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Syntrophic metabolism of a co-culture containing Clostridium cellulolyticum and Rhodopseudomonas palustris for hydrogen production
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Syntrophic metabolism of a co-culture containing Clostridium cellulolyticum and Rhodopseudomonas palustris for hydrogen production Yongqin Jiao a,*, Ali Navid a, Benjamin J. Stewart a, James B. McKinlay b, Michael P. Thelen a, Jennifer Pett-Ridge a a b
Physical and Life Sciences Directorate, Lawrence Livermore National Laboratory, 7000 East Ave, L-452, Livermore, CA 94550, USA Department of Biology, Indiana University, Bloomington, IN, USA
article info
abstract
Article history:
Several studies have explored combining fermentative and purple bacteria to increase
Received 21 April 2012
hydrogen yields from carbohydrates, but the metabolic interaction between these organ-
Received in revised form
isms is poorly understood. In an artificial co-culture containing Clostridium cellulolyticum
18 May 2012
and Rhodopseudomonas palustris with cellulose as the sole carbon source, we examined cell
Accepted 19 May 2012
growth kinetics, cellulose consumption, H2 production, and carbon transfer from C. cellu-
Available online 16 June 2012
lolyticum to R. palustris. When cultured alone, C. cellulolyticum degraded only 73% of the supplied cellulose. However, in co-culture C. cellulolyticum degraded 100% of the total
Keywords:
cellulose added (5.5 g/L) and at twice the rate of C. cellulolyticum monocultures. Concur-
Biohydrogen
rently, the total H2 production by the co-culture was 1.6-times higher than that by the C.
Fermentation
cellulolyticum monoculture. Co-culturing also resulted in a 2-fold increase in the growth rate
Photosynthesis
of C. cellulolyticum and a 2.6-fold increase in final cell density. The major metabolites
Cellulose degradation
present in the co-culture medium include lactate, acetate and ethanol, with acetate serving
Syntrophy
as the primary metabolite transferring carbon from C. cellulolyticum to R. palustris. Our
Clostridia
results suggest that the stimulation of bacterial growth and cellulose consumption under the co-culture conditions is likely caused by R. palustris’ removal of inhibitory metabolic byproducts (i.e., pyruvate) generated during cellulose metabolism by C. cellulolyticum. Copyright ª 2012, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved.
1.
Introduction
Cellulolytic microorganisms, such as cellulolytic clostridia, can break down cellulosic materials and ferment the sugars to hydrogen gas (H2), a promising transportation fuel [1e3]. As such, using cellulolytic clostridia may serve an economically relevant role in H2 production in industrial digestion processes, particularly in anaerobic digesters fed with municipal waste or where agricultural raw materials containing a high percentage
of cellulosic materials are used as feedstock [4,5]. However, fermentative H2 yields are low because electrons are lost in the obligate excretion of organic acids and alcohols, indicating a relatively inefficient pathway of carbon flow by cellulolytic clostridia [1,6e8]. A solution to this obstacle could be coculturing of cellulolytic clostridia with photosynthetic purple bacteria that can consume fermentation products and produce H2 via nitrogenase during nitrogen fixation [9e11]. Most anoxygenic photosynthetic purple bacteria cannot consume
* Corresponding author. Tel.: þ1 925 422 4482; fax: þ1 925 422 2099. E-mail address: [email protected] (Y. Jiao). 0360-3199/$ e see front matter Copyright ª 2012, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved. 10.1016/j.ijhydene.2012.05.100
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2.
Materials and methods
palustris CGA676 (kindly provided by Dr. C. Harwood at the University of Washington, Seattle, WA). CGA676 is a NifA mutant strain derived from wild type R. palustris CGA009 [21] that contains a 48 nucleotide deletion in nifA resulting in constitutive nitrogenase activity and H2 production [23]. Both strains were cultured in a modified CM3 medium [24], including (per liter of deionized water) 1.0 g (NH4)2SO4, 1.3 g KH2PO4, 2.9 g K2HPO4$3H2O, 0.2 g MgCl2$6H2O, 0.075 g CaCl2, 1.0 g cysteineeHCl, 25 ml of FeSO4 stock (5% in 50 mM of H2SO4), 1 ml mineral solution, and 10 ml vitamin solution. The mineral solution contained (per liter of deionized water) 1.5 g FeCl2 CaCl2$4H2O, 70 mg ZnCl2, 100 mg MnCl2, CaCl2$4H2O, 36 mg H3BO3, 190 mg, CoCl2$6H2O, 2.0 mg CuCl2$2H2O, 24 mg NiCl2$6H2O and 36 mg Na2MoO4$2H2O. The vitamin solution contained (per liter) 100 mg d-biotin, 250 mg para-aminobenzoic acid, 150 mg nicotinic acid, 250 mg riboflavin, 250 mg pantothenic acid, 250 mg thiamine, and 100 mg cyanocobalamin. The vitamin solution was sterilized by filtration through a 0.2-mm-pore-size filter. The media were adjusted to pH 7.2 using NaOH. In an anaerobic chamber (Coy Laboratory Products, Inc., Grass Lake, MI, USA), 120 ml serum bottles (Wheaton Scientific, Millville, NJ, USA) were each filled with 100 ml anaerobic modified CM3 medium (without carbon source) and sealed with a butyl rubber stopper (Bellco Glass, Vineland, NJ, USA) and aluminum crimp-seal. CM3 filled bottles were purged with argon gas outside the chamber afterward to ensure the headspace was free of H2 and N2. Concentrated cellulose (microcrystalline, powder size 20 mm, SigmaeAldrich) and cellobiose stocks were prepared similarly in separate bottles. After autoclaving all the bottles for 30 min, the carbohydrate stock solutions of appropriate volume were transferred aseptically into culture bottles to give a final concentration of approximately 5 g/L. Monocultures of C. cellulolyticum and R. palustris as well as co-cultures were grown anaerobically at 32 C in serum bottles placed 30 cm from a 65 W incandescent lamp. Before each batch experiment, aliquots of frozen stocks of C. cellulolyticum H10 were cultured in modified CM3 medium containing cellobiose (5 g/L) and used as inoculum when the cultures reached late exponential growth conditions. Inoculum of R. palustris CGA676 was grown in modified CM3 medium with sodium acetate (10 mM) as the carbon source, and cells were washed 3 times in modified CM3 medium before inoculation. For the co-culture, bottles were inoculated with both of the respective pure cultures, adjusting the inoculum volume to introduce approximately 2 107 cells/ml for each strain. Controls without inoculum were also included for each substrate and without carbon source. All tests were run in triplicate with results averaged. For the fermentation production inhibition experiment with C. cellulolyticum, modified CM3 medium with 5.0 g/L of cellulose was supplemented with different concentrations of sodium lactate, sodium acetate, sodium pyruvate or ethanol. Medium pH was checked after additions and adjusted to 7.2 when necessary. Controls without added organic acids were included.
2.1.
