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Metabolic shifts in Zymomonas mobilis in response to growth conditions
FEMS MicrobiologyReviews 54 (1988) 131-142 Published by Elsevier
131
FER 00088
Metabolic shifts in Zymomonas mobilis in response to growth conditions S. B r i n g e r - M e y e r a n d H. S a h m lnstitut fiir Biotechnologieder KernforschungsanlageJiilich GmbH, Jiilich, F.R.G.
Received 29 September1987 Accepted 3 November 1987 Key words: By-product; Oxygen toxicity; Ethanol tolerance; Osmotolerance
1. S U M M A R Y Extensive work on ethanol production with the Gram-negative bacterium Zymomonas mobilis has revealed that this is a promising microorganism for industrial use. Concise knowledge of the physiology and metabolism of this organism provides the basis for further improvements by genetic engineering and for the optimization of Zymomonas-specific fermentation processes.
2. I N T R O D U C T I O N Much attention has been paid to Zymomonas mobilis because of its unusual physiological and biochemical properties, as extensively reviewed by Swings and DeLey in 1977 [1]. It is unique among bacteria in fermenting sugar anaerobically by the Entner-Doudoroff pathway and in catabolizing pyruvate to ethanol over an unbranched route with pyruvate decarboxylase. During the past decade interest in this bacterium has broadened to include biotechnological aspects. Rogers and colCorrespondence to: H. Sabra, Institut far Biotechnologie 1 der Kernforschungsanlage Jiilich GmbH., Postfach 1913, D-5170 Jiilieh, F.R.G.
laborators published comparative data on the fermentation yields and kinetics of Z. mobilis and yeast at high sugar concentrations [2]. They showed that Z. mobilis rapidly converted glucose to ethanol. By comparison with Saccharomyces carlsbergensis, Z. mobilis had specific glucose uptake rates and specific ethanol productivities 3-4times greater than yeast and reached ethanol yields of up to 97% of the theoretical maximum value. The considerable amount of new research work on strain selection, methods of fermentation and genetic properties of Z. mobilis has frequently been reviewed [3-14]. This review focusses on some physiological properties of Z. mobilis that are of significance for its industrial use in ethanol production. In particular we shall discuss (1) possible catabolic diversions, depending on the sugar used, (2) oxygen tolerance, (3) ethanol tolerance, and (4) osmotolerante.
3. BY-PASSES O F T H E MAIN S U G A R CATABOLISM When glucose is used as carbon source with non-aerated cultures of Z. mobilis, only insignificant amounts of by-products are produced
0168-6445/88/$04.20 © 1988 Federation of European MicrobiologicalSocieties
132 (acetoin, glycerol, acetate, and lactate) [15-17], as is also evident from the high yields of ethanol obtainable under such conditions [6]. However, in fermentations of sucrose, mixtures of fructose plus glucose, or of fructose alone lower yields are observed, especially at high concentrations of these substrates [18-21]. With the exception of extracellular sugar polymerization reactions during growth on sucrose, reduced ethanol yields under these different growth conditions were found to be caused by the action of reductive enzymes. The following summarizes the present knowledge of these enzymes and of the sucrose-dependent polymerizations. 3.1. Transfructosylation reactions During the fermentation of sucrose by Z. mobilis three types of transfructosylation occur in which water, sucrose or levan can act as an acceptor. Such reactions lead, respectively, to the formation of free fructose, oligosaccharides, or higher polymers of the original levan molecule, a fl-(2,6)fructose polymer [22]. The rate of formation of levan slowed down when the glucose concentration was increased to 5-10 g/1 [22-24] and was substantially reduced at 3 5 ° C as compared to a growth temperature of 30 ° C [25,26]. The rates of sucrose hydrolysis and of transfructosylations were shown to be higher than the sugar uptake rate of Z. mobilis [27,28]. Consequently, the hydrolysis of sucrose was not the rate-limiting factor in the fermentation, as could be seen from the accumulation of residual glucose and fructose with increasing dilution rates in continuous cultures [27]. Whereas essentially no by-products were found under conditions of carbon limitation, a high molecular weight levan ( M r 20000 [22]) was formed at high dilution rates or at high sucrose feed concentrations. I n addition, non-precipitable oligosaccharides were detected in concentrations up to 10% of the initial sucrose concentration. These oligomers consisted of trimers, tetramers and pentamers, each containing 1 mol of glucose per 2, 3 or 4 mol of fructose [27]. The sucrose-hydrolyzing activity seems to be stimulated by sucrose and fructose [29]. Viikari [22] determined the activity of a levan sucrase (EC 2.4.1.10) (Fig. 1 (1)) by two methods. With sucrose as substrate the
formation of levan was monitored by measuring the turbidity, and the overall transfructosylations by the liberation of glucose. With both test methods the s a m e g m value for sucrose was found (60 mM) but the pH optimum for levan formation was at 5, whereas the pH optimum for overall transfructosylation was at 6. This may indicate the existence of two distinct enzymes. Approximately 75% of the total enzyme activity was in the cell fraction; the remainder was in the supernatant. Levan sucrase has been purified from the cells [24] as well as from the culture broth [29]; in both instances the liberation of glucose was followed for activity determinations. The reported values for p H optimum, Michaelis constant for sucrose and for the inhibitory concentrations of glucose, mannose and ethanol [22,24,29] show considerable differences. The interpretation of these results is complicated by the fact that the available data leave unresolved the question of whether a single levan sucrase can account for both sucrose hydrolysis and the transfructosylation reactions which lead to oligomers and levan. Thus, the existence of a separate invertase (EC 3.2.1.26) could not be excluded [22,29]. 3.2. Glucose-fructose oxidoreductase Z. mobilis produces sorbitol in amounts up to 11% of the initial carbon source when both glucose and fructose are present in the fermentation medium. These can be supplied either as a mixture of the two sugars or as hydrolysis products of sucrose [15,20,30-33]. Contrary to Barrow et al. [31], Viikari [15] also detected small amounts of sorbitol when the cells were grown on a high concentration of glucose alone or of fructose alone. Studies of the enzymatic mechansim of sorbitol formation showed that the reaction was independent of free NAD(P)H [22,33,34]. Crude extracts only formed sorbitol when both glucose and fructose were added, sorbitol was derived from fructose as was shown by (14C)-feeding experiments [31,34]. Zachariou and Scopes [35] isolated and characterized an enzyme catalyzing sorbitol formation, a glucose-fructose oxidoreductase (Fig. 1 (4)). This enzyme catalyzes the intermolecular oxidation-reduction of glucose and fructose to form gluconolactone and sorbitol. The oxidore-
133
r--
Sucrose
Fructose-6-P
Fructose
L
0
Glucose
I
Glucose-6-P
Glucose-6-P
6-P-Gluconolactone
I
0
2-Keto-3-desoxy6-P-gluconate 8 _: Glyceraldehyde-3-P
Dlhydroxyacetone-P
0 NADH
0 I
Glycerate-1,3-P
I----
ATP
@
NAD
@ 4
Glycerol-3-P
@
3-P-Glycerate 0 1 2-P-Glycerate NAD [Lactate]
+--
c3
ATP I
Pyruvate +
I
I
Phowhoenolwruvate
I
Fig. 1. Products of glucose, fructose and sucrose catabolism in Zymomonur mobilis. Broken lines: hypothetical pathways. Enzymes: 1, levansucrase; 2, invertase; 3, mannitol dehydrogenase; 4, glucose-fructose oxidoreductase; 5, fructokinase; 6, glucose-6-P isomerase; 7, glucose dehydrogenase; 8, ghrconolactonase; 9, ghtconatekinase; 10, ghrcokinase; 11, ghtcose-6-P dehydrogenase; 12, 6-P-ghtconolactonase; 13, 6-P-ghtconate dehydratase; 14, KDPG-aldolase; 15, glyceraldehyde-P dehydrogenase; 16, phosphoglycerate kinase; 17, Phosphoglycerate mutase; 18, enolase; 19, pyruvate kinase; 20, lactate dehydrogenase; 21, triose-P isomerase; 22, phosphatase; 23, glycerol-P dehydrogenase; 24, phosphatase; 25, pyruvate decarboxylase; 26, alcohol dehydrogenase; 27, pyruvate-formate lyase; 28, acetaldehyde dehydrogenase; 29, acetyl-CoA synthetase.
