African Journal of
Microbiology Research

  • Abbreviation: Afr. J. Microbiol. Res.
  • Language: English
  • ISSN: 1996-0808
  • DOI: 10.5897/AJMR
  • Start Year: 2007
  • Published Articles: 5233

Full Length Research Paper

Engineered microbial consortium for the efficient conversion of biomass to biofuels: A preliminary study

Ugochukwu Anieto
  • Ugochukwu Anieto
  • Department of Biological Sciences, University of North Texas, Denton 76203, TX, USA.
  • Google Scholar

  •  Received: 15 October 2016
  •  Accepted: 05 January 2017
  •  Published: 21 January 2017


This work showed ethanol production by a microbial consortium of Clostridium cellulolyticum and a recombinant Zymomonas mobilis (ZM4 pAA1). The ZM4 pAA1 and wild type ZM4 (ZM4 WT) were first tested on RM medium (ATCC 1341) containing 2% cellobiose as the carbon source. Ethanol production from ZM4 pAA1 was three times higher than that observed from the ZM4 WT. Concomitant with ethanol production was the reduction in OD from 2.00 to 1.580. The ZM4 pAA1 was then co-cultured with C. cellulolyticum using cellobiose and microcrystalline cellulose, respectively, as carbon sources. Results indicate that the ZM4 pAA1 with C. cellulolyticum utilized 2.0 g/L cellobiose, producing as much as 0.40 mM of ethanol, whereas only 0.20 mM ethanol was detected for the ZM4 WT co-cultured with C. cellulolyticum under similar conditions. A consortium of the ZM4 pAA1 and C. cellulolyticum using 7.5 g/L microcrystalline cellulose gave a far lower ethanol yield than when using cellobiose. In the latter case, ethanol production was detected within 5 days, whereas it took about 10 days for ethanol to be detectable for the ZM4 WT and C. cellulolyticum. Future efforts will concentrate on identifying suitable partners for the ZM4 pAA1, the correct concentration of feedstocks at which synergy will be observed, as well as optimize medium formulations and inoculation techniques.

Key words: Biofuel, ethanol, cellulosome, consortium, Zymomonas mobilis, Clostridium cellulolyticum.