Bacterial strains and culture conditions
2.2.
sugars but thrive and produce H2 when growing on another microbe’s fermentation waste products. Thus, purple bacteria are a natural choice co-culturing with cellulolytic clostridia to increase H2 production from carbohydrates. Previous work combining these bacterial metabolisms has focused on a two-stage process. In the first stage, fermentative bacteria are used to ferment carbohydrates to H2 and soluble products. The fermentative effluent is then transferred to a photobioreactor containing purple bacteria where the organic acids are further converted to H2 in the second stage [10e12]. However, efforts have also been made to form a consolidated bioprocess in which the same two groups of microbes, fermentative and purple bacteria, are co-cultured [13e18]. While modest H2 production has been reported, all but one of these studies used simple sugar feedstocks (such as glucose) rather than polymers such as cellulose. The one exception showed that purple bacteria could be used to produce H2 from cellulose by co-culturing with a cellulosedegrading bacterium, but in this case the cellulosedegrading bacterium itself did not produce H2 [19]. These studies showed that co-culturing can improve H2 production from sugars. However, there is still much to be understood about the key metabolic interactions and population dynamics in such co-cultures that are critical for improvement of syntrophic efficiency and H2 productivity [20]. Continued improvements in our ability to track metabolite fluxes will provide a more comprehensive picture of intermediary metabolism and the factors that control the flux of organic matter that result in H2 production. To obtain a better understanding of syntrophic metabolism, we examined the kinetics of cellular growth and carbon transfer in an artificial syntrophic co-culture containing Clostridium cellulolyticum H10 and purple bacterium Rhodopseudomonas palustris CGA676 [21e23] with the goal of increasing H2 yields from carbohydrates. Constitutive nitrogenase activity in CGA676 allows for H2 to be produced, even in media containing ammonia, which would normally inhibit H2 production by nitrogenase [23]. In this co-culture system, cellulose is the sole carbon and energy source for C. cellulolyticum. R. palustris utilizes the fermentation products secreted by C. cellulolyticum, and H2 is produced by both organisms. Because both organisms have been fully sequenced, their coculture is an ideal way to begin to understand the genomic underpinnings of syntrophic interactions. Both genomic information and an understanding of the physiology of the coculture could eventually lead to a robust metabolic model of the consortium, an important first step toward a systems biology understanding of microbial communities. We found that the presence of R. palustris in the co-culture greatly stimulated cellulose degradation and H2 production, which likely resulted from accelerated metabolism of C. cellulolyticum and consumption of acetate and pyruvate by R. palustris.
Two well-documented strains were used in this research: C. cellulolyticum H10 (ATCC# 35319, Manassas, VA, USA) [22] and R.
Analytical procedures
To microscopically monitor bacterial growth via cell counts, about 50 ml of culture solution was collected using a syringe. The
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solution was mixed with an equal volume of 4,6 diamidino-2phenylindole (DAPI, 10 mg/ml), vortexed vigorously, and filtered through Millipore membrane filter disks (0.22 mm in size, and 22 mm diameter). Dilutions were made before filtration when necessary. Cells in five different fields of view on the membrane were counted under a fluorescent microscope (Leica), and the total number of cells on the membrane was calculated based on area ratio. For the co-culture, the population of R. palustris was counted by its autofluorescence under a rhodamine filter without DAPI staining, and the accuracy of the counting was crosschecked with colony forming units (CFU) by plating onto YP agar plate (0.3% yeast extract and 0.3% bactopeptone (Difco) [25]) and incubated aerobically. The population of C. cellulolyticum H10 was derived by subtraction of the cell count of R. palustris from the total cell count. Fermentation products (acetate, lactate, pyruvate and ethanol) were quantified using high performance liquid chromatography (HPLC). Cell cultures were subsampled at multiple time points and filtered through a 0.2 mm membrane (Corning Costar Spin-X) and stored at 20 C. HPLC (Agilent 1100; Agilent Technologies, Santa Clara, CA, USA) was equipped with both a refractive index and a diode array detector. A fermentation column (Aminex HPX-87H Column, BioRad) was used for separation of organic acids and carbohydrates, with isocratic flow using 1 mM sulfuric acid as the mobile phase. Analysis conditions were as follows: injection volume, 20 mL; flow rate, 0.6 mL/ min; column thermostat, 60 C; and analysis time of 15 min. Acetate, lactate and pyruvate were measured with the diode array detector at 210 nm, and ethanol with the refractive index detector. Metabolites were identified by comparison of metabolite retention times with those of authentic standards, and metabolite concentrations were calculated using standard curves generated by measurement of the appropriate standards. The cellulose concentration in subsamples was determined as described by Huang and Forsberg [26]. About 500 ml of culture was withdrawn from the bottles, immediately after the bottles were shaken to achieve a homogenous suspension. Cells were lysed with the addition of 8% formic acid, and cellulose in samples was washed repeatedly (3 times) by sedimentation (10,000 g, 3 min) and suspension (by vigorous vortexing) in water. Sedimented cellulose was then solubilized in 67% (wt/vol) sulfuric acid as described by Updegraff [27] and quantified using the phenolesulfuric acid method for soluble carbohydrates [28]. Glucose was used as standard. Hydrogen gas in the bottle’s headspace was measured using a gas chromatograph (GC-2014, Greenhouse & Atmospheric Gases Analyzer, Shimadzu) equipped with a thermal conductivity detector (100 C, 150 mA), and Hayesep column (2.0 m, N 1/1600 80/100 mesh, set at 80 C) with argon as carrier gas (30 mL/min). For sampling, a gas tight syringe was used to withdraw 0.5 or 1.0 ml headspace gas, the volume was replaced in the bottle with high-purity argon gas. The amount of H2 removed in each sampling was considered when calculating total H2 production.
3.
Results
We developed a system for H2 production based on cellulose degradation using a co-culture of C. cellulolyticum H10 and R.
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palustris CGA676. These bacterial strains were chosen based on their compatibility of growth conditions. In addition, we determined that the metabolites released by C. cellulolyticum when grown on cellulose [29] can be used by R. palustris [21] under the conditions tested. R. palustris CGA676 was of particular interest because it is a NifA mutant strain with constitutive nitrogenase activity, allowing H2 to be produced in media containing ammonium, which normally prevents H2 production by nitrogenase [23].
3.1.
Cellulose degradation and H2 production
Compared with nearly complete consumption of cellulose (5.5 g/L) shortly after 10 days of incubation for the co-culture (Fig. 1), cellulose degradation was slower when C. cellulolyticum was cultured alone, with only about 50% (2.7 g/L) consumed during the same time period. Ultimately, cellulose was not depleted by the end of the incubation period in the C. cellulolyticum monoculture, with a final concentration of 1.5 g/ L remaining in the medium. The initial cellulose consumption rates doubled in the co-culture (50 mg cellulose/day)
Fig. 1 e Measurements of residual cellulose concentration in the medium and H2 produced over time when C. cellulolyticum was cultured alone on cellulose (A) versus cocultured with R. palustris (B). Error bars represent standard deviations from triplicate cultures.
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Fig. 2 e Comparison of cell growth when C. cellulolyticum and R. palustris were cultured individually on cellulose versus co-cultured together. The Y-axis is in logarithmic scale. (CC, diamonds) C. cellulolyticum in monoculture; (RP, squares), R. palustris in monoculture, (Co-CC, triangles) C. cellulolyticum in co-culture; (Co-RP, circles) R. palustris in coculture. Error bars represent standard deviations from triplicate cultures.
compared to that in the C. cellulolyticum monoculture (26 mg/ day). However, when we accounted for differences in cell growth under these conditions, the rates of cellulose consumption between these two cultures were quite similar, with 5 (2) 108 mg cellulose/day/cell in the co-culture, and 6 (2) 108 mg/day/cell in the C. cellulolyticum monoculture. No cellulose consumption was observed in abiotic controls. The total amount of H2 produced by the co-culture was 1.6times higher than that in the C. cellulolyticum monoculture (Fig. 1). From the 5.5 g of cellulose provided, C. cellulolyticum culture produced about 27 mmol of H2, whereas the co-culture
produced a total of 42 mmol of H2. However, when taking into account differences in the amount of cellulose consumed, the H2 yield was 1.2 0.2 mol of H2 per mole of glucose equivalent in the C. cellulolyticum monoculture, and 1.4 0.2 mol in the co-culture. No H2 was produced in abiotic controls or when cellulose was omitted. Besides total amount of H2 produced, the initial H2 production rate was significantly higher in the co-culture (3.5 mmol/day) compared to that in the C. cellulolyticum monoculture (2.5 mmol/day) (Fig. 1). In addition, the H2 production in the co-culture showed a biphasic pattern with a faster initial increase before day 10, followed by a much slower increase between day 10 and day 20. Since cellulose was already depleted at the later time period, the additional H2 produced was likely due to the metabolism of R. palustris, growing on metabolites secreted by C. cellulolyticum. In contrast, in the C. cellulolyticum monoculture, a relatively low amount of H2 was produced between day 10 and 20, fueled solely by a relatively slow rate of cellulose consumption.
