Abstract

Candida versatilis (halophila) CBS4019 was chosen to study the physiological reactions of long-term exposure to extremely high salt concentrations. In general, our results show a significant increase in enzyme expression during growth under stress conditions. Although glycerol and mannitol pathways are not under glucose repression, they were found to be metabolically regulated. Glycerol-3P-dehydrogenase used either of its cofactors NADPH or NADH, being in favor of NADPH during growth with high salt concentrations. This ability of interchanging cofactors, an increased fermentation rate, and the observed mannitol pathway activity are suggested to contribute to the yeasts’ redox stability. Enzymes per se were not salt-tolerant in vitro. Consistently, intracellular sodium was low and intracellular potassium, a requirement for growth, was high. The concept of halophily and its applicability to yeasts is discussed.

Introduction

Physiological mechanisms underlying extreme salt tolerance in yeasts are still poorly understood. Although some work has been done concerning Debaryomyces hansenii and Zygosaccharomyces rouxii, most of the studies involve Saccharomyces cerevisiae, which is a moderately salt-tolerant yeast. Yet, still no coherent picture of the main mechanisms of its response to salt stress has been obtained. Much more information is needed to understand how yeast cells can survive in high salt concentrations, especially near the limit of salt solubility. It has been extensively discussed in the literature that yeast cells have two main mechanisms for dealing with salt stress: (1) the activation of systems which are able to efficiently expel ions from the cell, and (2) the production and retention of osmolytes, thereby deviating cell components from growth to maintenance [1].

The aim of the present study was to acquire more knowledge about the physiological mechanisms of long-term resistance to salt stress in an extremely halotolerant yeast. Candida versatilis (syn. halophila) CBS4019 was chosen for its ability to grow in the presence of NaCl concentrations up to 5 M. When we started working with this yeast [2] it was the only strain of its species [3]. It was subsequently re-classified as a strain belonging to another yeast species, C. versatilis[4]. Since its type strain was proven to be different in several aspects from C. halophila CBS4019 used in this work [2,4,5], we decided to conserve the former designation.

Previous work [5] has revealed that C. halophila growing on glucose produces glycerol and mannitol. Glycerol was proven to be the osmolyte, though the quantities accumulated were smaller than in other yeasts under the same degree of salt stress [5]. The function of mannitol is still unknown. Their putative transport systems, which most probably contribute to the osmolytes’ retention under stress (as suggested before by Lages et al. [2]), have been characterized. They also appeared to have the more fundamental function of feeding the cells, since C. halophila grew equally well on glycerol, mannitol and glucose as the single carbon and energy sources. Intracellular pH values were constant, regardless of the salt concentration in the growth medium. The intracellular volume decreased during a salt shock, but recovered during growth, being even slightly higher as compared to cultivation in the absence of salt [5]. All these studies were performed in cells growing exponentially with NaCl concentrations up to 4.5 M, showing a surprising stability for cells cultivated under such extreme salt stress [5].

In this article we present further physiological studies, that, presumably, will help to better understand the mechanisms underlying long-term extreme salt tolerance during growth, thereby focussing on metabolism. Given the fact that this is a huge task to comprise in detail, we made some choices to address the issue. Our approach includes: (1) study of glycerol and mannitol pathways; (2) measurement of fermentative and respiratory fluxes; (3) in vitro assays of several enzymes from glycolysis and Krebs cycle; (4) measurement of intracellular bulk redox equivalents; and, finally, (5) quantification of intracellular sodium and potassium concentrations.

All experiments, as previously [5], were performed in cells cultivated with a range of salt concentrations up to 4 M NaCl. When considered necessary, a comparison was made with salt shock response under identical conditions. With these results in mind, a putative model of metabolic pathway regulation under stress is discussed.

Moreover, the applicability of the classification ‘halophily’ with respect to yeasts will be discussed. Discrepancies between measurements of intracellular sodium and potassium concentrations in yeasts can be found in the literature [6–13]. This stresses the influence of experimental methodology, and of the physiological state of the cells on these determinations, which is a major inconvenience, since the amount of ions that cells can tolerate inside without growth arrest has been the most common argument in this type of classification. Regardless, however, of the intracellular salt concentration, we still know too little concerning the intracellular environment and in particular concerning the intracellular compartmentation of soluble elements. There is no doubt, though, that it is substantially different from the environment created in in vitro enzyme assays. Unlike enzymes from halophilic bacteria [14], enzymes from halotolerant yeasts assayed in vitro do not tolerate more than some mM of salt [11,15]. Our results contribute to make a point in this discussion.

Physiological knowledge will help to elucidate long-term physiological and biochemical stress adaptation mechanisms [16]. It may also open some ways to enable the utilization of molecular approaches in extremely salt-tolerant eukaryotes. Finally, this type of information can be important for current industrial applications [17].

Materials and methods

Yeast strains and growth conditions

C. halophila CBS4019 as well as S. cerevisiae W303-1A [18], D. hansenii CBS767 and Pichia sorbitophila CBS7064 were conserved and cultivated in batch cultures as described by Silva-Graça and Lucas [5]. Cells were cultured in media containing a range of NaCl concentrations and collected in mid-exponential growth phase.

Measurement of respiratory and fermentative fluxes

Cells were grown, harvested in mid-exponential growth phase by centrifugation in a model 4K10 centrifuge (B. Braun, Melsungen, Germany), washed twice and resuspended in ice-cold distilled water containing the same amount of sodium chloride as the growth medium. The final cell suspension had a concentration of 25–30 (mg dry wt) ml−1. Respiration was monitored using an O2-selective electrode as described in Neves et al. [11]. The specific respiration rate was expressed as μmol O2 consumed min−1 (g dry wt)−1. CO2 production rates were determined using a CO2-selective electrode (Radiometer, Copenhagen, Denmark E5036-0). The electrode was coupled to a standard pH/mV meter (Radiometer, PHM 82), connected to a flat recorder (Kipp and Zonen, Delft, The Netherlands) and CO2 liberation was monitored in a chamber with temperature control and gentle magnetic stirring. The reaction mixture contained 4.5 ml of growth medium without carbon source and 0.25 ml of cellular suspension (25 mg ml−1). As soon as the mV value had stabilized, a starter of 0.25 ml of 2 M glucose was added. The concentration of CO2 released with time was determined within the linear slope of mV variation, using a logarithmic calibration curve made according to the electrode supplier instructions. The specific CO2 formation and the fermentation rate were estimated by indexing to biomass dry weight determinations: μmol CO2 released min−1 (g dry wt)−1.

Preparation of cell-free extracts

Cultures (100 ml) in mid-exponential growth phase, with an OD640 of 0.7–0.8, were harvested as stated above and resuspended in 1 ml 10 mM triethanolamine buffer (pH 7.5), containing 1 mM dithioerythritol and 1 mM ethylenediamine tetraacetic acid (EDTA). Cells were disrupted by at least 15 cycles of 1 min hand shaking on a vortex mixer, followed by a 1-min interval rest in ice, using 1 g of 0.5-mm diameter glass beads. Supernatants were separated by centrifugation at 15 000 rpm for 30 min at 4°C, and kept on ice for a short period of time prior to the assay.

Enzyme assays

The spectrophotometric assays to determine enzyme activity were performed in a Perkin-Elmer Lambda 2 UV-VIS spectrophotometer connected to an Epson FX-850 printer, in 1-ml polystyrene cuvettes. Protein concentration was determined by the method of Lowry, modified by Peterson [19], using bovine serum albumin as standard. The production of oxidized or reduced NAD and NADP was followed by measuring the absorbance at 340 nm. The absorption coefficients used to calculate enzyme activity were 6.22×103 M−1 cm−1 for reduced NADH, and 6.20×103 M−1 cm−1 for reduced NADPH. In each new assay, linear increase in activity with increasing amounts of cell extract was verified. Coupling enzymes, cofactors and substrates were obtained from Sigma (St. Louis, MO, USA). In all cases it was checked whether the correspondent quantities used were not limiting the reaction. Assays were performed at 25°C.

Glycerol-3-phosphate dehydrogenase/NAD+ (EC 1.1.1.8)/glycerol-3-phosphate dehydrogenase/NADP+ (EC 1.1.1.94). Activity was determined using the method described by Adler et al. [9], with some modifications. The assay mixture contained 20 mM imidazole buffer (pH 7.0), 1 mM MgCl2, 200 mM NaCl and 0.2 mM NADH or NADPH. The reaction was started with 5 mM dihydroxyacetone phosphate.

Glycerol-3P-phosphatase (EC 3.1.3.28) was determined by methods previously described [20–22], with some modifications. The assay mixture contained 20 mM Tricine buffer (pH 7.0) and 5 mM MgCl2. The reaction was started with 10 mM d,l-glycerol-3-phosphate. Samples of 90 μl were withdrawn with 20-s intervals, and immediately added to 10 μl of 50% (w/v) HClO4 to stop the reaction. These samples were kept on ice until phosphate quantification. This was performed using a calibration curve made with concentrations of inorganic phosphate ranging from 0.01 to 1 mM. Each sample was treated by the addition of 900 μl of a solution containing six parts of ammonium molybdate solution (4.2 g (NH4)6Mo7O24·4H2O and 1 l 0.5 M H2SO4) and one part of 10% (w/v) ascorbic acid solution freshly prepared. The samples were incubated at 45°C for 20 min and then the absorbance at 820 nm was read. The specific activity was determined using the slope of the curve of phosphate release per time.

Dihydroxyacetone kinase (EC 2.7.1.28) was measured using the method described by Babel et al. [23], with some modifications. The assay mixture contained 50 mM imidazole buffer (pH 7.5), 20 mM MgCl2, 0.13 mM NADH, 0.5 mM dihydroxyacetone, 10 U of glycerol-3-phosphate dehydrogenase. The reaction was started with 50 mM ATP.

Glycerol kinase (EC 2.7.1.30) assays were tried using the methodology described by Lin et al. [24], with some modifications. The assay mixture contained 100 mM glycine buffer (pH 9.5), 20 mM MgCl2, 300 mM hydrazine sulfate, 1.4 mM NAD+, 10 mM glycerol and 10 U of glycerol-3-phosphate dehydrogenase. The reaction was started with 100 mM ATP.

