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Applied and Environmental Microbiology, January 2009, p. 395-404, Vol. 75, No. 2
0099-2240/09/$08.00+0     doi:10.1128/AEM.01941-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.

Complete Fermentation of Xylose and Methylglucuronoxylose Derived from Methylglucuronoxylan by Enterobacter asburiae Strain JDR-1{triangledown}

Changhao Bi, John D. Rice, and James F. Preston*

Department of Microbiology and Cell Science, University of Florida, Gainesville, Florida 32611

Received 20 August 2008/ Accepted 8 November 2008


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ABSTRACT
 
Acid pretreatment is commonly used to release pentoses from the hemicellulose fraction of cellulosic biomass for bioconversion. The predominant pentose in the hemicellulose fraction of hardwoods and crop residues is xylose in the polysaccharide methylglucuronoxylan, in which as many as one in six of the β-1,4-linked xylopyranose residues is substituted with {alpha}-1,2-linked 4-O-methylglucuronopyranose. Resistance of the {alpha}-1,2-methylglucuronosyl linkages to acid hydrolysis results in release of the aldobiuronate 4-O-methylglucuronoxylose, which is not fermented by bacterial biocatalysts currently used for bioconversion of hemicellulose. Enterobacter asburiae strain JDR-1, isolated from colonized hardwood (sweetgum), efficiently ferments both methylglucuronoxylose and xylose, producing predominantly ethanol and acetate. 13C-nuclear magnetic resonance studies defined the Embden-Meyerhof pathway for metabolism of glucose and the pentose phosphate pathway for xylose metabolism. Rates of substrate utilization, product formation, and molar growth yields indicated methylglucuronoxylose is transported into the cell and hydrolyzed to release methanol, xylose, and hexauronate. Enterobacter asburiae strain JDR-1 is the first microorganism described that ferments methylglucuronoxylose generated along with xylose during the acid-mediated saccharification of hemicellulose. Genetic definition of the methylglucuronoxylose utilization pathway may allow metabolic engineering of established gram-negative bacterial biocatalysts for complete bioconversion of acid hydrolysates of methylglucuronoxylan. Alternatively, Enterobacter asburiae strain JDR-1 may be engineered for the efficient conversion of acid hydrolysates of hemicellulose to biofuels and chemical feedstocks.


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INTRODUCTION
 
The quest for alternatives to petroleum resources for production of fuels and chemicals has been motivated in part by economic incentives associated with limited and diminishing supplies and in part by the recognition that increasing levels of carbon dioxide derived from these resources have contributed to global warming (14). Microbial fermentation products derived from resources renewable through photosynthesis are particularly attractive alternatives that may be generated with existing technologies (17). The development of yeast and bacterial biocatalysts has been applied to the commercial production of ethanol as an alternative fuel from starch and sucrose derived from commodity crops, e.g., corn and sugarcane (3, 12). To expand production of ethanol and chemical feedstocks from renewable resources that do not economically impact these commodities, lignocellulosic resources, including forest and agricultural residues, have become targets for bioconversion of cellulose and hemicellulose to fermentable sugars (12, 29, 35).

Current industrial methods for pretreatment of lignocellulose for bioconversion to ethanol solubilize the hemicellulose fraction by dilute acid hydrolysis, releasing the pentoses for fermentation (15). The predominant structural polymer in the hemicellulose fraction of hardwoods and crop residues is methylglucuronoxylan (MeGAXn), a β-1,4-linked xylan in which xylose residues are periodically substituted with {alpha}-1,2-linked 4-O-methyl-glucuronic acid (MeGA). Variable substitutions on xylose residues may include 2'- and 3'-O-acetyl esters, as well as {alpha}-1,2- or {alpha}-1,3-linked L-arabinofuranosyl residues (21). Resistance of the {alpha}-1,2 glucuronosyl linkages to dilute acid hydrolysis results in the release of the aldobiuronate methylglucuronoxylose (MeGAX), which is not fermented by bacterial biocatalysts currently used to convert hemicellulose-derived xylose to ethanol, e.g., Escherichia coli KO11(11, 21, 25). The frequency of MeGA substitutions on the xylose residues of methylglucuronoxylan ranges from less than 1 in 10 in crop residues to 1 in 6 to 7 in hardwoods, e.g., sweetgum and yellow poplar, and as much as 18% to 27% of the carbohydrate may reside in this unfermentable fraction (primarily as MeGAX) following dilute acid pretreatment (25). A scheme for the release of xylose and MeGAX by dilute acid hydrolysis of sweetgum methylglucuronoxylan is depicted in Fig. 1.


Figure 1
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  		FIG. 1. Scheme for the release of xylose and MeGAX by dilute acid hydrolysis of sweetgum xylan.

In this report, we describe the isolation and characterization of a bacterium, Enterobacter asburiae strain JDR-1, that has the ability to ferment MeGAX as the sole carbon source. E. asburiae JDR-1 was defined with respect to its metabolic potential for the conversion of MeGAX, xylose, and glucose to specific fermentation products. As a gram-negative member of the Enterobacteriaceae, the genetic basis for this metabolic potential may be applied to the further development of bacterial biocatalysts for conversion of hemicellulose-derived pentoses to biobased products.