3.2.
Population dynamics
Along with faster rates of cellulose consumption and H2 production, C. cellulolyticum also grew faster in co-cultures (Fig. 2) and reached a higher cell density at the end of the 25-day incubation period. In C. cellulolyticum monocultures, the cells grew with a 36 h doubling time and bacterial growth ceased after 16 days (at about w3 108 cells/ml) when there was still about 1.5 g/L cellulose left, indicating that the early growth arrest is not caused by substrate depletion. In contrast, in the co-culture, C. cellulolyticum grew twice as fast (16 h doubling time) and the final cell density was 3 times higher at 1 109 cells/ml. The C. cellulolyticum growth yield in monoculture was 0.7 1011 cells per gram of cellulose but in coculture it was 2.6-times higher at 1.8 1011 cells per gram of cellulose. Thus, growing together with R. palustris has
Fig. 3 e Extracellular metabolite concentrations as measured by HPLC of subsamples from C. cellulolyticum monocultures and co-cultures with R. palustris. (CC, diamonds) C. cellulolyticum monoculture; (Co-culture, squares) co-culture containing C. cellulolyticum and R. palustris. Error bars represent standard deviations from triplicate cultures.
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Fig. 4 e Comparison of growth media pH from C. cellulolyticum monoculture and co-culture with R. palustris. (CC, diamonds) C. cellulolyticum monoculture; (Co-culture, squares) co-culture containing C. cellulolyticum and R. palustris. Error bars represent standard deviations from triplicate cultures.
a profound effect on the energetic efficiency of the C. cellulolyticum fermentation. R. palustris alone cannot grow on cellulose, whereas it showed steady growth in the co-culture reaching 5 109 cells/ml (Fig. 2). No growth was observed when no cellulose was added in the C. cellulolyticum monoculture or the co-culture.
3.3.
Metabolite transfer
In order to determine how co-culturing with R. palustris boosted cellulose degradation by C. cellulolyticum, we examined the fermentation products released into the medium when C. cellulolyticum was cultured alone versus under coculture conditions. The fermentation of cellulose by C.
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cellulolyticum in both monoculture and co-culture generated various soluble extracellular products: lactate, acetate, ethanol and pyruvate were the major compounds detected by HPLC (Fig. 3). These fermentation products could serve as growth substrates for R. palustris. Under both culture conditions, acetate, lactate, and ethanol were produced at concentrations an order of magnitude higher than pyruvate, which was only produced at sub-millimolar concentrations. All four compounds were produced sooner and/or at higher rates in the co-culture than the C. cellulolyticum monoculture. After approximately 10 days of co-culturing, significant amounts of acetate and pyruvate were removed as the incubation continued, suggesting that they were utilized by R. palustris. In comparison to pyruvate, there was approximately 50-fold more acetate removed, indicating that acetate was the major metabolite for carbon transfer between the two organisms in the co-culture.
3.4.
pH effects
Because it is well known that the release of fermentation products by fermentative bacteria can acidify the medium and slow cellular growth, we monitored the medium pH throughout the incubation period (Fig. 4). When C. cellulolyticum was cultured alone, the medium pH decreased from 7.2 to w5.5, during the initial 15 days of culturing in which cellulose was also depleted. The co-culture pH decrease was initially faster than the monoculture, but leveled off at w5.9 before the cellulose was depleted at day 8. Notably, this pH stabilization coincided with the onset of acetate consumption by R. palustris, suggesting that the removal of acetic acid helped stabilize the co-culture pH. There was no pH change throughout the incubation period in abiotic controls. To test whether the pH stabilization alone resulted in higher cellulose consumption (or rate of degradation) by C. cellulolyticum, we tested the effect of adding Na-HEPES (100 mM, pH 7.2) in the modified CM3 medium. Despite this extra buffering capacity there was still 1e1.5 g of residual cellulose at the end of incubation (data not shown).
Fig. 5 e The inhibition effect of four intermediate metabolites, at multiple concentrations, on cellulose-based growth of C. cellulolyticum. Growth of C. cellulolyticum on cellulose in the presence of test compounds is given in proportion to the control culture lacking additional compounds. In the control where C. cellulolyticum was grown in monoculture on cellulose in the absence of added organic acids or ethanol (the bar at far left, “cellulose alone”), the final cell density was w3 3 108 cells/ml, set at 100%. Error bars represent standard deviations from triplicate cultures.
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3.5.
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Growth inhibition of C. cellulolyticum by pyruvate
Because R. palustris is thought to grow on the fermentation products secreted by C. cellulolyticum, we were curious if the removal of these byproducts by R. palustris has a positive feedback on C. cellulolyticum growth. Accordingly, we tested the dominant intermediate metabolites for inhibitory effects on C. cellulolyticum growth on cellulose. In the presence of 25 mM acetate, lactate or ethanol, the final cell count was 73e82% of that in the control cultures with no additions (Fig. 5). In contrast, 25 mM pyruvate completely eliminated C. cellulolyticum growth (about 0.5% of the control). Due to the severe effect of pyruvate at 25 mM, we conducted further tests for pyruvate inhibition at lower concentrations and found that inhibition of C. cellulolyticum growth appeared linearly correlated with pyruvate concentration (Fig. 5). At 0.5 mM, a concentration similar to what C. cellulolyticum produces in co-cultures, C. cellulolyticum growth was 83% of the control culture.
4.