Zymomonas mobilis is a facultative anaerobic Gram-negative bacterium belonging to the alpha subdivision of the phylum Proteobacteria, class Alpha-Proteobacteria, order Sphingomonadales and family Sphingomonadaceae. It is rod shaped with dimensions 1.0-2.0 × 4.0-5.0 µm, motile, does not sporulate, does not produce capsules, intracellular lipids or glycogen, optimal pH  range  for  growth  is  6  to  7.0,  optimal  temperature range is 25 to 31°C, the G + C content of the cellular DNA is about 47.5 to 49.5% with an average Tm of 89.3 to 89.5°C (Gunasekeran et al., 1990). Z. mobilis uses the Entner-Doudoroff (ED) pathway which is found in microrganisms that are strictly aerobic, conducts fermentation with 50% less ATP produced relative to the Embden-Meyerhof-Parnas (EMP) pathway, which leads to improved ethanol yield (Yang et al., 2016).
Clostridium cellulolyticum ATCC 35319 formerly identified as strain H10 was isolated in the fall of 1982   from decayed grass compost packed for 3 to 4 months at the Université de Nancy, France. it is a Gram-positive, straight to slightly curved rods that are 3 to 6 µm long by 0.6 to 1.0 µm wide, with a mean G + C content of 41% and forms spores in cultures of cellulose media 3 or more days old (Petitdemange et al., 1984). C. cellulolyticum produces several cellulases, which are re-grouped into an extracellular enzymatic complex called cellulosomes and cellulolytic activities allow the release of soluble cellodextrins from cellulose, which in return permits microbial growth (Desvaux, 2005).
The wild-type Z. mobilis was primarily isolated from alcoholic liquids in natural environments containing fermentable sugars such as plant saps, and can only utilize a limited carbon source, including glucose, fructose and sucrose (Weir, 2016). This drawback is a major reason why it has not been used extensively for bioethanol production on an industrial scale. C. cellulolyticum is excellent at cellulose utilization due to the presence of cellulosomes, a complex mass of enzymes comprised of an array of cellulases and hemicellulases, but unlike Z. mobilis has a low yield of ethanol production due to its inability to utilize higher concentrations of pure cellulosic substrates. It has been suggested that there is also an overflow of pyruvate higher than the rate of procession of pyruvate ferridoxin oxidoreductase (PFO) and lactate dehydrogenase (LDH) (Senthikumar and Gunasekaran, 2005) in C. cellulolyticum.
Engineered microbial consortia and co-culturing of wild type bacteria have become pivotal tools in biotechnology and have been utilized in the production of a wide range of products from biofuels to pharmaceuticals. For examples, Abate et al. (1996) described ethanol production by a co-culture of Z. mobilis and Saccharomyces sp. with higher yields and production rates than with either microorganism in pure culture. Shin et al. (2010) genetically engineered two E. coli strains for xylan utilization, with one strain expressing two hemicellulases to hydrolyze xylan into xylooligosaccharides and another one importing the xylooligosaccharides to produce ethanol, with a 55% theoretical yield. Similarly, Shou et al. (2007) demonstrated a slightly more ideal cooperation using two engineered Saccharomyces cerevisiae strains. One strain required adenine and overproduced lysine while the other strain required lysine and overproduced adenine. Singh et al. (2014) co-cultured Pichia stipitis and Z. mobilis for bioethanol production from kans grass biomass with significant yields and Zhang et al. (2016a) employed C. cellulolyticum and hydrogen fermentation bacteria for enhanced biohydrogen production from corn stover with significant differences seen in the metabolites of the co-culture system  over  the  mono-cultures.  Other reports of successful bio-catalysis based on microbial consortia have equally been reported (Fu et al., 2009; He  et al., 2011; Ho et al., 2011; Li et al., 2011; Quinn et al., 2016; Reddy and Basappa, 1996; Yaun et al., 2016; Zhang et al., 2016b; Zhong et al., 2016).
In order to improve its industrial appeal in substrate utilization and sugar transport, Z. mobilis has been genetically modified extensively with significant improvements over the wild type, for example Luo and Bao (2015), expressed a heterologous β-D-glucosidase from Bacillus polymyxa in Z. mobilis, where the signal peptide ZMO 1086 facilitated its secretion. Other reported efforts include the works of Deanda et al. (1996), Dunn and Rao (2014), Yanase et al. (2005) and Yanase et al. (2012). To the best of the author’s knowledge, there has not been any reported microbial consortium involving Z. mobilis and C. cellulolyticum whether as wild types or engineered clones for the production of bioethanol. Given the immense potentials as previously outlined of these two bacteria, it became necessary to establish a testbed from which further research would be conducted in order to improve the process of bioethanol production. The present study aimed to study the effects of using a consortium of a recombinant Z. mobilis and C. cellulolyticum on the conversion of biomass to bioethanol and using this study as a Launchpad for further experimental studies and process improvement.


