Ethanol fermentation technology – Zymomonas mobilis

P. Gunasekaran and K. Chandra Raj

Department of Microbial Technology, School of Biological Sciences, Madurai Kamaraj University, Madurai 625 021, India

Due to dwindling of fossil fuel, microbial production of bio-fuel from organic byproducts has acquired significance in recent years. Ethanol has been trusted as an alternate fuel for the future. Even though several microorganisms, including Clostridium sp., have been considered as ethanologenic microbes, the yeast Saccharomyces cerevisiae and facultative bacterium Zymomonas mobilis are better candidates for industrial alcohol production. Z. mobilis possesses advantages over S. cerevisiae with respect to ethanol productivity and tolerance, thus encouraging researchers for exploiting Z. mobilis ability to utilize sucrose, glucose, and fructose by Entner–Deudoroff pathway. The bottlenecks in Z. mobilis are: (i) its inability to convert complex carbohydrate polymers like cellulose, hemicellulose, and starch to ethanol, (ii) its resulting in byproducts such as sorbitol, acetoin, glycerol, and acetic acid, and (iii) formation of extracellular levan polymer. To circumvent these problems, genetic manipulation of Z. mobilis has been attempted for broadening the utilizable range of Z. mobilis, i.e. genes encoding several hydrolytic enzymes from related bacterial species have been cloned, and transferred into Z. mobilis. Interestingly, a pet operon (production of ethanol) was constructed by combining pdc (pyruvate decarboxylase) and adhII (alcohol dehydrogenase) genes of Z. mobilis, and transferred to other bacterial strains to make them ethanologenic novel strains. Through classical mutation and selection approaches, mutants of Z. mobilis with improved fermentation characteristics and without byproduct formation have been obtained. In addition to ethanol, Z. mobilis has also been metabolically engineered to produce L-alanine and L-lactic acid. Genes encoding b -carotene synthesis have also been cloned and successfully expressed in Z. mobilis to enrich the fermented nutrients of farm animals. Several applications of levan in food and pharmaceutical industries provide an opportunity to exploit Z. mobilis for large-scale production of levan. The merits of Z. mobilis suggest the potential use of this organism in industrial production of various fermentation products.

The natural energy resources such as fossil fuel, petroleum and coal are being utilized at a rapid rate and these resources have been estimated to last over a few years. Therefore, alternative energy sources such as ethanol, methane, and hydrogen are being considered. Some biological processes have rendered possible routes for producing ethanol and methane in large volumes. A worldwide interest in the utilization of bio-ethanol as an energy source has stimulated studies on the cost and efficiency of industrial processes for ethanol production1. Intense research has been carried out for obtaining efficient fermentative organisms, low-cost fermentation substrates, and optimum environmental conditions for fermentation to occur. Traditionally, ethanol has been produced in batch fermentation with yeast strains that can- not tolerate high concentration of ethanol. This necessitated the strain improvement programme for obtaining alcohol-tolerant strains for fermentation process. Zymomonas mobilis, a gram-negative bacterium, is considered as an alternative organism in large-scale fuel ethanol production. Comparative laboratory- and pilot-scale studies on kinetics of batch fermentation of Z. mobilis versus a variety of yeast have indicated the suitability of Z. mobilis over yeasts due to the following advantages:

 

  1. its higher sugar uptake and ethanol yield,
  2. its lower biomass production,
  3. its higher ethanol tolerance,
  4. it does not require controlled addition of oxygen during the fermentation, and
  5. its amenability to genetic manipulations.

 

The only limitation of Z. mobilis compared to the yeast is that its utilizable substrate range is restricted to glucose, fructose, and sucrose. Z. mobilis was originally isolated from alcoholic beverages like the African palm wine, the Mexican ‘pulque’, and also as a contaminant of cider and beer in European countries. On the basis of evaluation using the modern taxonomic approaches, the genus Zymomonas2 has only one species with two subspecies, Z. mobilis subsp. mobilis and Z. mobilis subsp. pomaceae. The ability to utilize sucrose as a carbon source distinguishes Z. mobilis from Z. anaerobia3. It is one of the few facultative anaerobic bacteria which metabolizes glucose and fructose via the Entner–Deudoroff (E–D) pathway, which is usually present in aerobic microorganisms4. Under anaerobic conditions, Z. mobilis produces byproducts such as acetoin, glycerol, acetate, and lactate, which result in reduced production of ethanol from glucose. During growth of Z. mobilis in fructose, the formation of ace-
toin, acetic acid, and acetaldehyde was clearly more pronounced than when grown in glucose. However the cell yield was low during its growth in fructose.

In addition to ethanol fermentation, Z. mobilis has potential application in polymer production. Levan, a polymer of fructose units linked by b -2,6-fructosyl bond, is produced by Z. mobilis during its growth on sucrose medium. Microbial levan is of commercial importance and is used as a thickening, gelling, and suspending agent. In recent years, strategies to improve the yield of levan production by microorganisms attracted greater attention. In this review, the recent developments in the potential applications of Z. mobilis are discussed.

Biochemistry of Zymomonas mobilis

In Z. mobilis, D-glucose and D-fructose are transported by facilitated diffusion5. Concentrated glucose solutions are not inhibitory to the E–D pathway enzymes, since conversion of glucose to ethanol by this organism proceeds rapidly6. Thus, the extracellular osmotic pressure of the glucose solution may rapidly be balanced by corresponding intracellular sugar concentrations. High sugar concentrations decrease the total water potential, and exert osmotic pressures which are comparable to those of relatively strong salt solutions. The low-salt tolerance of Z. mobilis poses problems for the fermentation of molasses which usually contains a high-salt content4.

Study of metabolic intermediates showed that glucose-6-phosphate dehydrogenase and phosphoglycerolmutase are the limiting enzymes, and that phosphofructokinase is not present in the E–D pathway of Z. mobilis7. The pyruvate decarboxylase (Pdc) and alcohol dehydrogenase (Adh) are the key enzymes in ethanol formation. The Pdc is an unique enzyme in Z. mobilis which requires thiamin pyrophosphate for its activity8. While Adh I of Z. mobilis is zinc-dependent and is similar to Adh IV of yeast, Adh II is very unusual in containing iron and not zinc9. The Adh II appears to facilitate continuation of fermentation at high concentration of ethanol.

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In Z. mobilis, the E–D pathway enzymes are more tolerant to ethanol, as the cell-free system of Z. mobilis can rapidly consume glucose and produce ethanol more than 15% w/v (ref. 10). The cell membrane of Z. mobilis has acquired altered fatty acid content to counteract the adverse effects of ethanol. The major fatty acids occurring in Z. mobilis are myristic acid, palmitic acid, and cis-vaccenic acid. Among the phospholipids, phosphotidyl ethanolamine is the most abundantly present. The high concentration of cis-vaccenic acid and unusual hopanoids in the membrane are responsible for the high ethanol tolerance11.

Located in the cell membrane of Z. mobilis are NADH- and NADPH-oxidases which catalyse the oxidation of NAD(P)H (ref. 12). In addition, enzymes catalase, superoxide dismutase, and peroxidase are also present. Here, the transfer of electrons via the respiratory chain is not coupled with oxidative phosphorylation. Both glucose-fructose oxido-reductase and glucose dehydrogenase lead to the formation of gluconic acid which after phosphorylation enters the E–D pathway after the NAD(P)H formation. A mole of reduced co-enzyme formed is directly consumed by mannitol dehydrogenase and NAD(P)H oxidase, resulting in accumulation of byproducts, namely, acetaldehyde, acetoin, and acetic acid13. Production of acetaldehyde by Z. mobilis in the presence of oxygen is due to increased NADH oxidase activity resulting in the decreased availability of NADH for the reduction of acetaldehyde to ethanol by Adh. It does not have an aldehyde dehydrogenase to oxidize acetaldehyde to acetic acid. Adh mutants of Z. mobilis showing increased levels of acetaldehyde production have been isolated by using allyl alcohol as a selective agent14.

Z. mobilis possesses both acid and alkaline phosphatases. While acid phosphatase activity is highest in the presence of Mg2+, the alkaline phosphatase activity is highest with Zn(II) (ref. 15). The alkaline phophatase of Z. mobilis is associated with membranes and it occurs in two isoforms16.

The sucrose hydrolysing activity seems to be stimulated by sucrose and fructose17. The hydrolysis rate of sucrose and the rate of transfructosylation are shown to be higher than the sugar uptake rate of Z. mobilis. During fermentation of sucrose by Z. mobilis, three types of transfructosylation occur resulting in the formation of free fructose, oligosaccharides, and higher polymers of fructose and levan where water, sucrose, and levan of polyfructose respectively can act as an acceptor18.

