Laboratoire de Biochimie des Bactéries Gram +, Domaine Scientifique Victor Grignard, Université Henri Poincaré, Faculté des Sciences, BP 239, 54506 Vanduvre-lès-Nancy Cédex, France1
Author for correspondence: Henri Petitdemange. Tel: +33 3 83 91 20 53. Fax: +33 3 83 91 25 50. e-mail: hpetitde{at}lcb.uhp-nancy.fr
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ABSTRACT |
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Keywords: cellulolytic bacteria, flux analysis, environmental pH, cellulose degradation, chemostat
Abbreviations: AADH, acetaldehyde dehydrogenase; AK, acetate kinase; ADH, alcohol dehydrogenase; ATP-Eff, efficiency of ATP generation; Fd, ferredoxin; G1P, glucose 1-phosphate; G6P, glucose 6-phosphate; LDH, lactate dehydrogenase; meq C, milliequivalent of carbon; PFO, pyruvateferredoxin oxidoreductase; PTA, phosphotransacetylase; R, ratio of specific enzyme activity to metabolic flux
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INTRODUCTION |
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High concentrations of fermentation acids and, as a result, low pH conditions are often found in anaerobic habitats (Ljungdahl & Eriksson, 1985 ; Goodwin & Zeikus, 1987
). Yet due to their particular pattern of intracellular pH regulation (Huang et al., 1985
) compared with other low-G+C Gram-positive anaerobes, mainly the so-called lactic acid bacteria, the clostridial-type bacteria are generally considered as restricted to less acidic ecological niches (Russell et al., 1996
).
Cellulolytic clostridia digest cellulose through extracellular multienzyme complexes (Béguin & Lemaire, 1996 ; Bayer et al., 1998
). These cellulosomes are found at the surface of the bacteria and allow both cell adhesion to cellulose fibres (Bayer et al., 1996
) and very efficient degradative activity against crystalline cellulose due to a high synergism of the different cellulase components (Boisset et al., 1999
).
Clostridium cellulolyticum is a low-G+C Gram-positive nonruminal cellulolytic mesophilic bacterium belonging to the clostridial group III, and also classified in family 4, genus 2, of a new proposed hierarchical structure for clostridia (Collins et al., 1994 ). Using cellobiose, a soluble substrate, several advances in understanding of the metabolism of this bacterium have been made, such as (i) a better control of catabolism in a mineral salt-based medium (Payot et al., 1998
; Guedon et al., 1999b
), (ii) recognition of major differences in regulatory responses in cellobiose-limited and cellobiose-saturated chemostat cultures (Guedon et al., 2000b
), and (iii) the importance of glucose 6-phosphate (G6P) and glucose 1-phosphate (G1P) nodes in regulation of metabolic fluxes (Guedon et al., 2000a
). Earlier investigations using cellulose have been mainly devoted to cellulolytic performance and bacterial behaviour towards insoluble substrates (Giallo et al., 1985
; Gelhaye et al., 1993a
, b
). A few studies, however, have focused on the metabolism of this bacterium on cellulose: recent investigation of cellulose fermentation performed in batch culture (Desvaux et al., 2000
) indicated (i) variation of metabolite yields as a function of initial cellulose concentration, and (ii) an early growth inhibition related to pyruvate overflow as with cellobiose (Guedon et al., 1999a
). A study in a cellulose-limited chemostat indicated that bacterial metabolism was not as distorted as with cellobiose and C. cellulolyticum appeared well adapted to a cellulolytic lifestyle (Desvaux et al., 2001
).
Whereas the effects of acidic conditions on the growth of cellulolytic rumen bacteria have been the subject of considerable research (Russell & Dombrowski, 1980 ; Kalachniuk et al., 1994
; Russell & Wilson, 1996
; Russell & Diez-Gonzalez, 1998
), little is known of how these conditions affect the metabolism of cellulolytic clostridia (Duong et al., 1983
; Mitchell, 1998
). The aim of the present work was to investigate the cellulose degradation and metabolic changes of C. cellulolyticum on insoluble cellulose caused by environmental pH conditions. Since conditions in natural environments most likely resemble those somewhere between a closed batch culture and an open continuous culture system (Kovárová-Kovar & Egli, 1998
), both types of culture systems were used to study the effects of pH on growth and metabolism of C. cellulolyticum.