134 ductase was present in cells grown on glucose, fructose and mixtures of the two sugars; however, the highest activity was found in cells grown on 200 g/1 glucose. The enzyme contained tightly bound N A D P and did not require added cofactors for activity. The oxidoreductase had low affinities for its substrates but constituted up to 0.5% of the readily soluble protein in Z. mobilis [35]. The properties of this oxidoreductase are consistent with the observation that in vivo sorbitol is predominantly formed at high concentrations of glucose plus fructose [12,31]. Under these conditions the fructokinase which channels fructose into the ethanol-producing pathway is strongly inhibited by glucose [36-38]. The second product of the glucose-fructose oxidoreduction, gluconolactone, is quickly hydrolyzed by a gluconolactonase (EC 3.1.1.17) [35] and channelled into the EntnerDoudoroff pathway by a gluconate kinase (EC 2.7.1.12) [39]. A reversal of the enzymatic reaction for sorbitol formation seems improbable, since the lactone intermediate is highly unstable [34]; this would explain why Z. mobilis is unable to utilize sorbitol as a carbon source [1,31]. 3.3. Mannitol dehydrogenase Cells cultured in the presence of 150 g/1 fructose form small amounts of mannitol, corresponding to only 2.5% of the fructose metabolized [22]. In cell-free extracts, a NADP-dependent mannitol dehydrogenase activity (Fig. 1 (3)) was detected which was specific for fructose reduction and for mannitol oxidation [18]. It may be that this NADPH-dependent reduction of fructose is coupled to the dehydrogenation of glucose 6-phosphate, since the glucose-6-phosphate dehydrogenase present in Z. mobilis is active with N A D P as well as with N A D [38,40,41]. The high K m value of 180 mM for fructose of the crude mannitol dehydrogenase shows that detectable amounts of mannitol in culture supernatants would only be found at high fructose concentrations. The requirement for NADP(H) and the fact that the p H optimum for the reduction of fructose was at 8 but that for the oxidation of mannitol was at 10 [18] may help to explain why Z. mobilis cannot grow on mannitol in the p H range of 5 - 6 usually employed for fermentations [1,34]. It is not known
whether the mannitol dehydrogenase is constitutive or is induced by fructose. 3.4. Glucose dehydrogenase With ferricyanide or 2,6-dichlorophenolindophenol as artificial electron acceptor, a glucose oxidizing activity can be measured in cell-free extracts [34,35] (Fig. 1 (7)). Other aldohexoses and the sugar alcohols mannitol and sorbitol are also oxidized, although at lower rates [34] (Strohdeicher, M. et al. 1987, unpublished results). This enzyme activity is localized in the membrane fraction (Scopes, R.K., personal communication; Strohdeicher, M. et al. 1987, unpublished results) and seems to be similar to non-nucleotide-linked glucose dehydrogenases from other microorganisms. The latter enzymes have been identified as quinoproteins [42]. The in vivo end-acceptor of the electrons derived from glucose by the glucose dehydrogenase in Z. mobilis is not yet known.
4. E F F E C T OF O X Y G E N Aeration of cultures of Z. mobilis also leads to a diversion of reducing power and a consequent decrease in ethanol production [22,43-46]. Furthermore, the ability of Z. mobilis to grow in the presence of oxygen is not correlated with higher cell yields or with higher growth rates as compared to anaerobic conditions [22,43-46]. With increasing rates of aeration and concentrations of the carbon source, growth becomes strongly inhibited [44-46] and cell yields decrease [44,45]. 4.1. NADH-oxidase Z. mobilis was found to possess a N A D H - and NADPH-oxidase [44,45], located in the cell membrane, which catalyzes the oxidation of NAD(P)H with oxygen at a molar ratio of 2 : 1. Therefore, it can be assumed that oxygen is reduced to water by this reaction. The activity of N A D H oxidation accounted for the oxygen consumption rate of cell suspensions in the presence of a carbon source [44]. The cytochrome content of the cell membrane [43,45] and the inhibition by cyanide of both intracellular oxygen reduction by the NADH-oxidase and oxygen consumption by whole
135 Table 1 Influence of oxygen on growth and product formation of
Zymomonas mobilis Culture conditions *
Ethanol (g/l)
Acetaldehyde (g/l)
Growth yield g cell dry w e i g h t / mol glucose
Anaerobiosis Aerobiosis
63.4 20.1
0.24 1.31
2.3 1.4
* Continuous cultures were at D = 0.08 h -1 and with 14%
(w/v) glucose.
cells led to the suggestion that a respiratory chain is present in Z. mobilis [43,44]. Obviously, the transfer of electrons via this respiratory chain is not coupled with oxidative phosphorylation. In addition to the NAD(P)H-oxidase, catalase, superoxide dismutase and peroxidase were detected in Z. mobilis [1,44,45]. Thus, the observed toxicity of oxygen must be caused by a depletion in the pool of reduced coenzymes via the NAD(P)Hoxidase. As a consequence, acetaldehyde, a toxic intermediate of sugar catabolism accumulates (Table 1). In addition, acetic acid [43,44] and dihydroxyacetone [22] are formed as products during aerobic growth. A common feature of all the foregoing reactions, i.e., those catalyzed by glucose-fructose oxidoreductase, mannitol dehydrogenase, glucose dehydrogenase and NAD(P)H-oxidase, is that they cause a redox imbalance in the sugar catabolism of Z. mobilis. Glucose-fructose oxidoreductase and glucose dehydrogenase both lead to the formation of gluconic acid which is phosphorylated and enters the Entner-Doudoroff pathway after the NAD(P)H-yielding reaction catalyzed by glucose6-phosphate dehydrogenase. Thus only 1 mol of reduced coenzyme is formed by the glyceraldehyde-3-phopshate dehydrogenase for each mol of gluconic acid catabolized. Mannitol dehydrogenase and NAD(P)H-oxidase directly consume reducing power. These diversions of reducing power result in an accumulation of products which are more oxidized than ethanol, namely acetaldehyde, acetoin and acetic acid [18,22,33,43]. Surprisingly, dihydroxyacetone and glycerol were also formed, mainly in cultures growing at high fructose concentrations [18,22]. Dihydroxyacetone
is derived from glyceraldehyde 3-phosphate by the sequential action of triose phosphate isomerase [18,47] and a phosphatase, and it can be assumed that glycerol is formed by further reduction of dihydroxyacetone phosphate to glycerol phosphate followed by hydrolysis to glycerol [18]. The formation of both of these products, dihydroxyacetone and glycerol, increases the redox imbalance, since for each glyceraldehyde phosphate that is not converted to diphosphoglycerate, one NADH is lost. An additional reducing equivalent has to be consumed to form glycerol. The cause for the accumulation of dihydroxyacetone and glycerol is thus not readily apparent in a situation of NADH deficiency and poses an interesting problem for further investigation. Up to now, no physiological roles can be attributed to sorbitol and mannitol formation, or to the membranebound NAD(P)H-oxidase and glucose dehydrogenase of Z. mobilis.