Discussion and conclusions
Inter-species interactions in microbial communities play an important role in community fitness, and are an area of intense interest in both ecological and industrial research. It is well established that the majority of microbes in nature survive best in communities, most likely due to the partitioning of beneficial functions by individual microbes. Artificial communities, such as a co-culture of two or more microorganisms, are interesting to study because these interactions can be modeled using genomic and proteomic information, toward a comprehensive understanding of how microbial communities operate as a whole. In this study, we examined the physiology and carbon transfer in an artificial syntrophic co-culture containing C. cellulolyticum and R. palustris for cellulose degradation and H2 production. While the cellulose degrader C. cellulolyticum used in this study has low cellulolytic ability and cannot be used industrially without strain improvement [1,5], the focus of our study is for basic research in examining metabolic interactions in a syntrophic co-culture system. We found that the presence R. palustris in the co-culture stimulated C. cellulolyticum growth, cellulose degradation, and total H2 production. The higher H2 productivity in co-culture could be due to two reasons. First, R. palustris can utilize several of the fermentation products excreted by C. cellulolyticum to produce H2. In other words, some of electrons that would otherwise be lost in the obligate excretion of organic acids and alcohols in C. cellulolyticum monoculture could be further converted into H2 by R. palustris in the co-culture [4]. Second, the more complete consumption of cellulose under co-culture conditions should also contribute to the additional H2 produced, by allowing for H2 production by C. cellulolyticum to continue. This is consistent with our finding that the C. cellulolyticum monoculture produced about 27 mmol of H2, whereas the co-culture produced a total of 42 mmol of H2. However, the H2 yield was 1.2 0.2 mol of H2 per mole of glucose equivalent in the C. cellulolyticum monoculture, and 1.4 0.2 mol in the co-culture. Thus, while H2 production by R. palustris contributed to a minor increase in the overall H2 yield, it appears that most of
the additional H2 produced by the co-cultures was due to the more efficient utilization of cellulose. As a result, the main benefit that R. palustris appears to impart is an improvement of C. cellulolyticum metabolic efficiency. The total H2 yields in both the monoculture of C. cellulolyticum and the co-culture are lower than values previously reported for a co-culture consisting of Ethanoligenens harbinense and Rhodopseudomonas faecalis growing on glucose, where 1.8 mol H2 per mole of glucose were produced in single E. harbinense culture and 2.5 mol in the co-culture under similar conditions [15]. The lower H2 production in our system is likely due to the fact that additional energy input is required for cellulose breakdown, a requirement that was circumvented in the previous study [15] where glucose was provided as the carbon source. The examination of population dynamics of each species, the pH monitoring over the growth period, and the quantitative measurement of the intermediate metabolites were important to help make meaningful conclusions about the physiology of the co-culture. The stimulation of C. cellulolyticum growth in the co-culture could be partially due to enhanced buffering of the medium caused by consumption of organic acids by R. palustris. The pH of the co-culture ceased to decrease at w5.9 after 8 days of incubation when acetate in the medium reached maximum concentration (w15 mM). In the C. cellulolyticum monoculture, the pH continued to drop to w5.5 until cellular growth stopped. Nevertheless, the pH effect alone does not account for the growth enhancement of C. cellulolyticum on cellulose in the co-culture, because additional buffer added to the growth medium did not change the amount of cellulose consumed significantly. Therefore, acetate appeared to be the major carbon transfer metabolite between C. cellulolyticum and R. palustris, and acetate removal likely also contributed to growth stimulation of C. cellulolyticum in the co-culture and improved the thermodynamics of the fermentation. Additionally, the extra buffering of the environment provided by the metabolism of R. palustris consuming acetate likely meant that C. cellulolyticum could devote less ATP to maintaining a proper membrane potential and other maintenance processes. While the reduced acetate and pyruvate in the co-culture is likely to be the direct result of carbon transfer, the possibility that cellulolytic clostridia produce less intermediate metabolites in co-culture cannot be excluded. This is an inherited limitation of working with co-culture systems. However, from a mass balance prospective, R. palustris cell numbers increased by two orders of magnitude in the co-culture, therefore carbon was consumed, mostly likely according to the established substrate preference for R. palustris [9,10]. We compared the growth rate of R. palustris on acetate, pyruvate, ethanol and lactate, and found that acetate is the most favorable substrate (data not shown). Although media concentrations of pyruvate were much lower than the other fermentation products, its removal (ostensibly by R. palustris) likely plays an important role in alleviating product inhibition and boosting C. cellulolyticum growth under co-culture conditions. Previous studies have indicated that growth inhibition of C. cellulolyticum is caused by an inefficiently regulated carbon flow [6]. It has been shown that some intermediate products (such as NADH and
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pyruvate) of this metabolism accumulate intracellularly [6,30]. The release of pyruvate may represent an indicator of poorly regulated carbon flow, and the accumulation of the extracellular pyruvate suggests the onset of slowdown of glycolysis. Supporting these previous observations, we found that the highest levels of inhibition occurred when we tested for C. cellulolyticum growth at pyruvate concentrations equivalent to those of other dominant metabolites found in the media (Fig. 5). Since pyruvate was produced by C. cellulolyticum when growing on cellulose, the intracellular concentration is likely to be higher than the extracellular concentration. Conversely, when pyruvate is exogenously added, as in the inhibition experiments (Fig. 5), it likely takes a higher concentration to achieve a similar internal concentration leading to growth inhibition. At co-culture relevant concentrations (w0.5 mM), pyruvate addition caused a 17% reduction of C. cellulolyticum growth. While more study is needed to elucidate the mechanism of this inhibitory effect, our results suggest that pyruvate, albeit at low concentration, acted as an inhibitory compound for cell growth in C. cellulolyticum monocultures. The removal of pyruvate from the co-culture by R. palustris likely alleviated the inhibition effect, therefore boosting the cellular growth of C. cellulolyticum as well as cellulose consumption. In addition, acetate consumption by R. palustris could potentially increase the efficiency of pyruvate to acetate conversion by C. cellulolyticum, permitting it to re-consume and ferment the pyruvate. To our knowledge, this study is the first to use two-H2 producing microorganisms to produce H2 from cellulose degradation in a photosynthetic co-culture system [14,16]. The direct coupling of the metabolisms of the two species has implications for research in the ecology of microbial communities in natural and engineered systems. A systemlevel understanding of the interplay of multiple metabolic pathways relevant to carbon and energy flow will be useful for understanding the population dynamics and nutrient transfer within mixed communities, as well as in strategic bioengineering of microbes to improve overall community fitness and H2 production.
Acknowledgments This work was supported by the Genomic Science Program of the U.S. Department of Energy’s Office of Biological and Environmental Research under contract SCW1039, as part of the LLNL Biofuels Scientific Focus Area. Lawrence Livermore National Laboratory is operated by Lawrence Livermore National Security, LLC, for the U.S. Department of Energy, National Nuclear Security Administration under Contract DEAC52-07NA27344.
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[2] Desvaux M. Clostridium cellulolyticum: model organism of mesophilic cellulolytic clostridia. FEMS Microbiology Reviews 2005;29:741e64. [3] Leschine SB. Cellulose degradation in anaerobic environments. Annual Review of Microbiology 1995;49: 399e426. [4] Melis A, Melnicki MR. Integrated biological hydrogen production. International Journal of Hydrogen Energy 2006; 31:1563e73. [5] Lynd LR, Weimer PJ, van Zyl WH, Pretorius IS. Microbial cellulose utilization: fundamentals and biotechnology. Microbiology and Molecular Biology Reviews 2002;66:506. [6] Guedon E, Desvaux M, Payot S, Petitdemange H. Growth inhibition of Clostridium cellulolyticum by an inefficiently regulated carbon flow. Microbiology 1999;145(Pt 8):1831e8. [7] Giallo J, Gaudin C, Belaich JP. Metabolism and solubilization of cellulose by Clostridium cellulolyticum H10. Applied and Environmental Microbiology 1985;49:1216e21. [8] Desvaux M, Guedon E, Petitdemange H. Cellulose catabolism by Clostridium cellulolyticum growing in batch culture on defined medium. Applied and Environmental Microbiology 2000;66:2461e70. [9] Barbosa MJ, Rocha JMS, Tramper J, Wijffels RH. Acetate as a carbon source for hydrogen production by photosynthetic bacteria. Journal of Biotechnology 2001;85:25e33. [10] Chen CY, Lu WB, Liu CH, Chang JS. Improved phototrophic H2 production with Rhodopseudomonas palustris WP3-5 using acetate and butyrate as dual carbon substrates. Bioresource Technology 2008;99:3609e16. [11] Lee CM, Chen PC, Wang CC, Tung YC. Photohydrogen production using purple nonsulfur bacteria with hydrogen fermentation reactor effluent. International Journal of Hydrogen Energy 2002;27:1309e13. [12] Koku H, Eroglu I, Gunduz U, Yucel M, Turker L. Aspects of the metabolism of hydrogen production by Rhodobacter sphaeroides. International Journal of Hydrogen Energy 2002; 27:1315e29. [13] Vatsala TM, Raj SM, Manimaran A. A pilot-scale study of biohydrogen production from distillery effluent using defined bacterial co-culture. International Journal of Hydrogen Energy 2008;33:5404e15. [14] Ding J, Liu BF, Ren NQ, Xing DF, Guo WQ, Xu JF, et al. Hydrogen production from glucose by co-culture of Clostridium butyricum and immobilized Rhodopseudomonas faecalis RLD-53. International Journal of Hydrogen Energy 2009;34:3647e52. [15] Xie GJ, Feng LB, Ren NQ, Ding J, Liu C, Xing DF, et al. Control strategies for hydrogen production through co-culture of Ethanoligenens harbinense B49 and immobilized Rhodopseudomonas faecalis RLD-53. International Journal of Hydrogen Energy 2010;35:1929e35. [16] Sun Q, Xiao W, Xi D, Shi JP, Yan X, Zhou ZH. Statistical optimization of biohydrogen production from sucrose by a co-culture of Clostridium acidisoli and Rhodobacter sphaeroides. International Journal of Hydrogen Energy 2010; 35:4076e84. [17] Zhu HG, Wakayama T, Asada Y, Miyake J. Hydrogen production by four cultures with participation by anoxygenic phototrophic bacterium and anaerobic bacterium in the presence of NH4þ. International Journal of Hydrogen Energy 2001;26:1149e54. [18] Fang HHP, Zhu HG, Zhang T. Phototrophic hydrogen production from glucose by pure and co-cultures of Clostridium butyricum and Rhodobacter sphaeroides. International Journal of Hydrogen Energy 2006;31:2223e30. [19] Odom JM, Wall JD. Photoproduction of h(2) from cellulose by an anaerobic bacterial coculture. Applied and Environmental Microbiology 1983;45:1300e5.
