INTRODUCTION

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Ultramicrobacteria are major contributors to the world's biosphere in terms of biological cycling of carbon, nitrogen, and phosphorus (50). As reservoirs of nutrients in oligotrophic marine ecosystems, they interact with all trophic levels and control nutrient fluxes via mineralization, thus having an impact on the productivity of all marine life from microbial primary producers and plankton to whales. Because of predictions of increasing ocean oligotrophy as a consequence of global warming (32, 59), it is clearly important to understand the physiology of this class of bacteria in order to determine the impact that they have on life on earth.

The growth of virtually all microbial cells in nature is limited by the availability of one or more essential growth nutrients (22, 37, 50), and in many regions of the ocean carbon is the primary limiting substrate (1, 4, 5, 27). In the oligotrophic marine environment, bacteria generally adopt one of two different survival strategies; they are either copiotrophic organisms which form resting stage cells with spasmodic bursts of growth (e.g., Vibrio angustum S14) or oligotrophic organisms which grow slowly with intermittent periods of starvation or faster growth (e.g., Sphingomonas sp. strain RB2256) (10).

Despite our relatively good understanding of the physiology and genetics of marine copiotrophic bacteria, oligotrophic bacteria and the roles that they play in environmental processes are poorly understood. Knowledge about the physiology of oligotrophs is limited by the availability of environmental isolates. To date, most insight into the physiology of this class of marine bacteria has been obtained from Sphingomonas sp. strain RB2256 (8, 10, 11, 48, 49), which was isolated as a numerically dominant bacterium from Resurrection Bay, Alaska (3, 47). This strain has been formally described as Sphingomonas alaskensis RB2256 (57).

One of the characteristics that distinguish S. alaskensis RB2256 from V. angustum S14 is its high level of resistance to a variety of stress-inducing agents, including hydrogen peroxide (8). The ability to resist the damaging effects of hydrogen peroxide is an ecologically relevant characteristic because endogenous and exogenous oxidative stress is a common challenge for microorganisms in aquatic environments (6, 17, 42, 50). Reactive oxygen species, such as hydrogen peroxide, damage DNA, RNA, proteins, and lipids, and as a consequence, cells have evolved a broad range of mechanisms to cope with this type of stress (reviewed in reference 52).

Previous studies showed that S. alaskensis RB2256 grown in glucose-limited chemostats at a dilution rate of 0.027 h¹ was more resistant to hydrogen peroxide than logarithmic-phase or starved cells from batch cultures were (8). In view of the fact that slow, nutrient-limited growth is likely to be the type of growth most often exhibited by oligotrophic bacteria, we reasoned that the high degree of resistance observed with chemostat-grown cells may be triggered by nutrient-limited growth and that the precise level of resistance is controlled by the actual specific rate of growth under these conditions.

In order to determine the types of mechanisms and regulatory processes that S. alaskensis RB2256 has evolved to cope with hydrogen peroxide stress, in this study we examined the physiological and molecular responses of cells grown in nutrient-limited chemostats at different growth rates. Growth in chemostats permitted continued growth at a fixed rate, while it also permitted the use of different limiting nutrients (e.g., carbon or nitrogen).

While the growth rate still has been poorly studied, there is evidence that growth rate affects the physiology of Escherichia coli and Vibrio spp. The cell size, cellular composition, and starvation survival of Vibrio sp. strain ANT-300 are affected by growth rate (37, 38), and slow growth induces rpoS-dependent gene expression in E. coli (13, 39). In this study we extended assessment of the growth rate control of hydrogen peroxide resistance to include a marine copiotrophic bacterium (V. angustum S14). We found that growth rate played a critical role in the hydrogen peroxide resistance of S. alaskensis RB2256 and that slowly growing cells were more resistant than fast-growing cells. In contrast, in V. angustum nutrient availability caused an all-or-none type of response, and starved cells were more resistant than growing cells, regardless of the growth rate. These results indicate that the extent of nutrient limitation has fundamentally different consequences for the physiology of oligotrophic ultramicrobacteria and the physiology of copiotrophic bacteria.