5. EFFECTS OF ETHANOL Among bacteria, Z. mobilis has an exceptionally high ethanol tolerance and is capable of producing ethanol in concentrations up to 13% (w/v) [6]. Despite this fact, end-product inhibition by ethanol is clearly the main limiting factor for the overall efficiency of the fermentation process [48-50]. A search for the primary site of ethanol inhibition has led to a number of investigations on how ethanol affects catabolic enzymes and the cell membrane. Recent work of Scopes and coworkers [47,51] has shown that a cell-free system of Z. mobilis can rapidly consume glucose and produce ethanol concentrations exceeding 15% w/v. These results demonstrate that the Entner-Doudoroff enzymes are more resistant to ethanol than are the living cells [51]. Much evidence has been accumulated to support the hypothesis that the cell membrane is the primary target for ethanol inhibition. Ethanol is known to affect the integrity of biological membranes by intercalation into t h e lipid bilayer. This results in a disorganization of the cell membranes which is in some respects similar to the heat-induced increase of membrane fluidity. Leakage of cofactors and coenzymes
136 through the plasma membrane has been reported as a consequence of ethanol- and heat-damage [52,53]. The reviews of Ingrain and Buttke [54] and Ingrain [55] give a comprehensive overview of various aspects of the action of alcohols on microorganisms.
5.1. Fatty acid and phospholipid patterns In the case of Z. mobilis, specific mechanisms should then exist to counteract the adverse effects of ethanol on the cell membrane. As was shown by several groups, the major fatty acids occurring in Z. mobilis are myristic (14 : 0), palmitic (16 : 0) and cis-vaccenic acid (18 : 1). There is a very high level of vaccenic acid, accounting for up to 70% of the total fatty acids [56-60]. Among the phospholipids (phosphatidylethanolamine, phosphatidylglycerol, phosphatidylcholine, cardiolipin, and phosphatidylserine), phosphatidylethanolamine is the most abundant [57-59]. The fatty acid pattern of batch cultures did not change significantly during growth and alcohol accumulation, and only small shifts in the phospholipid pattern were observed [59,61].
4"'.
4"'.
©
@
G HO
OH
OH
T : i"
l
I
I
OH
NHX OH
tt~/~ 0
f
5.2. Ethanol- and temperature-dependent synthesis of hopanoids and proteins Another important class of lipids present in Z. mobilis are the hopanoids [57,58,60]. It has been suggested that these pentacyclic triterpenoids are the prokayrotic structural and functional equivalents of sterols in eukaryotes; sterols are known to be partly responsible for the high alcohol tolerance of yeast [62,63]. The biosynthesis of hopanoids, in contrast to that of sterols, does not require oxygen [64]. The hopanoid patterns of Z. mobilis ATCC 29191 and Z. mobilis ATCC 10988 are similar [60]; both strains contain hopene, hopanol [58], bacteriohopanetetrol [60], and the glycoside and ether derivatives of bacteriohopanetetrol [65] (Fig. 2). Bacteriohopanetetrol and its derivatives are not discriminated from each other when analyzed by gas-liquid chromatography, after derivatization [65]. A number of other triterpenoids, described by Tornabene et al. [57], probably occur only as minor components. Analysis by gas-liquid chromatography shows that the components analyzed as bacteriohopenetetrol represent the major hopanoids [60]. It is significant that the contribution of bacteriohopanetetrol to
~
OH
NH 2
2 Ho
[
OH
/ CH~OH ~°o.
@
~'.,
®
Fig. 2. Hopanoids from Zymomonasmobilis. 1, hopene; 2, diplopterol; 3, bacteriohopane-tetrol; 4, bacteriohopanetetrol ether; 5, glucosaminylbacteriohopanetetrol.
137 the total lipid fraction depends strongly on ethanol concentration, as was shown with cells grown under continuous or batch conditions [60,66]. In cells from continuous cultures, with an endogenous ethanol concentration of 5 g/l, the amount of bacteriohopanetetrol was 2.5% of the total lipids whereas, at an ethanol level of 63 g/l, this amount increased to 21% of the total lipids. Likewise, the amounts of hopene and hopanol increased from 0% and 0.9% to 1% and 2.9% of the total lipids, respectively [60]. A strong response of the hopanoid content of cells was also observed when cell suspensions were incubated in media with added ethanol. In cells exposed to 160 g/1 ethanol, bacteriohopanetetrol comprised 36.5% of the total lipids whereas in cells incubated without added ethanol, the corresponding value dropped to just 9.5% [66]. Besides ethanol, n-propanol and nbutanol also increased the relative hopanoid content [66]. Stimulation of the hopanoid biosynthesis seems to be mediated through changes in membrane fluidity, since a change in incubation temperature from 30 to 37°C leads to the same increase in hopanoid content as high concentrations of ethanol [66]. The large shifts in the contribution of hopanoids to the total lipid content raise the question of whether the amount of total lipids per cell is also variable, or whether it remains constant. In the first instance, the ethanol- and temperature-dependent synthesis of hopanoids could proceed irrespective of other membrane components, and consequently the amount of total lipids per cell would increase. Alternatively, the cell could compensate for increasing amounts of hopanoids by decreasing the synthesis of other membrane components. Investigations of Benschoter and Ingram [53] on the thermal tolerance of Z. mobilis support the second assumption. In their study the ratio of phospholipids to total cell protein and membrane protein strongly decreased with increasing growth temperature. The phospholipid-to-protein ratio of membranes from cells grown at 41°C was approximately half that of membranes from cells grown at 20 ° C. In addition it was shown by measurements of the fluorescence depolarization of diphenylhexatriene that, with increasing growth temperature, the membranes from Z. mobilis were progressively more rigid [53]. This
observation is in accordance with the properties predictable for membranes containing increasing amounts of hopanoids [63,67]. Even apart from this proposed partial substitution of hopanoids for phospholipids the increase in the relative protein content of the membranes [53,59] provides an additional means to counteract membrane fluidization at elevated temperature. Ethanol and temperature not only have parallel effects on the distribution of lipids but also on the protein pattern of Z. mobilis [68,69]. In cells incubated for 10 rain either in the presence of 5% (v/v) ethanol at 30°C, or without ethanol at 45°C, the synthesis of several polypeptides was induced or stimulated as compared to untreated ceils. Among these polypeptides, two were found associated with the envelope fraction. Interestingly, heat- and ethanol-shocked cells showed a significantly enhanced viability during subsequent heat treatment [69]. Although the physiological role of the proteins induced by heat and ethanol is not known at present, an involvement in the mechanism of ethanol tolerance of Z. mobilis can be assumed. 5.3. Significance of hopanoids A vital function, however, can be attributed to the hopanoids themselves. That hopanoids are indispensable for Z. mobilis was shown by Flesch and Rohmer [70] who used inhibitors of squalene hopene cyclase. These substances are squalene analogues that inhibit growth at concentrations of 1-3 #M. The squalene cyclase which catalyzes the cyclization of squalene into hopanoids [71-73] seems to be the main regulatory enzyme of hopanoid synthesis in Z. mobilis (Schmidt, A., unpublished). The mechanism by which ethanol tolerance is achieved in Z. mobilis appears to be a concerted shift in the relative amounts of phospholipds, hopanoids and proteins in the cell envelope. The large ethanol-dependent shifts in the hopanoid content suggest a major function of these sterollike substances for membrane stabilization. For further studies on the influence of the fatty acid composition of phospholipids, the incorporation of exogenous phospholipids by interaction of Z. mobilis cells with liposomes [74] may prove to be
138 useful. Attempts to alter the fatty acid composition of Z. mobilis by adding fatty acid supplements during growth have not yet been successful [59].