Glycerol dehydrogenase/NAD+ (EC 1.1.1.6)/glycerol dehydrogenase/NADP+ (1.1.1.72) was measured by the method of Lin and Magasanik [25], with some modifications. The assay contained 100 mM glycine buffer (pH 9.5), 35 mM (NH4)2SO4 and 0.2 mM NAD+ or NADP+. The reaction was started with 550 mM of glycerol.

Mannitol dehydrogenase (EC 1.1.1.67) was measured using the methodology described by Ueng et al. [26].

Mannitol-1-phosphate dehydrogenase (EC 1.1.1.17) was measured by the method of Boosaeng et al. [27], with some modifications. The reaction mixture contained 50 mM Tris buffer (pH 7.0) and 0.2 mM NADH. The reaction was started with 10 mM fructose-6-phosphate.

Hexokinase (EC 2.7.1.1) was measured using the method as described by Hirai et al. [28].

Glucose-6-phosphate dehydrogenase (EC 1.1.1.49) was measured by the method of Kuby and Noltmann [29].

Glyceraldehyde-3-phosphate dehydrogenase (EC 1.2.1.12) was measured according to Maitra and Lobo [30].

Alcohol dehydrogenase (EC 1.1.1.1) was measured following Postma et al. [31].

Malate dehydrogenase (EC 1.1.1.37) was measured by the method of Witt et al. [32].

Isocitrate dehydrogenase/NAD+ (EC 1.1.1.41)/isocitrate dehydrogenase/NADP+ (EC 1.1.1.42) was measured using the methodology described by Stueland et al. [33], with some modifications. The reaction mixture contained 40 mM MOPS buffer (pH 7.5), 5 mM MgCl2 and 1 mM NADP+. The reaction was started with 0.5 mM isocitrate.

Southern blot analysis

The probe was synthesized by polymerase chain reaction (PCR), using S. cerevisiae W303-1A genomic DNA as template and oligonucleotides designed for GUT1 sequence, available at the S.G.D. (Stanford University, CA, USA) with the following sequences: forward, 5′-CCTACAGTCTTGCCCTCCAC-3′; reverse, 5′-AACATTCCCGCAACACTTTC-3′. The resulting 756-bp fragment was gel-eluted, using high pure PCR product purification kit (Roche, Rotkreuz, Switzerland) and labelled with AlkPhos direct kit (Amersham Pharmacia Biotech, Sunnyvale, CA, USA). Southern blot was done using standard procedures [34]. C. halophila and S. cerevisiae genomic DNA were partially digested with Sau3AI to a nylon membrane, Gene Screen Plus (NEN Life Sciences Products), and allowed to hybridize overnight at different temperatures: 45, 55 and 60°C. Detection was performed using AlkPhos direct kit (Amersham Pharmacia Biotech).

Verification of the in vitro NaCl toxicity towards glyceraldehyde-3P-dehydrogenase

Salt toxicity in in vitro enzyme assays was performed according to Neves et al. [11]. The degree of inhibition of activity was quantified using the equation generally used to quantify ethanol inhibition for transport systems [35]. Accordingly, an exponential inhibition constant (Ki) for NaCl, KNaCl, was determined, as well as the minimum concentration above which the inhibition could be detected, Cmin.

Estimation of intracellular concentrations of reduced and oxidized enzyme cofactors

Cells were cultivated as described above. The cultures were divided into two parts and submitted to acid or alkaline extraction, after which the cell suspensions were centrifuged (13 000 rpm for 10 min at 4°C). The supernatants were used immediately for enzymatic determination of intracellular concentrations of NAD+, NADH, NADP+ and NADPH, using the methods described by Bergmeyer [36]. Results were presented using the following indices: PNF (phosphorylated nucleotide fraction), [NADP++NADPH]/[NAD++NADH]; CRC (catabolite reduction charge), NADH/[NAD++NADH]; ARC (anabolic reduction charge), NADPH/[NADP++NADPH] [37].

Estimation of intracellular concentrations of Na+ and K+

Cultures (500 ml) were harvested by centrifugation with a model 4K10 centrifuge (B. Braun, Germany) and washed three times for 2 min at 9000 rpm with 1 M ice-cold sorbitol solution containing 4 g l−1 MgCl2. Cells were digested with 10 ml of nitric acid (4 ml l−1), during 16 h, with gentle agitation. Cell debris was separated by centrifugation for 10 min at 4°C. Supernatant was filtered through 0.45-μm cellulose acetate filters (Schleicher and Schuell, Dassel, Germany). The samples were analyzed by atomic absorption spectroscopy (AAS) in a Varian Spectra AA.250 PLUS spectrometer in absorption mode. To avoid potassium and sodium ionization, 1 g l−1 CsCl was added to the samples for the determination of [K+] and 2 g l−1 KCl for the determination of [Na+]. Sample duplicates were used to confirm some of the results using an ion-coupled plasma (ICP) spectrometer, PU 7000 (Unicam), as in Neves et al. [11], as well as by 39K and 23Na nuclear magnetic resonance (NMR) (see Acknowledgements).

Results

Glycerol and mannitol metabolism

C. halophila growing exponentially on glucose produces approximately equal amounts of glycerol and mannitol [5]. Moreover, both compounds can be used as single carbon and energy sources, yielding very similar growth parameters [5]. As a first step for the characterization of glycerol and mannitol metabolism, cells were cultured on mixtures of equivalent molarities of glucose/glycerol (Fig. 1A), glucose/mannitol (not shown), and glycerol/mannitol (Fig. 1B), and substrate consumption was followed in time. Both glycerol and mannitol were consumed only after the glucose concentrations reached a level around 0.1–0.2%, indicating sequential growth. This suggests that glycerol and mannitol pathway enzymes might be under glucose regulation. Glycerol and mannitol were consumed simultaneously, although lagged behind (Fig. 1B).

1

Substrate consumption and culture growth (OD) of C. halophila cultivated at 30°C in MM with glucose/glycerol (A) and glycerol/mannitol (B) mixtures of similar molarity.

1

Substrate consumption and culture growth (OD) of C. halophila cultivated at 30°C in MM with glucose/glycerol (A) and glycerol/mannitol (B) mixtures of similar molarity.

All the enzymes of the glycerol and mannitol pathways were assayed in vitro in cell-free extracts from cells cultivated with glucose, glycerol, or mannitol. We basically made use of protocols previously established for the pathways to be determined, in S. cerevisiae for glycerol, in filamentous fungi for mannitol (see Section 2).

Specific activities of glycerol-3P-dehydrogenase (G3PD), glycerol-3P-phosphatase (G3P), glycerol dehydrogenase (GD) and dihydroxyacetone kinase (DK) were determined (Table 1). G3PD was able to use either NADH or NADPH as a cofactor with equal specific activity, which raises the question whether this is the result of one enzyme or two isoenzymes (Table 1). Mitochondrial glycerol-3P-dehydrogenase (G3PDmit) was hardly detectable, presenting an extremely low activity both in glucose- or glycerol-grown cells (not shown). Furthermore, we were not able to measure activity of glycerol kinase in cells grown on either substrate. Since previous references have reported the difficulty of measuring in vitro glycerol kinase activity [9,38], we decided to verify the existence of this enzyme by Southern blot. We constructed a probe based on the sequence of GUT1 (glycerol kinase) from S. cerevisiae. The probe gave, as expected, a good signal against S. cerevisiae DNA at all hybridization temperatures tested (not shown), but no signal against C. halophila DNA. Knowing that the kinase from another halotolerant yeast, D. hansenii, has been characterized and cloned (though no sequence is available for comparison) [38] we probed D. hansenii DNA, which also gave negative results. The same result was obtained with P. sorbitophila DNA. This suggests a rather high heterogeneity at nucleotide level of the glycerol kinase genes in different yeasts. These experiments did not provide conclusive results for the absence or presence of this enzyme in C. halophila.

1

Specific enzyme activities of the glycerol pathway in cells of C. halophila cultivated in MM with 2% (w/v) of either carbon source

Carbon source Glycerol-3P-dehydrogenase Glycerol-3P-phosphatase Dihydroxyacetone kinase Glycerol dehydrogenasea 
 NADPH NADH   NADP+ Glycerol consumption NADPH Glycerol production 
Glucose 65.8±10.3 (3) 56.8±12.1 (23) 79.4±26.0 (7) 35.2±8.5 (9) 13.3±9.1 (19) 52±19 (5) 
Glycerol n.d. 44.3±12.2 (16) 91.4±26.9 (8) 52.7±20.2 (8) 109±31.7 (17) 138±75 (6) 
Mannitol n.d. 89.7±25.4 (7) 102.7±27.6 (4) 72.4±1.5 (3) 228±77.8 (8) 238±92 (4) 
Carbon source Glycerol-3P-dehydrogenase Glycerol-3P-phosphatase Dihydroxyacetone kinase Glycerol dehydrogenasea 
 NADPH NADH   NADP+ Glycerol consumption NADPH Glycerol production 
Glucose 65.8±10.3 (3) 56.8±12.1 (23) 79.4±26.0 (7) 35.2±8.5 (9) 13.3±9.1 (19) 52±19 (5) 
Glycerol n.d. 44.3±12.2 (16) 91.4±26.9 (8) 52.7±20.2 (8) 109±31.7 (17) 138±75 (6) 
Mannitol n.d. 89.7±25.4 (7) 102.7±27.6 (4) 72.4±1.5 (3) 228±77.8 (8) 238±92 (4) 

Activity is expressed as mU (mg protein)−1 (U=μmol min−1).

n.d., not determined.

Number in parentheses, number of independent assays.

a

Activity of glycerol dehydrogenase was not detected using NAD(H).