Construction of ZM4 pAA1
Amplification of pBBR1 MCS-3
The vector backbone, pBBR1 MCS-3 (Kovach et al., 1995) was linearized with the restriction enzyme KpnI, the linearized vector verified for size correctness on electrophoresis gel to give a band size of 5.2 kb. The fragment was PCR amplified using Phusion DNA polymerase (NEB), with the PCR conditions set at 98°C for 1 min, 98°C for 30 seconds, 56°C for 30 s for annealing and 72°C for 90 s for extension. The cycle was repeated 35 times from the second to the fourth step, a final extension for 5 min at 72°C and a hold at 4°C.
Amplification of celZ and celY genes from Erwinia chrysenthemi
The celY endoglucanase gene and celZ endoglucanase gene with the ZM4 promoter from pLOI 2352 (kindly provided by Professor L. Ingram, University of Florida) were individually amplified using Phusion DNA polymerase (NEB). For the celY gene, the PCR condition was set at 98°C for 1 min, 98°C for 30 s, 72°C for 45 s (to include annealing and extension). The cycle was repeated 35 times from the second to the merged annealing and extension steps, a final extension for 5 min at 72°C and a hold at 4°C. Similar conditions were used for the celZ gene with the ZM4 promoter but with the annealing temperature set at 54°C for 30 s and extension time for 45 s. The amplicons were verified on the gel for size correctness and further sequenced for correctness.
Amplification of glucose-fructose oxidoreductase (GFOR) leader sequence of Z. mobilis and Β-glucosidase gene of Ruminococcus albus in Z. mobilis.
The β-glucosidase gene from R. albus was cloned and translationally fused to the glucose-fructose oxidoreductase (gfor) leader sequence of Z. mobilis for export; resulting in 61% secretion and 0.49 g ethanol yield per g cellobiose (Yanase et al., 2005). To amplify the 159 bp leader sequence of the glucose-fructose oxidoreductase gene, the reverse primer was designed to include 10 bp forward primer of the β-glucosidase gene. Similarly, the forward primer of the β-glucosidase gene was designed to include 10 bp of the complimentary sequence of the gfor leader reverse primer sequence. The β-glucosidase gene from the genomic DNA of R. albus (kindly provided by Professor P. Weimer, University of Wisconsin) and the leader sequence of the gfor gene of Z. mobilis, which was fused to the β-glucosidase gene, were amplified using synthetic oligonucleotide primers (Invitrogen). The PCR condition used was the same as previously described; however, the annealing temperature was 65°C. The amplicons were verified by DNA gel electrophoresis and sequenced for correctness (Table 1).
Cloning and assembly of plasmid pAA1
The cloning and expression of these three genes was to expand the substrate utilization range of Z. mobilis to include larger oligodextrins. These three fragments, ZM4 promoter with celZ, celY and gfor-betaglucosidase fusion were cloned into the KpnI site of the broad host range vector pBBR1MCS-3 (tcr). The cloning was performed using the Life Technologies Gene Art Seamless Cloning and Assembly kit (Life technologies). This kit was optimized to clone up to 4 DNA fragments with a total vector and insert size of 13 kb.
Escherichia coli NEB-10 Beta competent cells were transformed with the cloned vector pAA1 as described in New England Biolabs (NEB) manual. The transformants were plated out on lysogeny medium (LB) containing 40 µL of 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal) for blue/white screening and with 15 µg/ml of tetracycline. To select for the right clone, the plasmid pAA1 and the vector backbone pBBR1MCS-3 were extracted from their respective host cells and subsequently digested using KpnI and NotI HF restriction enzymes (NEB) which cut at different positions within the vector backbone. Furthermore, the vectors pAA1 and pBBR1MCS-3 were digested with the restriction enzyme NdeI (NEB). NdeI cut site CAˆTATG exists within the insert that produced pAA1 but not on the vector backbone pBBRIMCS-3.
The cloned pAA1 was completely sequenced by Eurofins MWG operon  using   the   Sanger   sequencing  method  and  verified  for correctness and thereafter used for the transformation of ZM4 WT using the Gene Pulser (Bio-Rad) as described by Liang and Lee (1998) to create ZM4 pAA1. The Z. mobilis cultures were grown in a stationary flask at 30°C to an absorbance (600 nm) of 0.3 to 0.4. The cells were harvested by centrifugation at 13000 g for 10 minutes at 4°C. The cells from an original 100-ml culture were suspended in 10 ml of sterile 10% glycerol (supplemented with 0.85% NaCl), centrifuged, and finally re-suspended in 2 to 3 ml of 10% glycerol. The plasmid pAA1 was extracted from E. coli NEB-10 Beta, suspended in water and concentrated to 3000 ng/µl DNA before electroporation.