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MATERIALS AND METHODS
 
Preparation of substrates and culture media.
Sweetgum methylglucuronoxylan (MeGAXn) was prepared from sweetgum stem wood (Liquidambar styraciflua) as previously described and characterized by 13C-nuclear magnetic resonance (NMR) (10, 13). Dilute acid hydrolysates of methylglucuronoxylan were prepared by hydrolysis with 0.1 N H2SO4 (4 g methylglucuronoxylan in 400 ml 0.1 N H2SO4) at 121°C for 60 min, followed by neutralization with BaCO3. Anion exchange resin (Bio-Rad AG2-X8) in the acetate form was used to adsorb the charged aldouronates; the uncharged xylose and xylooligosaccharides, mainly small amounts of xylobiose, were eluted with water. The aldouronates were then eluted with 20% (vol/vol) acetic acid. After concentration under vacuum at 50°C, aldouronates were separated on a 2.5-cm by 160-cm BioGel P-2 column (Bio-Rad, Hercules, CA) with 50 mM formic acid as the eluent. The formic acid was removed from the purified sugar sample fractions by lyophilization. MeGAX and MeGAX2 were identified by thin-layer chromatography analysis using MeGAX and MeGAX2 standards structurally defined by 13C- and 1H-NMR spectrometry (37). Xylobiose and xylotriose were obtained and purified from MeGAXn digested with the Paenibacillus sp. strain JDR-2 XynA1 catalytic domain (CD), a recombinant GH10 endoxylanase XynA1 CD overexpressed in E. coli (34). The substrate containing 30 mg/ml MeGAXn was prepared with 10 mM sodium phosphate buffer, pH 6.5. Digestions were initiated by the addition of 3.5 U of XynA1 CD into 50 ml substrate and incubated with rocking at 30°C for 24 h. An additional 1 U was added after 24 h and incubation was continued for 40 h. Aldouronates, xylobiose, and xylotriose were separated with the P2 column and identified by thin-layer chromatography. Total carbohydrate concentrations related to substrate preparations were determined by the phenol-sulfuric acid assay (4), with xylose as the reference. The conditions of acid hydrolysis generated mostly MeGAX and a small amount of MeGAX2 from MeGAXn, with no aldouronates larger than MeGAX2 detected. MeGAX3 was prepared from GH10 endoxylanase-catalyzed depolymerization of sweetgum MeGAXn and then purified by gel filtration on BioGel P4 (34).

Minimal medium containing the substrates described above was prepared upon mixing sterile substrate solutions (2x concentration) with the same volume of a 2x solution of Zucker and Hankin mineral salts (ZH salts) at pH 7.4 (36). Neutralized MeGAXn acid hydrolysate (0.5%, wt/vol) was also added to ZH salts directly as a growth substrate. Where indicated, some medium preparations were supplemented with 0.1% yeast extract.

Isolation and identification of E. asburiae JDR-1.
E. asburiae JDR-1 was isolated from discs of sweetgum stem wood (Liquidambar styraciflua) buried, soon after cutting, about 1 inch below the soil surface in a sweetgum stand for approximately 3 weeks. Discs were suspended in 50 ml sterile deionized water and sonicated in a 125-W Branson ultrasonic cleaner water bath for 10 min. The sonicate was inoculated into 0.2% (wt/vol) MeGAX yeast extract medium and incubated at 37°C. Cultures were streaked on MeGAX minimal medium agar plates. Isolated colonies were passed several times between MeGAX broths and agars until pure. Exponential-phase cultures growing on 0.2% MeGAX minimal medium were cryostored in 25% sterile glycerol at –70°C.

The purified isolate was submitted to MIDI Labs (http://www.midilabs.com) for partial 16S rRNA sequencing and fatty acid methyl ester analysis. BBL Enterotube II (Becton Dickinson and Company) inoculation was also used to identify the isolate based upon metabolic capability using the standard protocol. Differential interference contrast (DIC) micrographs of E. asburiae JDR-1 growing in MeGAX minimal medium at exponential phase were obtained with a Zeiss DIC microscope at 40x and 15-fold magnification. Negative stain electron micrographs were obtained with a Zeiss EM10A electron microscope.

Substrate utilization and fermentation product analysis.
Growth and substrate utilization analyses were performed in cultures aerated by shaking. For preparing inocula, cultures of E. coli B (ATCC 11303) and E. asburiae JDR-1 from cryostored samples were directly streaked on Luria-Bertani (LB) agar plates. After overnight incubation at 37°C, isolated colonies were picked to inoculate liquid medium specified for a particular experiment. Growth studies were performed at 37°C in 16-mm by 100-mm test tubes containing 6 ml medium. Optical densities of cultures were measured at 600 nm (OD600) with a Beckman DU500 series spectrophotometer. The relationship of cell density and OD600 was experimentally determined as grams of cell dry weight per liter with the following equation: cell dry weight per liter = (0.49 OD600) + 0.02. Sample dilutions were made to obtain OD600 readings between 0.2 and 0.8 absorbance units which, corrected for dilution factors, provided turbidity values for growth studies. Individual 6-ml cultures for study were inoculated with 12 µl (0.2% volume) of overnight cultures and maintained at 37°C with constant shaking (Eberbach shaker set at low).

