Consortia-mediated bioprocessing of cellulose to ethanol with a symbiotic Clostridium phytofermentans/yeast co-culture
© Zuroff et al.; licensee BioMed Central Ltd. 2013
Received: 19 October 2012
Accepted: 18 April 2013
Published: 29 April 2013
Lignocellulosic ethanol is a viable alternative to petroleum-based fuels with the added benefit of potentially lower greenhouse gas emissions. Consolidated bioprocessing (simultaneous enzyme production, hydrolysis and fermentation; CBP) is thought to be a low-cost processing scheme for lignocellulosic ethanol production. However, no single organism has been developed which is capable of high productivity, yield and titer ethanol production directly from lignocellulose. Consortia of cellulolytic and ethanologenic organisms could be an attractive alternate to the typical single organism approaches but implementation of consortia has a number of challenges (e.g., control, stability, productivity).
Ethanol is produced from α-cellulose using a consortium of C. phytofermentans and yeast that is maintained by controlled oxygen transport. Both Saccharomyces cerevisiae cdt-1 and Candida molischiana “protect” C. phytofermentans from introduced oxygen in return for soluble sugars released by C. phytofermentans hydrolysis. Only co-cultures were able to degrade filter paper when mono- and co-cultures were incubated at 30°C under semi-aerobic conditions. Using controlled oxygen delivery by diffusion through neoprene tubing at a calculated rate of approximately 8 μmol/L hour, we demonstrate establishment of the symbiotic relationship between C. phytofermentans and S. cerevisiae cdt-1 and maintenance of populations of 105 to 106 CFU/mL for 50 days. Comparable symbiotic population dynamics were observed in scaled up 500 mL bioreactors as those in 50 mL shake cultures. The conversion of α-cellulose to ethanol was shown to improve with additional cellulase indicating a limitation in hydrolysis rate. A co-culture of C. phytofermentans and S. cerevisiae cdt-1 with added endoglucanase produced approximately 22 g/L ethanol from 100 g/L α-cellulose compared to C. phytofermentans and S. cerevisiae cdt-1 mono-cultures which produced approximately 6 and 9 g/L, respectively.
This work represents a significant step toward developing consortia-based bioprocessing systems for lignocellulosic biofuels production which utilize scalable, environmentally-mediated symbiosis mechanisms to provide consortium stability.
KeywordsConsortia Consolidated bioprocessing Cellulosic ethanol Symbiosis Oxygen transport
In nature microbes rarely live in isolation, but rather exist in highly diverse and complex communities referred to as consortia . These consortia are often capable of tasks that are far too complex for any single organism to complete themselves including some of the most important global biogeochemical cycles . The organisms living in these communities interact in numerous ways ranging from cooperation to direct competition . Microbiologists and engineers have come to appreciate the diversity and capacity of natural microbial communities and large efforts have been undertaken to understand natural consortia and to engineer synthetic consortia for biotechnological purposes [4–7].
Two of the key challenges in society today are to reduce energy dependence on petroleum and reduce greenhouse gas emissions . The transportation sector is a prime candidate for addressing these two challenges because it relies on petroleum for approximately 93% of its energy and releases almost as much carbon dioxide as both the commercial and residential sectors combined . Lignocellulosic biomass has become an increasingly feasible source of carbohydrates for biological production of alternative fuels such as ethanol . Consolidated bioprocessing (the simultaneous biological hydrolysis and fermentation of biomass; CBP) is thought to be one of the most cost effective means of producing ethanol from lignocellulose [11, 12]. However, no single organism has been isolated or genetically engineered to reach high enough ethanol concentrations, yields and productivities from lignocellulose . Natural microbial consortia, on the other hand, are innately capable of high conversion of lignocellulosic biomass [see Table one in reference  but the resultant products (e.g., organic acids, CO2, CH4) are not suitable large-scale liquid transportation fuels.
An elegant example of a naturally occurring, lignocellulose degrading microbial consortium is the symbionts of the termite hindgut. Complex macromolecules are deconstructed in a series of steps that are facilitated by specific microbial species . Protists produce cellulases for cellulose hydrolysis, while nitrogen fixing bacteria (e.g., Clostridia) sequester atmospheric nitrogen to compensate for the low nitrogen content of wood. Fermentative organisms consume soluble sugars released from cellulose hydrolysis to produce organic acids, CO2 and H2 which are subsequently converted by methanogens to methane . This, along with other examples of natural microbial communities, has prompted interest in utilization of microbial consortia for lignocellulosic biofuels production [5, 7]. However, unlike these natural systems, scientists are currently limited in our ability to generate stable, productive microbial communities. In order to successfully implement large scale consolidated bioprocessing of lignocellulosic materials for fuel ethanol production we must develop stable microbial consortia with the necessary functionality, process control and efficiency.
