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a Deps. of Agronomy and Microbiology, 2537 Agronomy Hall, Iowa State Univ., Ames, IA 50011-1010 USA
b Dep. of Environmental Science, Policy, and Management, 151 Hilgard Hall, Univ. of California, Berkeley, CA 94720-3110 USA
larryh{at}iastate.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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Abbreviations: EDTA, ethylenediamine-tetraacetic acid • SEM, standard error of the mean • GMA, glucose minimal agar • TCA, trichloroacetic acid • TSA, trypticase soy agar
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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m) in dry soils and/or low solute water potential (s) in saline soils (Kieft et al., 1987; Harris, 1981). Since water moves freely through the cell membrane, the internal water potential of these microbes must be in equilibrium with the external environment. Many microorganisms accumulate intracellular organic and inorganic solutes such as K+, amino acids, carbohydrates, polyols, and quarternary amines, which are compatible with cellular metabolic processes (Harris, 1981; Killham and Firestone, 1984b; Measures, 1975; Yancey et al., 1982; Csonka, 1989; Le Rudulier et al., 1984; Vreeland, 1987), to achieve the intracellular water potential necessary for maintaining the proper cellular turgor pressure required for growth and survival. Some microbes respond to drying by forming desiccation-resistant spores or cysts (Gould and Measures, 1977), but many soil bacteria tolerate low water potentials as vegetative cells (Busse and Bottomley, 1989; Williams, 1985).
Soil water potential routinely fluctuates with time, declining gradually with soil drying (percolation, evaporation, evapotranspiration) then increasing rapidly upon wetting (irrigation, rainfall). The wetting of a dry soil can cause a rapid increase in water potential of surface soil from less than -20 MPa to almost zero (Evans et al., 1975) and, hence, may be the most severe environmental stress experienced by many surface soil organisms (Smith, 1979). Microorganisms subjected to sudden increases in water potential experience an immediate influx of water, which is driven by the difference in water potential between the cell cytoplasm ( low) and the cells' immediate environment ( high). This influx of water increases turgor pressure, which may cause physiological impairment, cell death, or even cell lysis (Brown, 1979; Harris, 1981; Luard, 1982; Salema et al., 1982). In an earlier study of two California soils, we showed that sudden wetting of dry soil resulting in water potential increases of 2.8 MPa caused the release of 17 to 27% of the soil microbial biomass C into the environment (Kieft et al., 1987). The mechanism causing this rapid release of soil microbial biomass C was not determined. This release of soil microbial biomass C could be the result of cell lysis or the rapid reduction in internal solute pools to maintain the proper cell turgor pressure. Possible mechanisms for reducing osmotically active solutes include (i) active or passive release of intracellular solutes to the environment (Britten and McClure, 1962; Christian, 1962; MacLeod et al., 1978; Reed et al., 1986; Smith, 1979), (ii) catabolism of compatible organic solutes to CO2, and (iii) polymerization of solutes to reduce osmotic activity (Avron and Ben-Amotz, 1979; Berrier et al., 1992; Reed and Stewart, 1983). The release of solutes into the environment could result in the loss of C in an environment in which C is the primary nutrient limiting microbial growth. Since many soils frequently experience sizeable fluctuations in water potential, soil bacteria may have evolved a variety of mechanisms for tolerating sudden increases in water potential. Considering the complexity of soil microbial community composition, different species of soil bacteria may have different mechanisms for responding to a sudden increase in water potential following the wetting of a dry soil.
The objective of this study was to characterize the physiological responses of four soil bacteria to a sudden increase in solution water potential (dilution stress) to better understand the mechanisms through which the microbial community responds to the wetting of a dry soil. Bacterial isolates from three genera commonly isolated from soil (Pseudomonas, Streptomyces, and Bacillus) were cultured at a low solute water potential and subjected to various magnitudes of dilution to determine how they respond to sudden increases in water potential. We examined the effects of dilution on cell culturability and on intracellular solute pools to determine whether solutes were released to the environment, catabolized to CO2, or polymerized into osmotically less active compounds. These data were compared with previously published data on bacteria from a range of habitats to assess patterns in mechanisms of acclimation to water potential increase.
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Materials and methods |
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TOPABSTRACTINTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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Media and Growth Conditions
For all experiments, cells were grown at 25°C in some form of medium 21-C, a peptone–glucose based medium (Smibert and Krieg, 1981). In the dilution stress experiments, cells were grown in 100 mL of 21-C medium supplemented with 32.1 g L-1 NaCl to reduce the water potential to -3.0 MPa in 500-mL, triple-baffled nepheloculture flasks (Bellco Biotechnology, Vineland, NJ). All isotonic solutions consisted of deionized water amended with NaCl to achieve the desired water potential. Water potentials of all solutions were confirmed by measurement with a psychrometer (HR 33T dewpoint microvoltmeter, Wescor C52 sample chamber, Decagon Devices, Pullman, WA) at 25°C.
Dilution Stress Protocol
All strains were grown in -3.0 MPa 21-C medium, harvested in log phase
by centrifugation at 10000 g and 5°C for 15 min, then resuspended in 26 mL of a -3.0 MPa isotonic solution. A 1-mL aliquot was removed for dilution plating before the culture was placed into a 25°C water bath. Fifty milliliters of the appropriate NaCl diluent was added to each culture flask and mixed with a magnetic stir bar. Depending on the NaCl concentration of the diluent, the cells were subjected to a rapid (
5 s) increase of 0.5 (25.7 g L-1 NaCl), 1.0 (19.3 g L-1 NaCl), 1.5 (12.8 g L-1 NaCl) or 2.0 (6.4 g L-1 NaCl) MPa in water potential. Control flasks received isotonic NaCl (38.4 g L-1) solution to maintain a -3.0 MPa water potential. After mixing, the diluted cultures were incubated at 25°C for 15 min to allow the cells to adjust to the new water potential. A 1-mL aliquot was then removed from each culture for plating prior to splitting the culture in half and harvesting the culture by centrifugation to generate two cell-pellets. The resulting supernatants and cell pellets were stored at -20°C. Using the protocol outlined above, we also used KCl or MgSO4 as the ion in the diluent in a series of experiments to assess whether amino acid release patterns of P. chlororaphis in response to dilution was ion specific.