The Gene Pulser (Bio-Rad) for generating exponential decay pulses were set at a peak voltage of 1.5 kV and 25 µF capacitance. A 200 µL aliquot of the Z. mobilis cultures were mixed with 10 µL of pAA1 in a chilled electroporation chamber with an electrode gap of 0.2 cm and held on ice for 5 min. Thereafter, the mixture of cells and DNA was pulsed and immediately after pulsing, the cells were mixed with 1 ml of RM medium for outgrowing at 30°C for 4 h. At the end of this outgrowth period, the cells were diluted with RM medium and plated on RM agar containing 15 µg/ml of tetracycline. The Z. mobilis cultures were also transformed with the original vector backbone pBBR1MCS-3 to create ZM4 pBBR1MCS-3 and plated on RM agar (15 µg/ml of tetracycline) to verify the expression of the plasmid with the antibiotic marker in a new host but without the inserts as seen in pAA1. ZM4 pAA1 and ZM4 pBBR1MCS-3 were inoculated into RM broth containing 15 µg/ml of tetracycline and 30 µg/ml of gentamicin and incubated at 30°C for 48 h. Gentamicin was added because Z. mobilis is naturally resistant to it up to 50 µg/ml, therefore most potential contaminants were not expected to survive under that condition. Thereafter, OD was determined at 600 nm using the spectrophotometer (Spectronic 20D+).
To further verify that the recombinants ZM4 pAA1 and ZM4 pBBR1MCS-3 were indeed Z. mobilis, the cells were once again grown in RM medium with 2% glucose as the carbon source and supplemented with 15 µg/ml of tetracycline. The ZM4 WT was also grown in RM medium with 2 percent glucose as the carbon source and supplemented with 30 µg/ml of gentamicin. Cells were harvested after 48 h and genomic DNA extraction was performed using the FastDNA spin kit (MP Biomedicals). The leader sequence of the gfor is unique to Z. mobilis and so this gene fragment was individually amplified from the genomic DNA extracted from ZM4 pAA1, ZM4 pBBR1MCS-3 and ZM4 WT. Band sizes of approximately 200 bp was observed for the 3 amplicons on the electrophoresis gel.
The recombinant ZM4 pAA1 (tetracycline resistant) was grown in RM medium containing 2% glucose supplemented with 15 µg/ml of tetracycline. The pAA1 was then extracted from ZM4 pAA1 using the  5  Prime  fast  plasmid extraction kit (5 Prime) and the inserts of ZM4celZ, celY and gfor-beta-glucosidase individually amplified from the plasmid pAA1 using the Q5 high fidelity DNA polymerase (New England Biolabs), verified for size correctness on gel electrophoresis and subsequently sequenced for correctness.
Ethanol production test
Ethanol production from cellobiose using recombinant ZM4 pAA1 and ZM4 WT
Cultures of ZM WT were grown in RM medium containing 2% glucose and supplemented with gentamicin (30 µg/ml). The recombinant ZM4 pAA1 were similarly grown in RM medium but were supplemented with 15 µg/ml of tetracycline in order to maintain the vector (pAA1) in addition to 30 µg/ml of gentamicin. Cells were harvested after 24 h, washed thoroughly in phosphate buffered saline (PBS, pH 7.2), re-suspended in RM medium containing 2% cellobiose to give an optical density (OD) of 2.0 and the appropriate antibiotics added in each medium as previously described. The cultures were incubated at 30°C in a shaking incubator (New Brunswick) at 150 rpm. The ODs were determined every twenty-four hours for 7 days and samples taken from the cultures for ethanol quantification starting from 48 h after original inoculation and every 24 h thereafter for three days.
Ethanol production from cellobiose and microcrystalline cellulose using ZM4 pAA1, ZM4 WT and Clostridium cellulolyticum
ZM4 pAA1 and ZM4 WT cultures were grown to OD of 0.4 to  0.6  in
RM medium containing glucose as carbon source. The cultures were washed in PBS (pH 7.2), re-suspended, and then kept on ice until further use (maximum time <20 min). C. cellulolyticum was grown in Clostridium medium (ATCC 1368) containing 7.5 g/L of microcrystalline cellulose as the carbon source to an OD of 0.4 to 0.6. The cultures were subsequently used to inoculate Clostridium medium containing cellobiose and microcrystalline cellulose as carbon sources, respectively. For the mono cultures of ZM4 pAA1, ZM4 WT and C. cellulolyticum, the medium was inoculated with 5% inoculum size (v/v) and for the consortium, the medium was inoculated with 2.5% each of the inoculum size. For the Clostridium medium containing cellobiose, 2 g/L of the oligosaccharide cellobiose was used, whereas for the regular Clostridium medium, 7.5 g/L of cellulose was used. 


