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Caldalkalibacillus thermarum strain TA2.A1 grew in pH-controlled batch culture containing a fermentable growth substrate (i.e., sucrose) from pH 7.5 to 10.0 with no significant change in the specific growth rate, suggesting that this bacterium was a facultative alkaliphile. However, when strain TA2.A1 was grown on a nonfermentable carbon source, such as succinate or malate, no growth was observed until the external pH was >9.0, suggesting that this bacterium was an obligate alkaliphile. Succinate transport and sucrose transport by strain TA2.A1 showed pH profiles similar to that of growth on these carbon sources, and the molar growth yield on sucrose was higher at pH 9.5 than at pH 7.5, despite the increased energy demands on the cell for intracellular pH regulation. Succinate transport, succinate-dependent oxygen consumption, and succinate dehydrogenase and F1Fo-ATPase specific activities were all significantly lower in cultures of strain TA2.A1 grown at pH 7.5 than in those cultured at pH 9.5. No significant ATP synthesis via the F1Fo-ATP synthase was detected until the external pH was >8.5. On the basis of these results, we propose that nonfermentative thermoalkaliphilic growth is specialized to function at high pH values, but not at pH values near neutral pH.
Alkaliphilic microorganisms have been isolated from a diverse range of environments and have traditionally been classified into two distinct groups based on their pH profile for growth (8). Bacteria that grow across a broad pH range from 7.0 to 11.0 have been classified as facultative alkaliphiles (e.g., Bacillus pseudofirmus OF4) (28), and those that are able to grow only above pH 9.0 have been classified as obligate alkaliphiles (e.g., Bacillus alcalophilus) (4). The reasons why obligate alkaliphiles fail to grow below pH 9.0 remain speculative.
While the classification of alkaliphilic bacteria based on pH profiles for growth has gained universal acceptance, it does not consider the nature of the carbon source that is used to grow the cells, and for aerobic alkaliphiles, this may have important consequences. For example, growth on succinate in aerobic bacteria is strictly coupled to oxidative phosphorylation and ATP is produced in the cell via the membrane-bound F1Fo-ATP synthase. Growth on fermentable carbon sources, such as glucose, allows the cells to bypass this machinery, as ATP can be produced via substrate-level phosphorylation and incomplete oxidation of glucose to acetate can occur.
A thermoalkaliphilic bacterium, Bacillus sp. strain TA2.A1, capable of optimal aerobic growth at a temperature of 65°C at pH 9.5 was isolated from an alkaline thermal bore at Mt. Te Aroha, New Zealand (19). The 16S rRNA gene sequence of strain TA2.A1, compared with those available in the EMBL database, shows 99.5% similarity to Caldalkalibacillus thermarum strain HA6T, an aerobic, heterotrophic, thermophilic bacterium isolated from an alkaline hot spring in China (30). On the basis of the similarity of its phenotypic and genotypic characteristics to those of strain HA6T, we assign strain TA2.A1 to the genus and species Caldalkalibacillus thermarum. C. thermarum strain TA2.A1 grows on sucrose, common C4-dicarboxylates, glutamate, pyruvate, and trehalose; however, glucose and fructose fail to support growth (19). We originally described strain TA2.A1 as a facultative alkaliphile based on its pH profile for growth on glutamate or sucrose (18-20); however, both are substrates whose metabolism is not strictly coupled to oxidative phosphorylation.
In this communication, we determine the pH profile for growth of C. thermarum strain TA2.A1 on nonfermentable (i.e., succinate and malate) and fermentable carbon sources (i.e., sucrose) using pH-controlled batch culture and demonstrate that strain TA2.A1 was unable to grow below pH 9.0 in pH-controlled batch culture on succinate but grew from pH 7.5 to 10 on sucrose. The physiological and biochemical bases for this phenomenon were investigated.