To transition from natural consortia which contain potentially hundreds of organisms to synthetic consortia containing several defined species, one must develop a mechanism for population control. Generating stable, controllable consortia has been a focus of recent work to genetically engineer mechanisms for establishing intra-species consortia [16–19]. These approaches range from growth-controlling genetic circuits based on quorum sensing compounds  to complimentary auxotrophic amino acid exchange . As proof-of-concept, these studies are quite elegant and encouraging but they may suffer on the large scale due to unstable and difficult genetic modifications in industrially relevant organisms. Others have demonstrated syntrophic interactions in a co-culture of Actinotalea fermentans with an engineered S. cerevisiae which produces methyl halides directly from cellulose . S. cerevisiae relieves acetate inhibition by converting it to methyl halides, which are precursors for various fuel compounds. However, synthetic syntrophic interactions may be unstable since at least one organism does not necessarily rely on the other for survival. Without an additional level of control (e.g., spatial structure [21–23]) the community may breakdown. Obligate mutualisms such as those described by You et al. 2004 and Shou et al. 2007 may be a more stable approach for consortia-mediated lignocellulosic ethanol production. With this in mind, symbiotic consortia of a wide range of organisms could be generated using various mechanisms based on both genetic and environmental control factors .
In this work we develop a symbiotic co-culture of the cellulolytic mesophile, Clostridium phytofermentans and a cellodextrin fermenting yeast, Candida molischiana or S. cerevisiae cdt-1 . We establish a symbiosis (obligate mutualism) between C. phytofermentans and the yeast species by controlling the volumetric transport rate of oxygen. Both yeasts are capable of providing respiratory protection to the obligate anaerobe, C. phytofermentans, in return for soluble carbohydrates released from cellulose hydrolysis. The yeast converts these soluble carbohydrates to ethanol. At high substrate loading we noted a decreased conversion of cellulose by C. phytofermentans therefore endoglucanase was added to further evaluate the potential for improvements in the co-culture approach. In this cellulase-assisted format, the co-culture produces over twice as much ethanol and degrades two and three times as much α-cellulose as the S. cerevisiae cdt-1 and C. phytofermentans mono-cultures, respectively. This work represents a significant step in utilizing a scalable environmental control mechanism (i.e. oxygen transport) to induce a stable symbiosis in a consortium of two diverse organisms. This general approach to symbiosis development is applicable to a variety of organisms, substrates and products and can be used to explore diverse consortia-mediated bioprocesses.
Results and discussion
Design and development of C. phytofermentans/yeast symbiosis
Oxygen introduction was proposed as a mechanism for establishing a symbiosis between mixtures of organisms in which one organism “protects” the other from oxygen inhibition in return for soluble carbohydrates released from cellulose . In this work we investigated this mechanism and the subsequent performance of a combination of the cellulolytic obligate anaerobic mesophile, C. phytofermentans, with one of two cellodextrin fermenting yeasts, C. molischiana or S. cerevisiae cdt-1. C. phytofermentans was chosen for its ability to hydrolyze and ferment pure cellulose and natural lignocellulosic materials. For example, under consolidated bioprocessing (CBP) conditions (i.e., simultaneous biological hydrolysis and fermentation), C. phytofermentans was shown to hydrolyze 76% of glucan and 88.6% of xylan in AFEX treated corn stover and ferment it to 2.8 g/L ethanol in 10 days . C. molischiana is naturally able to ferment cellodextrins with degree of polymerization 2 to 6 to ethanol with yields of approximately 0.40 g ethanol/g substrate in 44 hours . S. cerevisiae cdt-1 was engineered to import and cleave cellodextrins for subsequent fermentation (yield of approximately 0.44 g/g cellobiose in about 100 hours) through expression of cellodextrin transporters and β-glucosidase from Neurospora crassa. Although similar, the yeasts do have some important differences. For example, S. cerevisiae cdt-1 transports and cleaves cellodextrins internally  while C. molischiana does so externally with extracytoplasmic β-glucosidase . These yeasts were chosen for their unique ability to ferment cellodextrins (not only glucose) which was hypothesized to increase ethanol yield and relieve cellulase inhibition by the soluble sugar products. In addition, these two organisms provide a comparison across two genus and between engineered and naturally occurring strains. In this study we used α-cellulose (Sigma) as a model cellulosic substrate to study hydrolysis and fermentation since it avoids the presence of additional carbon sources (e.g., hemicellulose).