We tested mannitol-1P-dehydrogenase (M1PD), mannitol-1P-phosphatase (M1P), mannitol dehydrogenase (MD) and hexokinase (HxK) (Table 2). Hexokinase can function dually, either using glucose as a substrate in the first step of glycolysis, or converting fructose in the mannitol pathway to fructose-6P. For this reason, the enzyme was assayed using both substrates (Tables 2 and 3). Mannitol kinase was not assayed, in view of the lack of detection of glycerol kinase mentioned above, and M1P activity could not be detected. Although some protocol adjustments were made, the possibility could not be disregarded that further modification would have been necessary for better detection. As can be seen in Fig. 2, the mannitol cycle is similar to the glycerol cycle, concerning the type of enzymatic steps, the major difference being the junction with glycolysis.

2

Specific enzyme activities of the mannitol pathway in cells of C. halophila cultivated in MM with 2% (w/v) of either carbon source

Carbon source Mannitol-1P-dehydrogenase Mannitol dehydrogenasea Hexokinase 
 NADPH NADH Mannitol consumption Mannitol production Substrate fructose 
   NAD+a NADPH NADH  
Glucose 14.8 (1) 1.48 (1) 15.5±0.8 (2) 23.1 (1) 1.12 (1) 117±14 (3) 
Glycerol n.d. n.d. 46.0±2.6 (3) 35.2±8.1 (2) 5.76 (1) 102±9 (2) 
Mannitol n.d. n.d. 64.5±7.6 (3) 32.7±10.0 (2) 6.5±3.0 (2) 145 (1) 
Carbon source Mannitol-1P-dehydrogenase Mannitol dehydrogenasea Hexokinase 
 NADPH NADH Mannitol consumption Mannitol production Substrate fructose 
   NAD+a NADPH NADH  
Glucose 14.8 (1) 1.48 (1) 15.5±0.8 (2) 23.1 (1) 1.12 (1) 117±14 (3) 
Glycerol n.d. n.d. 46.0±2.6 (3) 35.2±8.1 (2) 5.76 (1) 102±9 (2) 
Mannitol n.d. n.d. 64.5±7.6 (3) 32.7±10.0 (2) 6.5±3.0 (2) 145 (1) 

Activity is expressed as mU (mg protein)−1 (U=μmol min−1).

n.d., not determined.

Number in parentheses, number of independent assays.

a

Mannitol dehydrogenase activity was not detected using NADP+ when mannitol was used as the substrate.

3

Specific enzyme activities in cells of C. halophila cultivated in MM with 2% (w/v) of either carbon source

Carbon source Hexokinase Substrate glucose Glucose-6P-dehydrogenase Glyceraldehyde-3P-dehydrogenase Alcohol dehydrogenase Malate dehydrogenase Isocitrate dehydrogenase 
Glucose 160±8 (4) 225 (1) 421±80.6 (6) 31.7±5.7 (5) 3170±450 (4) 116 (1) 
Glycerol 165±14 (3) n.d. 381±98.0 (7) 53.7±13.7 (4) 2790±90 (3) n.d. 
Mannitol 195 (1) n.d. 566±95.9 (3) 96.1±6.4 (3) 2990±110 (2) n.d. 
Carbon source Hexokinase Substrate glucose Glucose-6P-dehydrogenase Glyceraldehyde-3P-dehydrogenase Alcohol dehydrogenase Malate dehydrogenase Isocitrate dehydrogenase 
Glucose 160±8 (4) 225 (1) 421±80.6 (6) 31.7±5.7 (5) 3170±450 (4) 116 (1) 
Glycerol 165±14 (3) n.d. 381±98.0 (7) 53.7±13.7 (4) 2790±90 (3) n.d. 
Mannitol 195 (1) n.d. 566±95.9 (3) 96.1±6.4 (3) 2990±110 (2) n.d. 

Activity is expressed as mU (mg protein)−1 (U=μmol min−1).

Number in parentheses, number of independent assays.

n.d., not determined.

2

Scheme of glycerol and mannitol production and dissimilation pathways as they were assayed in cell-free extracts from C. halophila, and correspondent glycolysis branching.

2

Scheme of glycerol and mannitol production and dissimilation pathways as they were assayed in cell-free extracts from C. halophila, and correspondent glycolysis branching.

The results in Tables 1 and 2 show that all enzymes involved are active in cells cultivated on either carbon source, thus being constitutively expressed, though not without some degree of regulation, since their specific activity is mostly higher in cells cultivated on glycerol or mannitol. Another observation concerns the dehydrogenase activities for glycerol (GD) consumption and mannitol (MD) consumption. These enzymatic reactions show different cofactor specificity. Using the same reaction mixture for GD and MD determination, we verified that glycerol exclusively activated the enzyme with NADP+ and mannitol with NAD+. This result raises the question of whether this is due to a single enzyme, able to use these substrates with different cofactors, or whether two enzymes are counterparting.

Enzyme activities in NaCl-grown cells

Cells were cultivated in the presence of salt concentrations, ranging from 0.5 to 4 M, with glucose as the carbon and energy source. Enzymes were assayed without the addition of salt to the assay mixture, except for the G3PD protocol, which, according to literature, uses some NaCl in the reaction buffer (Section 2).

In general, the specific activities of the glycerol pathway enzymes (G3P, GD and DK) exhibited a very significant increase with increasing salt concentrations (∼70–230% increase at 4 M NaCl) (Fig. 3). On the other hand, G3PD activity decreased to about 50% using NADH as a cofactor, but increased around 100% at 4 M NaCl when NADPH was used (Fig. 3). All other enzymes were checked for cofactor specificity, but no differences from published results were found (not shown).

3

Specific activity of enzymes from glycerol pathways in cell-free extracts from C. halophila cultivated on 2% glucose (w/v), in the presence of a range of salt concentrations. Assays were performed in the absence of salt. Enzymes: glycerol-3P-dehydrogenase (G3PD) using either NADPH or NADH as cofactors; glycerol-3P-phosphatase (G3P); dihydroxyacetone kinase (DK) and glycerol dehydrogenase (GD) measured using NADP+.

3

Specific activity of enzymes from glycerol pathways in cell-free extracts from C. halophila cultivated on 2% glucose (w/v), in the presence of a range of salt concentrations. Assays were performed in the absence of salt. Enzymes: glycerol-3P-dehydrogenase (G3PD) using either NADPH or NADH as cofactors; glycerol-3P-phosphatase (G3P); dihydroxyacetone kinase (DK) and glycerol dehydrogenase (GD) measured using NADP+.

Likewise, mannitol pathway enzymes, HxK (fructose-driven reaction, Fig. 7) and M1PD/NADPH-dependent (data not shown), showed an increased activity in cells cultivated with salt. M1PD could also use NADH, but with a very low specific activity, which remained constant in cells grown at any salt concentration up to 4 M NaCl (data not shown). The same was observed for MD/NAD+-dependent, which maintained its specific activity at all salinities (not shown).

7

Specific enzyme activities measured in cell-free extracts from cells of C. halophila cultivated with 2% glucose (w/v) and a range of salt concentrations and collected in mid-exponential phase. Assays were performed in the absence of salt. Enzymes: hexokinase, using either glucose or fructose as reaction substrate (HxK); glyceraldehyde-3P-dehydrogenase (GldD); alcohol dehydrogenase (ADH); and malate dehydrogenase (MDH).

7

Specific enzyme activities measured in cell-free extracts from cells of C. halophila cultivated with 2% glucose (w/v) and a range of salt concentrations and collected in mid-exponential phase. Assays were performed in the absence of salt. Enzymes: hexokinase, using either glucose or fructose as reaction substrate (HxK); glyceraldehyde-3P-dehydrogenase (GldD); alcohol dehydrogenase (ADH); and malate dehydrogenase (MDH).

Enzyme resistance to salt stress

Considering the results above, we wondered if these enzymes, assayed in cells grown at such high salt concentrations, would behave like halophilic enzymes, or if, instead, this increase would be due to a generalized transcription and/or translation stimulation, leading to an increase in enzyme concentration. Glyceraldehyde-3P-dehydrogenase was exponentially inhibited by the addition of salt to the reaction buffer (Fig. 4A). At 200 mM NaCl the specific activity was already reduced to 40%. KNaCl was calculated (Fig. 4B). The same happened to glycerol-3P-dehydrogenase and dihydroxyacetone kinase, which activities, once incubated with 500 mM NaCl, were reduced to 32 and 38%, respectively (not shown).

4

A: Glyceraldehyde-3P-dehydrogenase, specific activity measured in cell-free extracts from C. halophila cultivated on 2% glucose (w/v) and collected in mid-exponential phase. Assays were performed in the presence of salt up to 200 mM. B: Activity presented as a percentage of the value in the absence of salt.

4

A: Glyceraldehyde-3P-dehydrogenase, specific activity measured in cell-free extracts from C. halophila cultivated on 2% glucose (w/v) and collected in mid-exponential phase. Assays were performed in the presence of salt up to 200 mM. B: Activity presented as a percentage of the value in the absence of salt.

Respiration and fermentation fluxes

C. halophila is a Crabtree-positive yeast. It ferments and respires glucose simultaneously in the presence of oxygen. O2 consumption and CO2 production rates were measured in the same batches of cells, cultivated with a range of NaCl concentrations. Cells cultivated in the presence of various salt concentrations were assayed with different salt concentrations, ranging from 0.5 to 4 M NaCl, with 0.5-M intervals (Fig. 5). Both O2 consumption and CO2 production rates, from cells grown in the absence of salt, were very sensitive to salt during the assay and decreased strongly with salt concentration. For comparison, we plotted the specific O2 consumption rates and the CO2 production rates of cells cultivated in and assayed with the same salt concentration (Fig. 6). At concentrations above 0.5 M, respiration rates increased in about 40%, and remained constant regardless of the increase in NaCl concentration. CO2 production rates, which yielded more scattered curves (Fig. 5), increased progressively once represented this way, reaching a maximum increase of 400% at 4 M NaCl (Fig. 6).

5

Specific CO2 production rates (A) and specific O2 consumption rates (B) of C. halophila cultivated on 2% glucose (w/v) with a range of salt concentrations and assayed in the presence of salt up to 4 M NaCl.

5

Specific CO2 production rates (A) and specific O2 consumption rates (B) of C. halophila cultivated on 2% glucose (w/v) with a range of salt concentrations and assayed in the presence of salt up to 4 M NaCl.