enzymes [51]. The view that substantial sugar concentrations exist intraceUularly in Z. mobilis at high external sugar concentrations is further supported by the occurrence of enzymes which have extremely low affinities for their substrates, e.g., glucose-fructose oxidoreductase [35].
6. OSMOTOLERANCE In their comprehensive, comparative study of about forty strains of Z. mobilis, Swings and DeLey [1] stated that all these strains were able to grow with 20% glucose and most strains even grew in media containing 30% or 40% glucose. Such high sugar concentrations decrease the total water concentration and exert osmotic pressures which are comparable to relatively strong salt solutions. For example, a 20% (w/v) glucose solution has approximately the same osmotic pressure as a 4.2% (w/v) solution of sodium chloride. However, the salt resistance of Z. mobilis is rather low. 71% of the strains tested by Swings and DeLey [1] grew in the presence of 1% NaC1, whereas none could grow in the presence of 2% NaCI. These results suggest that the salt intolerance of Z. mobilis is not a mere problem of increased osmotic pressure and decreased water activity, but is mainly due to effects specific for electrolyte solutes. 6.1. Tolerance towards highly concentrated sugar solutions A possible explanation for the tolerance of Z. mobilis towards highly concentrated glucose solutions may be derived from studies on glucose transport [75,76] and on the activity of a cell-free system at high glucose concentrations [47,51]. DGlucose is transported by a high-velocity, carriermediated facilitated diffusion system, whereby its intracellular concentration quickly reaches a plateau close to the external concentration [76]. Studies with a cell-free system showed that concentrated glucose solutions were not inhibitory to the Entner-Doudoroff enzymes, since conversion of 2 M (36%) glucose to ethanol by this system proceeded rapidly [47,51]. Thus, the extracellular osmotic pressure of glucose solution may rapidly be balanced by corresponding intracellular sugar concentrations. Reduced water activity seems to have only minor effects on the Entner-Doudoroff
6.2. Salt intolerance The low salt tolerance of Z. mobilis poses problems for the fermentation of molasses which usually have a high salt content [12,77,78]. Under normal batch conditions, Z. mobilis grew poorly on molasses due to salt inhibition [12,78]. It is likely, however, that this problem can be overcome by genetic improvement of Z. mobilis [7,11] and by optimized fermentation technologies for molasses [12,78].
7. CONCLUSIONS Z. mobilis performs a highly productive ethanol fermentation and offers a number of advantages over the traditional yeast fermentation. As was demonstrated by Rogers and collaborators [6] the specific rates of sugar uptake and ethanol production for Z. mobilis are 3-4-times faster than those for yeasts. In addition, higher ethanol yields and lower biomass yields are obtained with Z. mobilis, compared to yeasts, due to the differences in carbohydrate metabolism. Biosyntheses of membrane components, which have important functions in the exceptionally high ethanol tolerance of Z. mobilis, proceed independently of the presence of oxygen [64]. The ability of this bacterium to grow rapidly in the absence of oxygen simplifies the conditions of continuous fermentation processes. Thus, Z. mobilis is a promising microorganism for industrial ethanol production, especially from starch-based substrates [12,79-81]. Perhaps the greatest obstacle to commercialization is that, compared to yeast, much less practical experience exists for the Z. mobilis fermentation. The extensive work done in the field of evaluation and optimization of fermentation techniques (recent publications: [82-99]) reflects the neccessity for the development of Z. mobilis-specific processes, especially those that can operate stably
139 in a n o n - s t e r i l e e n v i r o n m e n t . T h e n a r r o w r a n g e of s u b s t r a t e s utilized b y Z. mobilis (i.e., glucose, fructose a n d sucrose), the f o r m a t i o n of b y - p r o d ucts in fructose a n d s u c r o s e - c o n t a i n i n g m e d i a a n d the low salt t o l e r a n c e are d i s a d v a n t a g e o u s for c o m m e r c i a l a p p l i c a t i o n s . However, this b a c t e r i u m is p e r h a p s m o r e r e a d i l y m o d i f i e d b y genetic eng i n e e r i n g techniques t h a n yeast (for reviews see [7,9,11]). R e c e n t p u b l i c a t i o n s c o n c e r n the cons t r u c t i o n of vectors a n d transfer o f p l a s m i d s in Z. mobilis [100-111]. F u r t h e r m o r e , a n u m b e r o f p r o teins have b e e n p u r i f i e d f r o m Z. mobilis, m o s t r e c e n t l y the E n t n e r - D o u d o r o f f e n z y m e s [112], the two alcohol d e h y d r o g e n a s e s [113] a n d the p y r u v a t e d e c a r b o x y l a s e [114-117]. T h e p y r u v a t e dec a r b o x y l a s e gene h a s b e e n c l o n e d a n d s e q u e n c e d [116-119]. It seems p r o b a b l e that b y genetic m a n i p u l a t i o n , the Z. mobilis f e r m e n t a t i o n c a n b e f u r t h e r i m p r o v e d with r e g a r d to the r a n g e of u t i l i z a b l e substrates, salt t o l e r a n c e a n d the byp r o d u c t s f o r m e d f r o m sucrose or invert sugar.
ACKNOWLEDGEMENTS T h e a u t h o r s g r a t e f u l l y a c k n o w l e d g e the critical r e a d i n g of the m a n u s c r i p t b y Professor R . K . F i n n .