Levansucrase has been purified from cells19 as well as from the culture broth20. Three different saccharolytic enzymes have been reported in Z. mobilis; an endocellular sucrase, an exocellular levansucrase, and an exocellular sucrase21. These enzymes are b -D-fructofuranosyl-transferases and are therefore able to hydrolyse sucrose. Moreover, levansucrase synthesizes levan and can in turn be partially hydrolysed. Sucrase and levansucrase do not hydrolyse inulin, a low molecular weight fructose polymer.

Molecular biology of Z. mobilis

The genome size of Z. mobilis strains is in the range of 1.53 ±  0.19 ´  109 Da, about 56% of the E. coli genome, and can accommodate about 1500 cistrons. The DNA base composition of Z. mobilis (48.5 ±  1.0% G + C) was determined by thermal denaturation22.

The pyruvate decarboxylase gene (pdc) has been cloned from Z. mobilis strains ATCC 31821 (ref. 23) and ATCC 29191 (refs 24, 25). The coding region is 1.7 kb-long and encodes a polypeptide of 567 amino acids with a subunit mass of 60.8 kDa (ref. 25). As the promoter region does not contain sequences homologous to the generalized promoter structure for E. coli, the promoter of pdc is not recognized in E. coli, although the cloned gene is expressed relatively at high levels under the control of alternative promoters. Comparison of the nucleotide sequence of the pdc gene from Z. mobilis strains ATCC 29191 and ATCC 31821 showed the existence of polymorphism in different isolates26.

The adh gene from Z. mobilis has been cloned using a novel indicator plate technique where mixtures of para rosaniline and bisulphite are incorporated. The DNA sequence for this gene contains an open reading frame (ORF) that encodes a polypeptide of 383 amino acids with mol wt of 40 kDa (ref. 27). The adhI gene is transcribed at low levels in E. coli from the P2 promoter of Z. mobilis, but is expressed well in E. coli under the control of lac promoter. This Adh I is found to exhibit very little homology with other known products of adh genes, but exhibits strong homology with Adh IV of S. cereviseae28.

The gene encoding for glyceraldehyde-3-phosphate dehydrogenase (gap) has been isolated from Z. mobilis by complementing a deficient strain of E. coli. The ORF for this gene encodes 337 amino acids with a mol wt of 36.1 kDa (ref. 29). Although the primary amino acid sequence of Gap has considerable functional homology and amino acid identity with same enzyme from other eukaryotes and bacteria, it appears to be more closely related to that of the thermophilic bacteria and to chloroplast isoenzymes. Comparison of this gene with other glycolytic genes from Z. mobilis reveals several common features of gene structure including a conserved pattern of codon bias. Other glycolytic genes that have been cloned and characterized are given in Table 1.

The Z. mobilis gene encoding phosphoglycerate kinase (pgk) has been cloned in E. coli and was sequenced34. This promoter-less pgk gene is located 225-bp downstream from the gap gene, and is part of the gap operon. The deduced amino acid sequence from the Z. mobilis pgk gene is less conserved than those of other known sequences for pgk, and also shows a high degree of conservation of structure. In general, the amino acid positions at the boundaries of a -sheet and b -helical regions, and those connecting these regions are more conserved than the amino acid positions within regions of secondary structure.

The phoC gene of Z. mobilis encodes for an acid phosphatase with a mol wt of 29 kDa (ref. 15). Its promoter comprises a – 35 pho box region, similar to that of E. coli genes, as well as the regulatory sequences of S. cerevisae acid phosphatase (pho5). The phoC gene
contains a 5¢ terminus which is AT-rich, has a weak ribosome-binding site, and has less biased codon usage than the highly expressed Z. mobilis genes. A comparison of the genes reveals that promoters of all these genes are similar in degree of conservation of spacing and identity with proposed Z. mobilis consensus sequence.

Five trp genes of Z. mobilis were identified by their ability to complement the trp mutants43. The organization of Z. mobilis trp genes is similar to that found in species of Rhizobium, Acinetobacter calcoaceticus, and Pseudomonas acidoovorans. The trpF, trpB, and trpA genes appear to be linked but they are not closely associated with trpD or trpC genes in Z. mobilis.

The sacA gene encoding an intracellular sucrase from Z. mobilis has been cloned, sequenced, and characterized44,45. The SacA protein is a monomer with a molecular weight of 58 kDa and its deduced amino acid sequence shows strong homology with the intracellular sucrase of B. subtilis and yeast invertase. The sacB encoding extracellular levansucrase and sacC encoding extracellular sucrase were cloned and expressed in E. coli38,41. Nucleotide sequence analysis of sacB gene revealed an ORF 1269-bp long encoding for a protein with mol wt of 46.7 kDa. The deduced amino acid sequence was identical to the N-terminal sequence of that deduced from the sacB gene of Z. mobilis Z6C (ref. 46.) The amino acid sequence of SacB showed very little similarity to those of other known sucrases, but was very similar to the levansucrase of Z. mobilis (61.5%), Erwinia amylovora (40.2%), and Bacillus subtilis (25.6%). The nucleotide sequence analysis of sacC revealed an ORF 1239-bp long encoding a 46-kDa protein. The first 30 deduced amino acids from this ORF were identical with those from the N-terminal sequence of the extracellular sucrase of Z. mobilis strain ZM4. This sacC gene is located 155-bp downstream of sacB gene forming a sucrase gene cluster. The SacB and SacC are the two extracellular enzymes of Z. mobilis sequenced which do not possess signal sequences38.

Studies have demonstrated the sequence requirements for membrane localization or protein transport in Z. mobilis. The lacZ fusion, which contains anchor sequences conferring membrane association, is used to isolate DNA from Z. mobilis containing promoter activity and amino terminal sequences47. Comparison of the sequences and transcription initiation sites showed that both E. coli and Z. mobilis recognize similar regions of DNA for transcription initiation. Five to eight consecutive hydrophobic amino acids in the amino terminus serve to anchor these hybrid proteins to the membrane in both E. coli and Z. mobilis.

Bioprocess potentials: Ethanol production

Ethanol production by Z. mobilis has been restricted to glucose, fructose, and sucrose substrates. Alternatively, crude sucrose substrates such as sugar beet, bagasse and molasses have also shown promise as substrates for direct fermentation to ethanol. Recombinant DNA technology could be exploited to construct microbial strains to produce ethanol from these low-cost agricultural substrates. To obtain ethanologenic strains utilizing the above- mentioned substrates, either Z. mobilis was transformed with genes of interest acquired from other organisms or gene of Z. mobilis involved in ethanol synthesis was transferred to other organisms with required characteristics. Ethanol production potential of Z. mobilis from a variety of substrates is discussed.

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Genetic manipulation of Z. mobilis

By genetic manipulation, Z. mobilis substrate range could be extended to industrially attractive agricultural byproducts such as whey, starch, and cellulose. This could be achieved by transfer of genes encoding appropriate hydrolases. Various genes encoding enzymes required for utilization of wide range of carbon sources transferred into Z. mobilis, are listed in Table 2.

Gene cloning in Z. mobilis is carried out using three kinds of vectors, viz. broad-host range plasmids, shuttle vectors, and modified broad-host range vectors. Though Z. mobilis strains can act as recipient for a number of broad-host range plasmids such as RP1, RP4, R68, etc.4,59,60 commonly used cloning vectors and phage vectors cannot be transferred or maintained in Z. mobilis. The most commonly used high-copy-number cloning vectors, derived from RSF1010, have been found to be more stable in Z. mobilis than in RP4-derived plasmids60. The occurrence of natural plasmids in Z. mobilis has been described and a high degree of homology has been demonstrated between them61,62. The native plasmids of low molecular weight present in Z. mobilis are used to develop cloning vehicles for Z. mobilis. They have been fused with small E. coli plasmids, such as pACYC184 and pBR325, to produce novel shuttle vectors63–65. The well-defined E. coli segments facilitate further plasmid maintenance, while the Z. mobilis portion which includes presently undefined sequences yields stability in Z. mobilis via the origin of replication or native plasmid66. Integrative shuttle vector plasmid, pZMOCP1, was constructed by ligating EcoRV digests of the cloning vector plasmid, pZMP1, with EcoRV-digested plasmid, pBluescript DNA (ref. 67).

Transfer of R-plasmids into Z. mobilis from either E. coli or Pseudomonas is usually achieved by conjugation. High conjugation rates have been reported for plasmid transferred in an antibiotic-sensitive mutant strain, CP4.45 (ref. 68). Efficient conjugation is routinely achieved by either a self-mobilizable or helper plasmid, such as RP4, pRK2013, etc., as the transformation system of Z. mobilis is in its rudimentary stage. Plasmid transfer mediated by donor cell chromosome mobilization, such as by E. coli S17.1, is very inefficient. To improve sucrose hydrolysis, extracellular sucrase genes, sacB and sacC, were subcloned in E. coli and Z. mobilis shuttle vector pZA22. This construct was transferred into Z. mobilis by conjugation using the helper plasmid pRK2013 (Gunasekaran et al., unpublished result). An alternate method for gene transfer in Z. mobilis is by spheroplast fusion. Yanase et al.69 isolated fusants on a raffinose medium by crossing fructose-assimilation negative strains, but the fusants could not assimilate fructose.