	MATERIALS AND METHODS

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Bacteria, media, and culture conditions. The bacterial strains used were the oligotrophic marine ultramicrobacterium strain S. alaskensis RB2256 (46, 47, 57) and V. angustum S14 (31). For cultivation in batch cultures, S. alaskensis RB2256 and V. angustum S14 were grown in an artificial seawater medium (ASW) (8) supplemented with 3 mM D-glucose. For chemostat cultivation, ASW was used with supplements. Glucose-limited feed medium contained D-glucose at a concentration of 3 mM. For ammonium-limited feed medium the concentration of NH₄Cl was 94 mM and 5 mM D-glucose was added. For mixed-amino-acid-limited feed medium the concentration of Casamino Acids was 0.5% (wt/vol) and glucose was omitted. The pH values of the media used for batch and chemostat cultures were maintained between 7.5 and 7.8 by addition of morpholinepropanesulfonic acid (MOPS) buffer (1.0 g liter¹) or by continuous automatic adjustment with sterile NaOH (0.25 M). The pH of each medium was adjusted to 7.8 prior to autoclaving. Batch cultures were grown at 30°C with orbital shaking at 120 rpm. Chemostat cultivation was carried out in glass culture vessels (working volume, 450 ml) magnetically stirred at 400 rpm and maintained at a constant temperature of 30°C. Small-scale chemostat cultivation was carried out in 100-ml glass culture vessels constructed from Erlenmyer flasks, which had small headspace volumes, were equipped with two stainless steel baffles, and were stirred magnetically at 400 rpm. The temperature was maintained by a continuous flow of water through a water jacket from a temperature-regulated water bath. Medium entered each vessel through a stainless steel needle (diameter, 0.5 mm) at the bottom of the vessel. Filter-sterilized, prehumidified air was supplied through the same needle. The specific growth rates imposed on chemostat cultures were alternated between high and low values, and hydrogen peroxide resistance was monitored as described below. In chemostat cultures, a steady state was assumed after growth for five to seven generations. The growth of batch and chemostat cultures was monitored by measuring the optical density at 433 nm.

Viability measurements and hydrogen peroxide exposure. Viable counts of S. alaskensis RB2256 and V. angustum S14 were determined by determining the number of CFU on marine nutrient agar (Bacto Marine Agar 2216) and on ASW-glucose solid medium consisting of ASW, 3 mM D-glucose, and 1.5% agar. A dilution series was prepared with ASW buffered with MOPS (1.0 g liter¹; pH 7.8). Colonies on drop plates (20) were counted with a binocular microscope (magnification, ×25) after 3 and 6 days of incubation at 30°C for S. alaskensis RB2256 and after overnight incubation at 30°C for V. angustum S14. At least five spots from duplicate plates were counted for each experiment. Experiments were performed at least twice. The survival fraction for hydrogen peroxide-treated samples was calculated for each sample separately as a percentage of the survival in untreated control samples. Cells were withdrawn from mid-exponential-phase batch cultures (optical density at 433 nm, 0.4) or steady-state chemostat cultures and exposed to hydrogen peroxide (2 to 75 mM; freshly prepared from a 30% commercial stock solution) at 30°C for up to 60 min. S. alaskensis RB2256 is inherently more resistant to hydrogen peroxide than V. angustum S14. To obtain equivalent levels of survival after 60 min, 25 and 2 mM hydrogen peroxide were used for S. alaskensis RB2256 and V. angustum S14, respectively.

Determination of catalase activity. Catalase activity was determined with a Clark oxygen electrode by the method of Rørth and Jensen (45). The concentration of oxygen in oxygen-saturated deionized water at 25°C was assumed to be 253 µM (60). The amount of enzyme activity that decomposed 1 µmol of H₂O₂ to 0.5 µmol of O₂ min¹ at 25°C was defined as 1 U of activity. Total catalase activity was determined by subtracting background respiration and oxygen production due to spontaneous decomposition of H₂O₂. The background activities were less than 10% of the measured oxygen production values. Catalase activities were determined from initial reaction rates from linear regression lines calculated over the first minute. Comparisons of the catalase activities of whole bacteria were made after the addition of H₂O₂ (final concentration, 1 mM) to 2.0 ml of culture. Enzyme activity was calculated from the initial reaction rate for at least five replicates from at least two independently prepared samples. Protein concentrations were determined by using a bicinchoninic acid kit (Sigma) with bovine serum albumin as the standard.

Two-dimensional polyacrylamide gel electrophoresis. Triplicate gels of total protein were prepared from at least two independent steady-state cultures of S. alaskensis RB2256 growing at rates of 0.026, 0.076, and 0.18 h¹. Sample preparation, two-dimensional polyacrylamide gel electrophoresis, silver staining, image acquisition, and analysis were carried out as previously described (12).

	RESULTS

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Effect of growth rate on hydrogen peroxide resistance in S. alaskensis RB2256. To determine whether growth rate affected the stress resistance of S. alaskensis RB2256, cells were grown at different dilution rates in a glucose-limited chemostat at 30°C, samples were removed, and survival was monitored after exposure to 25 mM hydrogen peroxide for 60 min. The specific growth rates used ranged from 0.02 to 0.18 h¹, which corresponded to approximately 10 to 90% of the maximum specific growth rate (0.21 h¹). In addition, resistance was determined for cells grown at the maximum rate in a batch culture.