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[20] Salimi F, Zhuang K, Mahadevan R. Genome-scale metabolic modeling of a clostridial co-culture for consolidated bioprocessing. Biotechnology Journal 2010;5:726e38. [21] Larimer FW, Chain P, Hauser L, Lamerdin J, Malfatti S, Do L, et al. Complete genome sequence of the metabolically versatile photosynthetic bacterium Rhodopseudomonas palustris. Nature Biotechnology 2004;22: 55e61. [22] Petitdemange E, Caillet F, Giallo J, Gaudin C. Clostridiumcellulolyticum sp-nov, a cellulolytic, mesophilic species from decayed grass. International Journal of Systematic Bacteriology 1984;34:155e9. [23] McKinlay JB, Harwood CS. Carbon dioxide fixation as a central redox cofactor recycling mechanism in bacteria. Proceedings of the National Academy of Sciences of the United States of America 2010;107:11669e75. [24] Weimer PJ, Zeikus JG. Fermentation of cellulose and cellobiose by Clostridium thermocellum in the absence of Methanobacterium thermoautotrophicum. Applied and Environmental Microbiology 1977;33:289e97.
[25] Weaver PF, Wall JD, Gest H. Characterization of Rhodopseudomonas capsulata. Archives of Microbiology 1975; 105:207e16. [26] Huang L, Forsberg CW. Cellulose digestion and cellulase regulation and distribution in Fibrobacter succinogenes subsp. succinogenes S85. Applied and Environmental Microbiology 1990;56:1221e8. [27] Updegraff DM. Semimicro determination of cellulose in biological materials. Analytical Biochemistry 1969;32:420. [28] Dubois M, Gilles KA, Hamilton JK, Rebers PA, Smith F. Colorimetric method for determination of sugars and related substances. Analytical Chemistry 1956;28:350e6. [29] Ren Z, Ward TE, Logan BE, Regan JM. Characterization of the cellulolytic and hydrogen-producing activities of six mesophilic Clostridium species. Journal of Applied Microbiology 2007;103:2258e66. [30] Guedon E, Payot S, Desvaux M, Petitdemange H. Carbon and electron flow in Clostridium cellulolyticum grown in chemostat culture on synthetic medium. Journal of Bacteriology 1999; 181:3262e9.
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Syntrophic metabolism of a co-culture containing Clostridium cellulolyticum and Rhodopseudomonas palustris for hydrogen production Yongqin Jiao a,*, Ali Navid a, Benjamin J. Stewart a, James B. McKinlay b, Michael P. Thelen a, Jennifer Pett-Ridge a a b
Physical and Life Sciences Directorate, Lawrence Livermore National Laboratory, 7000 East Ave, L-452, Livermore, CA 94550, USA Department of Biology, Indiana University, Bloomington, IN, USA
article info
abstract
Article history:
Several studies have explored combining fermentative and purple bacteria to increase
Received 21 April 2012
hydrogen yields from carbohydrates, but the metabolic interaction between these organ-
Received in revised form
isms is poorly understood. In an artificial co-culture containing Clostridium cellulolyticum
18 May 2012
and Rhodopseudomonas palustris with cellulose as the sole carbon source, we examined cell
Accepted 19 May 2012
growth kinetics, cellulose consumption, H2 production, and carbon transfer from C. cellu-
Available online 16 June 2012
lolyticum to R. palustris. When cultured alone, C. cellulolyticum degraded only 73% of the supplied cellulose. However, in co-culture C. cellulolyticum degraded 100% of the total
Keywords:
cellulose added (5.5 g/L) and at twice the rate of C. cellulolyticum monocultures. Concur-
Biohydrogen
rently, the total H2 production by the co-culture was 1.6-times higher than that by the C.
Fermentation
cellulolyticum monoculture. Co-culturing also resulted in a 2-fold increase in the growth rate
Photosynthesis
of C. cellulolyticum and a 2.6-fold increase in final cell density. The major metabolites
Cellulose degradation
present in the co-culture medium include lactate, acetate and ethanol, with acetate serving
Syntrophy
as the primary metabolite transferring carbon from C. cellulolyticum to R. palustris. Our
Clostridia
results suggest that the stimulation of bacterial growth and cellulose consumption under the co-culture conditions is likely caused by R. palustris’ removal of inhibitory metabolic byproducts (i.e., pyruvate) generated during cellulose metabolism by C. cellulolyticum. Copyright ª 2012, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved.
1.
Introduction
Cellulolytic microorganisms, such as cellulolytic clostridia, can break down cellulosic materials and ferment the sugars to hydrogen gas (H2), a promising transportation fuel [1e3]. As such, using cellulolytic clostridia may serve an economically relevant role in H2 production in industrial digestion processes, particularly in anaerobic digesters fed with municipal waste or where agricultural raw materials containing a high percentage
of cellulosic materials are used as feedstock [4,5]. However, fermentative H2 yields are low because electrons are lost in the obligate excretion of organic acids and alcohols, indicating a relatively inefficient pathway of carbon flow by cellulolytic clostridia [1,6e8]. A solution to this obstacle could be coculturing of cellulolytic clostridia with photosynthetic purple bacteria that can consume fermentation products and produce H2 via nitrogenase during nitrogen fixation [9e11]. Most anoxygenic photosynthetic purple bacteria cannot consume
* Corresponding author. Tel.: þ1 925 422 4482; fax: þ1 925 422 2099. E-mail address: [email protected] (Y. Jiao). 0360-3199/$ e see front matter Copyright ª 2012, Hydrogen Energy Publications, LLC. Published by Elsevier Ltd. All rights reserved. 10.1016/j.ijhydene.2012.05.100
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2.