A lactose operon has been introduced into Z. mobilis, using plasmid pGC91.14 carrying the transposon Tn951 encoding the operon70,71. Z. mobilis containing the lactose operon, Tn951, produced 0.4% ethanol from 4% lactose in 40 h (ref. 72). Since Z. mobilis can metabolize only the glucose portion of lactose and also generate only one mole of ATP per mole of glucose metabolized, there is no net gain of energy for cell growth as the same ATP is expended for each lactose molecule transported across the membrane via proton symport. To circumvent this problem, Buchhol et al.11 suggested the transfer of b -galatosidase gene into a leaky strain of Z. mobilis together with the galactose operon, so that the genes encoding for both uptake and complete utilization of lactose would be available.

A gal+ recombinant plasmid, pZG13, was constructed by the insertion of the galETK genes of E. coli, downstream to the Z. mobilis promoter in pZA22 and the plasmid was introduced into a Z. mobilis strain IFO133756 (ref. 44). The recombinant Z. mobilis could take up galactose and produce a small amount of ethanol.

In order to introduce the ability to catabolize raffinose, a plasmid, pRRL1, a resultant from the cointegration between pOD118 and R68.45 was introduced into Z. mobilis, but it was unstable73. Two recombinant plasmids; pZER193 containing a -galactosidase gene of E. coli, and pZY1 containing the lactose permease gene of E. coli were introduced into the strain Z6C of Z. mobilis by spheroplast transformation. Cells of the strain carrying both the plasmids could ferment raffinose and melibiose to ethanol54.

a -Amylase gene from Bacillus licheniformis was subcloned into pKT210 and the recombinant plasmid, pGNB6, was transferred into Z. mobilis50. Here, the enzyme was released by secretion than by cell lysis. It is also necessary to clone and transfer an amyloglucosidase so that the strain can grow directly on starch. Cloning of glucoamylase gene from Aspergillus niger was also attempted but stable transconjugants were not obtained in Z. mobilis49.

In order to convert cellulose directly to ethanol, cellulase genes have to be transferred into Z. mobilis. A CMCase gene of Cellulomonas uda, CB4, was cloned on pZA22, a cloning vector for Z. mobilis74. Z. mobilis carrying this gene synthesized cellulase immunologically identical with that of C. uda. Endoglucanase gene from B. subtilis51 and P. fluorescens52 were subcloned and introduced into Z. mobilis. In all these cases, no endogluconase activity was detected in the culture supernatant, and poor expression in Z. mobilis was obtained compared to E. coli. However, the cellulase gene from E. chrysanthemi coding for endoglucanase was subcloned into a broad-host-range vector pGSS33, and was conjugally transferred into Z. mobilis with the help of RP4 (ref. 50). In this case, most of the endoglucanase accumulated in the periplasmic space, suggesting an efficient export of this foreign protein in Z. mobilis. Here, the CM-cellulase activity was cell bound during exponential growth phase, and after 28 h up to 40% of the activity was recovered from the culture medium. A CM cellulase gene of Acetobacter xylinum was subcloned into Z. mobilis–E. coli shuttle vector plasmid pZA22 (ref. 75). The resulting recombinant plasmid, pZAAC21, was introduced into Z. mobilis IFO 13756 by electroporation. Though the expression of CM cellulase in Z. mobilis increased 10 fold, no attempt was made to use this strain in ethanol production from cellulose.

Transfer of the b -glucosidase gene of Xanthomonas albilineans to Z. mobilis was achieved by subcloning the b -glucosidase gene into pKT404 followed by triparental mating involving the helper plasmid pRK2013. The amounts of b -glucosidase produced by the recombinant strains ZM6901 and ZM6902 were 7.5 and 10%, respectively, of those expressed in E. coli. Z. mobilis ZM6901 produced 13.3 mM ethanol from 5 mM cellobiose after 3 days, using the whole cells53.

Hemicellulose, a major constituent of plant cell wall materials, makes up to 40% of many agricultural residues. Upon hydrolysis with acids and enzymes, hemicellulose is converted to a mixture of hexose sugars; D-xylose and D-arabinose. The microbial conversion of these pentose sugars to ethanol for use as a fuel additive has received considerable attention. Hence, D-xylose catabolic genes from Xanthomonas were also introduced into Z. mobilis57. Z. mobilis CP4 was incorporated together with two operons encoding xylose assimilation and pentose phosphate pathway enzymes such as xylose isomerase, xylulokinase, transketolase, and transaldolase58. This engineered strain yielded 0.44 g ethanol/g xylose corresponding to 86% of the theoretical yield. Lawford et al.76 have transferred plasmid pZB5 carrying genes encoding enzymes for xylose metabolism into Z. mobilis, and used this recombinant strain for ethanol production.

A Zymomonas sp. transformed with a gene encoding L-arabinose isomerase, L-ribulokinase, L-ribulose-5-phosphate- 4-epimerase, xylose-isomerase or xylulokinase under
the Z. mobilis glyceraldehyde-3-phosphate-dehydrogenase promoter. The substrate fermentation range of Z. mobilis ATCC 39676 was expanded to include L-arabinose by introduction of genes encoding L-arabinose-isomerase, L-ribulokinase, L-ribulose-phosphate-4-epimerase, transaldolase and transketolase of E. coli77. The engineered strain with plasmid pZB206, grew on arabinose as sole C-source and produced ethanol at 98% of theoretical yield. Thus, arabinose was metabolized more or less completely to ethanol as the sole fermentation product with little byproduct formation. Weisser et al.78 transferred E. coli K12 CA 8000 pmi gene encoding phosphomannose isomerase in Z. mobilis from a lacIq–ptac system on plasmid pZY507. The recombinant Z. mobilis utilized mannose as the sole C-source with a growth rate of 0.07/h. This recombinant Z. mobilis is useful for production of ethanol from mannose.

As pyruvate is the major intermediate in E–D pathway, researchers attempted to shift this catabolic pathway towards the production of L-alanine79. NAD-dependent L-alanine dehydrogenase of B. stearothermophilus carrying vector pZY50 showed poor transfer rates in Z. mobilis strains.

Z. mobilis genes transfer into other organisms

An artificial operon, pet operon, for the production of ethanol was constructed by combining pdc and adhII genes, and was placed under the control of lac promoter80. This pet operon was introduced into E. coli, where these two genes expressed at high levels, resulting in ethanol production from a variety of sugars81. A decreased feed-flow rate was used to optimize fuel ethanol production in recombinant E. coli which produced 1.8 g/l/h ethanol under the optimized conditions of pH 6.4, 34°C and 125 rpm (ref. 82). However, cells carrying cloned pdc gene grew only one-fourth compared to the wild type and tolerated only 2% ethanol83. Saucedo et al.84 demonstrated optimization of ethanol production by recombinant E. coli with plasmid pLOI297 containing Z. mobilis genes (pdc and adhII). Recombinant E. coli FBR1 and FBR2 were constructed by transformation of FMJ39 with pet operon plasmids pLOI295 and pLOI297, respectively, which produced 3.8% (FBR1) and 4.4% (FBR2) ethanol from 10% glucose in batch culture84. Both FBR1 and FBR2 strains showed no plasmid loss even after 60 generations. Introduction of pdc gene into E. chrysanthemi resulted in ethanol production from D-xylose and D-arabinose85.

The physiological influence of pet expression in E. coli ATCC11303 was investigated using glucose, xylose and mannose as the substrates86. Ethanol production was studied by introducing a plasmid containing the ethanol pathway from Z. mobilis into E. coli K12/FMJ39, the resulting strain grew well under anaerobic condition, and was genetically stable. This ethanologenic E. coli when grown in batch culture, an ethanol yield of 0.4–0.42 (w/w) was achieved. Two types of recombinants, one strain with pet expression via a multicopy plasmid, pLOI297, and another strain KO11 with chromosomal integration of pet operon were studied for ethanol production. Both the recombinants produced ethanol, while the parent strain produced exclusively lactic acid from glucose and mannose. Lawford et al.87 studied the effect of oxygen on ethanol production by recombinant E. coli ATCC11303 bearing pet plasmid pLOI297. They suggested that under anaerobic conditions, the pet plasmid channeled the flow of carbon to ethanol as the predominant end product of hexose and pentose catabolism. Ethanol was produced by recombinant E. coli KO11 using crude yeast autolysate as a nutrient supplement88. The recombinant E. coli KO11 produced 0.51 g ethanol/g sugar when grown on hemicellulose hydrolysates of agricultural residues89,90. Fermentation of an enzymatic hydrolysate of ammonia fibre explosion-pretreated corn fibre by recombinant E. coli strains, SL40 and KO11, and Klebsiella oxytoca strain P2 was examined91. Both E. coli strains efficiently utilized most of the sugars in the hydrolysate and produced a maximum of 26.6 and 27.1 g/l ethanol, respectively, equivalent to 90 and 92% of the theoretical yield. Fermentation of sugarcane bagasse or sugars by K. oxytoca P2-containing chromosomally integrated Z. mobilis ethanol pathway genes showed ethanol yield of 40 g/l and 33.3 g/l, respectively92,93. Gold et al.94 transferred pet genes of Z. mobilis into Lactobacillus casei using vector pRSG02; the recombinant strain produced 0.314 g ethanol/g glucose. York et al.95 have reported production of 44–45 g/l ethanol by an engineered E. coli KO11 grown on crude soybean hydrolysate from Spezyme FAN treatment.