From Figures 1 and 2, it can be seen that the ZM4 pAA1 utilized the cellobiose and produced three times as much ethanol after 5 days, respectively, than did the ZM4 WT. The ethanol produced and analyzed using the gas chromatography technique (Shimadzu) by the ZM4 pAA1 was significantly different from that produced by the ZM4 WT (p = 0.03761, SigmaPlot). The ethanol detected in the wild type could have come from the residual glucose from after wash transferred into the medium at inoculation. As ethanol was being produced, the optical density (OD) of the recombinant culture declined gradually to 1.58 after 7 days. Further decline in OD was observed but with no further change after 1.44. No change was detected for the ZM4 WT for the entire duration.
as seen in Figure 3, ZM4 pAA1 produced the most ethanol and was consistent across the three batches whereas ethanol production was not observed at all for ZM4 WT across the three batches. C. cellulolyticum produced ethanol once as can be seen from batch 1 (Figure 3) but none in subsequent batches.In Figure 4, ethanol production can be observed from the two conditions, with ZM4 pAA1 and C. cellulolyticum producing more ethanol than ZM4 WT and C. cellulolyticum. In a study by Payot et al. (1998) detailing the metabolism of cellobiose by C. cellulolyticum growing in continuous culture, it was reported that C. cellulolyticum was able to metabolize only a small quantity of soluble carbohydrates (3 g/L), with the molar growth yield reduced when the initial cellobiose concentration exceeded (2 g/L). In this work, the concentration of cellobiose utilized was originally set at 5 g/L and the results obtained (not shown) clearly indicated that such concentration negatively impacted growth of the C. cellulolyticum but the recombinant ZM4 pAA1 could have aided in rescuing the situation, with subsequent tests using 2 g/L cellobiose showing an improvement. The ZM4 WT however, was unable to do same, possibly due to the lack of the β-glucosidase gene and also by the use of a sugar source that could not adequately support its growth. Furthermore, as described by Payot et al. (1998), C. cellulolyticum growth was limited due to low rate of NADH re-oxidation leading to an intracellular accumulation of the reduced nucleotide and as described by Giallo et al. (1983), acetate was the main product for the continuous cultures of Clostridium. The acetate formation was found to increase with increasing carbon flow, leading to a high ATP production and to an insufficient rate of NADH regeneration (Giallo et al., 1983). They further described the ability of C. acetobutylicum to induce metabolic shifts to produce solvents such as ethanol, butanol and acetone and this shift was associated with high intracellular ATP and NAD(P)H. It does not appear that C. cellulolyticum is able to induce such metabolic shifts to produce reduced compounds such as ethanol. The result obtained from the consortium of recombinant ZM4 with C. cellulolyticum and ZM4 WT with C. cellulolyticum is consistent with previously observed findings, therefore the ethanol produced in the medium is likely as a result of the secretion of the fused β-glucosidase into the medium by ZM4 pAA1 and the ability of ZM4 WT to convert glucose released from cellobiose to ethanol, respectively. Figure 5 shows ethanol production from microcrystalline cellulose by C. cellulolyticum, ZM4 pAA1 and ZM4 WT in five days.
C. cellulolyticum showed ethanol production for only one batch but no ethanol production was detected from ZM4 pAA1 and ZM4 WT respectively. No ethanol production was detected in the first 5 days but was seen within 10 days for C. cellulolyticum. The amount of ethanol produced by C. cellulolyticum only was far lower than seen when C. cellulolyticum was co-cultured with ZM4 pAA1 using cellobiose as the carbon source.
In the consortium, Figures 6a and b, ethanol production was detected for C. cellulolyticum with ZM4 pAA1 for the three batches after 5 days and increased after 10 days while C. cellulolyticum with ZM4 WT produced ethanol in batches 1 and 2 but not batch 3 and only detected after 10 days. This observation suggested that the consortium involving recombinant ZM4 was more efficient than that involving the wild type. This pattern could be seen from both the time it took for ethanol to be detected and the quantity of ethanol detected. There appear to be only slight increases in ethanol production from the consortium involving the recombinant ZM4 pAA1 from the 10-day culture as compared to that produced after 5 days, suggesting that maximum ethanol yield could be possible in slightly over 5 days. The consortium involving the ZM4 WT produced less ethanol and that was detected only after 10 days, suggesting less efficiency in synergy.