Batch fermentations under anaerobic conditions at 37°C were conducted in 13-mm by 100-mm screw-cap tubes containing 3.0 ml medium. Inocula (0.5% [vol/vol]) were from overnight aerobic cultures grown in the same medium. After inoculation, nitrogen gas was used to flush and saturate the sealed batch culture. The tubes were set in a Glas-Col minirotator at 60 rpm.

For analysis of substrates and fermentation products, cells were removed by centrifugation and supernatants were passed through 0.22-µm filters and subjected to high-performance liquid chromatography (HPLC) analysis. Products were resolved on a Bio-Rad HPX-87H column with 0.01 N H2SO4 as the eluent at 65°C. Samples were delivered with a 710B WISP automated injector and chromatography controlled with a Waters 610 solvent delivery system at flow rate of 0.5 ml/min. Products were detected by differential refractometry with a Waters 2410 RI detector. Data analysis was performed with Waters Millennium software. To determine and quantify methanol, unfiltered supernatants from fermentation cultures were also analyzed by gas chromatography (GC; 6890N Network GC system; Agilent Technologies), using isopropanol as an internal standard. This detection method was used since diffusion during HPLC precluded quantitative detection of methanol by differential refractometry.

Determination of metabolic pathways by 13C-NMR.
The central metabolic pathways utilized by E. asburiae JDR-1 during glucose and xylose fermentation were evaluated with 13C-NMR (28). Cultures were grown in LB medium to mid-exponential phase at 37°C. Cultures (0.5 ml) were centrifuged and the cells washed with 2x ZH salts solution. The cell pellets were suspended in 1.0 ml 0.5% [2-13C]xylose (99% enrichment; Omicron Biochemicals Inc., IN) in ZH minimal medium. Similar fermentations were also prepared with 1.0 ml 0.5% [1-13C]glucose or 1.0 ml 0.5% [6-13C]glucose ZH minimal medium using D-[1-13C]glucose or D-[6-13C]glucose (99% enrichment; Cambridge Isotope Laboratories, Andover, MA). Fermentations were carried out under anaerobic conditions at 37°C for 8 h. Cells were removed by centrifugation, and the supernatants were analyzed by HPLC (after filtration) and 13C-NMR spectrometry. NMR spectra were obtained using a VXR300 NMR spectrometer (NMR facility of the Department of Chemistry, University of Florida) operating in the Fourier transform mode as follows: 75.46 MHz; excitation pulse width, 7.0 s; spectral width, 16,502; 256 acquisitions. Acetone (30 µl) containing 13C at natural abundance in 700 µl of sample was used as an internal reference at 31.07 ppm for the [13C]methyl carbon (13). Individual carbon atoms for fermentation products were identified by shift assignments and quantified by comparison with standards (13C at natural abundance) of known concentrations.

Determination of molar cell dry weight yield.
For molar growth yield experiments (1, 6, 33), anaerobic growth was performed in 50 ml minimal medium containing either 0.26% glucose, 0.36% xylose, 0.35% glucuronate, or 0.2% MeGAX as sole carbon source under the fermentation conditions described above. After 24 h of growth and complete utilization of the carbon source, cells were harvested by centrifugation and the resulting pellets were washed twice with deionized water. The pellets were dried to constant weight in a Sargent vacuum dryer at 60°C for up to 36 h. The culture supernatants were analyzed by HPLC to determine substrate consumption. The molar cell dry weight yield was calculated as cell dry weight (in grams) divided by consumed substrate (in moles).


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RESULTS
 
Identification and characteristics of E. asburiae JDR-1.
A novel bacterial strain able to grow on MeGAX minimal medium was obtained and subsequently identified with three tests. The partial 16S rRNA sequence (GenBank [NCBI] accession number EU117142), amplified using primers corresponding to E. coli 16S rRNA positions 005 and 531 (526 bp), provided an alignment with 99.5% identity within the sequence of Enterobacter asburiae (MIDI Aerobic Bacteria Database version 4.0; January 1999). Results of fatty acid methyl ester analysis indicated this strain had the greatest similarity index with Enterobacter asburiae species (0.766) compared with any other entry in the MIDI database. A biocode of 32061, obtained from the Enterotube II (BBL) test, also corresponded to Enterobacter asburiae species. Based upon these three criteria, the isolate was identified within Enterobacter asburiae species and designated as Enterobacter asburiae strain JDR-1. The strain has been deposited with the Agriculture Research Service Patent Culture Collection of the U.S. Department of Agriculture, Peoria, IL, under NRRL number NRRL B-S0074.

When exponential-phase cultures were observed by optical DIC microscopy, E. asburiae JDR-1 appeared as short motile rods. Negative-stain electron microscopy revealed 3-µm by 1-µm cells with peritrichous flagella. These morphological characteristics were similar to those of other isolates of Enterobacter asburiae (8). When grown on LB agar plates, colonies of E. asburiae JDR-1 were morphologically indistinguishable from E. coli colonies.