Materials and methods
palustris CGA676 (kindly provided by Dr. C. Harwood at the University of Washington, Seattle, WA). CGA676 is a NifA mutant strain derived from wild type R. palustris CGA009 [21] that contains a 48 nucleotide deletion in nifA resulting in constitutive nitrogenase activity and H2 production [23]. Both strains were cultured in a modified CM3 medium [24], including (per liter of deionized water) 1.0 g (NH4)2SO4, 1.3 g KH2PO4, 2.9 g K2HPO4$3H2O, 0.2 g MgCl2$6H2O, 0.075 g CaCl2, 1.0 g cysteineeHCl, 25 ml of FeSO4 stock (5% in 50 mM of H2SO4), 1 ml mineral solution, and 10 ml vitamin solution. The mineral solution contained (per liter of deionized water) 1.5 g FeCl2 CaCl2$4H2O, 70 mg ZnCl2, 100 mg MnCl2, CaCl2$4H2O, 36 mg H3BO3, 190 mg, CoCl2$6H2O, 2.0 mg CuCl2$2H2O, 24 mg NiCl2$6H2O and 36 mg Na2MoO4$2H2O. The vitamin solution contained (per liter) 100 mg d-biotin, 250 mg para-aminobenzoic acid, 150 mg nicotinic acid, 250 mg riboflavin, 250 mg pantothenic acid, 250 mg thiamine, and 100 mg cyanocobalamin. The vitamin solution was sterilized by filtration through a 0.2-mm-pore-size filter. The media were adjusted to pH 7.2 using NaOH. In an anaerobic chamber (Coy Laboratory Products, Inc., Grass Lake, MI, USA), 120 ml serum bottles (Wheaton Scientific, Millville, NJ, USA) were each filled with 100 ml anaerobic modified CM3 medium (without carbon source) and sealed with a butyl rubber stopper (Bellco Glass, Vineland, NJ, USA) and aluminum crimp-seal. CM3 filled bottles were purged with argon gas outside the chamber afterward to ensure the headspace was free of H2 and N2. Concentrated cellulose (microcrystalline, powder size 20 mm, SigmaeAldrich) and cellobiose stocks were prepared similarly in separate bottles. After autoclaving all the bottles for 30 min, the carbohydrate stock solutions of appropriate volume were transferred aseptically into culture bottles to give a final concentration of approximately 5 g/L. Monocultures of C. cellulolyticum and R. palustris as well as co-cultures were grown anaerobically at 32 C in serum bottles placed 30 cm from a 65 W incandescent lamp. Before each batch experiment, aliquots of frozen stocks of C. cellulolyticum H10 were cultured in modified CM3 medium containing cellobiose (5 g/L) and used as inoculum when the cultures reached late exponential growth conditions. Inoculum of R. palustris CGA676 was grown in modified CM3 medium with sodium acetate (10 mM) as the carbon source, and cells were washed 3 times in modified CM3 medium before inoculation. For the co-culture, bottles were inoculated with both of the respective pure cultures, adjusting the inoculum volume to introduce approximately 2 107 cells/ml for each strain. Controls without inoculum were also included for each substrate and without carbon source. All tests were run in triplicate with results averaged. For the fermentation production inhibition experiment with C. cellulolyticum, modified CM3 medium with 5.0 g/L of cellulose was supplemented with different concentrations of sodium lactate, sodium acetate, sodium pyruvate or ethanol. Medium pH was checked after additions and adjusted to 7.2 when necessary. Controls without added organic acids were included.
2.1.
Bacterial strains and culture conditions
2.2.
sugars but thrive and produce H2 when growing on another microbe’s fermentation waste products. Thus, purple bacteria are a natural choice co-culturing with cellulolytic clostridia to increase H2 production from carbohydrates. Previous work combining these bacterial metabolisms has focused on a two-stage process. In the first stage, fermentative bacteria are used to ferment carbohydrates to H2 and soluble products. The fermentative effluent is then transferred to a photobioreactor containing purple bacteria where the organic acids are further converted to H2 in the second stage [10e12]. However, efforts have also been made to form a consolidated bioprocess in which the same two groups of microbes, fermentative and purple bacteria, are co-cultured [13e18]. While modest H2 production has been reported, all but one of these studies used simple sugar feedstocks (such as glucose) rather than polymers such as cellulose. The one exception showed that purple bacteria could be used to produce H2 from cellulose by co-culturing with a cellulosedegrading bacterium, but in this case the cellulosedegrading bacterium itself did not produce H2 [19]. These studies showed that co-culturing can improve H2 production from sugars. However, there is still much to be understood about the key metabolic interactions and population dynamics in such co-cultures that are critical for improvement of syntrophic efficiency and H2 productivity [20]. Continued improvements in our ability to track metabolite fluxes will provide a more comprehensive picture of intermediary metabolism and the factors that control the flux of organic matter that result in H2 production. To obtain a better understanding of syntrophic metabolism, we examined the kinetics of cellular growth and carbon transfer in an artificial syntrophic co-culture containing Clostridium cellulolyticum H10 and purple bacterium Rhodopseudomonas palustris CGA676 [21e23] with the goal of increasing H2 yields from carbohydrates. Constitutive nitrogenase activity in CGA676 allows for H2 to be produced, even in media containing ammonia, which would normally inhibit H2 production by nitrogenase [23]. In this co-culture system, cellulose is the sole carbon and energy source for C. cellulolyticum. R. palustris utilizes the fermentation products secreted by C. cellulolyticum, and H2 is produced by both organisms. Because both organisms have been fully sequenced, their coculture is an ideal way to begin to understand the genomic underpinnings of syntrophic interactions. Both genomic information and an understanding of the physiology of the coculture could eventually lead to a robust metabolic model of the consortium, an important first step toward a systems biology understanding of microbial communities. We found that the presence of R. palustris in the co-culture greatly stimulated cellulose degradation and H2 production, which likely resulted from accelerated metabolism of C. cellulolyticum and consumption of acetate and pyruvate by R. palustris.
Two well-documented strains were used in this research: C. cellulolyticum H10 (ATCC# 35319, Manassas, VA, USA) [22] and R.
Analytical procedures
To microscopically monitor bacterial growth via cell counts, about 50 ml of culture solution was collected using a syringe. The
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solution was mixed with an equal volume of 4,6 diamidino-2phenylindole (DAPI, 10 mg/ml), vortexed vigorously, and filtered through Millipore membrane filter disks (0.22 mm in size, and 22 mm diameter). Dilutions were made before filtration when necessary. Cells in five different fields of view on the membrane were counted under a fluorescent microscope (Leica), and the total number of cells on the membrane was calculated based on area ratio. For the co-culture, the population of R. palustris was counted by its autofluorescence under a rhodamine filter without DAPI staining, and the accuracy of the counting was crosschecked with colony forming units (CFU) by plating onto YP agar plate (0.3% yeast extract and 0.3% bactopeptone (Difco) [25]) and incubated aerobically. The population of C. cellulolyticum H10 was derived by subtraction of the cell count of R. palustris from the total cell count. Fermentation products (acetate, lactate, pyruvate and ethanol) were quantified using high performance liquid chromatography (HPLC). Cell cultures were subsampled at multiple time points and filtered through a 0.2 mm membrane (Corning Costar Spin-X) and stored at 20 C. HPLC (Agilent 1100; Agilent Technologies, Santa Clara, CA, USA) was equipped with both a refractive index and a diode array detector. A fermentation column (Aminex HPX-87H Column, BioRad) was used for separation of organic acids and carbohydrates, with isocratic flow using 1 mM sulfuric acid as the mobile phase. Analysis conditions were as follows: injection volume, 20 mL; flow rate, 0.6 mL/ min; column thermostat, 60 C; and analysis time of 15 min. Acetate, lactate and pyruvate were measured with the diode array detector at 210 nm, and ethanol with the refractive index detector. Metabolites were identified by comparison of metabolite retention times with those of authentic standards, and metabolite concentrations were calculated using standard curves generated by measurement of the appropriate standards. The cellulose concentration in subsamples was determined as described by Huang and Forsberg [26]. About 500 ml of culture was withdrawn from the bottles, immediately after the bottles were shaken to achieve a homogenous suspension. Cells were lysed with the addition of 8% formic acid, and cellulose in samples was washed repeatedly (3 times) by sedimentation (10,000 g, 3 min) and suspension (by vigorous vortexing) in water. Sedimented cellulose was then solubilized in 67% (wt/vol) sulfuric acid as described by Updegraff [27] and quantified using the phenolesulfuric acid method for soluble carbohydrates [28]. Glucose was used as standard. Hydrogen gas in the bottle’s headspace was measured using a gas chromatograph (GC-2014, Greenhouse & Atmospheric Gases Analyzer, Shimadzu) equipped with a thermal conductivity detector (100 C, 150 mA), and Hayesep column (2.0 m, N 1/1600 80/100 mesh, set at 80 C) with argon as carrier gas (30 mL/min). For sampling, a gas tight syringe was used to withdraw 0.5 or 1.0 ml headspace gas, the volume was replaced in the bottle with high-purity argon gas. The amount of H2 removed in each sampling was considered when calculating total H2 production.
3.
Results
We developed a system for H2 production based on cellulose degradation using a co-culture of C. cellulolyticum H10 and R.