6

Specific CO2 production rates and specific O2 consumption rates of C. halophila cultivated on glucose 2% (w/v) with various concentrations of salt, and assayed in the presence of the same concentration of salt as in the growth medium.

6

Specific CO2 production rates and specific O2 consumption rates of C. halophila cultivated on glucose 2% (w/v) with various concentrations of salt, and assayed in the presence of the same concentration of salt as in the growth medium.

To establish whether CO2 production, beyond the respiratory CO2 produced, corresponded to alcoholic fermentation, we measured extracellular ethanol during growth, in the same batches of cells (collected at the end of exponential growth phase). We verified that ethanol concentrations increased linearly with the salt concentration (correlation 0.931) up to about 400% at 4 M NaCl (not shown).

These increased fluxes should have a counterpart in metabolism, i.e. in specific enzyme activities. In order to evaluate this, we chose the following enzymes: (1) hexokinase (Hxk, glucose-driven reaction), since it is the first enzyme in glycolysis in connection with the mannitol pathway; (2) glucose-6P-dehydrogenase (G6PD) from the pentose phosphate pathway; (3) glyceraldehyde-3P-dehydrogenase (Gld3PD) for its position in glycolysis, in connection with the glycerol pathway; (4) alcohol dehydrogenase (ADH) for its role in alcohol production during fermentation; (5) malate dehydrogenase (MDH) from the Krebs cycle and (6) cytoplasmic isocitrate dehydrogenase (IDH) for its involvement in NADPH supply. The specific activities of all these enzymes, from cells cultured on glucose, glycerol or mannitol (Table 3), were similar, suggesting that their expression is not significantly influenced by the carbon source.

When cells were cultivated in the presence of salt, a general increase in activity was observed (exemplified in Fig. 7). The highest increase in activity was found for alcohol dehydrogenase, reaching 400% of the reference value for cultures with 4 M NaCl. This increase is similar to the one observed for CO2 and ethanol production rates under identical experimental conditions (above).

Estimation of intracellular concentration of reduced and oxidized enzyme cofactors, NAD(H) and NADP(H)

The intracellular redox state of the cell is most dependent on the balance between the two co-substrate couples NADH/NAD+ and NADPH/NADP+. In S. cerevisiae, the regulation of the oxidized/reduced state of these two nucleotides, and the correspondent consequences for metabolism, are still poorly understood, in spite of the significant amount of work performed concerning this issue [39]. In an eventually simplistic way, a drastic unbalance in the concentration or redox state of one of these pairs can lead to termination of cell growth.

We measured the oxidized and the reduced forms of both cofactors in cells cultivated with different salt concentrations (Table 4). In addition, these results were analyzed using indices generally used in bacteria. This might provide more insight in the proportions in and between these oxidized/reduced pairs. PNF gives an indication of the proportion between NADP(H) and NAD(H), CRC and ARC give an indication of the percentage of oxidized forms in the total amount of each cofactor, and thus an idea of the reducing power available. Furthermore, from the data presented in Table 4, it is possible to see that the ratio between all the oxidized forms, against all the reduced forms, is approximately constant, indicating that there is no impasse or unbalance regarding the redox potential under salt stress in C. halophila, even at very high salt concentrations.

4

Intracellular reduced and oxidized nucleotide concentrations

Intracellular nucleotide concentration (μmol (g dry wt)−1
[NaCl] (M) NAD+ NADP+ NADH NADPH PNF CRC ARC 
1.44 0.44 1.97 1.18 0.48 0.49 0.73 
0.5 0.87 0.15 1.14 0.71 0.43 0.57 0.83 
1.0 0.94 0.15 1.04 0.81 0.49 0.53 0.84 
1.5 1.18 0.30 0.81 0.85 0.58 0.41 0.74 
2.0 0.96 0.29 1.14 0.93 0.58 0.54 0.76 
2.5 1.22 0.18 1.13 1.02 0.51 0.48 0.85 
3.0 0.91 0.38 1.04 0.89 0.65 0.53 0.70 
4.0 0.52 0.18 1.65 1.10 0.59 0.76 0.86 
Mean+S.D. 1.01±0.26 0.26±0.10 1.24±0.35 0.93±0.15 0.54±0.07 0.54±0.10 0.79±0.10 
Intracellular nucleotide concentration (μmol (g dry wt)−1
[NaCl] (M) NAD+ NADP+ NADH NADPH PNF CRC ARC 
1.44 0.44 1.97 1.18 0.48 0.49 0.73 
0.5 0.87 0.15 1.14 0.71 0.43 0.57 0.83 
1.0 0.94 0.15 1.04 0.81 0.49 0.53 0.84 
1.5 1.18 0.30 0.81 0.85 0.58 0.41 0.74 
2.0 0.96 0.29 1.14 0.93 0.58 0.54 0.76 
2.5 1.22 0.18 1.13 1.02 0.51 0.48 0.85 
3.0 0.91 0.38 1.04 0.89 0.65 0.53 0.70 
4.0 0.52 0.18 1.65 1.10 0.59 0.76 0.86 
Mean+S.D. 1.01±0.26 0.26±0.10 1.24±0.35 0.93±0.15 0.54±0.07 0.54±0.10 0.79±0.10 

PNF, phosphorylated nucleotide fraction; CRC, catabolite reduction charge; ARC, anabolic reduction charge (see Section 2).

Estimation of intracellular Na+ and K+ concentrations

According to the literature [4,5,17], the yeast C. halophila is able to maintain a very stable ion homeostasis, even in the presence of high ionic strength. We measured intracellular Na+ and K+ in cells cultivated as before. In view of discrepancies observed in the literature (mentioned in Section 1), we decided to use three experimental approaches: AAS, ICP and NMR. Only the results from AAS are presented, since no significant differences were observed using the other technological approaches.

As can be seen in Fig. 8, the potassium concentration remained approximately constant (around 300 mM) up to concentrations of about 2 M NaCl. This internal potassium concentration has frequently been designated as ‘normal’ in other yeasts in the absence of salt [13,40]. In the same cells, the sodium concentration also remained constant, although extraordinarily low (around 30 mM). Above the turning point of 2 M NaCl, sodium increased to another constant level (around 120 mM), while potassium decreased progressively, reaching a minimum of about 50 mM at the highest salt concentrations. Considering the total amount of the two ions (sodium plus potassium), this decreases from 300 to 200 mM, for cells grown in the absence of salt to cells cultivated at 4.5 M NaCl, respectively, a decrease of 1/3 in total ion concentration.

8

Intracellular sodium and potassium concentrations in C. halophila cultivated on 2% glucose (w/v) with salt and collected in mid-exponential growth phase.

8

Intracellular sodium and potassium concentrations in C. halophila cultivated on 2% glucose (w/v) with salt and collected in mid-exponential growth phase.

On the other hand, the observed decrease in intracellular potassium closely paralleled the decrease in specific growth rate on glucose (Fig. 9). Therefore, we suppose that in C. halophila, also under NaCl stress, the maintenance of intracellular potassium levels is an indispensable requirement for growth. Substitution of potassium in the growth medium by sodium (using H3PO4 instead of KH2PO4 and adding 0.5% NaCl) [12], resulted in impaired growth (data not shown).

9

Variation of specific growth rate and intracellular potassium concentration with salt concentration in the growth medium of C. halophila.

9

Variation of specific growth rate and intracellular potassium concentration with salt concentration in the growth medium of C. halophila.

As before [5], we compared the above results with cells cultivated without salt and submitted to a sudden shock using 1, 2 and 3 M NaCl (Fig. 10). During the experiments (6 h), an increase of [K+]in was observed, which at 3 M NaCl was only transient. In the same period of time, [Na+]in also transiently increased, but at 3 M NaCl, the subsequent decrease was slower than at 2 M or 1 M. The maximum concentration of sodium ions achieved during this transient increase, after about 1–2 h of incubation, was higher for high salt concentrations: 18 mM Na+ at 1 M NaCl; 150 mM Na+ at 2 M NaCl and 250 mM Na+ at 3 M NaCl. This increase followed a linear variation with extracellular salt concentration (correlation 0.997).

10

Variation of intracellular sodium and potassium concentrations with incubation time, at 30°C, in C. halophila cultivated on 2% glucose (w/v) without salt and transferred to growth media with a range of salt concentrations.

10

Variation of intracellular sodium and potassium concentrations with incubation time, at 30°C, in C. halophila cultivated on 2% glucose (w/v) without salt and transferred to growth media with a range of salt concentrations.

Discussion

Enzymes of the glycerol and mannitol pathways

As previously stated, the aim of this study was to acquire knowledge on the metabolism of cells growing with salt, by comparing cells grown with a range of salt concentrations to cells exposed to an immediate salt shock of identical molarity.

C. halophila grows equally well on either glucose, glycerol or mannitol as single carbon and energy sources, all yielding very similar specific growth rates [5]. Moreover, the inhibitory effect of salt on consumption of these substrates was the same. Remarkably, sequential growth was observed on glucose and glycerol or glucose and mannitol mixtures, in spite of the fact that all the enzymes for the glycerol and the mannitol pathways showed significant activity in glucose-grown cells. Besides, the correspondent transmembrane transport systems were also found to be constitutively expressed [5]. Enzymes leading to the production of glycerol and mannitol were expected to be present, in view of the amounts of these two compounds present in glucose-grown cells [5], but not expected were the enzymes responsible for the consumption of these substrates, neither the correspondent transmembrane transport systems. In view of these results, we consider that the regulation of these enzyme activities, from cells cultivated in the absence of stress, is unlikely to take place at the transcription level.

One problem though arose during our enzyme determinations: the inability to detect glycerol kinase. This can be explained by at least four hypotheses: (1) the overall absence of activity of the glycerol dissimilatory pathway in C. halophila; (2) the very low activity of this pathway compared to the alternative dihydroxyacetone pathway, since the mitochondrial glycerol-3P-dehydrogenase was detected at low levels; (3) the biochemical assay eventually not being optimal; or (4) the enzyme, although widely spread among a great variety of microorganisms, being much more different at the nucleotide level than at the protein level, making the molecular approach used inconclusive for its detection.