Caldalkalibacillus thermarum strain TA2.A1 was routinely grown at 65°C in an alkaline basal medium with the following composition (per liter): 0.5 g of Na2SO4, 0.1 g of (NH4)2SO4, 0.1 g of MgSO4·7H2O, 0.2 g of K2HPO4, 9.0 g of NaHCO3, 10 g of tryptone peptone (Difco), 5 ml of Dictyoglomus trace elements (26), and 2 g of l-glutamate. The pH of the medium was adjusted to 9.5 at 65°C. When l-glutamate was excluded from this medium, it was replaced with either l-succinate, l-malate, or sucrose to a final concentration of 50 mM. For pH-controlled batch culture, cells were grown in a 500-ml fermentor equipped with an automatic pH control unit (catalog no. 5656-05; Cole-Palmer), and growth was initiated with a 0.1% inoculum from a culture grown overnight. The pH value being studied was maintained throughout growth by the addition of sterile 1.0 N NaOH or 1.0 N HCl. Cells were grown with aeration by constant stirring at 250 rpm with a six-bladed Rushton-type impeller (5.2-cm diameter). To monitor culture growth, samples were withdrawn aseptically when required, and the optical density at 600 nm (OD600) was measured (1-cm light path length).
To calculate the molar growth yield (in milligrams [dry weight] of cells per millimole of substrate consumed) on growth substrates from pH-controlled batch culture experiments, cells (50 ml in triplicate) were harvested by filtration and the filters (0.45 μm; Millipore) were dried at 105°C until the same weight (dry weight) was achieved on consecutive days. Cell-free supernatants were assayed for succinate, malate, or sucrose. Succinate and malate concentrations were determined using a succinate and malate detection kits (Boehringer Mannheim; Vintessential) coupled to NADH oxidation or NAD+ reduction, respectively, at 340 nm. Sucrose concentration was determined by a series of coupled reactions to NADP+ reduction at 340 nm as described previously (3).
Succinate transport assays were performed as previously described (19, 20). Strain TA2.A1 was grown in alkaline basal medium containing either succinate or sucrose (final concentration of 50 mM). Cells were harvested by centrifugation (18,000 × g at 4°C, 20 min) during the mid-exponential phase of growth and washed twice in 50 mM Tris-HCl (pH 9.0) containing 50 mM NaCl. The 50 mM Tris-HCl buffer was replaced with 50 mM morpholinoethanesulfonic acid (MES)-morpholinepropanesulfonic acid (MOPS)-Tris-HCl, when the effects of different pH values on succinate transport activity were measured. The cells were resuspended in appropriate buffer to achieve a concentration of 0.3 to 0.4 mg protein/ml. Each assay mixture contained 200 μl of cell suspension that was preincubated at 65°C in 5-ml polystyrene test tubes. Transport was initiated by the addition of either 50 nCi l-[U-14C]succinate (55 mCi/mmol; Sigma) or 50 nCi l-[U-14C]sucrose (630 mCi/mmol; Sigma). After 0 to 180 s of incubation, reactions were stopped by the addition of 2 ml ice-cold 0.1 M LiCl and rapid filtration through 0.45-μm membrane filters (Millipore), using a vacuum manifold (Millipore) with an applied vacuum of approximately 80 lb/in2. The filters were washed again with 2 ml of 0.1 M LiCl, dried at 50°C in 4-ml scintillation vials, and covered in 2 ml scintillation fluid (Amersham). The amount of radioactivity taken up by the cells was determined with a 1214 Rackbeta liquid scintillation counter (LKB Wallac) using the 14C window and counting each vial for 1 min. The amount of substrate taken up by the cells was calculated from the counts on the filters of each time point relative to the count for a time zero control (i.e., simultaneous addition of 14C-labeled substrate and LiCl), the total counts initially added to the assay, and the excess of cold substrate over 14C-labeled substrate. 14C-labeled substrate uptake values were expressed as nanomoles of substrate transported per minute per milligram of protein. Controls for nonspecific binding of 14C-labeled substrate were prepared by preincubating cells with either nigericin and valinomycin (15 μM each) or 1% toluene for 30 min prior to adding 14C-labeled substrate. When conducting competition and metabolic inhibitor studies, the metabolic inhibitors monensin, carbonyl cyanide m-chlorophenylhydrazone (CCCP), and competitive substrates (e.g., 5 mM unlabeled dicarboxylates and amino acids) were added to the assay buffer 15 min before the addition of [14C]succinate. All water-insoluble inhibitors were dissolved in 100% ethanol and compared to cells treated with appropriate volumes of 100% ethanol. Vmax values of solute transport were determined from Michaelis-Menten plots, and Km values were determined from double-reciprocal plots of the same data.