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View larger version (15K): [in this window] [in a new window]   FIG. 1.   Percentages of survival of S. alaskensis RB2256 cells grown at different specific growth rates in glucose-limited chemostats after exposure to 25 mM hydrogen peroxide for 60 min. For the two trendlines the data were grouped. Symbols: , low specific growth rates (0.020, 0.023, 0.026, 0.058, 0.076, 0.095, 0.11, 0.12, and 0.13 h1); , high specific growth rates (0.14, 0.15, 0.16, 0.17, and 0.18 h1). The level of survival of logarithmic-phase cells in batch cultures () is also shown. Duplicate samples were taken from chemostats at every specific growth rate that was tested. Six individual chemostat runs were tested, and at least one sample was taken from a high specific growth rate and a low specific growth rate. The numbers of CFU were determined by using the drop plate method with marine nutrient agar and ASW-glucose solid medium. There was no difference in survival between ASW-glucose solid medium and marine nutrient agar for any time point. The standard deviation for each time point was less than 15%.

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View larger version (15K): [in this window] [in a new window]   FIG. 2.   Percentages of survival of nutrient-limited chemostat-grown S. alaskensis RB2256 cells following exposure to 25 mM hydrogen peroxide for up to 60 min. Experiments were performed twice, and CFU were counted by using the drop plate method with marine nutrient agar and ASW-glucose solid medium. There was no difference in survival between ASW-glucose solid medium and marine nutrient agar for any time point. The results of representative experiments are shown. The standard deviation for each time point was less than 20%. (A) Samples taken directly from steady-state glucose-limited chemostats at growth rates of 0.026 h1 (), 0.076 h1 (), 0.13 h1 (), 0.14 h1 (), 0.16 h1 (), and 0.18 h1 () and from a batch culture with a maximum specific growth rate of 0.21 h1 (). (B) Samples taken directly from ammonium-limited chemostats at growth rates of 0.15 h1 (*) and 0.050 h1 (). (C) Samples taken directly from mixed-amino-acid-limited steady-state chemostats at growth rates of 0.15 h1 () and 0.047 h1 ().

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Carbon limitation versus nitrogen limitation. To determine whether the link between growth rate and hydrogen peroxide resistance was affected by the nature of the limiting substrate in the chemostat, the responses of carbon-limited cultures (Fig. 1 and 2A) were compared to the responses of nitrogen-limited cultures (Fig. 2C). Ammonium-limited cultures were grown at low (0.050 h¹) and high (0.15 h¹) rates of growth, and samples taken from cultures after a steady state had been obtained were exposed to 25 mM hydrogen peroxide. These nitrogen-limited cultures were far more sensitive to hydrogen peroxide than any of the glucose-limited cultures were, exhibiting almost undetectable survival after 20 min of exposure. Furthermore, no difference in stress resistance was observed at the two specific growth rates used.

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Effect of growth rate on hydrogen peroxide resistance in V. angustum S14. To determine whether the rate of growth affected the ability of V. angustum S14 to survive after exposure to hydrogen peroxide, like S. alaskensis, cells were grown in glucose-limited chemostats at specific growth rates of 0.023, 0.20, and 0.60 h¹ and then exposed to stress in a preparation containing 2 mM hydrogen peroxide for up to 60 min (Fig. 3). In ASW defined minimal medium with 3 mM glucose as the sole carbon and energy source, growth rates of 0.023 and 0.60 h¹ corresponded to approximately 5 and 95% of the maximum specific growth rate (0.62 h¹), respectively. The resistance of the chemostat-grown cells was also compared with the resistance of cells grown at the maximum rate in batch culture and following 24 h of starvation (Fig. 3).

View larger version (24K): [in this window] [in a new window]   FIG. 3.   Percentages of survival of V. angustum S14 following exposure to 2 mM hydrogen peroxide for up to 60 min after growth at various rates in glucose-limited chemostats. Samples were taken directly from steady-state chemostats at growth rates of 0.023 h1 (), 0.20 h1 (), and 0.60 h1 () and from batch cultures in the logarithmic phase () and after 24 h of starvation (). Experiments were performed twice, and CFU were counted by using the drop plate method with marine nutrient agar and ASW-glucose solid medium for each time point. For each data point the standard deviation was less than 15%.

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Catalase activity and hydrogen peroxide resistance in S. alaskensis RB2256. When hydrogen peroxide was added to liquid cultures of S. alaskensis RB2256 cells, small bubbles developed after 5 to 30 min. This occurred irrespective of the growth rate and whether the cells were grown in batch or chemostat cultures. The production of bubbles may have resulted from production of O₂ due to decomposition of hydrogen peroxide by catalase.