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palustris CGA676. These bacterial strains were chosen based on their compatibility of growth conditions. In addition, we determined that the metabolites released by C. cellulolyticum when grown on cellulose [29] can be used by R. palustris [21] under the conditions tested. R. palustris CGA676 was of particular interest because it is a NifA mutant strain with constitutive nitrogenase activity, allowing H2 to be produced in media containing ammonium, which normally prevents H2 production by nitrogenase [23].
3.1.
Cellulose degradation and H2 production
Compared with nearly complete consumption of cellulose (5.5 g/L) shortly after 10 days of incubation for the co-culture (Fig. 1), cellulose degradation was slower when C. cellulolyticum was cultured alone, with only about 50% (2.7 g/L) consumed during the same time period. Ultimately, cellulose was not depleted by the end of the incubation period in the C. cellulolyticum monoculture, with a final concentration of 1.5 g/ L remaining in the medium. The initial cellulose consumption rates doubled in the co-culture (50 mg cellulose/day)
Fig. 1 e Measurements of residual cellulose concentration in the medium and H2 produced over time when C. cellulolyticum was cultured alone on cellulose (A) versus cocultured with R. palustris (B). Error bars represent standard deviations from triplicate cultures.
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Fig. 2 e Comparison of cell growth when C. cellulolyticum and R. palustris were cultured individually on cellulose versus co-cultured together. The Y-axis is in logarithmic scale. (CC, diamonds) C. cellulolyticum in monoculture; (RP, squares), R. palustris in monoculture, (Co-CC, triangles) C. cellulolyticum in co-culture; (Co-RP, circles) R. palustris in coculture. Error bars represent standard deviations from triplicate cultures.
compared to that in the C. cellulolyticum monoculture (26 mg/ day). However, when we accounted for differences in cell growth under these conditions, the rates of cellulose consumption between these two cultures were quite similar, with 5 (2) 108 mg cellulose/day/cell in the co-culture, and 6 (2) 108 mg/day/cell in the C. cellulolyticum monoculture. No cellulose consumption was observed in abiotic controls. The total amount of H2 produced by the co-culture was 1.6times higher than that in the C. cellulolyticum monoculture (Fig. 1). From the 5.5 g of cellulose provided, C. cellulolyticum culture produced about 27 mmol of H2, whereas the co-culture
produced a total of 42 mmol of H2. However, when taking into account differences in the amount of cellulose consumed, the H2 yield was 1.2 0.2 mol of H2 per mole of glucose equivalent in the C. cellulolyticum monoculture, and 1.4 0.2 mol in the co-culture. No H2 was produced in abiotic controls or when cellulose was omitted. Besides total amount of H2 produced, the initial H2 production rate was significantly higher in the co-culture (3.5 mmol/day) compared to that in the C. cellulolyticum monoculture (2.5 mmol/day) (Fig. 1). In addition, the H2 production in the co-culture showed a biphasic pattern with a faster initial increase before day 10, followed by a much slower increase between day 10 and day 20. Since cellulose was already depleted at the later time period, the additional H2 produced was likely due to the metabolism of R. palustris, growing on metabolites secreted by C. cellulolyticum. In contrast, in the C. cellulolyticum monoculture, a relatively low amount of H2 was produced between day 10 and 20, fueled solely by a relatively slow rate of cellulose consumption.
3.2.
Population dynamics
Along with faster rates of cellulose consumption and H2 production, C. cellulolyticum also grew faster in co-cultures (Fig. 2) and reached a higher cell density at the end of the 25-day incubation period. In C. cellulolyticum monocultures, the cells grew with a 36 h doubling time and bacterial growth ceased after 16 days (at about w3 108 cells/ml) when there was still about 1.5 g/L cellulose left, indicating that the early growth arrest is not caused by substrate depletion. In contrast, in the co-culture, C. cellulolyticum grew twice as fast (16 h doubling time) and the final cell density was 3 times higher at 1 109 cells/ml. The C. cellulolyticum growth yield in monoculture was 0.7 1011 cells per gram of cellulose but in coculture it was 2.6-times higher at 1.8 1011 cells per gram of cellulose. Thus, growing together with R. palustris has
Fig. 3 e Extracellular metabolite concentrations as measured by HPLC of subsamples from C. cellulolyticum monocultures and co-cultures with R. palustris. (CC, diamonds) C. cellulolyticum monoculture; (Co-culture, squares) co-culture containing C. cellulolyticum and R. palustris. Error bars represent standard deviations from triplicate cultures.
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Fig. 4 e Comparison of growth media pH from C. cellulolyticum monoculture and co-culture with R. palustris. (CC, diamonds) C. cellulolyticum monoculture; (Co-culture, squares) co-culture containing C. cellulolyticum and R. palustris. Error bars represent standard deviations from triplicate cultures.
a profound effect on the energetic efficiency of the C. cellulolyticum fermentation. R. palustris alone cannot grow on cellulose, whereas it showed steady growth in the co-culture reaching 5 109 cells/ml (Fig. 2). No growth was observed when no cellulose was added in the C. cellulolyticum monoculture or the co-culture.
3.3.
Metabolite transfer
In order to determine how co-culturing with R. palustris boosted cellulose degradation by C. cellulolyticum, we examined the fermentation products released into the medium when C. cellulolyticum was cultured alone versus under coculture conditions. The fermentation of cellulose by C.
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cellulolyticum in both monoculture and co-culture generated various soluble extracellular products: lactate, acetate, ethanol and pyruvate were the major compounds detected by HPLC (Fig. 3). These fermentation products could serve as growth substrates for R. palustris. Under both culture conditions, acetate, lactate, and ethanol were produced at concentrations an order of magnitude higher than pyruvate, which was only produced at sub-millimolar concentrations. All four compounds were produced sooner and/or at higher rates in the co-culture than the C. cellulolyticum monoculture. After approximately 10 days of co-culturing, significant amounts of acetate and pyruvate were removed as the incubation continued, suggesting that they were utilized by R. palustris. In comparison to pyruvate, there was approximately 50-fold more acetate removed, indicating that acetate was the major metabolite for carbon transfer between the two organisms in the co-culture.
3.4.
pH effects
Because it is well known that the release of fermentation products by fermentative bacteria can acidify the medium and slow cellular growth, we monitored the medium pH throughout the incubation period (Fig. 4). When C. cellulolyticum was cultured alone, the medium pH decreased from 7.2 to w5.5, during the initial 15 days of culturing in which cellulose was also depleted. The co-culture pH decrease was initially faster than the monoculture, but leveled off at w5.9 before the cellulose was depleted at day 8. Notably, this pH stabilization coincided with the onset of acetate consumption by R. palustris, suggesting that the removal of acetic acid helped stabilize the co-culture pH. There was no pH change throughout the incubation period in abiotic controls. To test whether the pH stabilization alone resulted in higher cellulose consumption (or rate of degradation) by C. cellulolyticum, we tested the effect of adding Na-HEPES (100 mM, pH 7.2) in the modified CM3 medium. Despite this extra buffering capacity there was still 1e1.5 g of residual cellulose at the end of incubation (data not shown).
Fig. 5 e The inhibition effect of four intermediate metabolites, at multiple concentrations, on cellulose-based growth of C. cellulolyticum. Growth of C. cellulolyticum on cellulose in the presence of test compounds is given in proportion to the control culture lacking additional compounds. In the control where C. cellulolyticum was grown in monoculture on cellulose in the absence of added organic acids or ethanol (the bar at far left, “cellulose alone”), the final cell density was w3 3 108 cells/ml, set at 100%. Error bars represent standard deviations from triplicate cultures.