Inverted membrane vesicles were prepared as previously described (2). Right-side-out (RSO) membrane vesicles were prepared from mid-log-phase cultures of strain TA2.A1 (OD600 of 0.5). Cells were harvested, washed, and resuspended to a concentration of 1 g/ml in 100 mM potassium-phosphate buffer (pH 7.8) containing 20% sucrose. Protoplasts were prepared by the addition of lysozyme to a final concentration of 20 mg/ml, and the suspension was gently stirred at room temperature for 45 min. Protoplasts were pelleted by centrifugation at 11,600 × g, resuspended with an 18-gauge needle in 100 mM potassium phosphate buffer (pH 7.8) containing 20 mM MgSO4, 20% sucrose, and 5 mg/ml DNase I and broken by osmotic shock by diluting the protoplasts 100-fold with 50 mM potassium phosphate buffer (pH 7.8) containing 5 mM ADP. The suspension was stirred for 15 min at room temperature before the addition of 10 mM EDTA and 15 mM MgSO4, with stirring for 15 min at room temperature between steps. The suspension was centrifuged at 22,300 × g, and the pellet was resuspended in precooled 100 mM potassium phosphate buffer (pH 7.8) containing 10 mM EDTA and 5 mM ADP. Cellular debris and unbroken protoplasts were removed by low-speed centrifugation (480 × g, 15 min). RSO membrane vesicles were harvested by centrifuging the supernatant at 180,000 × g for 45 min and resuspending the vesicles in 100 mM potassium phosphate buffer (pH 7.8) containing 2 mM MgCl2 and 10% glycerol. Estimates of the fraction of vesicles in the opposite orientation using these methods vary, but they generally range from 10 to 20% (1, 27). To confirm vesicle orientation, NADH oxidation activity was measured in RSO membrane vesicles and compared to the same activity in inverted membrane vesicles. NADH (final concentration of 0.25 mM) oxidation activity was monitored continuously at 340 nm using a Cary 50 (Varian) spectrophotometer at 45°C.
ATP hydrolysis activity was measured using a spectrophotometric ATP-regenerating assay at 45°C as previously described (2). ATP synthesis in ADP-loaded RSO membrane vesicles (energized by 20 mM potassium ascorbate and 0.1 mM phenazine methosulfate) was carried out as previously described (15). Succinate-2,6-dichlorophenolindophenol (DCPIP) oxidoreductase activity in inverted membrane vesicles was determined spectrophotometrically (600 nm) at 45°C using a millimolar extinction coefficient of 21 mM−1 cm−1 for DCPIP. The reaction mix in 50 mM sodium phosphate buffer (pH 8.0) contained (per ml): 2 mM MgCl2, 0.1% Triton X-100, 0.5 mM DCPIP, 40 mM sodium azide, and 200 mM sodium succinate. The reaction was started by the addition of 0.7 mg inverted membrane vesicles, the solution was mixed gently, and the reaction was allowed to proceed at 45°C. Succinate-dependent oxygen consumption by strain TA2.A1 was measured using a Rank Brothers Clark-type electrode calibrated at 65°C. Exponentially growing cells were harvested and washed twice in alkaline basal medium lacking peptone and any additional carbon source (pH 9.0) and then resuspended in 50 mM MES-MOPS-Tris-HCl buffer (pH 7.0 to 10.0) to give a final OD600 of 1.0 (0.34 mg protein/ml). Oxygen consumption was initiated by the addition of 20 mM sodium succinate. Determination of the cellular protein was performed by the method of Markwell et al. (13).