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Protein profiles for S. alaskensis RB2256 grown at different rates of growth in glucose-limited chemostat cultures. Approximately 1,500 spots can be resolved by high-resolution two-dimensional polyacrylamide gel electrophoresis of proteins from S. alaskensis RB2256 (12). To determine the changes in gene expression that are associated with high and low rates of growth and to identify the genes involved in the increased levels of hydrogen peroxide resistance at low rates of growth, two-dimensional polyacrylamide gel electrophoresis profiles were generated in triplicate for cells grown at rates of 0.026, 0.076, and 0.18 h¹ (Fig. 4). Protein spots for each preparation were separated evenly throughout the resolved pI range (pI 4 to 7) and molecular mass range (10 to 80 kDa). Up to 1,600 protein spots were detected on each silver-stained gel, and 1,049 spots conserved from triplicate gels obtained for each growth rate were analyzed. Relative spot intensity was calculated as the percentage of optical density by comparison with the total optical density of each gel. The intensities of spots ranged from 0.216 to 4.5%.

View larger version (141K): [in this window] [in a new window]   FIG. 4.   Two-dimensional gel showing proteins from S. alaskensis RB2256 grown in glucose-limited chemostats. Total protein was extracted from steady-state cultures growing at rates of 0.026, 0.076, and 0.18 h1. Protein profiles were visualized by silver staining. The protein profile shown is the profile from a culture growing at a rate of 0.18 h1, and the highlighted spots have relative intensities that were at least two-fold less or greater in at least one growth condition. Symbols: , spots unique to a growth rate of 0.026 h1; , spots unique to a growth rate of 0.076 h1; , spots unique to a growth rate of 0.18 h1; , spots found at growth rates of 0.026 and 0.076 h1; , spots found at growth rates of 0.076 and 0.18 h1. The molecular weight and pI values for specific spots were assigned by using Melanie II software (Bio-Rad) and are shown in Table 1.

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	DISCUSSION

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Growth rate control of hydrogen peroxide resistance. We previously found that cultures of S. alaskensis RB2256 grown at 25°C in a glucose-limited chemostat (growth rate, 0.027 h¹) were more resistant to hydrogen peroxide than cells grown in a batch culture (growth rate, 0.16 h¹) (8). In the present study we examined whether there was a link between the actual specific growth rate of a culture and hydrogen peroxide resistance and whether the resistance of cells was significantly affected by the mode of growth (e.g., carbon- or nitrogen-limited growth in chemostats and growth under nutrient-excess conditions in batch cultures). It is indeed apparent that hydrogen peroxide resistance in S. alaskensis RB2256 is strongly influenced by the specific growth rates of cultures when glucose or mixed amino acids are the limiting substrates (Fig. 1 and 2A and C); however, growth rate control of hydrogen peroxide stress resistance was not apparent under nitrogen limitation conditions (Fig. 2B). Thus, a comparison of stress survival of cells grown under carbon or energy limitation conditions with stress survival of ammonium-limited cells grown at comparable dilution rates in chemostats (approximately 0.03 to 0.05 and 0.14 to 0.16 h¹, respectively) clearly showed that the specific rate of growth is not the only major determinant of the level of stress resistance in S. alaskensis RB2256. The results indicate that hydrogen peroxide stress resistance in S. alaskensis RB2256 is influenced directly or indirectly both by the nature of the limiting substrate (carbon or energy versus nitrogen) and by the extent of nutrient limitation in carbon- and/or energy-limited chemostat cultures. In contrast, exponentially growing or starved cells from batch cultures had levels of resistance equivalent to those of carbon-limited, chemostat-grown cells at high dilution rates.

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Possible mechanisms of growth rate control of hydrogen peroxide resistance. The phenotypic responses of S. alaskensis RB2256 are consistent with a regulatory cascade in which very small changes in the concentration of effector molecules and/or proteins become amplified through the cascade. Such regulatory events may be mediated by global regulators, such as rpoS, oxyR, and soxRS, and other mechanisms, such as DNA methylation and stringent control, which are important in oxidative stress responses in E. coli (9, 52).

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Starvation versus low-growth-rate induction of peroxide stress resistance. In almost all cases starvation survival and in some cases starvation-induced cross-protection are dependent on the substrate for which the culture is starved. V. angustum starvation-induced survival, miniaturization, and stress resistance are specific for carbon starvation and not for nitrogen or phosphorus starvation (41). E. coli viability is more sensitive to phosphate starvation than to carbon or nitrogen starvation (7), while porin regulation in response to glucose limitation is not observed in nitrogen-limited cultures (29). An explanation based on the ecology of marine bacteria may be related to the fact that most pelagic heterotrophs are limited by the availability of carbon (1, 4, 5, 27) and adaptive responses are geared for carbon starvation.