Genetically engineered K. planticola, upon introduction of pdc gene of Z. mobilis, markedly increased the yield of ethanol to 1.3 mol/mol of xylose (25.1 g/l). Concurrently, the significant decrease in the yields of formate, acetate, lactate, and butanediol were observed96. The pyruvate- formate-lyase-defective strain of K. planticola harbouring the plasmid carrying pdc gene of Z. mobilis became an efficient ethanol producer97. This recombinant strain produced 387 mM ethanol from 275 mM xylose in 80 h, about 83% of the theoretical yield. Furthermore, this mutant consumed more than double the amount of xylose compared to the wild type, due to reduced production of inhibiting acids during growth98. An E. coli strain containing a recombinant plasmid encoding the pdc and adh genes from Z. mobilis, metabolized glucose and xylose to near theoretical yields of ethanol99. In aerobic condition, the natural expression of adh of E. coli resulted in less ethanol production from clones expressing only Z. mobilis pdc gene. The ethanol-producing genes of Z. mobilis were also incorporated into gram-positive bacteria, B. subtilis100.

 Fermentation: Process development

Batch fermentation

Generally polymeric carbohydrates are prehydrolysed with appropriate enzymes and the hydrolysate is used for fermentation in batch and continuous cultures. Mixed culture of different ethanologenic strains may be used in fermentation to obtain improved productivity. An ethanologenic strain is also mixed with ethanol nonproducer strain. The latter might hydrolyse carbohydrate polymer, facilitating the fermentation by Z. mobilis. In order to recycle the cells in fermentation, the cells are immobilized in a suitable matrix, and used in fermentation.

Starchy materials are attractive substrates for industrial production of alcohol and many reports are available on the production of ethanol from such materials. The different pathways for industrial ethanol production are depicted in Figure 1. Chay et al.101 reported that the best strains for ethanol production from saccharified syrups were strains of Z. mobilis and S. diastaticus. Toran-Diaz et al.102 investigated the effect of acid-hydrolysed substrate and enzyme-hydrolysed substrate on ethanol production by ZM4 and ZM4F strains of Z. mobilis. They obtained ethanol productivity of 4.8 g/g/h, with Z. mobilis grown on Jerusalem artichoke juice, which was higher than that reported for the yeast Kluyveromyces marxianus, by Duvnjak et al.103. Further, they observed that the juice of Jerusalem artichoke could be fermented without
the addition of any nutrients. Recently, Lawford et al.76 demonstrated that corn steep liquor, a by-product of maize wet-milling, as a cost-effective substrate for production of ethanol by Z. mobilis CP4. Torres and Baratti104 investigated the fermentation of wheat starch hydrolysate by Z. mobilis. They reported that in batch fermentation, sugar concentrations as high as 223 g/l could be fermented to 105 g/l ethanol in 70 h. The percentage theoretical yield was 92%. The fermentation pattern of Z. mobilis strains ATCC10988, ATCC12526 and NRRL B4286 on synthetic medium, cane juice and molasses showed that the strain NRRL B4286 produced maximum ethanol on synthetic medium, while the strains ATCC12526 and ATCC10988 performed well on cane juice. However, all the strains fermented molasses poorly105.

We have already reported the ethanol fermentation of cassava starch hydrolysate (CSH) by Z. mobilis106. Studies by Nellaiah et al.107 revealed the strain NRRL B-4286 of Z. mobilis to be superior to the already-established efficient strain, ZM4, in the fermentation of glucose, fructose, and sucrose up to a concentration of 200 g/l. NRRL B4286 also proved to be the best strain for fermentation of cassava starch hydrolysate. Our results showed that adaptation of the cells to the higher concentration of sugars in CSH could help to achieve maximal ethanol concentrations in relatively shorter period of time. With the culture adapted to the concentration of sugars in CSH, fermentation was completed in 28 h with a maximum concentration of 80.1 g/l ethanol. In contrast to this, a maximum concentration of alcohol of 78.5 g/l after 40 h of fermentation was obtained with the non-adapted culture. Supplementation of the CSH with various additives did not result in higher concentration of ethanol107.

When Z. mobilis and S. cerevisiae were compared for their efficiency to produce ethanol from glucose and starch hydrolysate, higher yield was observed for Z. mobilis (Table 3). Z. mobilis distillates showed 5-fold lower non-ethanol byproducts than S. cerevisiae when quality ethanol was produced from rye grain by high temperature extrusion cooking. Ethanol production using a hollow-fibre membrane to recycle amyloglucosidase (AMG) and Z. mobilis (ZM4) in a single-stage conti-nuous fermentor was described by Lee et al.108. Their findings showed that by using liquefied starch substrate, it was possible to obtain volumetric productivity of up to 60 g/l/h at ethanol concentrations of 60–65 g/l. A new method for converting cellulosic material to ethanol by cellulose hydrolysis using commercial cellulase and b -glucosidase, and glucose fermentation with Z. mobilis in a fluidized bed fermentor containing an ethanol adsorbant is reported109.

Continuous fermentation

Among various kinds of fermentation processes studied, a continuous process using co-immobilized AMG and cell was most favorable, with operational stability for over 40 days. Continuous production of ethanol from Jerusalem artichoke Juice using ZM4F of Z. mobilis was studied by Allais et al.111. Their results showed the volumetric productivity to be 67.2 g/l/h with a final ethanol concentration of 42 g/l from 100 g/l initial sugars. Doelle112 described a process for the continuous production of ethanol from hydrolysates of starch. This process made use of Z. mobilis in a single-stage fermentation. The author maintained that the quality of the starch hydrolysate was not crucial to the success of the fermentation, and reported a conversion efficiency of 92%. A process for the continuous production of ethanol on an industrial scale from hydrolysed wheat starch using Z. mobilis was described by Sahm and Bringer-Meyer113. These investigators reported that a strain of Z. mobilis that produced 60 g ethanol/l over a test period of 39 days was used for the industrial-scale fermentation. Hillary et al.114 studied continuous culture

using single- and double- fermentor systems, and reported that this system had higher productivity than immobilized enzyme systems. Biofilm reactors with polypropylene or plastic support were used for ethanol production115. Results showed that the ethanol production rate and concentration were greater in biofilm reactors than in suspension cultures116. In continuous fermentation using mixed cultures of Z. mobilis and S. cerevisiae, production of 54.3 g/l of ethanol was observed within 3 days. These authors reported that a high ethanol productivity of 70.7 g/l/h was obtained with a final ethanol concentration of 49.5 g/l and yield of 0.5 g/g. This amounted to 98% of the theoretical yield and 99% substrate conversion. Therefore this might be considered as right candidate for increasing the rate of the ethanol production in the existing industries.

Simultaneous saccharification and fermentation

In this process, along with Z. mobilis another organism capable of producing carbohydrate hydrolase is used to saccharify the polymeric substrate. The saccharified products are simultaneously utilized by Z. mobilis for ethanol production. Simultaneous saccharification and fermentation (SSF) of cassava starch using Z. mobilis or S. uvarum ATCC 26602 was investigated by Poosaran et al.117. They reported that Z. mobilis fermented considerably faster than S. uvarum, completing the fermentation in 20 h resulting in a yield 95% of the theoretical yield, while S. uvarum required a period of 33 h to complete fermentation resulting in a yield of 90% of the theoretical value. Rhee et al.118 investigated various SSF processes with sago starch using free enzyme and free cells of Z. mobilis, free enzyme and immobilized cells or co-immobilized enzyme and cells. They compared the results obtained in each of the above processes with those obtained with a system using pre-saccharified sago starch. Improvement of ethanol production from sweet sorghum was achieved to 29.7 g ethanol/100 g dry sorghum stalks by using Fusarium oxysporum mixed culture with Z. mobilis119.