REFERENCES [1] Swings, J. and DeLey, J. (1977) The biology of Zyraomonas. Bacteriol. Rev. 41, 1-46. [2] Rogers, P.L., Lee, K.J. and Tribe, D.E. (1979) Kinetics of alcohol production by Zymomonas mobilis at high sugar concentrations. Bioteclmol. Lett. 1, 165-170. [3] Rogers, P.L., Lee, K.J. and Tribe, D.E. (1980) High productivity ethanol fermentations with Zymomonas mobilis. Proc. Biochem., August/September, 7-11. 14] Rogers, P.L., Lee, K.J., Skotnicki, M.L. and Tribe, D.E. (1981) Ethanol fermentation by highly productive strains of Zymomonas raobilis. Adv. Bioteehnol. 2, 189-194. [5] Stokes, H.W., Dally, E., Spaniel, D.T., Williams, R.L., Montenecourt, B.S. and Eveleigh, D.E. (1981) Zymomonas: The phantom ethanol producer. Biomass Digest 3, 124-132. [6] Rogers, P.L., Lee, K.J., Skotnicki, M.L. and Tribe, D.E. (1982) Ethanol production by Zymomonas mobilis. Adv. Biochem. Eng. 23, 37-84. [7] Skotnicki, M.L., Lee, K.J., Tribe, D.E. and Rogers, P.L. (1982) Genetic alteration of Zymomonas raobilis for ethanol production. Basic Life Sei. 19, 271-290.
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131
FER 00088
Metabolic shifts in Zymomonas mobilis in response to growth conditions S. B r i n g e r - M e y e r a n d H. S a h m lnstitut fiir Biotechnologieder KernforschungsanlageJiilich GmbH, Jiilich, F.R.G.
Received 29 September1987 Accepted 3 November 1987 Key words: By-product; Oxygen toxicity; Ethanol tolerance; Osmotolerance
1. S U M M A R Y Extensive work on ethanol production with the Gram-negative bacterium Zymomonas mobilis has revealed that this is a promising microorganism for industrial use. Concise knowledge of the physiology and metabolism of this organism provides the basis for further improvements by genetic engineering and for the optimization of Zymomonas-specific fermentation processes.
2. I N T R O D U C T I O N Much attention has been paid to Zymomonas mobilis because of its unusual physiological and biochemical properties, as extensively reviewed by Swings and DeLey in 1977 [1]. It is unique among bacteria in fermenting sugar anaerobically by the Entner-Doudoroff pathway and in catabolizing pyruvate to ethanol over an unbranched route with pyruvate decarboxylase. During the past decade interest in this bacterium has broadened to include biotechnological aspects. Rogers and colCorrespondence to: H. Sabra, Institut far Biotechnologie 1 der Kernforschungsanlage Jiilich GmbH., Postfach 1913, D-5170 Jiilieh, F.R.G.
laborators published comparative data on the fermentation yields and kinetics of Z. mobilis and yeast at high sugar concentrations [2]. They showed that Z. mobilis rapidly converted glucose to ethanol. By comparison with Saccharomyces carlsbergensis, Z. mobilis had specific glucose uptake rates and specific ethanol productivities 3-4times greater than yeast and reached ethanol yields of up to 97% of the theoretical maximum value. The considerable amount of new research work on strain selection, methods of fermentation and genetic properties of Z. mobilis has frequently been reviewed [3-14]. This review focusses on some physiological properties of Z. mobilis that are of significance for its industrial use in ethanol production. In particular we shall discuss (1) possible catabolic diversions, depending on the sugar used, (2) oxygen tolerance, (3) ethanol tolerance, and (4) osmotolerante.
3. BY-PASSES O F T H E MAIN S U G A R CATABOLISM When glucose is used as carbon source with non-aerated cultures of Z. mobilis, only insignificant amounts of by-products are produced
0168-6445/88/$04.20 © 1988 Federation of European MicrobiologicalSocieties
132 (acetoin, glycerol, acetate, and lactate) [15-17], as is also evident from the high yields of ethanol obtainable under such conditions [6]. However, in fermentations of sucrose, mixtures of fructose plus glucose, or of fructose alone lower yields are observed, especially at high concentrations of these substrates [18-21]. With the exception of extracellular sugar polymerization reactions during growth on sucrose, reduced ethanol yields under these different growth conditions were found to be caused by the action of reductive enzymes. The following summarizes the present knowledge of these enzymes and of the sucrose-dependent polymerizations. 3.1. Transfructosylation reactions During the fermentation of sucrose by Z. mobilis three types of transfructosylation occur in which water, sucrose or levan can act as an acceptor. Such reactions lead, respectively, to the formation of free fructose, oligosaccharides, or higher polymers of the original levan molecule, a fl-(2,6)fructose polymer [22]. The rate of formation of levan slowed down when the glucose concentration was increased to 5-10 g/1 [22-24] and was substantially reduced at 3 5 ° C as compared to a growth temperature of 30 ° C [25,26]. The rates of sucrose hydrolysis and of transfructosylations were shown to be higher than the sugar uptake rate of Z. mobilis [27,28]. Consequently, the hydrolysis of sucrose was not the rate-limiting factor in the fermentation, as could be seen from the accumulation of residual glucose and fructose with increasing dilution rates in continuous cultures [27]. Whereas essentially no by-products were found under conditions of carbon limitation, a high molecular weight levan ( M r 20000 [22]) was formed at high dilution rates or at high sucrose feed concentrations. I n addition, non-precipitable oligosaccharides were detected in concentrations up to 10% of the initial sucrose concentration. These oligomers consisted of trimers, tetramers and pentamers, each containing 1 mol of glucose per 2, 3 or 4 mol of fructose [27]. The sucrose-hydrolyzing activity seems to be stimulated by sucrose and fructose [29]. Viikari [22] determined the activity of a levan sucrase (EC 2.4.1.10) (Fig. 1 (1)) by two methods. With sucrose as substrate the
formation of levan was monitored by measuring the turbidity, and the overall transfructosylations by the liberation of glucose. With both test methods the s a m e g m value for sucrose was found (60 mM) but the pH optimum for levan formation was at 5, whereas the pH optimum for overall transfructosylation was at 6. This may indicate the existence of two distinct enzymes. Approximately 75% of the total enzyme activity was in the cell fraction; the remainder was in the supernatant. Levan sucrase has been purified from the cells [24] as well as from the culture broth [29]; in both instances the liberation of glucose was followed for activity determinations. The reported values for p H optimum, Michaelis constant for sucrose and for the inhibitory concentrations of glucose, mannose and ethanol [22,24,29] show considerable differences. The interpretation of these results is complicated by the fact that the available data leave unresolved the question of whether a single levan sucrase can account for both sucrose hydrolysis and the transfructosylation reactions which lead to oligomers and levan. Thus, the existence of a separate invertase (EC 3.2.1.26) could not be excluded [22,29]. 3.2. Glucose-fructose oxidoreductase Z. mobilis produces sorbitol in amounts up to 11% of the initial carbon source when both glucose and fructose are present in the fermentation medium. These can be supplied either as a mixture of the two sugars or as hydrolysis products of sucrose [15,20,30-33]. Contrary to Barrow et al. [31], Viikari [15] also detected small amounts of sorbitol when the cells were grown on a high concentration of glucose alone or of fructose alone. Studies of the enzymatic mechansim of sorbitol formation showed that the reaction was independent of free NAD(P)H [22,33,34]. Crude extracts only formed sorbitol when both glucose and fructose were added, sorbitol was derived from fructose as was shown by (14C)-feeding experiments [31,34]. Zachariou and Scopes [35] isolated and characterized an enzyme catalyzing sorbitol formation, a glucose-fructose oxidoreductase (Fig. 1 (4)). This enzyme catalyzes the intermolecular oxidation-reduction of glucose and fructose to form gluconolactone and sorbitol. The oxidore-
133
r--
Sucrose
Fructose-6-P
Fructose
L
0
Glucose
I
Glucose-6-P
Glucose-6-P
6-P-Gluconolactone
I
0
2-Keto-3-desoxy6-P-gluconate 8 _: Glyceraldehyde-3-P
Dlhydroxyacetone-P
0 NADH
0 I
Glycerate-1,3-P
I----
ATP
@
NAD
@ 4
Glycerol-3-P
@
3-P-Glycerate 0 1 2-P-Glycerate NAD [Lactate]
+--
c3
ATP I
Pyruvate +
I
I
Phowhoenolwruvate
I
Fig. 1. Products of glucose, fructose and sucrose catabolism in Zymomonur mobilis. Broken lines: hypothetical pathways. Enzymes: 1, levansucrase; 2, invertase; 3, mannitol dehydrogenase; 4, glucose-fructose oxidoreductase; 5, fructokinase; 6, glucose-6-P isomerase; 7, glucose dehydrogenase; 8, ghrconolactonase; 9, ghtconatekinase; 10, ghrcokinase; 11, ghtcose-6-P dehydrogenase; 12, 6-P-ghtconolactonase; 13, 6-P-ghtconate dehydratase; 14, KDPG-aldolase; 15, glyceraldehyde-P dehydrogenase; 16, phosphoglycerate kinase; 17, Phosphoglycerate mutase; 18, enolase; 19, pyruvate kinase; 20, lactate dehydrogenase; 21, triose-P isomerase; 22, phosphatase; 23, glycerol-P dehydrogenase; 24, phosphatase; 25, pyruvate decarboxylase; 26, alcohol dehydrogenase; 27, pyruvate-formate lyase; 28, acetaldehyde dehydrogenase; 29, acetyl-CoA synthetase.
