Cellvibrio gilvus, Ruminococcus sp. and Clostridium phytofermentans. Although co-fermentation has not been shown with these organisms, they may have evolved means of avoiding the production of GX. Previous efforts to construct an anaerobic cellobiose and xylose co-fermentation system in S. cerevisiae using CBP and xylose isomerase from Ruminococcus flavefaciens were only partly successful. This may be due to the inefficient xylose isomerase conversion step, resulting in high intracellular concentrations of xylose that negatively impact CBP-mediated cellobiose consumption.
We also explored whether CBP-mediated cellobiose conversion in the presence of xylose could be augmented by the use of a hydrolytic enzyme to cleave GX after its formation. We found that the intracellular β-glucosidase GH1-1 from N. crassa was capable of GX hydrolysis to glucose and xylose (Figure 5). Thus, low levels of GH1-1 co-expressed with CBP might be used to reduce GX and its associated burdens on the cellobiose consumption pathway (Figure 6). However, cellobiose was preferred as a substrate for GH1-1 in comparison to GX (Table 3) and the catalytic efficiency of GH1-1 for cellobiose is higher than that of CBP for cellobiose. Hence, co-expression of GH1-1 with CBP would likely result in most of the cellobiose being hydrolyzed to glucose instead of following the phosphorolytic pathway. This effect would defeat the purpose of using CBP for its energetic advantage, because G1P generation would be replaced by glucose production. To circumvent this challenge, the intracellular β-glucosidase would need to have an increased substrate specificity for GX and lower activity for cellobiose. Protein engineering of an intracellular β-glucosidase with these properties may be feasible, because xylose is smaller than glucose. Thus, the enzyme active site of β-glucosidase could be engineered to be more bulky, allowing the binding of GX while eliminating that of cellobiose. Successes in similar protein engineering challenges have been reported[30–32].
Alternatively, CBP could be engineered to reduce or eliminate GX production. This approach is advantageous because it addresses the GX complications directly. In contrast to the use of engineered GH1-1 for GX cleavage, this approach allows the system to fully harvest the energetic advantages of the phosphorolytic pathway. However, CBP protein engineering may be challenging because xylose is a mixed-inhibitor of CBP activity (Figure 3), and therefore may require random mutagenesis, multiple site saturation mutagenesis or evolutionary engineering approaches to achieve the necessary cellobiose specificity[30, 33, 34].
We have shown that xylose can have negative impacts on anaerobic cellobiose fermentation mediated by CBP in S. cerevisiae. Xylose can serve as a substrate along with G1P in a favorable reverse reaction to form the byproduct GX dimer. We have provided evidence that GX is likely exported out of cells and imported back by the exogenous cellodextrin transporter before being cleaved by SdCBP, exhausting resources that could have been reserved for cellobiose fermentation. Additionally, we identified xylose as a mixed-inhibitor of CBP activity, possibly due to the arrangement of enzyme active sites in the CBP homodimer. Cellobiose and xylose co-fermentation by the efficient xylose-utilizing SR8-a strain increased the cellobiose fermentation rate and decreased GX formation, likely by maintaining a low intracellular xylose concentration. The intracellular β-glucosidase GH1-1 from N. crassa was also capable of cleaving GX, and could be used to augment the CBP-mediated cellobiose consumption pathway. However, the use of an intracellular β-glucosidase alongside CBP may require further protein engineering to improve the β-glucosidase specificity for GX over cellobiose.
Plasmids constructed and used in this study are listed in Table 1. Plasmids containing a codon-optimized CBP gene from S. degradans (SdCBP) [GenBank: 90020965] and cellodextrin transporter mutant from N. crassa cdt-1 (F213L) were used as templates for combining cdt-1 (F213L) and SdCBP expression cassettes in pRS426 (pCS). SdCBP was cloned into E. coli expression plasmid pET302 with an N-terminal His6 tag to create pET-Sd. The In-Fusion HD Cloning Kit (Clontech, Mountain View, CA, USA) was used for all plasmid construction. Primers used are listed in Additional file1: Table S1.
Yeast strains and media
S. cerevisiae background strains used in this study were D452-2 (MATα leu 2 his 3 ura 3 can 1) and SR8-a (ura 3) (Table 1). Plasmids were transformed into these strains using a standard lithium acetate yeast transformation protocol. Transformants were selected on synthetic defined medium plates, which contained DOBA (MP Biomedicals, Santa Ana, CA, USA) mixed with two-fold appropriate CSM dropout mixture.
Single colonies from synthetic defined plates were selected and re-streaked. Re-streaked colonies were inoculated in optimal minimal medium (oMM) supplemented with 20 g/L of cellobiose to prepare seed cultures. The oMM contained 1.7 g/L YNB Y1251 (Sigma, Saint Louis, MO, USA), two-fold appropriate CSM dropout mixture, 10 g/L (NH4)2SO4, 1 g/L MgSO4.7H2O, 6 g/L KH2PO4, 100 mg/L adenine hemisulfate, 10 mg/L inositol, 100 mg/L glutamic acid, 20 mg/L lysine, 375 mg/L serine and 0.1 M 2-(N-morpholino) ethanesulfonic acid (MES) pH 6.0. Seed cultures were harvested at mid- to late-exponential phase and washed twice with sterile water. Washed seed cultures were inoculated at an initial optical density at 600 nm of 20 in 200 mL serum flasks containing 50 mL of media. The flasks were closed with butyl rubber stoppers, sealed with aluminum crimps, and purged with nitrogen gas to obtain strict anaerobic fermentations. The fermentation media contained oMM supplemented with 80 g/L cellobiose, with or without 40 g/L xylose. The flasks were incubated at 30°C, 220 rpm. Extracellular concentrations of cellobiose, xylose, xylitol and ethanol were determined by high performance liquid chromatography on a Prominence HPLC (Shimadzu, Kyoto, Japan) equipped with Rezex RFQ-FastAcid H 10 × 7.8 mm column. The column was eluted with 0.01 N of H2SO4 at a flow rate of 1 mL/min, 55°C. Quantification of GX was performed using an ICS-3000 Ion Chromatography System (Dionex, Sunnyvale, CA, USA) equipped with a CarboPac® PA200 carbohydrate column. The column was eluted with a NaOAc gradient in 100 mM NaOH at a flow rate of 0.4 mL/min, 30°C.