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Use of a mixed culture of Saccharomycopsis fibuligera and Z. mobilis, for simultaneous saccharification and fermentation process resulted in 29 g/l ethanol. Ethanol yield could be improved by growing S. fibuligera on liquefied starch for 12 h, followed by addition of Z. mobilis for fermentation. Recently, by this process, steam-pretreated willow was fermented using Z. mobilis, S. cerevisiae, and cellulase119. They could achieve over 85% of the theoretical ethanol yield based on the glucan available in the raw material in three days. Z. mobilis was also co-immobilized with an industrial glucoamylase within beads of kappa-carrageenan, and fermentation of maltodextrin was carried out to produce ethanol120. Co-immobilized Z. mobilis and glucoamylase were also used as a biocatalyst for fuel ethanol production in a three-phase fluidized bed reactor. Various antimicrobial agents were supplemented to SSF of paddy malt mash using a mixed culture of Z. mobilis and S. cerevisiae121 that resulted in 10.1% v/v ethanol which was more compared to the ethanol produced by using boiled and fermented mash (9.3%).

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Studies on co-immobilization of Z. mobilis and AMG in k-carrageenan for this process were carried out by Kim et al.122 using sago starch. These workers observed the conversion of 10–20% sago starch to ethanol with 93–97% of theoretical yields. The authors stated that the combined co-immobilized system showed higher ethanol-producing activity compared to free enzyme and cells or separately immobilized cells and enzyme. Batch fermentation of Cassava starch hydrolysate by immobilized cells of Z. mobilis showed that while a maximum ethanol concentration of 59 g/l and productivity of 3.57 g/l/h could be obtained, the final ethanol concentration obtained with free cells was 66 g/l with a productivity of 2.75 g/l/h. The immobilized cells were reported to be stable for seven cycles.

Since the fermentation of lactose to ethanol is slower than glucose fermentation, considerable interest has been directed towards improving the rate of ethanol production from lactose. Gunasekaran et al.123 have described the increased ethanol production to 72 g/l from lactose using co-immobilized yeast and Z. mobilis in alginate gel. Tanaka et al.124 reported ethanol production by a co-immobilized mixed-culture system of A. awamori and Z. mobilis. The production of ethanol from Manioc (cassava) flour by strains of Z. mobilis was investigated by De Franca et al.125. According to these investigators, strain CP3 proved to be the best in their studies with an ethanol yield of 0.48 g/g and ethanol productivity of 4.14 g/l/h. Ho et al.126 reported the production of ethanol from cassava starch by this process using co-immobilized Z. mobilis and immobilized glucoamylase. John et al.127 improved ethanol production in co-immobilized cultivation of Z. mobilis and A. awamori by increasing oxygen concentration for 24 h, followed by using a mixture of 20% air + 80% nitrogen. Z. mobilis was entrapped into PVA-cryogel carrier in the presence of polyol cryoprotectants and the resultant biocatalyst was examined for fermentation of glucose to ethanol128.

Flocculant strains of Z. mobilis can aid cell recycle in continuous fermentation129,130 or increase the cell density of immobilized cell system and thus increase ethanol production131. Varieties of immobilization cell techniques, including attaching a flocculant strain to glass fibre pad, have been investigated to overcome the cost and complexities associated with maintaining high cell concentrations (70–80 g/l) (refs 132,133), and to reduce ethanol toxicity associated with cell recycle in CSTR. An economic analysis shows that a reduction in the production costs of ethanol134 is expected with immobilized cells of Z. mobilis and Saccharomyces bayanus135, Zymomonas produced ethanol at more than twice the rate of Saccharomyces from glucose with essentially no residual sugar present in the effluent system. Since Z. mobilis forms levan in sucrose medium, it poses instability to the cells during fermentation by immobilization. A mutant lacking the property of levan synthesis has been shown to work well in immobilized fermentation by Kannan et al.136. They reported alcohol production of 73.5 g/l from 150 g sucrose/l. Lee et al.137 demonstrated that addition of immobilized invertase to Z. mobilis culture improved the ethanol production and reduced the byproduct formation. When they added 2,100 U/g of
b -fructofuranosidase immobilized on a methacrylamide base polymer (0.2 g), the ethanol yield was improved from 0.29 to 0.40 g/g. The coupled microorganism-immobilized system was reported to be better than the two-stage (hydrolysis and fermentation) system. Co-immobilization of fermentative cells and hydrolytic enzyme was also demonstrated in agar138, pectin bead139, and loofa sponge140. The effect of these on various parameters of ethanol fermentation using various processes is given in Table 4.

Coculture fermentation

Substrates like cellulose require more complex co-culture or extensive pre-treatment to permit efficient ethanol production by Z. mobilis. Ethanol production from lactose was studied using an adapted initial inoculum of K. fragilis NRRL 665 in monoculture and in co-culture with strains NRRL B 4286 and NRRL B 1960 of Z. mobilis145. The monoculture of adapted K. fragilis produced more ethanol than the non-adapted cultures. In co-culture with Z. mobilis NRRL B 4286, K. fragilis produced 61.3 g/l ethanol. After 72 h, K. fragilis 665 produced 53.5 and 55.1 g/l ethanol from 200 and 250 g/l lactose, respectively, and in co-culture with Z. mobilis 4286, produced 64.4 and 66.0 g/l ethanol. However on increasing the substrates concentration from 100 to 250 g/l, parameters related to growth and sugar uptake were affected in both the cases146. When Z. mobilis was co-cultured with yeast, both the specific ethanol productivity and volumetric productivity increased since both the strains produced ethanol. Though K. fragilis utilized glucose more rapidly than galactose, parameters relating to growth and ethanol productivity were more affected when galactose alone was supplied, than was glucose. Moreover at high ethanol concentrations, growth and ethanol productivity of K. fragilis 665 were greatly affected. Direct fermentation of cassava starch to ethanol using mixed cultures of Endomycopsis fibuligera and Z. mobilis, resulted in more ethanol (10.5%) production compared to other mixed cultures tested as well as monocultures147. When glucoamylase was supplemented to mixed culture, ethanol yield increased from 88% to 98% of the theoretical yield. A mixed culture of Z. mobilis and S. cerevisiae produced 0.5 g ethanol/g sugar consumed with a volumetric productivity of 15 g ethanol/h.

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Solid state fermentation

In recent years, considerable interest has been shown in using agricultural byproducts such as sweet sorghum, corn, apple, grape, sugar cane, sugar beets, fodder beets, and Jerusalem artichoke tubers for fuel ethanol production. Due to the complex composition and insolubility of these agro-substrates, solid-state fermentation of these sources would be economical. Very few reports are available regarding the production of ethanol by solid state fermentation. Amin148 has described ethanol fermentation in solid state by Z. mobilis grown on sugar-beet. Ethanol yield of 0.48 g/g sugar, volumetric productivity of 12 g/l/h and final ethanol concentration of 130 g/l showed good performance of Z. mobilis in a solid-state fermentation. Furthermore, Amin149 reported that during solid-state fermentation fewer by-products were produced, compared to conventional submerged fermentation. At optimum fermentation temperature of 35°C, an ethanol yield of up to 95% of the theoretical value with final ethanol concentration of 142 g/l were obtained.

Improved strains for fermentation

Mutagenesis is particularly useful for manipulation of an organism to improve its tolerance to toxic compounds. Many mutagens have been used to develop high level of ethanol-yielding mutants of Z. mobilis150. Nitrosoguanidine (NTG) was found to be effective for Z. mobilis151 and thus mutants tolerant to 15% ethanol and high temperature were isolated. Mutants have been selected to improve the utilization of feedstock; for example a strain that could efficiently produce ethanol from 25% sugar cane juice or sugar cane syrup without the need for nutritional supplementation or sterilization of the medium has been reported. The growth and ethanol production of Z. mobilis are poor on molasses due to the high concentration of salts. By mutation, the ethanol production from molasses was enhanced by two-fold152. Mutants showing tolerance to salt, low pH (ref. 153) and ability to grow on mannitol154, flocculent mutants155,156, and sucrose or fructose non-utilizers157 have also been reported. A stable thermotolerant mutant of Z. mobilis ZM4 was isolated that showed better ethanol tolerance at 42°C. Growth of Z. mobilis in continuous culture for extended periods results in the selection of spontaneous flocculant variants, and selection for or against depends both on the nutrition and fermenter design158. Skotnicki et al.159 have developed a system for transposon mutagenesis of Z. mobilis. Kannan et al.160 isolated mutants Ls1 and Ls2 of Z. mobilis B-806 lacking extracellular levansucrase. The ethanol yield of the mutants increased from 0.48 g/g to 0.50 g/g on sucrose medium. Lindsay et al.161 isolated glucose-negative fosfomycin-resistant mutants of recombinant E. coli KO11 containing ethanol pathway of Z. mobilis. These mutants (SL31 and SL142) retained the ability to ferment sugars and were used to ferment pentose sugars to ethanol selectively in the presence of high concentrations of glucose. Some hyper-productive strains (SL28 and SL40) were also isolated which completed fermentation rapidly and produced 60 g/l ethanol from 120 g/l xylose in 60 h, 20% more ethanol than KO11 under identical conditions. Glucose-negative and fructose-negative mutants of Z. mobilis were isolated by D-cycloserine-mediated enrichment method162. Recently, a mutant of Z. mobilis was isolated which could use mannitol as the sole carbon source due to the presence of a newly evolved mannitol dehydrogenase. This was shown to have evolved from a mutation in the adhA gene (ZADH-1), thus expanding the substrate range of the wild type to include mannitol163.