Although they were present in glucose-grown cells, the activities of the glycerol and mannitol consumption enzymes were several times lower than the activities measured when cells were growing on the correspondent substrate as a carbon and energy source. This type of results has already been presented for Candida albicans[41]. In this yeast, mannitol transport was under glucose repression and consequently growth on mixtures of both substrates was sequential. Nevertheless, NAD+-mannitol dehydrogenase was expressed in the presence of glucose, though at very low levels compared to the levels achieved after glucose exhaustion. This means that in C. halophila, as in C. albicans, enzymes might be subject to glucose regulation, although they are not under glucose repression. So far, biochemical control seems to be the most likely candidate responsible for impairment of glycerol and mannitol metabolism during growth on glucose.

The result that in glucose-grown cells of C. halophila mannitol production occurs simultaneously with glycerol production suggests substantially different roles for these compounds in C. halophila, as compared to S. cerevisiae[39] or non-conventional yeasts [42].

The function of mannitol

The role of mannitol in plants and fungi has been extensively studied. This compound can be an osmolyte during salt stress, accumulated in high amounts inside fruiting bodies [43]. As in the mushroom Agaricus bisporus[43], it can serve as a supply of NADP+ for the oxidation reactions of the pentose phosphate shunt through mannitol NADPH dehydrogenase, thus controlling growth. As an a-cyclic hexitol, it can also serve as a scavenger. It has been suggested that, during the proliferation process in mammalian tissue, it protects pathogenic yeasts like Cryptococcus neoformans against lethal oxidative stress mediated by host phagocytes [44]. C. albicans produces and accumulates high amounts of mannitol in the absence of glucose through an NAD+-mannitol dehydrogenase [41]. Pathogenicity of C. albicans has been associated with cytosolic levels of NADPH, in response to oxidative stress [45]. Apparently, enzymes responsible for protection against free radicals are NADPH-dependent [46]. The function of mannitol as an a-cyclic hexitol scavenger has been demonstrated in S. cerevisiae growing under oxidative stress [47]. Within this species, several strains can produce mannitol through an NAD+-mannitol dehydrogenase instead of an NADP+ dehydrogenase (like in filamentous fungi) [48]. Furthermore, in recent articles it has been suggested that in yeasts, mannitol can substitute for glycerol as an osmolyte, comparable to what happens in plants [47,49].

In C. halophila growing under salt stress, glycerol production was proportional to external salt concentration [5], confirming that glycerol plays the role of the osmolyte, as has been shown before for other yeasts [50–54]. On the other hand, cells growing on mannitol under salt stress also produced glycerol, at the same concentrations as cells growing on glucose (not shown), though mannitol concentrations were higher than in glucose-grown cells. Cells cultivated in the absence of salt produced mannitol, but this production decreased progressively as the salt concentration in the growth medium increased [5]. This corresponds with previous results from Onishi and Suzuki [55], who have demonstrated that in several mannitol-producing yeasts the intracellular mannitol concentrations were considerable, but decreased when the cells were cultivated with high levels of NaCl or KCl. Two of the yeasts studied by these authors were Torulopsis halophila and Torulopsis versatilis[55]. These have been re-named in 1990 by Barnett and co-workers [3] as C. halophila and C. versatilis. Our results thus indirectly confirm these findings.

As mentioned above, mannitol can act as a scavenger. Nevertheless, when C. halophila, which resists oxidative stress as well, was grown in the presence of 200 mM H2O2, intracellular mannitol concentrations did not present a significant increase in comparison to glucose-grown cells, to justify this role. Results were thus not conclusive. Thus, in C. halophila, mannitol is not the osmolyte and cannot substitute for glycerol in that role. Furthermore, it apparently also does not work as a scavenger. The role of mannitol thus remains unclear.

The function of glycerol

The role of glycerol production during growth on glucose has been extensively discussed in S. cerevisiae. In a simplistic way we can say that, during anaerobic growth, S. cerevisiae produces excess NADH in the cytoplasm, due to the formation of biomass and several fermentation byproducts [39,56]. Glycerol formation restores the redox equilibrium through conversion of dihydroxyacetone-P to glycerol-3P by NAD+-dependent glycerol-3P-dehydrogenase [57]. Aerobically, this regeneration is achieved by an NADH dehydrogenase localized in the outer face of the mitochondrial membrane [58] and the glycerol-3P shuttle [57], in addition to several other possible mechanisms [39]. In the system NADPH/NADP+ reducing equivalents follow autonomous cycles: glucose consumption requires NADPH for assimilation and biomass production. The reduction of NADP+ is achieved through the pentose phosphate pathway, NADP+-isocitrate dehydrogenase and acetaldehyde dehydrogenase [59–61]. Interconversion between the two coenzyme systems NADH/NAD+ and NADPH/NADP+ is not possible in S. cerevisiae, because there is no transhydrogenase [62,63]. Any unbalance in the redox state of their two systems can lead to growth termination. The above applies to cells growing on glucose in the absence of stress and confirms the role of the glycerol pathway as an important intervenient in the redox balance in S. cerevisiae.

In C. halophila, the reason for the counterbalance between glycerol and mannitol concentrations under stress can only be suggested. To our knowledge, there is no experimental evidence of salt-growing cells suffering from oxidative stress. Nevertheless, very recently it has been suggested that both osmotic and heat stress can interfere with the electron transport chain, thus enhancing the production of reactive oxygen species [64]. It has also been suggested that stress conditions demand higher quantities of NADPH for protection against these reactive oxygen species. Under stress in S. cerevisiae, the NADPH-generating enzymes from the pentose phosphate pathway are enhanced [64]. Furthermore, the complete glycerol cycle can convert NADH to NAD+ while producing glycerol and reduce NADP+ to NADPH via dihydroxyacetone. If this hypothesis is correct, the glycerol cycle would compensate to some extent for the lack of a transhydrogenase in S. cerevisiae[62], constituting a futile cycle regulated at several levels [64,65].

In C. halophila, mannitol-1P-dehydrogenase (M1PD), the enzyme catalyzing the first step of the mannitol production pathway, can apparently use NADPH or NADH indistinctively as cofactors. Furthermore, unlike the equivalent enzymes Gpd1p and Gpd2p of the glycerol pathway in S. cerevisiae, [66], glycerol-3P-dehydrogenase from C. halophila can also use either NADPH or NADH. This type of glycerol-3P-dehydrogenase versatility had already been published in another salt-tolerant yeast, Z. rouxii[67]. This creates a whole new vision on redox balance regulation, because the versatility of these two enzymes might substitute for the possible role of a transhydrogenase. No further enzymatic steps would be needed, besides a simple choice of cofactors according to metabolic convenience. Furthermore, GD and MD also have the possibility of using different cofactors, dependent on the production of glycerol or the consumption of mannitol. This alternative cofactor utilization reinforces once more the flexibility of the two cycles mentioned above. Thus, the lack of NADPH in C. halophila, even under stress, might not be so pronounced as in S. cerevisiae and the versatility of these enzymes would preclude an NADPH unbalance. We can only speculate whether the glycerol and mannitol pathways work together as futile cycles in order to regenerate cofactors in C. halophila.

NADPH is prefered as co-substrate for G3PD and glycerol production under stress and glycerol is the osmolite, accumulating increasingly more with heavier stress conditions [5]. Nevertheless, it is possible that switching the cofactor utilization, in the case of mannitol and glycerol dehydrogenases, is not the only way by which C. halophila controls its redox balance. The specific activities of isocitrate dehydrogenase/NADP+-dependent (cytosolic or peroxisomal) and glucose-6P-dehydrogenase remained the same with increasing salt stress, suggesting that they might neither cope with the increasing necessity for NADPH regeneration, nor interfere with the NADP(H) redox balance, unlike what is known for S. cerevisiae. This way, the production of acetate through acetaldehyde dehydrogenase remains as one of the few ways to regenerate NADPH. Indeed, in C. halophila, we observed that acetate production was much higher in cells grown at higher salt concentrations, though we were not able to detect the activity of the enzyme (not shown).

Patterns of enzyme activities

An almost general increase in enzyme activity was observed in cell-free extracts from salt-grown cells of C. halophila. The increase might contribute to: (1) the production of more glycerol; (2) the production of more ATP in order to stimulate H+-ATPase, as in C. versatilis[68] or Z. rouxii[69], thereby maintaining the intracellular pH [5]; and (3) the production of biomass. In accordance with previous results [5], less biomass is produced since more glycerol is needed for osmoprotection and more ATP is consumed for ion homeostasis. Moreover, fermentation rates of C. halophila increased significantly in the presence of salt, as did the activity of alcohol dehydrogenase and the amounts of extracellular ethanol measured. Altogether, these results and considerations might help to explain the surprising result of the redox indicators PNF, CRC and ARC, remaining constant in cells grown in salt up to 4 M NaCl. However, this interpretation might be discussable, since it comprises the necessity of an increase in ATP in order to maintain the internal pH balance through ATPase activity, which is supposedly stimulated. In S. cerevisiae it has been postulated otherwise [65]. According to the model proposed by Blomberg [65], the glycerol and trehalose pathways function as futile cycles, so-called ‘safety valves’ of glycolysis, depleting excess ATP caused by stress. Our results do not fit into this model, except maybe from the point of view that glycerol and mannitol cycles could be working together, creating a large flexibility on redox and ATP balance.