Low level oxygen transfer promotes symbiosis while maintaining ethanol production
Using this OTR, C. molischiana and S. cerevisiae cdt-1 mono-cultures grew and fermented cellobiose with a lag phase of approximately 150–200 and 50–75 hours, respectively (Additional file 1B and F). C. phytofermentans mono-cultures displayed only slightly inhibited growth and ethanol production compared to no oxygen controls (Additional file 1D). Without oxygen, growth and/or cellobiose consumption were not observed in C. molischiana mono-cultures while gradual cellobiose consumption was observed in S. cerevisiae cdt-1 mono-cultures (Additional file 1A and E). In fact, even with the addition of Ergosterol and Tween 80 (two common anaerobic growth factors for yeast ) and replacement of cysteine with glutathione  we were unable to achieve growth of C. molischiana on cellobiose under completely anaerobic conditions. S. cerevisiae cdt-1, on the other hand, displayed growth under these conditions (Additional file 2). This observation agrees well with other reports that indicated only a small number of yeast are able to grow in the complete absence of oxygen .
As expected, in anaerobic cellobiose co-culture C. molischiana and S. cerevisiae cdt-1 were outcompeted; the viable cell counts fell below detection (< 100 CFU/mL) after about 100–200 hours (Additional file 3). On the other hand, co-cultures with oxygen transfer fermented all the cellobiose and supported the growth of both C. phytofermentans and C. molischiana or S. cerevisiae cdt-1 (Additional file 3). The ethanol yield of the co-culture was slightly lower than that of the yeast mono-cultures since a small amount of acetate was produced early in the fermentation. In contrast to Figure 1, C. phytofermentans benefited early in the culture since the media was initially anaerobic while the increase in yeast viable cell counts occurred after approximately 100–200 hours during which time oxygen was continuously diffused into the culture (Additional file 3). This OTR appeared suitable for inducing and maintaining a co-culture of C. phytofermentans and either C. molischiana or S. cerevisiae cdt-1 on cellobiose while allowing for high ethanol yield. Therefore, the same OTR was implemented in cellulose fermentations. In contrast to a recent co-culture CBP study which required sequential culturing conditions for complimentary function while selectively inhibiting growth , this approach allows simultaneous growth and fermentation of both organisms which drastically simplifies culturing techniques.
Stable co-culture cellulose fermentation
To investigate the scalability and attempt to improve performance of the co-culture, the fermentations were scaled up ten-fold (from 50 mL to 500 mL) to an Infors Sixfors bioreactor system (Figure 3C) with enhanced mixing, pH control and an α-cellulose concentration of 100 g/L. The OTR was maintained at approximately 0.4 μmol O2/hour by increasing the submerged tubing length by a factor of ten from 10 to 100 cm. Under these conditions, the population dynamics were essentially identical to the small scale reactors with only a slightly longer delay in S. cerevisiae cdt-1 re-growth (Figure 4C). Even with pH control, mixing and higher initial substrate concentration, the fermentation performance was not significantly improved compared to the small scale fermentations (data not shown). It is none-the-less encouraging that the population control mechanism based on oxygen transport rates could be readily scaled. In addition to increasing tubing length it is easy to envision that the OTR could be maintained by altering tubing thickness, internal oxygen concentration or with minimal gas sparging at a larger scale.
The long term population stability of the co-culture was determined by growing C. phytofermentans and S. cerevisiae cdt-1 in co-culture with and without oxygen for 50 days. Serum bottles with 25 mL GS2 medium and 150 g/L α-cellulose were inoculated with approximately equal cell concentration (about 105 CFU/mL) of C. phytofermentans and S. cerevisiae cdt-1. At 30, 40 and 50 days the individual cell concentrations were determined (Additional file 5A and B). The co-culture with added oxygen maintained C. phytofermentans and S. cerevisiae cdt-1 concentrations of 105 to 107 CFU/mL at every time point whereas, without oxygen, the yeast population fell below detection prior to the 30 day sample. The presence of yeast in the oxygen positive co-culture was also apparent in that no glucose was accumulated compared to almost 3 g/L glucose accumulation in the co-culture without oxygen (Additional file 5C and D).