Other applications

Misawa et al.164 reported the successful cloning of the b -carotene biosynthesis gene crtB, crtE, crtI, and crtR in pZA22 and transferring them by conjugation into Z. mobilis. The transconjugants of Z. mobilis strain accumulating b -carotene after fermentation process may be utilized as a nutrient for farm animals. Z. mobilis was also used to produce sorbitol and gluconic acid by control of ethanol production165. Belay et al.166 demonstrated ethane production from glucose or starch by co-cultures of methanogens and ethanol producers. Cultures of Methanosarcina barkeri or Thermoanaerobater ethanolica or Candida psuedotropicalis in combination with Z. mobilis, produced ethane on glucose medium. A lactate dehydrogenase gene was introduced into recombinant E. coli K12/FMJ39 carrying plasmid containing genes of ethanol pathway of Z. mobilis. The resulting strain is expected to produce L-lactic acid from mixed sugars.

Conclusion

In recent years, attention has been focussed on effective utilization of agro-byproducts to produce fuel using Zymomonas mobilis. As seen from previous reviews, a thorough investigation of molecular biology and biochemistry of ethanol production by Z. mobilis has been accomplished. Transformation of E. coli with genes (pet) from Z. mobilis for alcohol production has been successfully carried out. However, Z. mobilis only utilizes glucose or fructose or sucrose for ethanol production. In order to broaden the capability of Z. mobilis to utilize other sugars, hydrolytic and isomerase genes from recombinant E. coli have been transferred to Z. mobilis, resulting in utilization of xylose, mannose, lactose and arabinose as the carbon source by Z. mobilis. At present, there exists a considerable literature on transformation of Z. mobilis to produce ethanol from lignocellulose. In addition to this, researchers are also interested in using mixed cultures to convert complex sugars to ethanol. Owing to the genetic amenability of Z. mobilis, it is possible to make use of this organism in industrial production of ethanol. Thus, it is time that the industrialists collaborate with academicians to translate the laboratory findings in science for the benefit of society.

 

 