Utilization of acid hydrolysates of methylgluronoxylan by E. asburiae JDR-1.
The unique ability of E. asburiae JDR-1 to grow on the aldobiuronate MeGAX as the sole carbon source suggested a potential for the complete metabolism of the carbohydrates generated by the dilute acid pretreatment currently applied for the release and fermentation of xylose in hemicellulose fractions. To evaluate this potential, E. asburiae JDR-1 was grown aerobically in minimal medium comprised of neutralized MeGAXn acid hydrolysate and Zucker and Hankin mineral salts. Based upon HPLC analysis of medium samples taken at different stages of growth, E. asburiae JDR-1 utilized MeGAX completely in minimal medium containing MeGAXn hydrolysate after it depleted xylose (Fig. 2A). Biphasic growth occurred as E. asburiae JDR-1 switched from utilizing xylose to MeGAX. In contrast to E. asburiae JDR-1, E. coli B consumed only the free xylose, with the MeGAX concentration in the medium remaining constant. Concentrations of xylose and MeGAX in MeGAXn hydrolysate medium, as determined by HPLC, were 0.206% (wt/vol) and 0.036% (wt/vol), respectively. Therefore, E. asburiae JDR-1 utilized 17.5% more substrate (mass amount) than E. coli B, which was unable to utilize MeGAX (Fig. 2B). Under aerobic conditions, both E. asburiae JDR-1 and E. coli B formed acetic acid during the exponential growth phase which was metabolized upon complete utilization of the carbon sources in the MeGAXn hydrolysates. E. asburiae JDR-1 was also able to grow in xylobiose and xylotriose minimal medium, which E. coli B could not utilize. However, E. asburiae JDR-1 was unable to utilize MeGAX2 and MeGAX3 (data not shown).


Figure 2
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  		FIG. 2. Aerobic growth, substrate utilization, and formation of products from acid hydrolysates of MeGAXn by E. asburiae JDR-1 (A) and E. coli B (B). Xylose (diamonds), MeGAX (squares), and acetic acid (triangles) were determined in media by HPLC. Growth was determined by measuring turbidity as the OD600 (open circles).

Substrate preference of E. asburiae JDR-1.
E. asburiae JDR-1 was found to grow aerobically in minimal media containing different sole carbon sources, such as glucose, xylose, mannitol, maltose, rhamnose, mannose, glucuronate, and glycerol. As noted above, it was able to quantitatively metabolize MeGAX but was unable to utilize MeGAX2 generated by acid hydrolysis or MeGAX3 generated by a GH10 endoxylanase. When growing in a minimal medium containing an equimolar mixture of glucose and xylose, E. asburiae JDR-1 displayed a diauxic growth pattern typical of species of Enterobacteriaceae (Fig. 3A). Glucose (8 mM) was consumed within approximately 8 h, while xylose utilization began when glucose was almost entirely consumed and was depleted in 14 h.


Figure 3
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  		FIG. 3. Aerobic growth of E. asburiae JDR-1 on different combinations of sugar substrates. Concentrations of substrates and acetic acid as a product were determined by HPLC. Growth was determined as turbidity (OD600). (A) Growth on glucose (7.5 mM) and xylose (7.5 mM). Closed circles, glucose; diamonds, xylose; triangles, acetic acid; OD600 values are shown by the open circles. (B) Growth on glucuronic acid (10 mM) and xylose (10.5 mM). Open squares, glucuronic acid; diamonds, xylose; open circles show OD600 values. (C) Growth on MeGAX (6.5 mM). Squares, MeGAX; open circles, OD600 values.

To study the process by which MeGAX was utilized, E. asburiae JDR-1 was grown in minimal medium containing both xylose and glucuronate, products that might be generated from MeGAX. A single-phase growth curve was observed in which both substrates were consumed by 15 h (Fig. 3B). This is similar to its single-phase growth curve on MeGAX, in which the 6.5 mM substrate was depleted in about 11 h (Fig. 3C). The similarity in growth pattern with MeGAX and the combination of xylose and glucuronate as carbon sources supports the possibility that free glucuronate and free xylose may be released during the metabolism of MeGAX.

Fermentation characteristics.
Fermentation experiments were performed to evaluate the potential of E. asburiae JDR-1 as a biocatalyst for the production of biobased products and define the processes involved in the metabolism of MeGAX. Using limiting amounts (0.25% [wt/vol]) of substrates and cultivation under anaerobic standing conditions, E. asburiae JDR-1 was able to ferment all major sugars constituting hemicellulose, including D-glucose, D-xylose, D-mannose, L-arabinose, and D-galactose. The major products from xylose and galactose fermentation were acetic acid and ethanol, present in similar molar quantities. Acetic acid, ethanol, and small amounts of lactic acid were produced from glucose, mannose, and arabinose (Table 1). Small amounts of formic acid and very small amounts of fumaric and succinic acids were detected in most fermentation mixtures. The HPLC profiles indicated that E. asburiae JDR-1 performs mixed acid fermentation as does E. coli, but with preferential formation of acetate and ethanol over lactate.