The recombinant Z. mobilis bearing the plasmid vector pAA1 supported the findings of Yanase et al. (2005) in which ZM4 re-engineered with β-glucosidase gene from Ruminococcus albus was able to secrete 61% through the cytoplasmic membrane which resulted in the production of 0.49 g ethanol per gram of cellobiose. From this work, the highest ethanol concentration determined for recombinant ZM4 pAA1 was approximately 0.06 mM, whereas the wild type showed no evidence of ethanol production after 5 days on C. cellulolyticum medium containing cellobiose. Despite the tagging of the β-glucosidase gene to gfor, only about 36% of the total activity was reported to be located on the cell surface fraction, with 20% of the activity on the periplasmic fraction (Yanase et al., 2005). With the introduction of celZ and celY genes, the recombinant ZM4 with C. cellulolyticum was able to indicate ethanol production from microcrystalline cellulose within 5 days whereas the consortium of the ZM4 WT took twice that amount of time and produced significantly less ethanol. The recombinant ZM4 and ZM4 WT did not produce ethanol using cellulose as the carbon source as expected but C. cellulolyticum alone  did  after  10 days,  twice the  time  it took to make ethanol for the consortium.
This preliminary study was initiated to study the effect of the microbial consortium on ethanol production. It is far from exhaustive, did not establish a convincing case for the consortium using the two microorganisms and more investigation is required in order to optimize the desired synergy between  the  ZM4  pAA1  and  C. cellulolyticum. The concentration of microcrystalline cellulose used was 7.5 g/L, but further investigation is needed to determine the best concentration at which a synergy can be clearly observed. Furthermore, the culture growth conditions and incubation times chosen could also be a major factor in contributing to the lack of a clear evidence of synergy; therefore,  different   culture   conditions  and  time  points would be tested to better investigate the consortium.
Based on the wide differences in the genetics and physiology of the two bacteria used, the low yields could simply have been as a result of metabolic bottlenecks, which were not investigated in this preliminary study. Batch culture conditions were investigated and the need to investigate same conditions using the continuous culture conditions cannot be overemphasized. Using pure cellobiose and pure microcrystalline cellulose only, were insufficient, other carbon sources to reflect natural conditions would be investigated
Finally, the recombinant ZM4 pAA1 would be partnered with other known cellulose degraders other than the one used here to further understand its efficiency in a consortium. 


There is no conflict of interest in preparation of the manuscript.


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