  1. Cysewski, G. R. and Wilke, C. R., Biotechnol. Bioeng., 1978, 20, 1421–1427.
  2. Swings, J. and De Ley, J., Bacteriol. Rev., 1977, 41, 1–46.
  3. Swings, J. and De Ley, J., in Bergies Manual of Systematic Bacteriology (eds Krieg, N. R. and Holt, J. S.), William and Wilkins, Baltimore, USA, 1984, vol. 2, pp. 576.
  4. Montenecourt, B. S., in Biology of Industrial Organisms (eds Demain, A. L. and Soloman, N. A.), Benjamin–Cummings Publishing Co., California, 1985, pp. 261–189.
  5. Parker, C., Barnell, W. O., Snoep, J. L., Ingram, L. O. and Conway, T., Mol. Microbiol., 1995, 15, 795–802.
  6. Scopes, R. K. and Griffiths-Smith, Biotechnol. Lett., 1986, 8, 653–656.
  7. Barrow, K. D., Collins, J. G., Leigh, D. A., Rogers, P. L. and Warr, R. G., Appl. Microbiol. Biotechnol., 1984, 20, 225–232.
  8. Dawes, E. A., Ribbons, D. W. and Large, P. J., Biochem. J., 1966, 98, 795–803.
  9. Wills, C., Kratofil, P., Londo, D. and Martin, T., Arch. Biochem. Biophys., 1981, 210, 775–785.
  10. Algar, E. M. and Scopes, R. K., J. Bacteriol., 1985, 2, 275–287.
  11. Buchholz, S. E., Dooley, M. M. and Eveleigh, D. E., Trends. Biotechnol., 1987, 5, 199–204.
  12. Pankova, L. M., Shvinka, Y. E., Beker, M. E. and Salva, E. E., Microbiologia, 1985, 54, 141–145.
  13. Viikari, L. and Korhola, M., Appl. Microbiol. Biotechnol., 1986, 24, 471–476.
  14. Wecker, M. S. A. and Zall, R. R., Appl. Microbiol. Biotechnol., 1987, 53, 2815–2820.
  15. Pond, J. L., Eddy, C. K., Mac Kenzie, K. F., Conway, T., Borecky, D. J. and Ingram, L. O., J. Bacteriol., 1989, 171, 767–774.
  16. Michel, G. P. F. and Baratti, J., J. Gen. Microbiol., 1989, 135, 453–460.
  17. Mortatte, M. P. L., Sato, H. H. and Park, Y. K., Biotechnol. Lett., 1983, 5, 518–526.
  18. Viikari, L. and Linko, M., Biotechnol. Lett., 1986, 8, 139–144.
  19. Lyness, E. W. and Doelle, H. W., Biotechnol. Lett., 1983, 5, 345–350.
  20. Liegh, D., Scopes, R. K. and Rogers, P. L., Appl. Microbiol. Biotechnol., 1985, 20, 413–415.
  21. O’Mullen, P., Szakacs-Dobogi, M. and Eveliegh, D. E., Biotech. Lett., 1991, 13, 137–142.
  22. Swings, J. and De Ley, J., Int. J. Syst. Bacteriol., 1975, 25, 321–328.
  23. Brau, B and Sahm, H., Arch. Microbiol., 1986, 144, 296–301.
  24. Conway, T., Osman, Y. A., Konnan, J. I., Hoffman, E. M. and Ingram, L. O., J. Bacteriol., 1987, 169, 949–954.
  25. Neale, A. D., Scopes, R. K., Wattenhall, R. E. H. and Hoogenraad, N. J., J. Bacteriol., 1987, 169, 1024–1028.
  26. Renyen, M and Sahm, H., J. Bacteriol., 1988, 170, 3310–3313.
  27. Conway, T., Sewell, G. W., Osman, Y. A. and Ingram, L. O., J. Bacteriol., 1987, 169, 2591–2597.
  28. Keshav, K. F., Yomano, L. P., An, H. and Ingram, L. O.,
    J. Bacteriol., 1990, 172, 2491–2497.
  29. Conway, T., Sewell, G. W. and Ingram, L. O., J. Bacteriol., 1987, 169, 5653–5662.
  30. Barnell, W. O., Yi, K. C. and Conway, T., J. Bacteriol., 1990, 172, 7227–7240.
  31. Zembruski, B., Chilco, P., Liu, S. L., Conway, T. and Scopes,
    R. K., J. Bacteriol., 1992, 174, 3455–3460.
  32. Hesman, T. L., Barnell, W. O. and Conway, T. J. Bacteriol. 1991, 173, 3215–3223.
  33. Conway, T., Fliege, R., Jones-Kilpatrick, D., Liu, J., Barnell,
    W. O. and Egan, S. E., Mol. Microbiol., 1991, 5, 2901–2911.
  34. Conway, T. and Ingram, L. O., J. Bacteriol., 1988, 170, 1926–1933.
  35. Yomano, P. L., Scopes, R. K. and Ingram, L. O., J. Bacteriol., 1993, 175, 3926–3933.
  36. Burnett, M. E., Liu, J. and Conway, T., J. Bacteriol., 1992, 174, 6548–6553.
  37. Gunasekaran, P., Karunakaran, T., Cami, B., Mukundan, A. G., Preziosi, L. and Baratti, J., J. Bacteriol., 1990, 172, 6727–3223.
  38. Gunasekaran, P., Mukundan, G., Kannan, T. R., Velmurugan, S., Ait-Abdelkader, N., Alvarez-Macarie, E. and Baratti, J., Biotechnol. Lett., 1995, 17, 635–642.
  39. Kanagasundaram, V. and Scopes, R. K., Biochem. Biophys. Acta, 1992, 1171, 198–200.
  40. Kanagasundaram, V. and Scopes, R. K., J. Bacteriol., 1992, 174, 1439–1447.
  41. Kannan, R., Mukundan, G., Ait-Abdelkader, N., Augier-Magro, V., Baratti, J. and Gunasekaran, P., Arch. Microbiol., 1995, 163, 195–204.
  42. Neveling, V., Klasen, R., Bringer-Meyer, S. and Sham, H., J. Bacteriol., 1998, 180, 1540–1548.
  43. Eddy, C. K., Smith, O. H. and Dale-Noel, K., J. Bacteriol., 1988, 170, 3158–3163.
  44. Yanase, H., Kotani, T., Yasuda, M., Matsuzawa, A. and Tonomura, K., Appl. Microbiol. Biotechnol., 1991, 35, 364–368.
  45. Yanase, H., Fukushi, H., Veda, N., Maeda, Y., Toyoda, A. and Tonomura, K., Agric. Biol. Chem., 1991, 55, 1383–1390.
  46. Yanase, H., Iwata, M., Nakahiagashi, R., Kita, K., Kato, N. and Tonomura, K., Biosci. Biotech. Biochem., 1992, 56, 1335–1337.
  47. Conway, T., Osman, Y. A. and Ingram, L. O., J. Bacteriol., 1987, 169, 2327–2335.
  48. Brestic-Goachet, N., Gunasekaran, P., Cami, B. and Baratti, J., Arch. Microbiol., 1990, 153, 219–225.
  49. Skotnicki, M. L., Warr, R. G., Goodman, A. G., Lee, K. J. and Rogers, P. L., Biochem. Soc. Symp., 1983, 48, 53–86.
  50. Brestic-Goachet, N., Gunasekaran, P., Cami, B. and Baratti, J., J. Gen. Microbiol., 1989, 135, 893–902.
  51. Yoon, K. H., Park, S. H. and Pack, M. Y., Biotechnol. Lett., 1988, 10, 213–216.
  52. Lejeune, A., Eveleigh, D. E. and Colson, C., FEMS Microbiol. Lett., 1988, 49, 363–366.
  53. Su, P., Delaney, S. F. and Rogers, P. L., J. Biotechnol., 1989, 9, 139–152.
  54. Yanase, H., Masuda, M., Tamaki, T. and Tonomura, K., J. Ferm. Bioeng., 1990, 70, 1–16.
  55. Buchholz, S. E., Dooley, M. M. and Eveleigh, D. E. J. Ind. Microbiol, 1989, 4, 19–27.
  56. Lodge, J., Fear, J., Busby, S., Gunasekaran, P. and Kamini, N. R., FEMS Microbiol. Lett., 1992, 95, 271–276.
  57. Liu, C. Q., Goodman, A. E. and Dunn, N. W., J. Biotechnol., 1988, 7, 61–70.
  58. Zhang, M., Eddy, C., Deanda, K., Finkelstein, M. and Picataggio, S., Science, 1995, 267, 240–243.
  59. Dally, E. L., Stokes, H. W. and Eveleigh, D. E., Biotechnol. Lett., 1982, 4, 91–96.
  60. Brestic-Goachet, N., Gunasekaran, P., Cami, B. and Baratti, J., Biotechnol Lett., 1987, 9, 13–18.
  61. Skotnicki, M. L., Lee, K. J., Tribe, D. E. and Rogers, P. L., Microbios., 1984, 39, 189–192.
  62. Scordaki, A. and Drainas, C., J. Gen. Microbiol., 1987, 133, 2547–2556.
  63. Tonomura, K., Okamoto, T., Yasui, M. and Yanase, H., Agric. Biol. Chem., 1986, 50, 805–808.
  64. Byun, M. O. K., Kaper, J. B. and Ingram, L. O., J. Ind. Microbiol., 1986, 1, 9–15.
  65. Yoon, K. H. and Pack, M. Y., Biotechnol. Lett., 1987, 9, 163–
    168.
  66. Conway, T., Byun, M. O. K. and Ingram, L. O., Appl. Environ. Microbiol., 1987, 5, 235–241.
  67. Delgado, O. D., Abate, C. M., Sineriz, F., FEMS Microbiol. Lett., 1995, 132, 1–2, 23–26.
  68. Buchholz, S. E. and Eveleigh, D. E., Appl. Environ. Microbiol., 1986, 52, 366–370.
  69. Yanase, H., Yasui, M., Miyasaki, T. and Tonomura, K., Agric. Biol. Chem., 1985, 49, 133–140.
  70. Carey, V. C. and Ingram, L. O., J. Bacteriol., 1983, 154, 1291–
    1300.
  71. Goodman, A. E., Strzelecki, A. T. and Rogeres, P. L., J. Biotechnol., 1984, 1, 219–228.
  72. Chun, U. H. and Rogers, P. L., Biotechnol. Lett., 1986, 8, 807–810.
  73. Kosugi, N., Okamoto, T., Yanase, H. and Tonomura, K., Agric. Biol. Chem., 1986, 50, 209–219.
  74. Misawa, N., Okamoto, T. and Nakamura, K., J. Biotechnol., 1988, 7, 167–178.
  75. Okamoto, T., Yamano, S., Ikeaga, H, and Nakamura, K., Appl. Microbiol. Biotechnol., 1994, 42, 563–568.
  76. Lawford, H. G., Rousseau, J. D., and McMillan, J. D. Appl. Biochem. Biotechnol., 1997, 63, 269–286.
  77. Deanda, K. and Zhang, M., Eddy, C. and Picataggio, S., Appl. Environ. Microbiol., 1996, 62, 4465–4470.
  78. Weisser, P., Kraemer, R. and Sprenger, G. A., Appl. Environ. Microbiol., 1996, 62, 4155–4161.
  79. Uhlenbusch, I., Sahm, H. and Sprenger, G., Appl. Microbiol. Biotechnol., 1991, 57, 1360–1366.
  80. Ingram, L. O., Conway, T., Clark, D. P., Sewell, G. W., Preston,
    J. F., Appl. Environ. Microbiol., 1987, 53, 2420–2425.
  81. Takahashi, D. F., Carvalhal, M. L. and Alterthum, F., Biotechnol. Lett., 1994, 16, 747–750.
  82. Lawford, H. G. and Rousseau, J. D., Energy Biomass Wastes, 1993, 16, 559–597.