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METHODS |
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Growth conditions.
Clostridium cellulolyticum was grown either in batch or in continuous culture with cellulose as sole carbon and energy source. All experiments were performed in a 1·5 l working volume fermenter (LSL Biolafitte) as previously described (Guedon et al., 1999a , b
); the temperature was maintained at 34 °C and the pH was controlled by automatic addition of 3 M NaOH or 1 M HCl as specified in Results. The inoculum was 10% (v/v) from an exponentially growing culture.
Batch cultures were prepared as previously described (Desvaux et al., 2000 ). The chemostat system used was a segmented gasliquid continuous culture device as described by Weimer et al. (1991b)
with some modifications (Desvaux et al., 2001
). The cultures were maintained for a period of eight to nine residence times (Desvaux et al., 2001
); for each condition the data were the mean of at least three samples.
Analytical procedures.
Biomass, cellulose concentration, gases, extracellular proteins, amino acids, glucose, soluble cellodextrins, glycogen, acetate, ethanol and lactate extracellular pyruvate were assayed as described by Desvaux et al. (2001) . Pyridine nucleotides, coenzyme A (CoA), acetyl-CoA, G1P and G6P were extracted and fluorimetric determination performed as previously described (Desvaux et al., 2001
). ATP and ADP were measured using the luciferinluciferase luminescence system (Microbial Biomass Test Kit, Celsis Lumac) (Guedon et al., 2000a
).
Enzyme assays.
Cell extracts were prepared and enzyme assays performed as previously described (Guedon et al., 2000a ). Pyruvate-ferredoxin (Fd) oxidoreductase (PFO) (EC 1 . 2 . 7 . 1), lactate dehydrogenase (LDH) (EC 1 . 1 . 1 . 27), phosphotransacetylase (PTA) (EC 2 . 3 . 1 . 8), acetate kinase (AK) (EC 2 . 7 . 2 . 1), acetaldehyde dehydrogenase (AADH) (EC 1 . 2 . 1 . 10) and alcohol dehydrogenase (ADH) (EC 1 . 1 . 1 . 1) were assayed as described by Desvaux et al. (2001)
.
Calculations and carbon flux analysis.
The metabolic pathways and equations of cellulose fermentation by C. cellulolyticum, expressed as n hexose equivalents (hexose eq) corresponding to n glucose residues of the cellulose chain, were previously reported (Desvaux et al., 2000 ).
The qcellulose is the specific rate of hexose residue fermented in mmol (g cells)-1 h-1. qacetate, qethanol and qlactate are the specific rates of product formation in mmol (g cells)-1 h-1. qextracellular pyruvate is the specific rate of extracellular pyruvate formation in µmol (g cells)-1 h-1. qNADH produced and qNADH consumed are the specific rates of NADH production and NADH consumption respectively in mmol (g cells)-1 h-1 and were calculated as follows: qNADH produced=qpyruvate and qNADH consumed=2 qethanol+qlactate. The specific rate of acid production (OSullivan & Condon, 1999 ), was calculated as follows: qH+=qacetate+qlactate+qextracellular pyruvate.
The molar growth yield (Yx/s) is expressed in g cells (mol hexose eq fermented)-1. The energetic yield of biomass (YATP) was (Desvaux et al., 2000 ): YATP=concnbiomass/(1·94 concnacetate+0·94 concnethanol+0·94 concnlactate+ 0·94 concnextracellular pyruvate). YATP is expressed in g cells (mole ATP produced)-1. qATP is the specific rate of ATP generation in mmol (g cells)-1 h-1, calculated by the following equation (Desvaux et al., 2000
): qATP=1·94 qacetate+0·94 qethanol+0·94 qlactate+0·94 qextracellular pyruvate. The energetic efficiency (ATP-Eff) corresponding to the ATP generation in cellulose catabolism is given by the ratio of qATP to qcellulose (Miyagi et al., 1994
).