These results demonstrate the ability to effectively generate a stable, symbiotic co-culture of C. phytofermentans and S. cerevisiae cdt-1 via diffusion of oxygen at a rate of approximately 8 μmol O2/L hour. The oxygen transfer mechanism was scalable from 50 mL to 500 mL. However, the fermentations were still limited in their hydrolytic capacity meaning the co-culture would not significantly out-perform C. phytofermentans mono-cultures.
Co-culture simultaneous saccharifcation and fermentation
The lack of increased performance of the co-culture relative to C. phytofermentans mono-culture appears to be due to an inability of C. phytofermentans to hydrolyze high solids loading cellulose to provide soluble substrate for the yeast partner. C. phytofermentans was repeatedly unable to fully utilize insoluble substrate concentrations in excess of approximately 30 g/L (unpublished observation). To explore the potential for improving the co-culture approach we sought to increase soluble sugar yield by fermenting α-cellulose with and without added endoglucanase from Trichoderma viride (18.104.22.168). This specific approach was taken to simulate additional hydrolysis capacity of C. phytofermentans while acknowledging the differences that exist between fungal and Clostridial cellulases.
These results demonstrate that co-culture with C. phytofermentans significantly enhances simultaneous saccharification and fermentation of α-cellulose with S. cerevisiae cdt-1. It is important to note in Figure 5 that the maximum productivity was observed for the consortium relative to the mono-culture even when exogenous cellulase was provided. This suggests there is considerable room for optimizing the productivity, and contrary to concerns of complexity, the symbiotic consortium is inherently stable. This synergistic interaction produced a final ethanol concentration that was 30% greater than the sum of the mono-culture titers (22 g/L versus 15 g/L). The improvements shown here are likely due to the combination of endoglucanse and the enzyme repertoire of C. phytofermentans relative to the single enzyme (endoglucanase) used in S. cerevisiae cdt-1 mono-cultures. The relief of feedback inhibition on T. viride endoglucanase by consumption of accumulated sugars and decreased pH caused by C. phytofermentans acetate production also likely contributed to the observed improvements. It is important, however, to note that the media conditions are optimized for co-culture growth and not necessarily mono-culture SSF productivity so changes in culturing conditions could lead to improved mono-culture performance. Independent of these subtle interpretations, these results unambiguously demonstrate the power of symbiotic microbial consortia in cellulosic ethanol production. The results also suggest that with improvements in C. phytofermentans hydrolytic capacity, the co-culture has the potential to reach commercially relevant ethanol concentrations from α-cellulose.
This work represents a significant advancement in establishing and controlling a microbial consortium of diverse organisms for biochemical production. In contrast to other approaches for generating symbiotic consortia that require genetic engineering, oxygen introduction is a simple, scalable technique that could be applied to organisms where genetic engineering tools are not yet available. However, both approaches have significant merit and genetic modification combined with an environmental control mechanism, like oxygen tension, could be an extremely powerful approach to consortia-mediated bioprocessing. The successful development of controllable, stable and productive engineered microbial consortia is likely to be a key advancement in many aspects of biotechnology. This approach combines complex functionality that is distributed across numerous organisms that can be independently optimized for a specific function. To gain better understanding, control and predictive capabilities, community mass and energy balance and metabolic models [39–41] may prove quite useful. Modeling combined with ecological theory could be applied to predict the fate of the populations in a consortium and the relative fitness and/or productivity of the individuals within the overall community.
Oxygen-facilitated symbiosis should be pursued as a mechanism for generating stable consortia of various other organisms including other cellulolytic bacteria and fungi as well as pentose and hexose fermenting yeasts and bacteria. In addition, other products, interactions and processes can be explored by incorporating other naturally occurring or genetically engineered organisms that have unique metabolic properties such as the production of biofuel molecules with improved energy density. As a model system, this symbiotic consortium can be used to explore the structural and functional aspects of cellulose degrading microbial communities. The work described herein used pure cellulose as a substrate but more complex lignocellulosic substrates must be investigated. The appropriate combination of organisms in a symbiotic consortium could result in an efficient consortia-mediated bioprocess which could improve the economic feasibility of lignocellulosic biofuels.