In our opinion, the most interesting result obtained in C. halophila concerns the increase in specific activity of most of the enzymes measured. This almost general pattern can be due to two main processes: de novo synthesis, by stimulation of transcription through signal transduction; or a biochemical adjustment, comprising in vivo activation/inhibition of pre-existing enzymes. This last process has been suggested for the redox balance of S. cerevisiae in aerobic chemostat cultures [58]. This is probably not the case with C. halophila, since biochemical regulation functions on a much shorter time scale not comparable to the extended lag phases presented in salt-growing cells [5]. In S. cerevisiae, though, in vivo activation or inhibition of activities of external mitochondrial NADPH dehydrogenase and G3P shuttle enzymes via biochemical processes, rather than by molecular regulation, has been suggested by Påhlmann et al. [58]. Changes in genetic expression patterns, mentioned probably for the first time in 1969 by Norkrans and Kylin [7], have been exhaustively addressed in S. cerevisiae[64]. We have not performed any experiment in order to investigate which of the high salt-responding signal transduction pathways presently known in S. cerevisiae could possibly be active in C. halophila. But we cannot avoid a comment on the fact that GPP (glycerol-3P-phosphatase, G3P) and GPD (glycerol-3P-dehydrogenase, G3PD) genes from S. cerevisiae glycerol consumption and production pathways have been shown to be required for oxidative stress protection and adaptation of cells to anaerobiosis [70,71]. This is apparently achieved by differential expression and cross-talk between the HOG pathway [70] and a Ca2+/calmodulin-dependent phosphatase (calcineurin-mediated pathway) [72]. The latter seems to be required for Na+, Li+, Mg2+ and OH resistance via a direct involvement of the H+-ATPase [72], for proton homeostasis and consequently for the maintenance of the electrochemical gradient across the plasma membrane [73]. Yet, in C. halophila, H+-ATPase activity was not affected by salt [5]. Van Wuytswinkel et al. [74] have suggested that the response to severe osmotic stress, comparable to the type we describe here for C. halophila, corresponds to a transient defect in nucleo-cytoplasmic trafficking, which might create a delay in transcriptional response: a delay in Hog1p (first enzyme from High Osmolarity Signal transduction Pathway) phosphorylation, and induction of stress-responsive genes, and/or cell wall and cytoskeleton reorganization [75].

The concept of halophily

Finally, we address the concept of halophily. To our knowledge, four very different yeasts have been classified as halophilic: the black yeast Hortea werneckii[15], D. hansenii[12], Candida halonitratophila[3] and C. versatilis[3,68]. The reasons for this classification are in each case of a rather different nature. In the first case, H. werneckii, the authors claim that this basidiomycete responds to salt via a change in membrane properties, in particular membrane fluidity, dependent on sterol synthesis. The key enzyme for sterol synthesis, HMG R (3-hydroxy-3-methylglutaryl coenzyme-A reductase) shows maximum activity at 0.5% NaCl in in vitro assays, whereas cells of H. werneckii can live in the presence of NaCl up to 30%. In the case of D. hansenii[1,12], the authors have shown that the organism improves growth at NaCl concentrations around 0.5–1 M, just like C. halophila and P. sorbitophila[5]. But, more important, they claim that the metabolism of this yeast is intrinsically less sensitive to salt, being able to exchange equally K+ for Na+ without poisoning the cell. Accordingly, intracellular Na+ concentrations published by the same authors are very high [12]. Nevertheless, taking into account the extreme heterogeneity of intracellular ion concentrations found in the literature for D. hansenii, as well as for other yeasts, these results could eventually be illusive [6,7,10,11,13,76]. Enzymes from D. hansenii assayed in vitro presented their optimum activity around 0.5 M NaCl [11], while the yeast can grow up to 3.5 M NaCl [11,12]. Finally, C. halonitratophila and C. versatilis, considered halophilic yeasts by Barnett et al. [3], were classified as such because, within their experimental frame, these yeasts could only grow when salt was added to the growth medium. They have though been shown to grow well in the absence of salt by Lages et al. [2].

As discussed above, C. halophila glyceraldehyde-3P-dehydrogenase (localized in the cell wall like in S. cerevisiae[77]) did not endure the presence of high salt concentrations in the in vitro assay. Its activity decreased exponentially and was 40% of the initial value at 200 mM NaCl. Its exponential inhibition constant (KNaCl) was smaller, but still of the same order of magnitude, as the value published for the same enzyme from D. hansenii[11]. Also C. halophila could not grow in the absence of potassium in the growth medium.

This way, we can discuss the true meaning of living in the presence of very high salt concentrations. On the one hand, there are fast membrane fluidity adaptations [15], and on the other hand, the efficiency of the transport mechanisms able to expel ions which drastically varies [1]. The cell has to deal with a delicate total ionic balance between the two sides of the membrane, in a way that simultaneously sustains cell growth. This means maintaining low intracellular ionic concentration, since cytosolic enzymes per se do not cope with very high salinity. This is the case of moderately halophilic bacteria [78]. In these bacteria, enzymes secreted into the medium or located on the outer cell surface are much more salt-tolerant than cytosolic enzymes, which show a very moderate resistance to salt in in vitro assays [78]. In our case, we consider C. halophila to be an exceptional yeast, since it is able to simultaneously maintain very low sodium concentrations and potassium levels compatible with growth. Potassium concentrations never decreased below 50 mM, being the growth limit described so far for yeasts of 0.2 mM [79]. According to the classification proposed by Ramos for yeasts [1] (concerning Na+/K+ ratios and thereby distinguishing sodium excluders and sodium includers), C. halophila, like S. cerevisiae[1], will have to be classified as an excluder, while D. hansenii apparently behaves as an includer [12]. In spite of this, Ramos [1] has also stated that the kinetics of Na+ exclusion are similar in S. cerevisiae and D. hansenii, concluding that this criterion is certainly not enough to explain the higher salt tolerance of one yeast compared to the other. In this context, and in view of our results, C. halophila appears to be a much more efficient excluder than other yeasts, since the levels of intracellular sodium never exceed about 150 mM.

So, what eukaryote can be called a halophilic organism? Apparently one that, besides growing better in the presence of salt, is (nearly) unable to grow without it [80], or one that tolerates intracellular concentrations of salt and depends on those to survive, therefore possessing halophilic enzymes working exclusively at high salinities, like in halophilic bacteria [14,77,81,82]. This is not the case in yeasts. They grow well without salt stress and, furthermore, yeast enzymes do not cope with high concentrations of salt. While halotolerance is a clear, well established concept, halophily in yeasts may be a concept defining a transient state of very limited nature and thus of doubtful use. Yeasts have been classified halophilic in spite of the significance of the term in itself, i.e. yeasts may be halotolerant, eventually salt-loving microorganisms, which can nevertheless grow very well in the absence of salt. Therefore, it is our opinion that, for the moment, no strong reason supports such classification for any yeast.

Final conclusions

As a conclusion, we recapitulate the main results obtained in C. halophila, before [5] as well as in this article, supporting all speculations presented: (i) the ability to grow up to 5 M NaCl; (ii) the improvement in growth rate ranging from 0.5 and 2 M NaCl; (iii) the maintenance of a constant intracellular volume up to the highest salt concentrations; (iv) the lower need for accumulating glycerol as an osmolyte than other yeasts; (v) the constitutive expression of active transport systems for glycerol and mannitol, which are not significantly affected by salt; (vi) the constitutive expression of enzymes from the glycerol and mannitol pathways, though apparently under strict metabolic regulation; (vii) the eventual role of the mannitol pathway for maintaining the redox balance; (viii) the ability of the enzymes of the mannitol and glycerol pathways (G3PD, M1PD, GD and MD) for alternative NADP(H)/NAD(H) utilization; (ix) the moderate increase in respiratory flux, accompanied by a very high increase in alcoholic fermentation rates; (x) the consistency of intracellular PNF, CRC and ARC, up to the highest salt concentrations, indicating a balanced, interchangeable redox regulation between cofactors; (xi) the consistency of the intracellular volume and pH constant, up to the highest salt concentrations; (xii) the maintenance of the intracellular low [Na+] and high [K+], up to 2 M NaCl; (xiii) the maintenance of intracellular [K+] at levels that allow growth up to the highest salt concentrations; (xiv) the general pattern of stimulation of enzyme activities in the presence of increasing salt concentrations; and finally, (xv) the sensitivity of enzymes to low salt concentrations once assayed in vitro. These results, altogether, provide substantial evidence for to support C. halophila as an archetype representing the limits of extreme halotolerance in yeasts and put into evidence the reasons why, in our opinion, even salt-tolerant yeasts should not, so far, be classified or considered true halophilic microorganisms.

Acknowledgements

This experimental work as well as the PhD grant of M.S.-G. was supported by the European Commission, Project BIOTECH PL95-0161. We thank Prof. H. Santos and Dr. L. Martins from the I.T.Q.B. Research Institute, Oeiras (Portugal) and Dr. C. Ribeiro from the Earth Sciences Department, Minho University, for NMR and ICP ion determinations. We also want to address a special thank to Prof. T. Tavares and Dr. A. Ferraz from the Biological Engineering Department from Minho University for allowing AAS ion determinations. Finally, we thank our colleague R. Oliveira for designing the primers used for Southern blot experiments.

References

[1]
Ramos
J.
(
1999
)
Contrasting salt tolerance mechanisms in Saccharomyces cerevisiae and Debaryomyces hansenii
.
Recent Res. Dev. Microbiol.
 
3
,
377
390
.
[2]
Lages
F.
Silva-Graça
M.
Lucas
C.
(
1999
)
Active glycerol uptake is a mechanism underlying halotolerance in yeasts: a study of 42 species
.
Microbiology
 
145
,
2577
2586
.
[3]
Barnett
J.A.
Payne
R.W.
Yarrow
D.
(Eds.) (
1990
)
Yeasts: Characteristics and Identification
 ,
2nd
Edn.
Cambridge University Press
,
Cambridge
.
[4]
Barnett
J.A.
Payne
R.W.
Yarrow
D.
(Eds.) (
2000
)
Yeasts: Characteristics and Identification
 ,
3rd
Edn.
Cambridge University Press
,
Cambridge
.
[5]
Silva-Graça
M.
Lucas
C.
(
2002
)
Physiological studies on long-term adaptation to salt stress in the extremely halotolerant yeast Candida versatilis CBS 4019 (syn. C. halophila)
.
FEMS Yeast Res.
 
1535
,
1
14
.
[6]
Norkrans
B.
(
1969
)
The sodium and potassium contents of yeasts differing in halotolerance, at various NaCl concentrations in the media. Antonie van Leeuwenhoek 35, Supplement: Yeast Symposium 1969
  , G31.
[7]
Norkrans
B.
Kylin
A.
(
1969
)
Regulation of the potassium-to-sodium ratio and of the osmotic potential in relation to salt tolerance in yeasts
.
J. Bacteriol.
 