Distribution of the carbon flow by stoichiometric flux analysis (Papoutsakis, 1984 ; Desai et al., 1999a
, b
) was determined by adapting the model developed by Holms (1996)
to C. cellulolyticum metabolism (Desvaux et al., 2001
).
At steady state, the carbon flux through each enzyme of the known metabolic pathway, as indicated in Fig. 1, was calculated in milliequivalents of carbon (meq C) (g cells)-1 h-1, as follows:
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qglucose=0·37 qcellulose
qbiosynthesis=qbiomass-qglycogen+qamino acid+qextracellular protein
qG6P=qbiosynthesis+qpyruvate
qphosphoglucomutase=qG6P-qglucose
qexopolysaccharide=qG1P-qphosphoglucomutase-qglycogen
qacetyl-CoA=qacetate+qethanol
qcarbon dioxide=(qacetate+qethanol)/2
qpyruvate=
qacetate+qethanol+qlactate+qextracellular pyruvate+qcarbon dioxide
The turnover of a pool (h-1) was calculated from specific rate and pool size expressed in moles or in carbon equivalents (Holms, 1996 ). The ratio R corresponded to the ratio of specific enzyme activity to metabolic flux (Holms, 1996
; Desvaux et al., 2001
).
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RESULTS |
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The kinetic profile for individual soluble glucide accumulation indicated that it was a non-growth-associated event (Figs 2Id and 2IId
). A higher level of soluble cello-oligosaccharides was achieved in fermentation without pH control: up to 1·56 mM cellobiose was detected as well as 0·36 mM cellotriose. With pH control, the sugar level in the broth medium was very limited since only 0·16 mM cellobiose was found and no cellotriose could be detected. In both culture conditions, longer cellodextrins could not be detected either by HPLC or by TLC techniques. Inasmuch as cellulose was degraded to a greater extent in the pH-controlled culture, the difference in accumulation of sugars may reflect a difference in the cellulose substrate, e.g. its surface area (Weimer et al., 1990
, 1991a
; Fields & Russell, 2000
), at the time cultures entered stationary phase, and/or differences in metabolism of stationary-phase cells.
Effect of pH on the growth and metabolite production of C. cellulolyticum in cellulose continuous culture
Growth parameters, notably end products measured at each steady state, as a function of the pH value are compiled in Table 1. C. cellulolyticum was grown in continuous culture under cellulose-limited conditions and at a constant dilution rate of 0·053 h-1 with pH values ranging from 7·4 to 6·4. The primary metabolic end products of the cellulose fermentation were acetate, ethanol, lactate, H2 and CO2. In addition to carbon conversion into biomass, amino acids and extracellular proteins were also detected in the supernatant (Table 1
). Exopolysaccharides were readily observable by microscopic examination but could not be measured as previously described (Payot et al., 1998
) due to the significant interference with cellulose fibres leading to erroneous estimation of their concentration. Taking into account amino acids, proteins, fermentative end products and biomass concentration, the carbon balance ranged between 91·4 and 95·3% (Table 1
).
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Metabolic flux analysis of cells grown on cellulose in an acid environment
The metabolism of C. cellulolyticum when grown on cellulose is depicted in Fig. 1. With acidification of the growth medium from pH 7·4 to 6·4, the rate of cellulose consumption declined from 11·74 to 10·13 meq C (g cells)-1 h-1 (Table 3
). The proportion of the cellulose converted into biomass, free amino acids and extracellular proteins, i.e. qbiosynthesis, varied from 24·2 to 27·4% with this pH decline (Table 3
); considering each, the biomasss formation increased from 17·9 to 20·7% of the original carbon while cellulose conversion into amino acid and extracellular protein accounted for around 5·3 and 1·6% of the carbon uptake respectively. Another part of the carbon flow was directed towards metabolite fermentation, i.e. acetate, ethanol, CO2, extracellular pyruvate and lactate, which as a whole decreased from 70·7 to 65·2% of the cellulose consumed (Table 3
). Metabolism distributed carbon differently over known catabolic routes as the pH declined. Carbon flow through the CO2 and acetate formation pathways declined from 23·4 to 19·8% and from 33·1 to 20·4% respectively. However, ethanol formation increased from 13·8 to 19·2% of the carbon consumed; lactate formation also increased, from only 0·3% at pH 7·4 to 4·8% at pH 6·4. At the same time, the proportion of carbon flowing towards extracellular pyruvate rose from 0·1 to 1% with lower environmental pH.