134 ductase was present in cells grown on glucose, fructose and mixtures of the two sugars; however, the highest activity was found in cells grown on 200 g/1 glucose. The enzyme contained tightly bound N A D P and did not require added cofactors for activity. The oxidoreductase had low affinities for its substrates but constituted up to 0.5% of the readily soluble protein in Z. mobilis [35]. The properties of this oxidoreductase are consistent with the observation that in vivo sorbitol is predominantly formed at high concentrations of glucose plus fructose [12,31]. Under these conditions the fructokinase which channels fructose into the ethanol-producing pathway is strongly inhibited by glucose [36-38]. The second product of the glucose-fructose oxidoreduction, gluconolactone, is quickly hydrolyzed by a gluconolactonase (EC 3.1.1.17) [35] and channelled into the EntnerDoudoroff pathway by a gluconate kinase (EC 2.7.1.12) [39]. A reversal of the enzymatic reaction for sorbitol formation seems improbable, since the lactone intermediate is highly unstable [34]; this would explain why Z. mobilis is unable to utilize sorbitol as a carbon source [1,31]. 3.3. Mannitol dehydrogenase Cells cultured in the presence of 150 g/1 fructose form small amounts of mannitol, corresponding to only 2.5% of the fructose metabolized [22]. In cell-free extracts, a NADP-dependent mannitol dehydrogenase activity (Fig. 1 (3)) was detected which was specific for fructose reduction and for mannitol oxidation [18]. It may be that this NADPH-dependent reduction of fructose is coupled to the dehydrogenation of glucose 6-phosphate, since the glucose-6-phosphate dehydrogenase present in Z. mobilis is active with N A D P as well as with N A D [38,40,41]. The high K m value of 180 mM for fructose of the crude mannitol dehydrogenase shows that detectable amounts of mannitol in culture supernatants would only be found at high fructose concentrations. The requirement for NADP(H) and the fact that the p H optimum for the reduction of fructose was at 8 but that for the oxidation of mannitol was at 10 [18] may help to explain why Z. mobilis cannot grow on mannitol in the p H range of 5 - 6 usually employed for fermentations [1,34]. It is not known
whether the mannitol dehydrogenase is constitutive or is induced by fructose. 3.4. Glucose dehydrogenase With ferricyanide or 2,6-dichlorophenolindophenol as artificial electron acceptor, a glucose oxidizing activity can be measured in cell-free extracts [34,35] (Fig. 1 (7)). Other aldohexoses and the sugar alcohols mannitol and sorbitol are also oxidized, although at lower rates [34] (Strohdeicher, M. et al. 1987, unpublished results). This enzyme activity is localized in the membrane fraction (Scopes, R.K., personal communication; Strohdeicher, M. et al. 1987, unpublished results) and seems to be similar to non-nucleotide-linked glucose dehydrogenases from other microorganisms. The latter enzymes have been identified as quinoproteins [42]. The in vivo end-acceptor of the electrons derived from glucose by the glucose dehydrogenase in Z. mobilis is not yet known.
4. E F F E C T OF O X Y G E N Aeration of cultures of Z. mobilis also leads to a diversion of reducing power and a consequent decrease in ethanol production [22,43-46]. Furthermore, the ability of Z. mobilis to grow in the presence of oxygen is not correlated with higher cell yields or with higher growth rates as compared to anaerobic conditions [22,43-46]. With increasing rates of aeration and concentrations of the carbon source, growth becomes strongly inhibited [44-46] and cell yields decrease [44,45]. 4.1. NADH-oxidase Z. mobilis was found to possess a N A D H - and NADPH-oxidase [44,45], located in the cell membrane, which catalyzes the oxidation of NAD(P)H with oxygen at a molar ratio of 2 : 1. Therefore, it can be assumed that oxygen is reduced to water by this reaction. The activity of N A D H oxidation accounted for the oxygen consumption rate of cell suspensions in the presence of a carbon source [44]. The cytochrome content of the cell membrane [43,45] and the inhibition by cyanide of both intracellular oxygen reduction by the NADH-oxidase and oxygen consumption by whole
135 Table 1 Influence of oxygen on growth and product formation of
Zymomonas mobilis Culture conditions *
Ethanol (g/l)
Acetaldehyde (g/l)
Growth yield g cell dry w e i g h t / mol glucose
Anaerobiosis Aerobiosis
63.4 20.1
0.24 1.31
2.3 1.4
* Continuous cultures were at D = 0.08 h -1 and with 14%
(w/v) glucose.