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3.5.
i n t e r n a t i o n a l j o u r n a l o f h y d r o g e n e n e r g y 3 7 ( 2 0 1 2 ) 1 1 7 1 9 e1 1 7 2 6
Growth inhibition of C. cellulolyticum by pyruvate
Because R. palustris is thought to grow on the fermentation products secreted by C. cellulolyticum, we were curious if the removal of these byproducts by R. palustris has a positive feedback on C. cellulolyticum growth. Accordingly, we tested the dominant intermediate metabolites for inhibitory effects on C. cellulolyticum growth on cellulose. In the presence of 25 mM acetate, lactate or ethanol, the final cell count was 73e82% of that in the control cultures with no additions (Fig. 5). In contrast, 25 mM pyruvate completely eliminated C. cellulolyticum growth (about 0.5% of the control). Due to the severe effect of pyruvate at 25 mM, we conducted further tests for pyruvate inhibition at lower concentrations and found that inhibition of C. cellulolyticum growth appeared linearly correlated with pyruvate concentration (Fig. 5). At 0.5 mM, a concentration similar to what C. cellulolyticum produces in co-cultures, C. cellulolyticum growth was 83% of the control culture.
4.
Discussion and conclusions
Inter-species interactions in microbial communities play an important role in community fitness, and are an area of intense interest in both ecological and industrial research. It is well established that the majority of microbes in nature survive best in communities, most likely due to the partitioning of beneficial functions by individual microbes. Artificial communities, such as a co-culture of two or more microorganisms, are interesting to study because these interactions can be modeled using genomic and proteomic information, toward a comprehensive understanding of how microbial communities operate as a whole. In this study, we examined the physiology and carbon transfer in an artificial syntrophic co-culture containing C. cellulolyticum and R. palustris for cellulose degradation and H2 production. While the cellulose degrader C. cellulolyticum used in this study has low cellulolytic ability and cannot be used industrially without strain improvement [1,5], the focus of our study is for basic research in examining metabolic interactions in a syntrophic co-culture system. We found that the presence R. palustris in the co-culture stimulated C. cellulolyticum growth, cellulose degradation, and total H2 production. The higher H2 productivity in co-culture could be due to two reasons. First, R. palustris can utilize several of the fermentation products excreted by C. cellulolyticum to produce H2. In other words, some of electrons that would otherwise be lost in the obligate excretion of organic acids and alcohols in C. cellulolyticum monoculture could be further converted into H2 by R. palustris in the co-culture [4]. Second, the more complete consumption of cellulose under co-culture conditions should also contribute to the additional H2 produced, by allowing for H2 production by C. cellulolyticum to continue. This is consistent with our finding that the C. cellulolyticum monoculture produced about 27 mmol of H2, whereas the co-culture produced a total of 42 mmol of H2. However, the H2 yield was 1.2 0.2 mol of H2 per mole of glucose equivalent in the C. cellulolyticum monoculture, and 1.4 0.2 mol in the co-culture. Thus, while H2 production by R. palustris contributed to a minor increase in the overall H2 yield, it appears that most of
the additional H2 produced by the co-cultures was due to the more efficient utilization of cellulose. As a result, the main benefit that R. palustris appears to impart is an improvement of C. cellulolyticum metabolic efficiency. The total H2 yields in both the monoculture of C. cellulolyticum and the co-culture are lower than values previously reported for a co-culture consisting of Ethanoligenens harbinense and Rhodopseudomonas faecalis growing on glucose, where 1.8 mol H2 per mole of glucose were produced in single E. harbinense culture and 2.5 mol in the co-culture under similar conditions [15]. The lower H2 production in our system is likely due to the fact that additional energy input is required for cellulose breakdown, a requirement that was circumvented in the previous study [15] where glucose was provided as the carbon source. The examination of population dynamics of each species, the pH monitoring over the growth period, and the quantitative measurement of the intermediate metabolites were important to help make meaningful conclusions about the physiology of the co-culture. The stimulation of C. cellulolyticum growth in the co-culture could be partially due to enhanced buffering of the medium caused by consumption of organic acids by R. palustris. The pH of the co-culture ceased to decrease at w5.9 after 8 days of incubation when acetate in the medium reached maximum concentration (w15 mM). In the C. cellulolyticum monoculture, the pH continued to drop to w5.5 until cellular growth stopped. Nevertheless, the pH effect alone does not account for the growth enhancement of C. cellulolyticum on cellulose in the co-culture, because additional buffer added to the growth medium did not change the amount of cellulose consumed significantly. Therefore, acetate appeared to be the major carbon transfer metabolite between C. cellulolyticum and R. palustris, and acetate removal likely also contributed to growth stimulation of C. cellulolyticum in the co-culture and improved the thermodynamics of the fermentation. Additionally, the extra buffering of the environment provided by the metabolism of R. palustris consuming acetate likely meant that C. cellulolyticum could devote less ATP to maintaining a proper membrane potential and other maintenance processes. While the reduced acetate and pyruvate in the co-culture is likely to be the direct result of carbon transfer, the possibility that cellulolytic clostridia produce less intermediate metabolites in co-culture cannot be excluded. This is an inherited limitation of working with co-culture systems. However, from a mass balance prospective, R. palustris cell numbers increased by two orders of magnitude in the co-culture, therefore carbon was consumed, mostly likely according to the established substrate preference for R. palustris [9,10]. We compared the growth rate of R. palustris on acetate, pyruvate, ethanol and lactate, and found that acetate is the most favorable substrate (data not shown). Although media concentrations of pyruvate were much lower than the other fermentation products, its removal (ostensibly by R. palustris) likely plays an important role in alleviating product inhibition and boosting C. cellulolyticum growth under co-culture conditions. Previous studies have indicated that growth inhibition of C. cellulolyticum is caused by an inefficiently regulated carbon flow [6]. It has been shown that some intermediate products (such as NADH and
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pyruvate) of this metabolism accumulate intracellularly [6,30]. The release of pyruvate may represent an indicator of poorly regulated carbon flow, and the accumulation of the extracellular pyruvate suggests the onset of slowdown of glycolysis. Supporting these previous observations, we found that the highest levels of inhibition occurred when we tested for C. cellulolyticum growth at pyruvate concentrations equivalent to those of other dominant metabolites found in the media (Fig. 5). Since pyruvate was produced by C. cellulolyticum when growing on cellulose, the intracellular concentration is likely to be higher than the extracellular concentration. Conversely, when pyruvate is exogenously added, as in the inhibition experiments (Fig. 5), it likely takes a higher concentration to achieve a similar internal concentration leading to growth inhibition. At co-culture relevant concentrations (w0.5 mM), pyruvate addition caused a 17% reduction of C. cellulolyticum growth. While more study is needed to elucidate the mechanism of this inhibitory effect, our results suggest that pyruvate, albeit at low concentration, acted as an inhibitory compound for cell growth in C. cellulolyticum monocultures. The removal of pyruvate from the co-culture by R. palustris likely alleviated the inhibition effect, therefore boosting the cellular growth of C. cellulolyticum as well as cellulose consumption. In addition, acetate consumption by R. palustris could potentially increase the efficiency of pyruvate to acetate conversion by C. cellulolyticum, permitting it to re-consume and ferment the pyruvate. To our knowledge, this study is the first to use two-H2 producing microorganisms to produce H2 from cellulose degradation in a photosynthetic co-culture system [14,16]. The direct coupling of the metabolisms of the two species has implications for research in the ecology of microbial communities in natural and engineered systems. A systemlevel understanding of the interplay of multiple metabolic pathways relevant to carbon and energy flow will be useful for understanding the population dynamics and nutrient transfer within mixed communities, as well as in strategic bioengineering of microbes to improve overall community fitness and H2 production.
Acknowledgments This work was supported by the Genomic Science Program of the U.S. Department of Energy’s Office of Biological and Environmental Research under contract SCW1039, as part of the LLNL Biofuels Scientific Focus Area. Lawrence Livermore National Laboratory is operated by Lawrence Livermore National Security, LLC, for the U.S. Department of Energy, National Nuclear Security Administration under Contract DEAC52-07NA27344.
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