Media and chemical sources
GS2 medium  was used in all experimental cultures with the following composition per liter: 6 g Yeast Extract (BD), 2.9 g K2HPO4 (JT Baker), 2.1 g Urea (Sigma), 1.5 g KH2PO4 (JT Baker), 3 g Tri-Na Citrate H2O (Fischer), 2.33 g Cysteine HCl H2O (MP Biomedical), 1 g MOPS (Sigma), 0.1 g MgCl2 6H2O (JT Baker), 0.0113 g CaCl2 Annhydrous (Fischer), and 0.000125 g FeSO4 7H2O (Fischer) and 0.01% Resazurin (Aldrich). The salts solution (MgCl2 6H2O, CaCl2 Annhydrous and FeSO4 7H2O) and resazurin solution were made and sterilized separately and added after autoclave. The pH of the basal components was adjusted to 7 prior to sterilization. The concentration of α-Cellulose (Sigma) was as described in the text. For inoculum cultures α-Cellulose was replaced with 3 g/L Cellobiose. The GS2 medium was modified for α-cellulose co-cultures to contain 10 mg/L Ergosterol and 420 mg/L Tween 80 and 4.09 g/L Glutathione in place of 2.33 g/L Cysteine HCl H2O (termed ETGtGS2). Either GS2 medium without cysteine and/or glutathione or YPC (5 g/L yeast extract, 10 g/L peptone and 3 g/L cellobiose) was used for cultivation of C. molischiana and S. cerevisiae cdt-1. Half strength Sabouraud medium contained (per liter) 5 g Casien Digest (Sigma), 5 g Dextrose (EM Sciences) and 10 g Agar (BD).
Initial cultures of Clostridium phytofermentans ISDgT were prepared from cryogenically (−80°C) frozen stocks in 7.5 mL screw cap tubes with 5 mL GS2 medium. Oxygen was removed from C. phytofermentans culture medium by degassing (until de-pinked) in a Coy Anaerobic Chamber with a 1.5% H2/98.5% N2 atmosphere. C. phytofermentans inoculum cultures were allowed to grow at 30°C to an OD600 of about 0.3. Cultures were then restarted by placing 100 μL of the initial culture into 4.9 mL of fresh GS2 medium. Experiments were inoculated by injecting approximately 2–3 mL of restarted cultures (OD600 of about 0.2-0.3) to give an initial OD600 of approximately 0.0125-0.025. The amount of substrates and products in the inoculum was determined by HPLC and was subtracted from the final concentrations in the cultures to determine true yields.
For cellobiose co-culture experiments and filter paper degradation experiments yeast cultures were grown in GS2 medium without cysteine and/or glutathione under aerobic conditions at 30°C until reaching an OD600 of at least 0.3 then restarted. The cultures were then restarted and the restarted cultures were used as inoculum to obtain an initial OD600 of approximately 0.0125-0.025. For the α-cellulose experiments yeast cultures were grown in YPC medium at 30°C until reaching an OD600 of at least 0.5, were restarted in YPC and allowed to grow until reaching an OD600 of approximately 1 and were then concentrated and used to inoculate with an initial OD600 of about 0.05.
Fermentations were conducted in 100 mL serum vials sealed with butyl rubber stoppers and incubated at 30°C with 200 rpm agitation or in the 500 mL Infors Sixfors bioreactor system at 30°C with 300 rpm stirring. In the Sixfors bioreactor system, pH was controlled using addition of 1 N sodium hydroxide. Cultures were degassed in an anaerobic chamber to remove oxygen at the beginning of the experiment as described previously. A loop of neoprene tubing (OR = 3.175 mm, IR = 1.45 mm) was submerged in the culture medium and attached at each end to stainless steel tubing inserted through the butyl stopper. In oxygen positive cultures, air was allowed to flow through the neoprene tubing at a flow rate of about 100 L/hr using an aquarium pump. The stainless steel tubing was fashioned in a loop for oxygen negative cultures so that no oxygen could enter the culture.
Static hydrolysis experiments were conducted in 30 mL flat-bottom test tubes and incubated at 30°C with periodic agitation. Experimental media was not degassed prior to inoculation and residual carbon was not removed from the inoculum. Resazurin was not included in the media.
Simultaneous saccharification and fermentation experiments were conducted in 100 mL serum vials sealed with butyl rubber stoppers and incubated at 30°C with 200 rpm agitation as previously described. Endoglucanase (22.214.171.124 from Trichoderma viride) was suspended in phosphate buffered saline, degassed and injected to reach a final concentration of 400 mg/L. Cultures without endoglucanase received degassed, phosphate buffered saline only. One way valves attached to needles were inserted through the butyl stopper to release the gas produced via fermentation.