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  		TABLE 1. Fermentation products formed by E. asburiae JDR-1 from monosaccharides derived from hemicellulosea

With sweetgum MeGAXn hydrolysate as substrate, E. asburiae JDR-1 consumed 99% of the substrate when the pH was maintained above 5, giving the major products acetic acid and ethanol (Table 2). With glucuronic acid as carbon source, acetic acid was the major fermentation product. To study the process of MeGAX metabolism, the presumed degradation products of MeGAX, xylose and glucuronate, both at 11 mM, were used as substrates. The predominant products were 20.4 mM acetate and 5.25 mM ethanol. Major fermentation products from 4.0 mM MeGAX were 8.1 mM acetic acid, 1.2 mM ethanol, and 4.3 mM methanol (Table 2).


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  		TABLE 2. Fermentation products of E. asburiae JDR-1 derived from MeGAXna

Central metabolic pathways determined by 13C-NMR.
The total quantities of ethanol, acetate, and lactate were determined by HPLC, and the quantities of 13C-labeled products were quantified from integration of differentially labeled compounds detected in the 13C-NMR spectra. This allowed determination of the fraction of each fermentation product that was differentially labeled with 13C, which helped to illustrate the central metabolic pathways E. asburiae JDR-1 uses. The quantities of each product and the fractions labeled with 13C are presented in Table 3.


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  		TABLE 3. Distribution of 13C in fermentation products formed in anaerobic cultures of E. asburiae JDR-1 and E. coli strain B grown with differentially 13C-labeled xylose and glucosea

To determine the primary pathway of xylose metabolism by E. asburiae JDR-1, comparisons were made for the fermentation of [2-13C]xylose with cultures of E. coli B. For E. coli B, employing only the pentose phosphate pathway to metabolize xylose, the prominent shift signals in the 13C-NMR spectrum of the fermentation products were assigned to [1-13C]ethanol at 57.6 ppm, [2-13C]lactate at 68.8 ppm, and [1-13C]acetate at 181.0 ppm. Shift signals at 71.0 ppm and 74.5 ppm were assigned to the {alpha}- and β- anomers of unused [2-13C]xylose, and the signal at 30.6 ppm was assigned to the methyl carbons of the acetone standard (Fig. 4B). Fractions of labeled versus total acetate, ethanol, and lactate with E. coli B were 0.26, 0.27, and 0.31, respectively, which was slightly less than the theoretical fraction of 0.4 expected for metabolism through the pentose phosphate pathway (Table 3). The lower quantities of labeled products as fractions of the total found for E. coli may reflect accuracy limitations for integration against the [13C]acetone standard, as these products all showed that similar fractions (0.26 to 0.31) were labeled.


Figure 4
Figure 4
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  		FIG. 4. Pathway determination for the metabolism of xylose and glucose by E. asburiae JDR-1. Media from anaerobic cultures of E. asburiae JDR-1 and E. coli B grown with xylose or glucose enriched with 13C in specific carbons were analyzed by 75.5 MHz 13C-NMR spectrometry. (A) [2-13C]xylose fermented by E. asburiae JDR-1; (B) [2-13C]xylose fermented by E. coli B; (C) [1-13C]glucose fermented by E. asburiae JDR-1; (D) [6-13C]glucose fermented by E. asburiae JDR-1.

When E. asburiae JDR-1 fermented [2-13C]xylose, a 13C-NMR spectrum for fermentation products was obtained with prominent signals for [1-13C]ethanol, [2-13C]lactate, and [1-13C]acetate at expected shift positions of 57.8 ppm, 68.8 ppm, and 181.0 ppm, respectively (Fig. 4A). The fractions of labeled ethanol, labeled acetate, and labeled lactate to their total amounts were 0.43, 0.4, and 0.45, respectively (Table 3), and nearly identical to the theoretical fraction of 0.4. Moreover, the fractions of labeled acetate and ethanol were not higher than the fraction of labeled lactate (Table 3). These results establish that the pentose phosphate pathway is the main metabolic pathway for xylose utilization in E. asburiae JDR-1.

To determine the primary pathway that E. asburiae JDR-1 utilizes to metabolize glucose, [1-13C]glucose and [6-13C]glucose were used as fermentation substrates. Similar 13C-NMR spectra of fermentation products were obtained from [6-13C]glucose and [1-13C]glucose (Fig. 4C and D). Shift signals at 92.4 and 96.2 ppm were assigned to the {alpha}- and β-anomers of unused [1-13C]glucose (Fig. 4C); signals at 60.9 and 60.1 ppm were assigned to the {alpha}- and β-anomers of unused [6-13C]glucose (Fig. 4D). The signal at 30.6 ppm was assigned to the methyl carbons of the acetone standard. Excepting the shift signals for reference and unused substrates, the prominent signals in both spectra were for [2-13C]ethanol at 17.1 ppm, [2-13C]acetate at 22.2 ppm, and [3-13C]lactate at 20.3 ppm with similar distributions for both substrates. The absence of [1-13C]lactate indicates that no [1-13C]glucose was metabolized through the Entner-Douderoff (ED) pathway. Moreover, the fractions of all labeled products of their total amounts were similar for fermentation of [6-13C]glucose and [1-13C]glucose, and the fractions for [6-13C]glucose were not higher than those found for [1-13C]glucose (Table 3), indicating little or no [1-13C]glucose went through the pentose phosphate pathway. Collectively, these results establish that the Embden-Meyerhof (EM) pathway is the main metabolic pathway for glucose utilization in E. asburiae JDR-1.