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Enzymic activities as a function of environmental pH
The influence of environmental pH on the specific activities of the enzymes studied is compiled in Table 4. In vitro, PFO, PTA, AK and AADH activities were higher under conditions giving higher in vivo specific production rates (Table 4
). When the carbon flow was expressed as µmol (mg protein)-1 h-1 from previously calculated values (Table 3
), R (the ratio of the specific enzyme activity to metabolic rate) could be calculated (Holms, 1996
). In the metabolic branch leading to acetate production through PTA and AK, R fluctuated between 18·8 and 24·0, and between 20·6 and 12·9, respectively, as pH decreased (Table 4
). R for the enzymes of the ethanol pathway varied between 6·3 and 12·4, and between 15·7 and 12·3 for AADH and ADH, respectively (Table 4
). At each step in the central metabolic pathways, the intracellular concentration of substrates, products, cofactors or effector molecules as well as intracellular ionic strength, redox potential or pH can influence the partition and regulation of the carbon flux (Holms, 1986
). Nevertheless, the fact that fluxes were much less than the available enzyme activity indicated that the carbon flows were determined by the concentration of substrate available rather than the enzyme activity (Holms, 1996
). Despite the variation of enzyme biosynthesis, the amount of these enzymes was always sufficient to catabolize the flowing carbon since R was much higher than 1. At pH 7·4, for the lactate formation pathway, R was very high, i.e. 529·5. This indicated that although LDH was readily available, little carbon was catabolized by this metabolic pathway. As the pH decreased, however, R declined to 46·1, indicating that the LDH was more and more implicated in carbon conversion (Table 4
). As for the metabolic route through PFO, R was in the same range as PTA, AK, AADH or ADH for environmental pH values between 7·4 and 7·0, i.e. R between 6·3 and 11·2 (Table 4
), indicating that the fluxes were much less than the available enzyme activity. Yet for a pH lower than 7·0, R markedly decreased and reached 1·8 at pH 6.4 (Table 4
). Then both biosynthesis and catalytic efficiency of PFO declined with pH since the specific enzyme activity and qacetyl-CoA decreased (Tables 3
and 4
).
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DISCUSSION |
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In continuous culture, maximum cell density was obtained at pH 7·0, but as the pH declined from 7·0 to 6·4 at constant D, biomass was lowered more than fourfold. At the same time, the specific rate of cellulose consumption, however, decreased only from 1·84 to 1·69 meq C (g cells)-1 h-1. Thus environmental acidification influenced chiefly the biomass formation rather than cellulose degradation and assimilation. C. cellulolyticum did not show depressed yields and the transition to wash-out appeared abrupt. This result would be more consistent with a direct effect on a cellular constituent, such as the negative effect of acid on an enzyme or transport protein (Russell & Dombrowski, 1980 ; Russell & Diez-Gonzalez, 1998
).
During cellulose catabolism by C. cellulolyticum, soluble ß-glucans are first converted into G1P and G6P (Desvaux et al., 2000 ). With environmental acidification, the G1P pool increased, since the proportion of carbon flowing via phosphoglucomutase varied between 57·9 and 55·5%. The remaining G1P was directed towards exopolysaccharides (up to 9·9% of the G1P) rather than glycogen synthesis (3·0% maximum of the G1P), both allowing dissipation of carbon surplus (Guedon et al., 2000b
). As the culture pH was lowered, the flow through glycolysis decreased while carbon directed to biosynthesis increased; as a result, the G6P pool was between 15·9 and 18·1 µmol (g cells)-1. Compared with conditions of uncoupling between catabolism and anabolism encountered during ammonium-limited chemostat performed with cellobiose (Guedon et al., 2000a
), the excess of carbon at the G1PG6P branch point was here limited; in fact, exopolysaccharides and glycogen could represent up to 16·0 and 21·4% respectively of the cellobiose consumed and cellotriose was detected extracellularly (Guedon et al., 2000a
).