Dry weight measurements
Cellulose dry weight samples (1 mL) were removed from reactors with a syringe and placed into pre-tared 1.7 mL eppi-tubes (VWR). The samples were centrifuged at 14,000 x g for 10 minutes and the supernatant was removed (used as HPLC sample as described in the following section). The cellulose pellet was washed with DI H2O and again centrifuged at 14,000 x g for 10 minutes. The supernatant was removed and the tube was placed, open, in a drying oven at 70°C. The samples were dried until reaching a constant mass and then weighed. The difference between the final and initial weight of the tube was assumed to be the dry mass of cellulose neglecting cellular mass.
Fermentation product determination
Cellobiose, glucose, ethanol and acetate concentrations were determined from supernatant (as described previously) taken from each culture and frozen immediately at −20°C. Following the completion of the experiment the samples were thawed and filtered into 1.5 mL screw cap vials (Agilent) using 0.45 μm nylon syringe filters (Whatman). Samples were analyzed on an Agilent 1100 HPLC with a Jasco RI-1531 refractive index detector (RID) using an Aminex HPX-87H Cation exchange column (BioRad) with the appropriate guard column. Filtered (through 0.22 um filter) 0.01 M sulfuric acid was used as the mobile phase, the column temperature was set at 65°C and the RID was set at 30°C. Samples were injected at a volume of 25 uL and the operating flow rate was 0.6 mL/min. Product formation reflects correction to the initial concentration in the medium immediately following inoculation. Therefore, carryover of low concentrations of fermentation products and subsequent consumption of these products (e.g., ethanol) may lead to negative production values.
Soluble carbohydrate determination
Soluble carbohydrates were determined using the anthrone-sulfuric acid colorimetric assay of Dreywood (Dreywood, 1946) adapted to a 96-well plate (Leyva et al. 2008). Breifly, 50 μL of appropriately diluted sample was mixed with 150 μL anthrone reagent (2 g/L anthrone in 98% sulfuric acid) in a polypropylene 96-well plate and covered with a nylon adhesive cover. The 96-well plates were incubated at 4°C for 10 minutes, 100°C for 20 minutes and room temperature for 20 minutes. The nylon cover was then removed and the plate was read in a Spectramax 384 Plus spectrophotometer at 620 nm. The soluble carbohydrate concentration was determined using a known concentration standard curve that was run with each 96-well plate used.
Colony forming unit determination
C. phytofermentans and yeast populations were determined by performing serial dilutions of experimental samples in 96-well plates and subsequent drop plating of each dilution on the appropriate medium. C. phytofermentans was plated on GS2 medium with 6 g/L lactose and was incubated at 30°C in an anaerobic chamber. C. molischiana and S. cerevisiae cdt-1 were plated on ½ Sabouraud medium and incubated at 30°C under aerobic conditions. Co-culture samples were plated on both types of plates and incubated under their respective conditions. Colonies were counted on dilutions that contained approximately 3 to 30 colonies per drop. Due to the non-soluble nature of cellulose which may act to trap, attach to or reject cells, colony counts often displayed significant variability but trends were found to be consistent across experiments.
TZ and WC conceived and designed the study. TZ carried out the majority of the experiments and drafted the manuscript. SX conducted the C. phytofermentans/C. molischiana static hydrolysis experiments and analysis as well as participated in media design. TZ and WC edited the draft. All authors read and approved the final manuscript.
Ammonia fiber expansion
Optical density at 600 nm
Oxygen transfer rate
High performance liquid chromatography
Colony forming unit
Simultaneous saccharificaiton and fermentation
We are grateful to Dr. Jamie Cate at the University of California, Berkeley for providing Saccharomyces cereivisae strain cdt-1, Dr. Susan Leschine at the University of Massachusetts, Amherst for Clostridium phytofermentans ISDg10 and the USDA NRRL for Candida molischiana (Y-2237). We thank Patrick Hillery, Jessica Bigham and Taylor Maher who assisted in sample collection and analysis and Mark Signs who supported analytical work. We would like to thank Thomas K. Wood at The Pennsylvania State University for his comments on the research and manuscript. This material is based upon work done by TR Zuroff supported by the National Science Foundation Graduate Research Fellowship under Grant No. DGE-0750756. TR Zuroff also received funding for this work from the Pennsylvania State University Department of Chemical Engineering John and Jeanette McWhirter Graduate Research Fellowship. Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the National Science Foundation.
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