  83. Hilaly, A. K., Karim, M. N. and Linden, J. C., J. Ind. Microbiol., 1994, 13, 83–89.
  84. Saucedo, V. M., Karim, M. N., Biotechnol. Lett., 1996, 18, 1055–
    1060.
  85. Tolan, J. S. and Finn, R. K., Appl. Environ. Microbiol., 1987, 53, 2033–2038.
  86. Hespell, R. B., Wyckoff, H., Dien, B. S. and Bothast, R. J., Appl. Environ. Microbiol., 1996, 62, 4594–4597.
  87. Lawford, H. G., Rousseau, J. D., Appl. Biochem. Biotechnol., 1996, 57/58, 277–292.
  88. Lawford, H. G., Rousseau, J. D., Appl. Biochem. Biotechnol., 1994, 45/46, 349–366.
  89. York, S. W., Ingram, L. O., Biotechnol. Lett., 1996, 18, 683–
    688.
  90. Asghari, A., Bothast, R. J., Doran, J. B. and Ingram, L. O., J. Ind. Microbiol., 1996, 16, 42–47.
  91. Asghari, A., Gerber, J. F. and Ingram, L. O., Abstr. Gen. Meet. Am. Soc. Microbiol., 1994, 94, 361.
  92. Moniruzzaman, M., Dien, B. S., Ferrer, B., Hespell, R. B., Dale, B. E., Ingram, L. O. and Bothast, R. J., Biotechnol. Lett., 1996, 18, 985–990.
  93. Doran, J. B., Aldrich, H. C. and Ingram, L. O., Biotechnol. Bioeng., 1994, 44, 240–247.
  94. Gold, R. S., Meagher, M. M., Hutkins, R. W. and Conway, T., Abstr. Gen. Meet. Am. Soc. Microbiol., 1995, 95, 330.
  95. York, S. W., Ingram, L. O., J. Ind. Microbiol., 1996, 16, 374–
    376.
  96. Tolan, J. S. and Finn, R. K., Appl. Environ. Microbiol., 1987, 53, 2039–2044.
  97. Feldmann, S., Sprenger, G. A. and Sahm, H., Appl. Microbiol. Biotechnol., 1989, 31, 152–156.
  98. Neale, A. D., Scopes, R. K. and Kelly J. M., Arch. Microbiol., 1988, 29, 162–167.
  99. O'Mullan, P. J. and Hespell, R. B., Abstr. Gen. Meet. Am. Soc. Microbiol., 1994, 94, 361.
  100. Barbosa, M. D. S. and Ingram, L. O., Curr. Microbiol., 1994, 28, 279–282.
  101. Chay, P. B., Chonvel, H., Cheftel, J. C., Ghomidh, C. and Navarro, J. M., Lebensm–Wiss Technol., 1984, 17, 257–267.
  102. Toran-Diaz, I., Jain, V. K., Allais, J. J. and Baratti, J., Biotechnol. Lett., 1985, 7, 527–530.
  103. Duvnjak, Z., Kosaric, N. and Hayes, R. D., Biotechnol. Lett., 1985, 3, 589–594.
  104. Torres, E. F. and Baratti, J., J. Ferm. Technol., 1988, 66, 67–172.
  105. Gunasekaran, P., Karunakaran, T and Kasthuribai, J. Biosci., 1986, 10, 181–186.
  106. Nellaiah, H., Karunakaran, T. and Gunasekaran, P., Biomass, 1988, 15, 201–207.
  107. Nellaiah, H., Karunakaran, T. and Gunasekaran, P., J. Ferment. Technol., 1988, 66, 219–223.
  108. Lee, J. H., Pagan, J. and Rogers, P. L., Biotechnol. Bioengg., 1983, 25, 659–669.
  109. Scott, C. D., Davison, B. H., Scott, T. C., Woodward, J., Dees, C. and Rothrock, D. S., Appl. Biochem. Biotechnol., 1994, 45/46, 641–653.
  110. Nellaiah, H. and Gunasekaran, P., Indian J. Microbiol., 1992, 15, 201–207.
  111. Allais, J. J., Torres, E. F. and Baratti, J., Biotechnol. Bioeng., 1987, 24, 778–782.
  112. Doelle, H. W., PCT Int. Appl. WO 8702, 702 (C1, C12 P7/10)07, March 1987.
  113. Sahm, H. and Bringer-Meyer, S. Acta Biotechnol., 1987, 7, 307–313.
  114. Hillary, Z. D. and Ishizaki, Biotechnol. Tech., 1997, 11, 537–541.
  115. Reddy Kunduru, M. and Pometto, III, A. L., J. Ind. Microbiol., 1996, 16, 241–248.
  116. Reddy-Kunduru-M and Pometto, III. A. L., J. Ind. Microbiol., 1996, 16, 249–256.
  117. Poosaran, N., Heyes, R. H. and Rogers, P. L., Biomass, 1985, 7, 171–183.
  118. Rhee, S. K., Lee, G. M., Kim, C. H., Abidin, Z. and Han, M. H., Biotechnol. Bioeng. Symp., 1986, 17, 481–493.
  119. Lezinou, V., Christakopoulos, P., Kekos, D. and Macris, B. J., Biotechnol. Lett., 1994, 16, 983–988.
  120. Eklund, R. and Zacchi, G., Enzyme Microb. Technol., 1995, 17, 255–259.
  121. Agrawal, R. and Basappa, S. C., Biotechnol. Lett., 1996, 18, 673–678.
  122. Kim, C. H., Lee, G. M., Han, M. H. and Rhee, S. K., Sanop Misaengmul Hakhoechi, 1987, 15, 55.
  123. Gunnasekaran, P. and Kamini, N. R., World J. Microbiol. Biotechnol., 1991, 7, 552–556.
  124. Tanaka, H., Kurosawa, H. and Murakami, H., Biotechnol. Bioeng., 1986, 28, 1761–1768.
  125. De Franca, E. P., Mano, D. S. and Leite, S. G. F., Rev. Latinoam Microbiol., 1986, 28, 313–316.
  126. Ho, Y. C. and Ghazali, H. M., Pertanika, 1986, 9, 235–240.
  127. John, G., Sisak, C. S., Komaromi, P., Hellendoorn, L. and Schugerl, K., Prog. Biotechnol., 1996, 11, 257–623.
  128. Lozinsky, V. I., Zubov, A. L., and Makhalis, T. A., Prog. Biotechnol., 1996, 11, 112–117.
  129. Toran-Diaz, I., Jain, V. K. and Baratti, J., Biotechnol. Lett., 1984, 6, 389–394.
  130. Prince, I. G. and Barford, J. P., Biotechnol. Lett., 1984, 8, 525–530.
  131. Scott, C. D., Ann. NY Acad. Sci, 1983, 413, 448–456.
  132. Arcuri, E. J., Biotechnol. Bioeng., 1982, 24, 595–604.
  133. Klein, J. and Kressdorf, B., Biotechnol. Lett., 1983, 5, 497–502.
  134. Amin, G., Van den Eynde, E. and Verachtert, H., Eur. J. Appl. Microbiol. Biotechnol., 1983, 18, 1–5.
  135. Luong, J. H. T., Biotechnol. Bioeng., 1985, 27, 1652–1661.
  136. Kannan, T. R., Sangiliyandi, G. and Gunasekaran, P., Enzyme Microb. Technol., 1998, 22, 179–184.
  137. Lee, W. C., Huang, C. T., Enzyme Microb. Technol., 1995, 17, 79–84.
  138. Hellendoorn, L., van-den-Heuvel, J. C., Ottengraf S. P. P., Prog. Biotechnol., 1996, 11, 479–485.
  139. Siva-Kesava, S., Panda, T., and Rakshit, S. K., Process Biochem., 1996, 31, 449–456.
  140. Ogbonna, J. C., Tomiyama, S. and Tanaka, H., Process Biochem., 1996, 31, 737–744.
  141. Amutha, R. and Gunasekaran, P., J. Microbial Biotechnol., 1994, 9, 22–34.
  142. Kannan, T. R., Sangiliyandi, G. and Gunasekaran, P., Biotechnol. Lett., 1992, 19, 661–664.
  143. Rogers, P. L., Lee, K. J., Skotnicki, M. L. and Tribe, D. E., Adv. Biochem. Eng. Biotechnol., 1982, 23, 37–84.
  144. Gunasekaran, P., Karunakaran, T., Nellaiah, H., Kamini, N. R. and Mukundan, A. G., Ind. J. Microbiol., 1990, 30, 107–133.
  145. Kamini, N. R. and Gunasekaran, P., J. Ferment. Bioeng., 1989, 68, 305–309.
  146. Kamini, N. R. and Gunasekaran, P., Curr. Microbiol., 1987, 16, 153–157.
  147. Reddy, O. V. S. and Basappa, S. C., Biotechnol. Lett., 1996, 18, 1315–1318.
  148. Amin, G., Biotechnol. Lett., 1992, 14, 499–504.
  149. Amin, G. and Allah-A. M. R., Biotechnol. Lett., 1992, 14, 1187–1192.
  150. Skotnicki, M. L., Lee, K. J., Tribe, D. E. and Rogers, P. L., in Genetic Engineering of Microorganism for Chemicals (ed. Hollaender, A.), Plenum Press, New York, 1982, pp. 271–290.
  151. Sreekumar, O. and Basappa, S. C., Biotechnol. Lett., 1991, 13, 365–370.
  152. Eveleigh, D. E., Stokes, H. W. and Dally, E. L., Biotechnol. Ser., 1983, 4, 69–91.
  153. Ingram, L. O., Garey, V. C., Dombeck, K. M., Holt, H. S., Holt,
    W. A., Osman, Y. A. and Walia, S. K., Biomass., 1984, 6, 131–143.
  154. Buchholz, S. E., O’Mullan, P. and Eveleigh, D. E., Appl. Microbiol. Biotechnol., 29, 275–281.
  155. Doelle, H. W. and Greenfield, P. F., Appl. Microbiol. Biotechnol., 1985, 22, 405–410.
  156. Lawford, G. R., Lavers, B. H., Good, D., Charley, R., Fein, J. and Lawford, H. G., in Ethanol from Biomass (eds Duckworth, H. E. and Thomson, E. A.), Royal Society of Canada, Ottawa, 1983, pp. 482–507.
  157. Suntinanalert, A. E., Rogers, P. L. and Skotinicki, M. L., Biotechnol. Lett., 1986, 8, 351–356.
  158. Stranberg, G. W., Donaldson, T. L. and Arcuri, E. J., Biotechnol. Lett., 1982, 6, 347–352.
  159. Skotnicki, M. L., Tribe, D. E. and Rogers, P. L., Appl. Environ. Microbiol., 1980, 40, 7–12.
  160. Kannan, T. R., Mukundan, A. G. and Gunasekaran, P., J. Ferment. Bioeng., 1993, 75, 265–270.
  161. Lindsay, S. E., Bothast, R. J. and Ingram, L. O., Appl. Microbiol. Biotechnol., 1995, 43, 70–75.
  162. Ait-Abdelkader, N., Pencreach, G., Joset, F. and Baratti, J. C., Appl. Environ. Microbiol., 1996, 62, 1096–1098.
  163. O’Mullan, P., Stein, D., Chase Jr, T., Eveleigh, D. E., Abstr. Gen. Meet. Am. Soc. Microbiol., 1995, 95, 381.
  164. Misawa, N., Yamano, S. and Ikenaga, H. Appl. Environ. Microbiol., 1991, 57, 1847–1849.
  165. Sahm, H., Loos, H., Rehr, B. and Sprenger, G., Meded. Fac. Landbouwwet. Rijksuniv. Gent., 1994, 59, 2403–2410.
  166. Belay, N., Daniels, L., Abstr. Gen. Meet. Am. Soc. Microbiol., 1995, 95, 331.

ACKNOWLEDGEMENTS. We thank CSIR, UGC and DST, New Delhi, for financial support.