Growth and projected ATP yields with different substrates.
To understand the bioenergetics in the process of MeGAX fermentation by E. asburiae JDR-1, molar cell dry weight yields were determined after 24 h of growth with glucose, xylose, glucuronate, and MeGAX as sole carbon sources in Zucker-Hankin minimal medium. The experiment was performed three times, and the average approximate YM values were about 10 g per mole of substrate for growth on xylose and glucuronate, 20 for growth on glucose, and 30 for growth on MeGAX (Table 4). The experimental YATP in anaerobic growth has been reported to be in the range of 8 to 12 g cell dry weight per mole of ATP for bacteria (26). An estimated YATP value at the lower end of this range, 8, was used here since this is for anaerobic growth in batch cultures in minimal medium with a relatively low concentration of carbon source (1, 6). The apparent ATP yields per mole of substrate were calculated based on the estimated YATP of 8 as 1.3 mol of ATP produced from either xylose or glucuronate, 2.6 from glucose, and 4.0 from MeGAX (Table 4). These apparent ATP yields allow an estimate of the relative ATP yields obtained for the different substrates without considerations of maintenance energy or overflow metabolism (26), providing insight into the metabolism of MeGAX. The ratios of the molar growth yields obtained with xylose, glucuronate, and MeGAX as carbon sources are 1.0:1.0:3.2 (Table 4), indicating that the requirement for MeGAX transport is less than that for separate transport of xylose and glucuronate.


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  		TABLE 4. Anaerobic molar cell dry weights and ATP yieldsa


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DISCUSSION
 
Fermentation of MeGAX by E. asburiae JDR-1.
E. asburiae JDR-1 is the first bacterium that has been described to ferment MeGAX. The complete utilization of MeGAX by E. asburiae JDR-1 clearly indicates a metabolic potential not shared with E. coli B, as well as the ethanologenic strains E. coli KO11 (25) and Klebsiella oxytoca P-2 (23).

Based upon the studies presented here, E. asburiae JDR-1 uses the EM glycolytic pathway to metabolize glucose and the pentose phosphate pathway to metabolize xylose. This is the first definitive study to determine the role of these pathways in any Enterobacter species, although the findings are not unexpected, as the role of these pathways has been well defined in other Enterobacteriaceae (18). The roles of these pathways in central metabolism are supported by genome annotations in Enterobacter sp. strain 638 (complete sequence of chromosome of Enterobacter sp. strain 638; NC_009436; NCBI genome database).

The formation of equal molar amounts of ethanol and acetic acid from xylose under anaerobic conditions suggests that pyruvate serving as a precursor for acetic acid and ethanol might be converted to acetyl coenzyme A by pyruvate formate lyase (18). The ability of E. asburiae JDR-1 to completely convert MeGAX to acetic acid, ethanol, and methanol suggests that demethylation may occur as an early event in the metabolic processing. The metabolic potential of E. asburiae JDR-1 to ferment mannose, galactose, and glucose, as well as xylose and arabinose, defines its potential to convert all of the pentoses and hexoses that comprise hemicellulosic biomass to fermentation products.

Central metabolic pathways used by E. asburiae JDR-1.
In E. coli, xylulose-5-phosphate enters the pentose phosphate pathway, yielding fructose-6-phosphate and glyceraldehyde-3-phosphate (18). Fructose-6-phosphate can be further metabolized to glyceraldehyde-3-phosphate. In the phosphoketolase pathway, [2-13C]xylose metabolism is expected to yield [1-13C]acetate or [1-13C]ethanol and unlabeled pyruvate, with no label found in lactate (5). However, if organisms use the pentose phosphate pathway, the [1,2-13C] pyruvate produced is expected to account for one-fifth of the total pyruvate, with [2-13C]pyruvate accounting for another one-fifth of the total pyruvate. The remaining three-fifths of the pyruvate do not carry the 13C label (20). [1,2-13C]pyruvate can be reduced to [1,2-13C]lactate or converted to [1-13C]ethanol or [1-13C]acetate by pyruvate formate lyase and subsequent enzymes; [2-13C]pyruvate can be reduced to [2-13C]lactate or converted to [1-13C]ethanol and [1-13C]acetate. Therefore, if an organism uses only the pentose phosphate pathway to metabolize [2-13C]xylose, both [1-13C]ethanol and [1-13C]acetate would account for two-fifths of total ethanol and acetate produced, whereas [1,2-13C]lactate and [2-13C]lactate together would account for two-fifths of total lactate. The fraction of labeled C-2 in lactate would be two-fifths of the total C-2 in lactate, the same fraction as labeled acetate and ethanol to total acetate and ethanol. If both the phosphoketolase and pentose phosphate pathways were used, label would be found in the lactate, but the ratio of labeled ethanol or acetate to total ethanol or acetate would be higher than the ratio of labeled lactate to total lactate. The labeling patterns obtained in this study indicate that the pentose phosphate pathway is the main if not exclusive metabolic pathway for xylose utilization in E. asburiae JDR-1.