100
,
836
845
.
[8]
Adler
L.
Gustafsson
L.
(
1980
)
Polyhydric alcohol production and intracellular amino acid pool in relation to halotolerance of the yeast Debaryomyces hansenii
.
Arch. Microbiol.
 
124
,
123
130
.
[9]
Adler
L.
Blomberg
A.
Nilsson
A.
(
1985
)
Glycerol metabolism and osmoregulation in the salt tolerant yeast Debaryomyces hansenii
.
J. Bacteriol.
 
162
,
300
306
.
[10]
Burke
R.M.
Jennings
D.H.
(
1990
)
Effect of sodium chloride on growth characteristics of the marine yeast Debaryomyces hansenii in batch and continuous culture under carbon and potassium limitation
.
Mycol. Res.
 
94
,
378
388
.
[11]
Neves
L.
Oliveira
R.
Lucas
C.
(
1997
)
Metabolic flux response to salt-induced stress in the halotolerant yeast Debaryomyces hansenii
.
Microbiology
 
143
,
1133
1139
.
[12]
Prista
C.
Almagro
A.
Loureiro-Dias
M.C.
Ramos
J.
(
1997
)
Physiological basis for high salt tolerance of Debaryomyces hansenii
.
Appl. Environ. Microbiol.
 
63
,
4005
4009
.
[13]
Thomé-Ortiz
P.E.
Peña
A.
Ramirez
J.
(
1998
)
Monovalent cation fluxes and physiological changes of Debaryomyces hansenii grown at high concentrations of KCl and NaCl
.
Yeast
 
14
,
1355
1371
.
[14]
Eisenberg
H.
(
1995
)
Life in unusual environments: progress in understanding the structure and function of enzymes from extreme halophilic bacteria
.
Arch. Biochem. Biophys.
 
318
,
1
5
.
[15]
Petrovic
U.
Gunde-Cimerman
N.
Plemenitas
A.
(
1999
)
Salt-stress affects sterol biosynthesis in the halophilic black yeast Hortaea werneckii
.
FEMS Microbiol. Lett.
 
180
,
325
330
.
[16]
Causton
H.C.
Ren
B.
Koh
S.S.
Harbison
C.T.
Kanin
E.
Jennings
E.G.
Lee
T.I.
True
H.L.
Lander
E.S.
Young
R.A.
(
2001
)
Remodelling of yeast genome expression in response to environmental changes
.
Mol. Biol. Cell
 
12
,
323
337
.
[17]
van der Sluis
C.
Stoffelen
C.J.P.
Castelein
S.J.
Engbers
G.H.M.
ter Schure
E.G.
Tramper
J.
Wijffels
R.H.
(
2001
)
Immobilized salt-tolerant yeasts: application of a new polyethylene-oxide support in a continuous stirred-tank reactor for flavour production
.
J. Biotechnol.
 
88
,
129
139
.
[18]
Thomas
B.J.
Rothstein
K.D.
(
1989
)
Elevated recombination rates in transcriptionally active DNA
.
Cell
 
56
,
619
630
.
[19]
Peterson
G.L.
(
1977
)
A simplification of the protein assay method of Lowry et al. which is more generally applicable
.
Anal. Biochem.
 
83
,
346
356
.
[20]
Ames
B.N.
(
1966
)
Assay of inorganic phosphate, total phosphate and phosphatases
. In:
Methods in Enzymology
  (
Neufeld
E.F.
Ginsburg
V.
Eds.), pp.
115
117
.
Academic Press
,
London
.
[21]
Sussman
I.
Avron
M.
(
1981
)
Characterization and partial purification of dl-glycerol-1-phosphatase from Dunaliella salina
.
Biochim. Biophys. Acta
 
661
,
199
204
.
[22]
Larsson
C.
Gustafsson
L.
(
1993
)
The role of physiological state in osmotolerance of the salt tolerant yeast Debaryomyces hansenii
.
Can. J. Microbiol.
 
39
,
603
609
.
[23]
Babel
W.
Hofmann
K.H.
(
1982
)
The relation between the assimilation of methanol and glycerol in yeasts
.
Arch. Microbiol.
 
132
,
179
184
.
[24]
Lin
E.C.C.
Koch
J.P.
Chused
T.M.
Jorgensen
S.E.
(
1962
)
Utilization of l-α-glycerophosphate by Escherichia coli without hydrolysis
.
Biochemistry
 
48
,
2145
2150
.
[25]
Lin
E.C.C.
Magasanik
B.
(
1960
)
The activation of glycerol dehydrogenase from Aerobacter aerogenes by monovalent cations
.
J. Biol. Chem.
 
235
,
1820
1823
.
[26]
Ueng
S.T.-H.
Hartanowicz
P.
Lewandoski
C.
Keller
J.
Holick
M.
McGuinness
E.T.
(
1976
)
d-mannitol dehydrogenase from Absidia glauca. Purification, metabolic role and subunit interactions
.
Biochemistry
 
15
1743
1749
.
[27]
Boonsaeng
V.
Sullivan
P.A.
Shepherd
M.
(
1976
)
Mannitol production in fungi during glucose catabolism
.
Can. J. Microbiol.
 
22
,
808
816
.
[28]
Hirai
M.E.
Ohtani
E.
Fukui
S.
(
1977
)
Glucose-phosphorylating enzymes of Candida yeasts and their regulation in vivo
.
Biochim. Biophys. Acta
 
480
,
357
366
.
[29]
Kuby
S.A.
Noltmann
E.A.
(
1966
)
Glucose-6-phosphate dehydrogenase from brewers yeast
.
Methods Enzymol.
 
9
,
116
125
.
[30]
Maitra
P.K.
Lobo
Z.
(
1971
)
A kinetic study of glycolytic enzyme synthesis in yeast
.
J. Biol. Chem.
 
246
,
475
489
.
[31]
Postma
E.
Kuiper
A.
Tomasouw
W.F.
Scheffers
W.A.
van Dijken
J.P.
(
1989
)
Competition for glucose between the yeasts Saccharomyces cerevisiae and Candida utilis
.
Appl. Environ. Microbiol.
 
55
,
3214
3220
.
[32]
Witt
J.
Kronau
R.
Holzer
H.
(
1968
)
Repression of alcohol dehydrogenase, malate dehydrogenase, isocitrate lyase and malate synthase in yeast by glucose
.
Biochim. Biophys. Acta
 
118
,
522
537
.
[33]
Stueland
C.S.
Gorden
K.
LaPorte
D.C.
(
1988
)
The isocitrate dehydrogenase phosphorylation cycle – Identification of the primary rate-limiting step
.
J. Biol. Chem.
 
263
,
19475
19479
.
[34]
Ausubel
F.M.
Brent
B.
Kingston
R.E.
Moore
D.D.
Seidman
J.G.
Smith
J.A.
Struhl
K.
(Eds.) (
1995
)
Current Protocols in Molecular Biology
 .
Massachusetts General Hospital/Harvard University Wiley,
,
New York
.
[35]
van Uden
N.
(
1989
)
Effects of alcohols on membrane transport in yeasts
. In:
Alcohol Toxicity in Yeasts and Bacteria
  (
van Uden
N.
Ed.), pp.
135
145
.
CRC Press
,
Boca Raton, FL
.
[36]
Bergmeyer
H.U.
(
1985
)
Methods of Enzymatic Analysis Vol VII
 ,
3rd
Edn.
VCH Publishers
,
Germany
.
[37]
Führer
L.
Kubicek
C.P.
Röhr
M.
(
1979
)
Pyridine nucleotide levels and ratios in Aspergillus niger
.
Can. J. Microbiol.
 
26
,
405
408
.
[38]
Nilsson
A.
Thomson
K.S.
Adler
L.
(
1989
)
Purification and characterization of glycerol kinase in the salt-tolerant yeast Debaryomyces hansenii
.
Biochim. Biophys. Acta
 
991
,
296
302
.
[39]
Bakker
B.M.
Overkamp
K.M.
van Maris
A.J.A.
Kötter
P.
Luttik
M.A.H.
van Dijken
J.P.
Pronk
J.T.
(
2001
)
Stoichiometry and compartmentation of NADH metabolism in Saccharomyces cerevisiae
.
FEMS Microbiol. Rev.
 
25
,
15
37
.
[40]
Garcia
M.J.
Ríos
G.
Ali
R.
Bellés
J.M.
Serrano
R.
(
1997
)
Comparative physiology of salt tolerance in Candida tropicalis and Saccharomyces cerevisiae
.
Microbiology
 
143
,
1125
1131
.
[41]
Niimi
M.
Tokunaga
M.
Nakayama
H.
(
1986
)
Regulation of mannitol catabolism in Candida albicans: evidence for cyclic AMP-independent glucose effect
.
J. Med. Vet. Micol.
 
24
,
211
217
.
[42]
Flores
C.-L.
Rodríguez
C.
Petit
T.
Gancedo
C.
(
2000
)
Carbohydrate and energy-yielding metabolism in non-conventional yeasts
.
FEMS Microbiol. Rev.
 
24
,
507
529
.
[43]
Stoop
J.M.H.
Mooibroek
H.
(
1998
)
Cloning and characterization of NADP-mannitol dehydrogenase cDNA from the bottom mushroom, Agaricus bisporus, and its expression in response to NaCl stress
.
Appl. Environ. Microbiol.
 
64
,
4689
4696
.
[44]
Chaturvedi
V.
Flynn
T.
Niehaus
W.G.
Wong
B.
(
1996
)
Stress tolerance and pathogenic potential of a mannitol mutant of Cryptococcus neoformans
.
Microbiology
 
142
,
937
943
.
[45]
Jamieson
D.J.
Stephen
D.W.S.
Terrière
E.C.
(
1996
)
Analysis of the adaptative oxidative stress response of Candida albicans
.
FEMS Microbiol. Lett.
 
138
,
83
88
.
[46]
Minard
K.I.
McAlister-Henn
L.
(
2001
)
Antioxidant function of cytosolic sources of NADPH in yeast
.
Free Radic. Biol. Med.
 