SdCBP protein purification
pET-Sd (pET302-NT/His6-SdCBP) was transformed into the BL21 (DE3) E. coli strain and induced by isopropyl β-D-1-thiogalactopyranoside at a final concentration of 0.2 mM. E. coli cells were lysed and protein purified by His.Bind Resin (Novagen, Darmstadt, Germany) according to supplied protocols. Purified SdCBP was stored in 20 mM MES, pH 6.0 and quantified using a NanoDrop 1000 spectrophotometer, assuming an extinction coefficient of 1.79 × 105 M-1 cm-1, 280 nm.
In vitro synthesis and purification of glucopyranosyl-xylose
We incubated 10 mM xylose and 10 mM G1P with and without 20 nM purified SdCBP in 20 mM MES, pH 6.0 at 37°C for 12 hours. The reaction was stopped by dilution with 0.1 M NaOH at a ratio of 1:200. Signals of components in the solutions were detected using an ICS-3000 Ion Chromatography System (Dionex) with the same conditions described above.
The synthesized GX was purified by ÄKTA Purifier (GE Healthcare Life Sciences, Munich, Germany) equipped with a Supelclean™ ENVI-Carb™ column. The column was eluted with a gradient of acetonitrile at a flow rate of 3.0 mL/min at room temperature. Purified fractions were verified using an ICS-3000 Ion Chromatography System (Dionex) with the same conditions described above.
Mass spectrometry and tandem mass spectrometry
MS of the GX in the in vitro synthesis solution was performed on an LTQ XL ion trap instrument (Thermo Fisher Scientific, San Jose, CA, USA) with an electrospray ionization source operated in negative mode. The sample was introduced into the mass spectrometer by direct injection into a flow of 50% water/0.1% formic acid and 50% acetonitrile/0.1% formic acid set at a flow rate of 0.2 mL/min. The MS settings were capillary temperature 350°C, ion spray voltage 4 kV, sheath gas flow 60 (arbitrary units), auxiliary gas flow 10 (arbitrary units), sweep gas flow 5 (arbitrary units). The scan rate for full scan and MS/MS product ion scan was m/z 95 to m/z 500. The compound at m/z 357 was isolated with a m/z 2 isolation width (±1 Da) and fragmented with a normalized collision-induced dissociation energy setting of 35%. The activation time and the activation Q were 30 ms and 0.250, respectively. The mass measurement accuracy was < 3 ppm root mean square.
Competition assay and kinetic parameters
We incubated 10 nM of purified SdCBP, 5 mM of inorganic phosphate and varying cellobiose concentrations at 30°C in 20 mM MES, pH 6.0 with 0, 2.5 or 5 mM xylose. All experiments were carried out in duplicate. G1P concentrations were detected continuously using a G1P Colorimetry Assay Kit (Sigma-Aldrich), according to the provided protocol. Initial rates at each cellobiose concentration were calculated from the rate of G1P production. Apparent kinetic parameters were determined by non-linear regression.
In vitro glucopyranosyl-xylose hydrolytic activity assay
GX synthesized as described above was used at a concentration of 1 mM. The substrate was incubated with 0.5 μM of purified β-xylosidase (NCU01900) or β-glucosidase (NCU00130) in 1× PBS, pH 7.4 at 30°C for 2 hours. β-xylosidase (NCU01900) (unpublished observations by Dr. Xin Li) and β-glucosidase (NCU00130) were expressed and purified as described. To stop the reactions, they were diluted with 0.1 M NaOH. Signals of components in the solutions were detected using an ICS-3000 Ion Chromatography System (Dionex) with the same conditions described above.
For kinetic parameter comparisons, 20 nM of purified GH1-1 and varying concentrations of cellobiose and purified GX were incubated at 30°C in 1× PBS, pH 7.4. All reactions were carried out in duplicate. The reactions were stopped at 0, 5, 10 and 15 minutes by addition of 0.1 M NaOH. Initial rates at each cellobiose and GX concentration were calculated from the rate of glucose production. The ICS-3000 Ion Chromatography System (Dionex) equipped with CarboPacTM PA20 column was eluted with 3 mM KOH at a flow rate of 0.4 mL/min, 30°C, for glucose and xylose separation and to determine glucose concentrations in the reactions. Apparent kinetic parameters were determined by non-linear regression.
cellodextrin transporter from Neurospora crassa
- CDT-1 (F213L):
mutant of cellodextrin transporter from N. crassa
2-(N-morpholino) ethanesulfonic acid
tandem mass spectrometry
optimal minimal media
open reading frame
phosphate buffered saline
cellobiose phosphorylase from Ruminococcus albus NE1
cellobiose phosphorylase from Saccharophagus degradans.
This work was supported by funding from the Energy Biosciences Institute to YSJ and JHDC. The authors thank Dr Soo Rin Kim for generously providing S. cerevisiae SR8-a, and Dr Yuping Lin, Dr Ligia Acosta-Sampson, and Dr Owen W Ryan for helpful discussions.
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