Three major pathways, the EM, pentose phosphate, and ED pathways, are used by bacteria to catabolize glucose or other sugars into pyruvate (18). To determine the primary pathway E. asburiae JDR-1 utilizes to metabolize glucose, [1-13C]glucose and [6-13C]glucose were used as fermentation substrates. For either the EM or ED pathway, labeled carbon derived from [6-13C]glucose would be found in C-3 of the intermediate pyruvate. Following subsequent metabolic processing, [2-13C]ethanol, [2-13C]acetate, and [3-13C]lactate would be produced. No labeled carbons would be found at the other carbon positions of these compounds. However, different pathways give different labeled fermentation products for [1-13C]glucose. If [1-13C]glucose were utilized by the EM pathway, all labeled carbon would also be found at the C-3 position of pyruvate, and the amounts of labeled products and their ratios to total products would be the same as those derived from that of [6-13C]glucose. If the pentose phosphate pathway were utilized, C-1 of [1-13C]glucose would be oxidized to the carboxyl group in 6-phosphogluconate, which would then be oxidized to CO2 by glucose-6-phosphate dehydrogenase (18). After the two oxidation steps, no other labeled fermentation products would be obtained except for labeled CO2. If the ED pathway is used, C-1 of [1-13C]glucose would become C-1 in one of the two pyruvate molecules generated from 2-keto-3-deoxy-6-phosphogluconate. In the subsequent fermentation process, [1-13C]pyruvate would be cleaved by pyruvate formate lyase, yielding acetate or ethanol without the 13C label, and [1-13C]pyruvate would be reduced to [1-13C]lactate. The HPLC and NMR data (Fig. 3C and D; Table 3) showed that both [6-13C]glucose and [1-13C]glucose gave similar products fermented by E. asburiae JDR-1 with similar distributions of labeled carbons. These results establish that the EM pathway is the main metabolic pathway for glucose utilization in E. asburiae JDR-1.

Possible pathway of MeGAX metabolism.
The processing of aldouronates formed by the action of GH10 endoxylanases, e.g., MeGAX3, has been shown to involve the removal of the {alpha}-1,2-linked 4-O-methylglucuronosyl moiety from the reducing terminus of β-1,4-linked xylotriose by the action of a GH67 {alpha}-glucuronidase (31). {alpha}-Glucuronidases (GH67) from some bacteria have been shown to act on MeGAX as well as MeGAX3 (19, 31). Consensus primers based on GH67 family genes (http://www.cazy.org/fam/GH67.html) may be used to amplify aguA genes from bacterial genomic DNA prior to partial sequencing and cloning. The application of this strategy was not successful in identifying a gene encoding a GH67 {alpha}-glucuronidase in E. asburiae JDR-1 (G. Nong and C. Bi, unpublished data). Either this strain synthesizes an {alpha}-glucuronidase completely different from the GH67 that has been well defined or a novel metabolic process may be used in which the cleavage of the glycosidic bond follows other reactions.

The fermentation products derived from xylose and glucuronate were similar to those obtained from MeGAX, although the ethanol yield from MeGAX was lower. This result suggested that xylose and glucuronate might be released in the process of MeGAX utilization. This possibility was also supported by the similarity in the growth patterns of E. asburiae JDR-1 growing in MeGAX minimal medium and the glucuronate plus xylose minimal medium (Fig. 2).

Due to the small amount of methanol observed relative to system noise of the GC system, accurate quantitative data for methanol as a fermentation product were not obtained in media containing acid hydrolysates of MeGAXn. Methanol was quantitatively detected in stoichiometric equivalence with MeGAX during the metabolism of MeGAX, indicating methanol is derived from the methyl group released from methylglucuronate as an early event in the metabolism of MeGAX (Table 2). Based on the results obtained from this study, the process of MeGAX catabolism by E. asburiae JDR-1 may involve the release of methanol, glycosidic bond cleavage to release glucuronate and xylose, and catabolism of these carbohydrates to generate ethanol and acetic acid as predominant fermentation products. An elimination reaction for the release of methanol prior to glycosidic bond cleavage is a possibility, in which case the product of glycosidic bond cleavage would be delta-4,5-hexuronate.

Bioenergetics of E. asburiae JDR-1.
Glucose was shown to be metabolized in E. asburiae JDR-1 by the EM pathway. If the PTS system is used to import glucose as in E. coli (32), the net formation of 3 mol of ATP results from 1 mol of glucose when the major fermentation products are ethanol and acetate. For xylose utilization via the pentose phosphate pathway, presumably requiring 1 mole of ATP to import xylose with an ABC transporter, as in E. coli (16), 1.5 mol of ATP would be produced per mol of xylose when the fermentation products are ethanol and acetate (7). Glucuronate has been shown to be transported into cells with the UxuT transporter in E. coli. This transporter bears significant homology to ExuT in Erwinia chrysanthemi, which has been defined as responsible for the transport of glucuronate as well as galacturonate through an active process in which energy is consumed. Considering energy consumed in active transport, the maximal ATP yield in the process of glucuronate fermentation may be 2 moles or less per mole of glucuronate when the major fermentation product is acetic acid (9, 24, 27). The theoretical ATP yields from glucose, xylose, and glucuronate were close to the calculated ATP yields based on the estimated YATP of 8, suggesting the transportation systems and metabolism of these substrates are correctly interpreted. The ratios of the molar growth yields obtained with xylose, glucuronate, and MeGAX as carbon sources are 1.0:1.0:3.2 (Table 4), clearly indicating that the bioenergetic requirement for MeGAX transport is less than that for separate transport of xylose and glucuronate.