When strain TA2.A1 was grown in non-pH-controlled batch culture at an initial pH of 7.5 on a nonfermentable carbon source, such as sucrose, glutamate, or succinate, the final pH was always >9.0 regardless of the carbon source (data not shown). These data demonstrated that the medium used to grow strain TA2.A1, even though strongly buffered (i.e., 100 mM NaHCO3), lacked the necessary buffering capacity to hold the external pH near neutral pH. Uninoculated medium was tested in parallel, and the pH remained unchanged over the same time period of incubation. On the basis of these observations, all growth experiments were performed in pH-controlled batch culture.
The pH limits for growth of strain TA2.A1 were examined using alkaline basal medium supplemented with either succinate, malate, or sucrose under pH-controlled conditions. Strain TA2.A1 grew at a doubling time (td) of 1.5 h in basal medium with no added carbon source, and the final OD600 was less than 0.2 over the entire pH range studied (Fig. (Fig.1A).1A). Strain TA2.A1, like the majority of other alkaliphilic microorganisms (9), has an obligate growth requirement for tryptone and is unable to grow in defined medium (minimal medium); therefore, the growth observed in the absence of an added carbon source most likely reflects the utilization of tryptone. When basal medium was supplemented with either succinate or malate at pH 7.5, the final OD600 was comparable to that of basal medium (Fig. (Fig.1A).1A). Robust growth on either malate (td = 1.83 h) or succinate (td = 1.95) was observed only when the external pH was >9.0 (Fig. (Fig.1A),1A), and no growth was observed at pH 10.5 (data not shown). Unlike growth on malate and succinate, growth on sucrose was comparable over the pH range 7.5 to 9.5 (td = 1.62 h to 1.5 h, respectively) (Fig. (Fig.1A)1A) with no growth at pH 10.5 (data not shown). To validate the premise that the final OD600 was an accurate indicator of cell growth, the dry weight of the cells from these experiments was measured (Fig. (Fig.1B).1B). The dry weight of cells produced/liter of culture showed the same trend as that obtained for OD600, supporting the observation that the growth of strain TA2.A1 on sucrose was pH independent and growth on malate and succinate was pH dependent (Fig. (Fig.1B).1B). The amount of substrate (i.e., sucrose, succinate, or malate) consumed in the growth experiments was measured to determine the molar growth yield on the respective carbon sources (Table (Table1).1). The molar growth yield on sucrose increased as a function of pH, demonstrating that sucrose was utilized more efficiently at higher pH (Table (Table1).1). The molar growth yields on succinate and malate were comparable at high pHs, indicating that both substrates were utilized efficiently under these conditions (Table (Table11).
These data suggest that consideration should be given to the nature of the carbon source when describing alkaliphilic bacteria as obligate or facultative on the basis of their pH profile for growth. For example, strain TA2.A1 could be termed a facultative alkaliphile (i.e., pH 7.5 to 10.0) during growth on fermentable carbon sources but an obligate alkaliphile during growth on nonfermentable carbon sources (i.e., pH > 9.0). However, the question remains as to why obligate alkaliphiles fail to grow below pH 9.0 even on fermentable carbon sources.
We hypothesize that the molecular machinery required for growth on succinate or malate is adapted to function at high pH values and not at lower pH values. To investigate this hypothesis, succinate transport and sucrose transport in strain TA2.A1 were monitored as a function of external pH. Cells grown at pH 9.5 on succinate, washed, and resuspended in 50 mM Tris-HCl (pH 9.0) containing 50 mM NaCl exhibited [14C]succinate uptake that was linear over a period of approximately 180 s (Fig. (Fig.2A).2A). Succinate uptake under these conditions was inhibited by either 1% toluene (vol/vol) (Fig. (Fig.2A),2A), monensin (10 μM), or CCCP (100 μM) (Fig. (Fig.2B),2B), demonstrating that succinate transport was an energy-dependent process. Succinate uptake was examined as a function of external succinate concentration at pH 9.0 and 50 mM NaCl (Fig. (Fig.2C).2C). Cells demonstrated Michaelis-Menten saturation kinetics for succinate uptake, and the apparent Km and Vmax for [14C]succinate transport were 19 μM and 0.45 nmol succinate·min−1 mg protein−1, respectively (Fig. (Fig.2C,2C, inset). As sodium is the predominant ion for solute transport in alkaliphiles that have been studied (11, 19, 20), further investigations were made into the requirement for extracellular sodium and the influence of pH on [14C]succinate uptake. The kinetics of succinate uptake was determined at pH 9.0 in the presence of increasing NaCl concentrations (Fig. (Fig.2D).2D). Succinate transport exhibited Michaelis-Menten saturation kinetics with increasing concentrations of NaCl; the apparent Km for sodium was 14 mM, and the apparent Vmax was 0.42 nmol succinate·min−1 mg protein−1 (Fig. (Fig.2D,2D, inset). The transport of succinate was dependent on NaCl, and Na+ ions could not be substituted by other monovalent cations, i.e., NH4Cl, KCl, CaCl2, LiCl, or CsCl (data not shown).