cells led to the suggestion that a respiratory chain is present in Z. mobilis [43,44]. Obviously, the transfer of electrons via this respiratory chain is not coupled with oxidative phosphorylation. In addition to the NAD(P)H-oxidase, catalase, superoxide dismutase and peroxidase were detected in Z. mobilis [1,44,45]. Thus, the observed toxicity of oxygen must be caused by a depletion in the pool of reduced coenzymes via the NAD(P)Hoxidase. As a consequence, acetaldehyde, a toxic intermediate of sugar catabolism accumulates (Table 1). In addition, acetic acid [43,44] and dihydroxyacetone [22] are formed as products during aerobic growth. A common feature of all the foregoing reactions, i.e., those catalyzed by glucose-fructose oxidoreductase, mannitol dehydrogenase, glucose dehydrogenase and NAD(P)H-oxidase, is that they cause a redox imbalance in the sugar catabolism of Z. mobilis. Glucose-fructose oxidoreductase and glucose dehydrogenase both lead to the formation of gluconic acid which is phosphorylated and enters the Entner-Doudoroff pathway after the NAD(P)H-yielding reaction catalyzed by glucose6-phosphate dehydrogenase. Thus only 1 mol of reduced coenzyme is formed by the glyceraldehyde-3-phopshate dehydrogenase for each mol of gluconic acid catabolized. Mannitol dehydrogenase and NAD(P)H-oxidase directly consume reducing power. These diversions of reducing power result in an accumulation of products which are more oxidized than ethanol, namely acetaldehyde, acetoin and acetic acid [18,22,33,43]. Surprisingly, dihydroxyacetone and glycerol were also formed, mainly in cultures growing at high fructose concentrations [18,22]. Dihydroxyacetone
is derived from glyceraldehyde 3-phosphate by the sequential action of triose phosphate isomerase [18,47] and a phosphatase, and it can be assumed that glycerol is formed by further reduction of dihydroxyacetone phosphate to glycerol phosphate followed by hydrolysis to glycerol [18]. The formation of both of these products, dihydroxyacetone and glycerol, increases the redox imbalance, since for each glyceraldehyde phosphate that is not converted to diphosphoglycerate, one NADH is lost. An additional reducing equivalent has to be consumed to form glycerol. The cause for the accumulation of dihydroxyacetone and glycerol is thus not readily apparent in a situation of NADH deficiency and poses an interesting problem for further investigation. Up to now, no physiological roles can be attributed to sorbitol and mannitol formation, or to the membranebound NAD(P)H-oxidase and glucose dehydrogenase of Z. mobilis.
5. EFFECTS OF ETHANOL Among bacteria, Z. mobilis has an exceptionally high ethanol tolerance and is capable of producing ethanol in concentrations up to 13% (w/v) [6]. Despite this fact, end-product inhibition by ethanol is clearly the main limiting factor for the overall efficiency of the fermentation process [48-50]. A search for the primary site of ethanol inhibition has led to a number of investigations on how ethanol affects catabolic enzymes and the cell membrane. Recent work of Scopes and coworkers [47,51] has shown that a cell-free system of Z. mobilis can rapidly consume glucose and produce ethanol concentrations exceeding 15% w/v. These results demonstrate that the Entner-Doudoroff enzymes are more resistant to ethanol than are the living cells [51]. Much evidence has been accumulated to support the hypothesis that the cell membrane is the primary target for ethanol inhibition. Ethanol is known to affect the integrity of biological membranes by intercalation into t h e lipid bilayer. This results in a disorganization of the cell membranes which is in some respects similar to the heat-induced increase of membrane fluidity. Leakage of cofactors and coenzymes
136 through the plasma membrane has been reported as a consequence of ethanol- and heat-damage [52,53]. The reviews of Ingrain and Buttke [54] and Ingrain [55] give a comprehensive overview of various aspects of the action of alcohols on microorganisms.
5.1. Fatty acid and phospholipid patterns In the case of Z. mobilis, specific mechanisms should then exist to counteract the adverse effects of ethanol on the cell membrane. As was shown by several groups, the major fatty acids occurring in Z. mobilis are myristic (14 : 0), palmitic (16 : 0) and cis-vaccenic acid (18 : 1). There is a very high level of vaccenic acid, accounting for up to 70% of the total fatty acids [56-60]. Among the phospholipids (phosphatidylethanolamine, phosphatidylglycerol, phosphatidylcholine, cardiolipin, and phosphatidylserine), phosphatidylethanolamine is the most abundant [57-59]. The fatty acid pattern of batch cultures did not change significantly during growth and alcohol accumulation, and only small shifts in the phospholipid pattern were observed [59,61].
4"'.
4"'.
©
@
G HO
OH
OH
T : i"
l
I
I
OH
NHX OH
tt~/~ 0
f
5.2. Ethanol- and temperature-dependent synthesis of hopanoids and proteins Another important class of lipids present in Z. mobilis are the hopanoids [57,58,60]. It has been suggested that these pentacyclic triterpenoids are the prokayrotic structural and functional equivalents of sterols in eukaryotes; sterols are known to be partly responsible for the high alcohol tolerance of yeast [62,63]. The biosynthesis of hopanoids, in contrast to that of sterols, does not require oxygen [64]. The hopanoid patterns of Z. mobilis ATCC 29191 and Z. mobilis ATCC 10988 are similar [60]; both strains contain hopene, hopanol [58], bacteriohopanetetrol [60], and the glycoside and ether derivatives of bacteriohopanetetrol [65] (Fig. 2). Bacteriohopanetetrol and its derivatives are not discriminated from each other when analyzed by gas-liquid chromatography, after derivatization [65]. A number of other triterpenoids, described by Tornabene et al. [57], probably occur only as minor components. Analysis by gas-liquid chromatography shows that the components analyzed as bacteriohopenetetrol represent the major hopanoids [60]. It is significant that the contribution of bacteriohopanetetrol to
~
OH
NH 2
2 Ho
[
OH
/ CH~OH ~°o.
@
~'.,
®
Fig. 2. Hopanoids from Zymomonasmobilis. 1, hopene; 2, diplopterol; 3, bacteriohopane-tetrol; 4, bacteriohopanetetrol ether; 5, glucosaminylbacteriohopanetetrol.
137 the total lipid fraction depends strongly on ethanol concentration, as was shown with cells grown under continuous or batch conditions [60,66]. In cells from continuous cultures, with an endogenous ethanol concentration of 5 g/l, the amount of bacteriohopanetetrol was 2.5% of the total lipids whereas, at an ethanol level of 63 g/l, this amount increased to 21% of the total lipids. Likewise, the amounts of hopene and hopanol increased from 0% and 0.9% to 1% and 2.9% of the total lipids, respectively [60]. A strong response of the hopanoid content of cells was also observed when cell suspensions were incubated in media with added ethanol. In cells exposed to 160 g/1 ethanol, bacteriohopanetetrol comprised 36.5% of the total lipids whereas in cells incubated without added ethanol, the corresponding value dropped to just 9.5% [66]. Besides ethanol, n-propanol and nbutanol also increased the relative hopanoid content [66]. Stimulation of the hopanoid biosynthesis seems to be mediated through changes in membrane fluidity, since a change in incubation temperature from 30 to 37°C leads to the same increase in hopanoid content as high concentrations of ethanol [66]. The large shifts in the contribution of hopanoids to the total lipid content raise the question of whether the amount of total lipids per cell is also variable, or whether it remains constant. In the first instance, the ethanol- and temperature-dependent synthesis of hopanoids could proceed irrespective of other membrane components, and consequently the amount of total lipids per cell would increase. Alternatively, the cell could compensate for increasing amounts of hopanoids by decreasing the synthesis of other membrane components. Investigations of Benschoter and Ingram [53] on the thermal tolerance of Z. mobilis support the second assumption. In their study the ratio of phospholipids to total cell protein and membrane protein strongly decreased with increasing growth temperature. The phospholipid-to-protein ratio of membranes from cells grown at 41°C was approximately half that of membranes from cells grown at 20 ° C. In addition it was shown by measurements of the fluorescence depolarization of diphenylhexatriene that, with increasing growth temperature, the membranes from Z. mobilis were progressively more rigid [53]. This
observation is in accordance with the properties predictable for membranes containing increasing amounts of hopanoids [63,67]. Even apart from this proposed partial substitution of hopanoids for phospholipids the increase in the relative protein content of the membranes [53,59] provides an additional means to counteract membrane fluidization at elevated temperature. Ethanol and temperature not only have parallel effects on the distribution of lipids but also on the protein pattern of Z. mobilis [68,69]. In cells incubated for 10 rain either in the presence of 5% (v/v) ethanol at 30°C, or without ethanol at 45°C, the synthesis of several polypeptides was induced or stimulated as compared to untreated ceils. Among these polypeptides, two were found associated with the envelope fraction. Interestingly, heat- and ethanol-shocked cells showed a significantly enhanced viability during subsequent heat treatment [69]. Although the physiological role of the proteins induced by heat and ethanol is not known at present, an involvement in the mechanism of ethanol tolerance of Z. mobilis can be assumed. 5.3. Significance of hopanoids A vital function, however, can be attributed to the hopanoids themselves. That hopanoids are indispensable for Z. mobilis was shown by Flesch and Rohmer [70] who used inhibitors of squalene hopene cyclase. These substances are squalene analogues that inhibit growth at concentrations of 1-3 #M. The squalene cyclase which catalyzes the cyclization of squalene into hopanoids [71-73] seems to be the main regulatory enzyme of hopanoid synthesis in Z. mobilis (Schmidt, A., unpublished). The mechanism by which ethanol tolerance is achieved in Z. mobilis appears to be a concerted shift in the relative amounts of phospholipds, hopanoids and proteins in the cell envelope. The large ethanol-dependent shifts in the hopanoid content suggest a major function of these sterollike substances for membrane stabilization. For further studies on the influence of the fatty acid composition of phospholipids, the incorporation of exogenous phospholipids by interaction of Z. mobilis cells with liposomes [74] may prove to be
138 useful. Attempts to alter the fatty acid composition of Z. mobilis by adding fatty acid supplements during growth have not yet been successful [59].