Role of E. asburiae JDR-1 in soil ecology and bioprocessing.
Most Enterobacter asburiae species have been identified and characterized as human pathogens (2). However, strains of Enterobacter asburiae have been isolated from soil environments and implicated in the mobilization of phosphate for plant nutrition from calcium phosphate (30). The ability of E. asburiae JDR-1 to utilize MeGAX, as well as xylobiose and xylotriose, may reflect its evolution in a soil environment with hemicellulose providing a carbon resource. With the exception of Erwinia species, the Enterobacteriaceae are not known for their ability to secrete endoxylanases and endoglucanases. For Erwinia spp., the secreted endoxylanases are members of glycohydrolase family 5 that generate oligosaccharides, all of which are replaced with MeGA residues, and are not further metabolized by the cell (10, 22). The role of these enzymes in Erwinia species may be related to their colonization and maceration of plant tissues and thus may serve a virulence function related to pathogenesis. Enterobacter spp. have been found associated with plant roots and may provide commensal, if not symbiotic, relationships that contribute to positive development of the plant (30). However, none has been identified which secretes endoxylanases. E. asburiae JDR-1 is unable to utilize the aldotetrauronate product of GH10 endoxylanase, MeGAX3, which serves as a substrate for several gram-positive bacteria that secrete GH10 endoxylanases. At present, the generation of the aldobiuronate MeGAX is only known to result from the acid hydrolysis of methylglucuronoxylans. Since MeGAX is the only aldouronate this bacterium will use, this metabolic potential has yet to be linked to the biological generation of aldouronates from hemicellulosic biomass.

Biotechnological applications.
The predominant structural polymer in the hemicellulose fraction of hardwoods and crop residues is methylglucuronoxylan (21), and depending on the source, as much as 27% of the carbohydrate may occur in the form of MeGAX following dilute acid pretreatment (25). Capturing the xylose, including that found in MeGAX, in acid hydrolysates of hemicellulose will increase the efficiency of converting lignocellulosic biomass to targeted products. Our current research on the metabolism of E. asburiae JDR-1 has established its potential for the engineering of gram-negative bacteria for bioconversion of aldouronates as well as xylose and other pentoses and hexoses that may be derived from the hemicellulose fraction of any lignocellulosic source. The definition of the genetic basis for MeGAX metabolism may allow its application to the engineering of established bacterial biocatalysts, e.g., E. coli KO11, for the efficient conversion of cellulosic biomass to biofuels and chemical feedstocks.


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ACKNOWLEDGMENTS
 
We are indebted to Donna Williams and the Laboratory for Electron Microscopy and Imaging, Department of Microbiology and Cell Science, for providing assistance with microscopy that defined flagellar structures and the basis for motility. Franz St. John was particularly helpful in providing his technical expertise and advice. We also thank K. T. Shanmugam and L. O. Ingram for helpful discussions and review of the manuscript.

This research was supported by U.S. Department of Energy grants DE FC36-99GO10476 and DE FC36-00GO10594, The Consortium for Plant Biotechnology research project GO12026-198 (DE FG36-02GO12026), and the Institute of Food and Agricultural Sciences, University of Florida Experiment Station, as CRIS Project MCS 3763.


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FOOTNOTES
 
* Corresponding author. Present address: University of Florida, Department of Microbiology and Cell Science, Bldg. 981, Museum Rd., Gainesville, FL 32611-0700. Phone: (352) 392-5923. Fax: (352) 392-5922. E-mail: jpreston{at}ufl.edu Back

{triangledown} Published ahead of print on 14 November 2008. Back


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Applied and Environmental Microbiology, January 2009, p. 395-404, Vol. 75, No. 2
0099-2240/09/$08.00+0     doi:10.1128/AEM.01941-08
Copyright © 2009, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Bi, C., Zhang, X., Ingram, L. O., Preston, J. F. (2009). Genetic Engineering of Enterobacter asburiae Strain JDR-1 for Efficient Production of Ethanol from Hemicellulose Hydrolysates. Appl. Environ. Microbiol. 75: 5743-5749 [Abstract] [Full Text]  
  • Nong, G., Rice, J. D., Chow, V., Preston, J. F. (2009). Aldouronate Utilization in Paenibacillus sp. Strain JDR-2: Physiological and Enzymatic Evidence for Coupling of Extracellular Depolymerization and Intracellular Metabolism. Appl. Environ. Microbiol. 75: 4410-4418 [Abstract] [Full Text]  

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