The effect of external pH on the transport of succinate and sucrose was studied (Fig. (Fig.3).3). When cells were grown on succinate at pH 9.5, the pH profile of succinate transport by whole cells exhibited a pH optimum of 9.0 to 9.5 (0.14 nmol·min−1 mg protein−1), with a sharp decrease in the rate of succinate transport observed below pH 9.0 and above pH values of 9.5 (Fig. (Fig.3A).3A). In contrast, sucrose transport by the same cells showed a broad pH profile (Fig. (Fig.3A).3A). Cells grown at pH 9.5 on sucrose had equivalent rates of sucrose and succinate uptake at pH values of 9.0 (Fig. (Fig.3B).3B). As the external pH of the transport assay was lowered to pH 7.2, succinate and sucrose uptake decreased, but the decrease for succinate was significantly higher than for sucrose. When cells were grown on sucrose at pH 7.5, the level of succinate uptake was reduced by >90% (0.01 nmol·min−1 mg protein−1), even when the growth medium also included 50 mM succinate (Fig. (Fig.3C).3C). Sucrose uptake showed a pH profile similar to that observed in cells grown at pH 9.5. The specificity of succinate transport was determined in whole cells of strain TA2.A1 at pH 9.0 by measuring the uptake of [14C]succinate in the presence of a 50-fold excess of competing substrate, i.e., a variety of C4-dicarboxylates and Krebs cycle intermediates. The ability of a competitor substrate to inhibit succinate transport was defined as a >20% inhibitory effect on the initial rate of [14C]succinate transport. [14C]succinate transport was strongly inhibited by C4-dicarboxylates, i.e., malate, fumarate, and aspartate (Fig. (Fig.3D),3D), but not by α-ketoglutarate, glutamate, and pyruvate, suggesting that a general C4-dicarboxylate permease is present in strain TA2.A1.
On the basis of these results, both succinate permease activity and succinate permease synthesis were regulated by pH. In contrast, sucrose transport was constitutively expressed, and transport activity was not significantly regulated by external pH. Sucrose and succinate transport systems were dependent on sodium and membrane potential, and these driving forces are maintained at constant values over the pH range 7.5 to 10 (18). This implies that the inability of cells to transport succinate at low pH does not reflect a lack of appropriate driving force but suggests that the succinate permease is specialized to function over a narrow pH range. This is backed up at a molecular level by the inability of strain TA2.A1 to express the genes for succinate transport when grown at pH 7.5 on sucrose, even when succinate (inducer) is included in the growth medium. Succinate transport was via a general C4-dicarboxylate permease, and inspection of the genome sequence for strain TA2.A1 reveals the presence of a dctPQM operon, which encodes all three subunits of a tripartite ATP-independent periplasmic (TRAP) transporter. The strain TA2.A1 TRAP-type transporter has 30 to 40% similarity to other C4-dicarboxylate TRAP transport systems previously described that commonly use ion gradients (e.g., Na+ and H+) to drive solute transport (10, 17).