enzymes [51]. The view that substantial sugar concentrations exist intraceUularly in Z. mobilis at high external sugar concentrations is further supported by the occurrence of enzymes which have extremely low affinities for their substrates, e.g., glucose-fructose oxidoreductase [35].
6. OSMOTOLERANCE In their comprehensive, comparative study of about forty strains of Z. mobilis, Swings and DeLey [1] stated that all these strains were able to grow with 20% glucose and most strains even grew in media containing 30% or 40% glucose. Such high sugar concentrations decrease the total water concentration and exert osmotic pressures which are comparable to relatively strong salt solutions. For example, a 20% (w/v) glucose solution has approximately the same osmotic pressure as a 4.2% (w/v) solution of sodium chloride. However, the salt resistance of Z. mobilis is rather low. 71% of the strains tested by Swings and DeLey [1] grew in the presence of 1% NaC1, whereas none could grow in the presence of 2% NaCI. These results suggest that the salt intolerance of Z. mobilis is not a mere problem of increased osmotic pressure and decreased water activity, but is mainly due to effects specific for electrolyte solutes. 6.1. Tolerance towards highly concentrated sugar solutions A possible explanation for the tolerance of Z. mobilis towards highly concentrated glucose solutions may be derived from studies on glucose transport [75,76] and on the activity of a cell-free system at high glucose concentrations [47,51]. DGlucose is transported by a high-velocity, carriermediated facilitated diffusion system, whereby its intracellular concentration quickly reaches a plateau close to the external concentration [76]. Studies with a cell-free system showed that concentrated glucose solutions were not inhibitory to the Entner-Doudoroff enzymes, since conversion of 2 M (36%) glucose to ethanol by this system proceeded rapidly [47,51]. Thus, the extracellular osmotic pressure of glucose solution may rapidly be balanced by corresponding intracellular sugar concentrations. Reduced water activity seems to have only minor effects on the Entner-Doudoroff
6.2. Salt intolerance The low salt tolerance of Z. mobilis poses problems for the fermentation of molasses which usually have a high salt content [12,77,78]. Under normal batch conditions, Z. mobilis grew poorly on molasses due to salt inhibition [12,78]. It is likely, however, that this problem can be overcome by genetic improvement of Z. mobilis [7,11] and by optimized fermentation technologies for molasses [12,78].
7. CONCLUSIONS Z. mobilis performs a highly productive ethanol fermentation and offers a number of advantages over the traditional yeast fermentation. As was demonstrated by Rogers and collaborators [6] the specific rates of sugar uptake and ethanol production for Z. mobilis are 3-4-times faster than those for yeasts. In addition, higher ethanol yields and lower biomass yields are obtained with Z. mobilis, compared to yeasts, due to the differences in carbohydrate metabolism. Biosyntheses of membrane components, which have important functions in the exceptionally high ethanol tolerance of Z. mobilis, proceed independently of the presence of oxygen [64]. The ability of this bacterium to grow rapidly in the absence of oxygen simplifies the conditions of continuous fermentation processes. Thus, Z. mobilis is a promising microorganism for industrial ethanol production, especially from starch-based substrates [12,79-81]. Perhaps the greatest obstacle to commercialization is that, compared to yeast, much less practical experience exists for the Z. mobilis fermentation. The extensive work done in the field of evaluation and optimization of fermentation techniques (recent publications: [82-99]) reflects the neccessity for the development of Z. mobilis-specific processes, especially those that can operate stably
139 in a n o n - s t e r i l e e n v i r o n m e n t . T h e n a r r o w r a n g e of s u b s t r a t e s utilized b y Z. mobilis (i.e., glucose, fructose a n d sucrose), the f o r m a t i o n of b y - p r o d ucts in fructose a n d s u c r o s e - c o n t a i n i n g m e d i a a n d the low salt t o l e r a n c e are d i s a d v a n t a g e o u s for c o m m e r c i a l a p p l i c a t i o n s . However, this b a c t e r i u m is p e r h a p s m o r e r e a d i l y m o d i f i e d b y genetic eng i n e e r i n g techniques t h a n yeast (for reviews see [7,9,11]). R e c e n t p u b l i c a t i o n s c o n c e r n the cons t r u c t i o n of vectors a n d transfer o f p l a s m i d s in Z. mobilis [100-111]. F u r t h e r m o r e , a n u m b e r o f p r o teins have b e e n p u r i f i e d f r o m Z. mobilis, m o s t r e c e n t l y the E n t n e r - D o u d o r o f f e n z y m e s [112], the two alcohol d e h y d r o g e n a s e s [113] a n d the p y r u v a t e d e c a r b o x y l a s e [114-117]. T h e p y r u v a t e dec a r b o x y l a s e gene h a s b e e n c l o n e d a n d s e q u e n c e d [116-119]. It seems p r o b a b l e that b y genetic m a n i p u l a t i o n , the Z. mobilis f e r m e n t a t i o n c a n b e f u r t h e r i m p r o v e d with r e g a r d to the r a n g e of u t i l i z a b l e substrates, salt t o l e r a n c e a n d the byp r o d u c t s f o r m e d f r o m sucrose or invert sugar.
ACKNOWLEDGEMENTS T h e a u t h o r s g r a t e f u l l y a c k n o w l e d g e the critical r e a d i n g of the m a n u s c r i p t b y Professor R . K . F i n n .
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