The increase of the acetyl-CoA pool was corroborated by the analysis of carbon flux; the proportion of cellulose consumed flowing through PFO diminished as the pH declined, as did the ratio of specific enzymic activity to metabolic flux (R), and the flux was rerouted away from acetate production. The acetyl-CoA/CoA ratio decrease was paralleled by decreases in H2/CO2 and qNADH produced/qNADH used. Despite the variation of the NADH balance, calculated from catabolic pathways producing and consuming reducing equivalents, the intracellular NADH/NAD+ ratio was well regulated. Such a result is in good agreement with the model of Decker et al. (1976) , where the NADH-Fd reductase is activated by the acetyl-CoA and inhibited by CoA and which underlines that the fates of NADH and acetyl-CoA regulation are interwined. From acetyl-CoA, acetate was mainly formed but the flux split differently as the environment was acidified, favouring ethanol production. In addition, as pH declined, the level of lactate production rose and coincided with the pyruvate leak, indicating that PFO could no longer support carbon flowing from glycolysis, R decreasing to 1·8 at pH 6·4. Whatever the pH, LDH was always biosynthesized. This enzyme operated mainly as the pH declined but always remained in excess since even at pH 6·4, R was 46·1. In these experimental conditions, LDH allowed draining off part of the pyruvate surplus. At high pH values, H2/CO2 ratios higher than 1 suggested that H2 was produced via NADH-Fd reductase and hydrogenase activites in addition to pyruvate-Fd oxidoreductase and hydrogenase activities. With lower pH values, this ratio decreased and was compensated by the increase of ethanol production until washout occurred.
Reinvestigation of cellulose degradation by C. cellulolyticum (Desvaux et al., 2000 ) showed marked differences in the catabolism of this bacterium as compared with the first investigations carried out (Giallo et al., 1985
). The present paper demonstrates that the inhibition of growth first observed with batch culture performed in penicillin flasks sealed with butyl rubber stoppers and without shaking of the medium (Giallo et al., 1983
, 1985
) is mainly the result of low pH due to acid production in the course of fermentation. The range of pH allowing maximum cell density is restricted; strict control of pH is therefore necessary to obtain the optimum cellulolytic performance in biotechnological processes using C. cellulolyticum. Cellulolytic bacteria so far investigated cannot grow at pH values significantly less than 6·0 (Stewart, 1977
; Russell & Dombrowski, 1980
; Russell & Diez-Gonzalez, 1998
). However, it is well established that in anaerobic habitats, particularly in the natural environment, high fermentation acid concentrations and, as a result, low pH values are often encountered (Ljungdahl & Eriksson, 1985
; Goodwin & Zeikus, 1987
). Since these bacteria have not developed resistance to low pH environments, this implies that they have evolved in an ecological niche where competition for efficient metabolism in acidic conditions is not crucially important. In the same way that growth of C. cellulolyticum under an excess of nutrients (Guedon et al., 1999b
) or with an easily available carbon source, such as soluble glucides, appeared as aberrations considering the natural bacterial ecosystem (Desvaux et al., 2000
, 2001
), cultures without pH control have been shown to be detrimental for optimum growth of this bacterium. These data from monospecies laboratory culture must be extrapolated to microbial ecosystems to explain the maintenance of C. cellulolyticum in natural environments. Clearly much remains to be learned about the complex interactions in which this bacterium takes part in microbiota (Kuznetsov et al., 1979
; Ljungdahl & Eriksson, 1985
; Leschine, 1995
; Costerton et al., 1995
).
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ACKNOWLEDGEMENTS |
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The authors thank G. Raval for technical assistance and E. McRae for correcting the English and for critical reading of the manuscript.
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Received 6 November 2000;
revised 29 January 2001;
accepted 12 February 2001.