The observation that C4-dicarboxylate transport was pH regulated prompted an investigation into what other components of the oxidative phosphorylation machinery were also regulated by external pH. Succinate-dependent oxygen consumption was examined in whole cells of strain TA2.A1 and showed a pH profile that had an optimum at pH values of 9.0 to 9.5 (Fig. (Fig.4A).4A). The level of succinate dehydrogenase activity (pH optimum 7.5) was determined in cells grown at either pH 7.5 or pH 9.5 (Fig. (Fig.4B).4B). Cells grown at pH 7.5 on sucrose or sucrose-containing medium plus succinate exhibited low levels of succinate dehydrogenase activity (Fig. (Fig.4B).4B). When cells were grown on either sucrose or succinate at pH 9.5, the specific activity increased three- to fourfold (Fig. (Fig.4B).4B). A similar pattern of enzyme activity (i.e., ATP hydrolysis) was shown for the F1Fo-ATP synthase of strain TA2.A1 (Fig. (Fig.4C).4C). Recently, we demonstrated that ATP synthesis by the F1Fo-ATP synthase of strain TA2.A1 (TA2F1Fo) expressed in Escherichia coli membranes has a pH optimum of 9.0 to 9.5 with no measurable ATP synthesis at pH 7.5 (15). Stirred and aerated ADP- plus Pi-loaded RSO membrane vesicles of strain TA2.A1 were energized with 20 mM potassium ascorbate and 0.1 mM phenazine methosulfate, and ATP synthesis was measured at various pH values (Fig. (Fig.4D).4D). The optimum pH for ATP synthesis was 9.0 to 9.5, showing that in its native lipid membrane environment, the TA2F1Fo functions over a very narrow pH range (Fig. (Fig.4D),4D), as previously shown for the recombinant TA2F1Fo in E. coli membranes (15). ATP synthesis activity was inhibited by DCCD over this pH range, demonstrating that the observed ATP synthesis was due to membrane-bound F1Fo-ATP synthase activity and not substrate-level phosphorylation (Fig. (Fig.4D4D).
The pH-dependent expression of several enzymes and cofactors involved in oxidative phosphorylation has been previously investigated in alkaliphiles (23-25, 31). The results of these studies demonstrate that components of the oxidative phosphorylation machinery, i.e., cytochrome caa3 oxidase (23, 24), b- and c-type cytochromes (31), and succinate:quinone oxidoreductase (25), all show a higher level of activity and expression at pH values near 10.0 than in cells grown at pH 7.5. Moreover, vesicles from cells of B. pseudofirmus OF4 grown at pH 10.5 have higher rates of ATP synthesis at pH 10.5 than those from cells grown at pH 7.5 (5). These data would also appear to be congruent with reported molar growth yields that are higher at pH 10.5 than at pH 7.5 (28), despite the increased energy demand for pH homeostasis. In addition, the generation times of the facultative alkaliphile B. pseudofirmus OF4 are 54 min at pH 7.5 and 38 min at pH 10.6 in malate-containing medium (28).
Taken together, these data demonstrate that the levels of expression of various components involved in energy generation of alkaliphilic bacteria are greater at alkaline pH values than at low pH values, suggesting that the oxidative phosphorylation machinery may have special adaptations that allow it to function optimally at alkaline pH values but less efficiently at neutral pH. The pH optima of both sodium-dependent succinate transport and ATP synthesis by the F1Fo-ATP synthase are in the pH range 9.0 to 9.5, supporting this contention. The ATP synthase itself from alkaliphilic Bacillus species exhibits several features that are unique, namely, latent ATPase activity (2, 6, 7), a-subunit modifications (15, 29), and a larger oligomeric c ring (14, 16). Enlarged c rings are also found in alkaliphilic cyanobacteria (21, 22). The observation in B. pseudofirmus OF4 that the caa3 terminal oxidase interacts with the F1Fo-ATP synthase in a 1:1 stoichiometry suggests that these two complexes may be coordinately regulated to achieve optimal ATP synthesis coupled to proton pumping of the respiratory chain to generate the membrane potential (12). Current studies are aimed at determining the transcriptional response of C. thermarum strain TA2.A1 to external pH to identify the gene complement required for thermoalkaliphilic growth.
This work and D.G.G.M. and S.K. were supported by a Marsden grant from the Royal Society of New Zealand.
Published ahead of print on 23 October 2009.