31
,
832
843
.
[47]
Chaturvedi
V.
Bartiss
A.
Wong
B.
(
1997
)
Expression of bacterial mtlD in Saccharomyces cerevisiae results in mannitol synthesis and protects a glycerol-defective mutant from high-salt and oxidative stress
.
J. Bacteriol.
 
179
,
157
162
.
[48]
Quain
D.E.
Boulton
C.A.
(
1987
)
Growth and metabolism of mannitol by strains of Saccharomyces cerevisiae
.
J. Gen. Microbiol.
 
133
,
16785
16840
.
[49]
Shen
B.
Hohmann
S.
Jensen
R.G.
Bohnert
H.
(
1999
)
Roles of sugar alcohols in osmotic stress adaptation replacement of glycerol by mannitol and sorbitol in yeast
.
Plant Physiol.
 
121
,
45
52
.
[50]
Gustafsson
L.
Norkrans
B.
(
1976
)
On the mechanism of salt tolerance. Production of glycerol and heat during growth of Debaryomyces hansenii
.
Arch. Microbiol.
 
110
,
177
183
.
[51]
Blomberg
A.
Adler
L.
(
1989
)
Roles of glycerol and glycerol-3-phosphate dehydrogenase (NAD+) in acquired osmotolerance of Saccharomyces cerevisiae
.
J. Bacteriol.
 
171
,
1087
1092
.
[52]
Reed
R.H.
Chudeck
J.A.
Foster
R.
Gadd
G.M.
(
1987
)
Osmotic significance of glycerol accumulation in exponentially growing yeasts
.
Appl. Environ. Microbiol.
 
53
,
2119
2123
.
[53]
van Eck
J.H.
Prior
B.A.
Brandt
E.V.
(
1993
)
The water relations of growth and polyhydroxy alcohol production by ascomycetous yeasts
.
J. Gen. Microbiol.
 
139
,
1047
1054
.
[54]
Tokuoka
K.
(
1993
)
Sugar- and salt-tolerant yeasts
.
J. Appl. Bacteriol.
 
74
,
101
110
.
[55]
Onishi
H.
Suzuki
T.
(
1968
)
Production of d-mannitol and glycerol by yeasts
.
Appl. Microbiol.
 
16
,
1847
1952
.
[56]
van Dijken
J.P.
Scheffers
W.A.
(
1986
)
Redox balances in the metabolism of sugars by yeasts
.
FEMS Microbiol. Rev.
 
32
,
199
224
.
[57]
Larsson
C.
Påhlman
I.-L.
Ansell
R.
Rigoulet
M.
Adler
L.
Gustafsson
L.
(
1998
)
The importance of the glycerol-3-phosphate shuttle during aerobic growth of Saccharomyces cerevisiae
.
Yeast
 
14
,
347
357
.
[58]
Påhlman
I.-L.
Gustafsson
L.
Rigoulet
M.
Larsson
C.
(
2001
)
Cytosolic redox metabolism in aerobic chemostat cultures of Saccharomyces cerevisiae
.
Yeast
 
18
,
611
620
.
[59]
Bruinenberg
P.M.
de Bot
P.H.M.
van Dijken
J.P.
Scheffers
W.A.
(
1983
)
The role of redox balances in the anaerobic fermentation of xylose in yeasts
.
Eur. J. Appl. Microbiol. Biotechnol.
 
18
,
287
292
.
[60]
Bruinenberg
P.M.
van Dijken
J.P.
Scheffers
W.A.
(
1983
)
A theoretical analysis of NADPH production and consumption in yeasts
.
J. Gen. Microbiol.
 
129
,
953
964
.
[61]
Nissen
T.L.
Anderlund
M.
Nielsen
J.
Villadsen
J.
Kielland-Brandt
M.C.
(
2001
)
Expression of a cytoplasmic transhydrogenase in Saccharomyces cerevisiae results in formation of 2-oxoglutarate due to depletion of the NADPH pool
.
Yeast
 
18
,
19
32
.
[62]
Minard
K.I.
Jennings
G.T.
Loftus
T.M.
Xuan
D.
McAlister-Henn
L.
(
1998
)
Sources of NADPH and expression of mammalian NADP+-specific isocitrate dehydrogenases in Saccharomyces cerevisiae
.
J. Biol. Chem.
 
273
,
31486
31493
.
[63]
Nissen
T.L.
Hamann
C.W.
Kielland-Brandt
M.C.
Nielsen
J.
Villadsen
J.
(
2000
)
Anaerobic and aerobic batch cultivations of Saccharomyces cerevisiae mutants impaired in glycerol synthesis
.
Yeast
 
16
,
463
474
.
[64]
Hohmann
S.
(
2002
)
Osmotic stress signalling and osmoadaptation in yeasts
.
Microbiol. Mol. Biol. Rev.
 
66
,
300
372
.
[65]
Blomberg
A.
(
2000
)
Metabolic surprises in Saccharomyces cerevisiae during adaptation to saline conditions: questions, some answers and a model
.
FEMS Microbiol. Lett.
 
182
,
1
8
.
[66]
Albertyn
J.
van Tonder
A.
Prior
B.A.
(
1992
)
Purification and characterization of glycerol-3-phosphate dehydrogenase of Saccharomyces cerevisiae
.
FEBS Lett.
 
308
,
130
132
.
[67]
Ohshiro
K.
Yagi
T.
(
1996
)
Regulation of intracellular osmotic pressure and some factors that influence the promotion of glycerol synthesis in a respiration-deficient mutant of the salt-tolerant yeast Zygosaccharomyces rouxii during salt-stress
.
J. Gen. Appl. Microbiol.
 
42
,
201
212
.
[68]
Watanabe
Y.
Sanemitsu
Y.
Tamai
Y.
(
1993
)
Expression of plasma membrane proton-ATPase in salt-tolerant yeast Zygosaccharomyces rouxii is induced by sodium chloride
.
FEMS Microbiol. Lett.
 
114
,
105
108
.
[69]
Watanabe
Y.
Yamaguchi
M.
Sakemoto
J.
Tamai
Y.
(
1993
)
Characterization of plasma membrane H+-ATPase from salt tolerant yeast Candida versatilis
.
Yeast
 
9
,
213
220
.
[70]
Påhlman
A.K.
Granath
K.
Ansell
R.
Hohmann
S.
Adler
L.
(
2001
)
The yeast glycerol-3-phosphatases Gpp1p and Gpp2p are required for glycerol biosynthesis and differentially involved in the cellular responses to osmotic, anaerobic, and oxidative stress
.
J. Biol. Chem.
 
276
,
3555
3563
.
[71]
Ansell
R.
Granath
K.
Hohmann
S.
Thevelein
J.M.
Adler
L.
(
1997
)
The two enzymes for yeast NAD+-dependent glycerol-3-phosphate dehydrogenase encoded by GPD1 and GPD2 have distinct roles in osmoadaptation and redox regulation
.
EMBO J.
 
16
,
2179
2187
.
[72]
Withee
J.L.
Sen
R.
Cyert
M.S.
(
1998
)
Ion tolerance of Saccharomyces cerevisiae lacking the Ca2+/CaM-dependent phosphatase (calcineurin) is improved by mutations in URE2 or PMA1
.
Genetics
 
119
,
865
878
.
[73]
Serrano
R.
Kielland-Brandt
M.
Fink
G.R.
(
1986
)
Yeast plasma membrane ATPase is essential for growth and has homology with (Na+, K+), K+ and Ca2+ ATPases
.
Nature
 
319
,
689
693
.
[74]
van Wuytswinkel
O.
Reiser
V.
Siderius
M.
Kelders
M.
Ammerer
C.
Ruis
H.
Mager
W.H.
(
2000
)
Response of Saccharomyces cerevisiae to severe osmotic stress: evidence for a novel activation mechanism of the HOG MAP Kinase pathway
.
Mol. Microbiol.
 
37
,
382
397
.
[75]
Slaninová
I.
Sesták
S.
Svoboda
A.
Farkas
V.
(
2000
)
Cell wall and cytoskeleton reorganization as the response to hyperosmotic shock in Saccharomyces cerevisiae
.
Arch. Microbiol.
 
173
,
245
252
.
[76]
Larsson
C.
Morales
C.
Gustafsson
L.
Adler
L.
(
1990
)
Osmoregulation of the salt-tolerant yeast Debaryomyces hansenii grown in a chemostat at different salinities
.
J. Bacteriol.
 
172
,
1769
1774
.
[77]
Delgado
M.L.
O'Connor
J.E.
Azorín
I.
Renau-Piqueras
J.
Gil
M.L.
Gozalbo
D.
(
2001
)
The glyceraldehyde-3-phosphate dehydrogenase polypeptides encoded by the Saccharomyces cerevisiae TDH1, TDH2 and TDH3 genes are also cell wall proteins
.
Microbiology
 
147
,
411
417
.
[78]
Ventosa
A.
Nieto
J.J.
Oren
A.
(
1998
)
Biology of moderately halophilic aerobic bacteria
.
Microbiol. Mol. Biol. Rev.
 
62
,
504
544
.
[79]
Camacho
M.
Ramos
J.
Rodríguez-Navarro
A.
(
1981
)
Potassium requirements of Saccharomyces cerevisiae
.
Curr. Microbiol.
 
6
,
295
299
.
[80]
Singleton
P.
Sainsbury
D.
(Eds.) (
1989
)
Dictionary of Microbiology and Molecular Biology
 ,
2nd
Edn.
Wiley
,
New York
.
[81]
Cendrin
F.
Chroboczek
J.
Zaccai
G.
Eisenberg
H.
Mevarech
M.
(
1993
)
Cloning, sequencing, and expression in Escherichia coli of the gene coding for malate dehydrogenase of the extremely halophilic Archaebacterium Haloarcula marismortui
.
Biochemistry
 
32
,
4310
4313
.
[82]
Obón
J.M.
Manjón
A.
Iborra
J.L.
(
1996
)
Comparative thermostability of glucose dehydrogenase from Haloferax mediterranei. Effects of salts and polyols
.
Enzymol. Microb. Technol.
 
19
,
352
360
.