Culturability Measurements
Effect of dilution on cell culturability was determined by comparing culture plate counts before and after dilution. Samples were serially diluted in isotonic NaCl solutions and aliquots were plated onto TSA and/or glucose minimal agar (GMA). The GMA medium contained 9.17 g L-1 K2 HPO4·3H20, 2.0 g L-1 KH2PO4, 0.5 g L-1 sodium citrate, 0.1 g L-1 MgSO4, 1.0 g L-1 (NH4)2SO4, 1.0 g L-1 glucose, 15.0 g L-1 Bacto-Agar (Difco, Detroit, MI). Streptomyces griseus and B. pumulis did not grow on GMA and were cultured on TSA only. Plates were incubated at 25°C for 3 to 5 d before counting.
Protein, Amino Acid, and DNA Analyses
One of the two cell pellets from each dilution stress treatment was resuspended in 1 mL of 1 M NaOH and incubated at 100°C for 10 min to solubilize cell proteins (Hanson and Phillips, 1981). Cellular and extracellular protein contents were measured by the Bradford method (Bio-Rad, Hercules, CA) with bovine serum albumin (BSA) as the standard; pH of cellular protein samples were neutralized prior to protein determinations. Intracellular amino acids were extracted from the second pellet by resuspending in 1 mL of 20% (200 g L-1) trichloroacetic acid (TCA) and incubating the mixture overnight at 5°C (Killham and Firestone, 1984b). Insoluble cell material was removed by centrifugation (40 000 g at 5°C for 30 min), with the resulting supernatant containing the amino acids. To extract cellular DNA, cell pellets were mixed with 5% (50 g L-1) trichloroacetic acid and incubated for 1 h at 100°C. Intracellular (pellet) and extracellular (supernatant) amino acids and DNA concentrations were quantified by the ninhydrin (Rosen, 1957) and diphenylamine (Clark and Switzer, 1977) methods, respectively. The standard for amino acid determinations included equimolar concentrations of arginine, histidine, threonine, and glutamic acid. The standard for DNA determinations was calf thymus DNA (Sigma Chemical, St. Louis, MO).
Carbohydrate Analyses
For determining the cellular pool size of low and high molecular weight carbohydrates, cell pellets were resuspended in 1 M NaOH, heated for 15 min at 70°C, and then centrifuged at 35000 g for 30 min at 5°C (Breedveld et al., 1991, 1990). High molecular weight cellular carbohydrates in the alkaline supernatant were precipitated by mixing the supernatant with one volume ice-cold absolute ethanol, incubating overnight at -20°C, and centrifuging as above. The high molecular weight cellular carbohydrates (pellets) were resuspended in 1 M NaOH. The low molecular weight cellular carbohydrates remaining in the alkaline, ethanolic supernatant were concentrated by roto-evaporation. For determining the extracellular pool size of low and high molecular weight carbohydrates, the high molecular weight carbohydrates present in the original supernatants of cultures from control or dilution stress treatments were ethanol precipitated by adding three volumes ice-cold ethanol and incubating the ethanolic supernatants overnight at -20°C. The ethanolic supernatant was then centrifuged at 35000 g for 30 min at 5°C, the supernatant decanted, and the high molecular weight extracellular carbohydrates in the pellet were resuspended in deionized water. The low molecular weight extracellular carbohydrates remaining in the ethanolic supernatant were concentrated by roto-evaporation. Uronic acids were measured by the meta-hydroxydiphenyl method (Blumenkrantz and Osboe-Hansen, 1973) and neutral hexoses were measured by the phenol-sulfuric acid method (Ashwell, 1966). Standards were glucuronic acid and glucose for the uronic acid and neutral hexoses determinations, respectively.
Toluene and Lysozyme–EDTA Treatments
The effects of increased membrane permeability (toluene treatment) and cell wall removal (lysozyme–EDTA treatment) on protein and amino acid release were examined. Cultures were harvested by centrifugation as described earlier, and the cell pellets were resuspended in 15 mL of either a -3.0 MPa isotonic solution (toluene treatments) or a buffer solution (lysozyme–EDTA treatments). For the toluene treatments, 120 µL of toluene was added to the resuspended cell suspensions and mixed (Dobrogosz, 1981). Two different buffer solutions were used in the lysozyme–EDTA experiments. For S. griseus and B. pumulis, the buffer solution consisted of 0.1 M K2HPO4 buffer (pH 8.0) containing 21 µM lysozyme (Sigma Chemical Co.), and the cell suspensions were incubated at 25°C for 1 h (Koenigs et al., 1973). For the Pseudomonas spp., the buffer solution consisted of 100 mM Tris buffer (pH 8.0) containing 445 µM EDTA and 0.7 µM lysozyme, and the cell suspensions were incubated for 30 min at 25°C (Repaske, 1956). After incubation, cultures were split into two fractions and harvested by centrifugation (14 000 g and 5°C for 20 min). The supernatants and cell pellets were analyzed for amino acid and protein content as described earlier.
Carbon-14 Labeling of Cell Constituents
Previous work indicated that S. griseus accumulates proline as its main compatible solute (Killham and Firestone, 1984a), while P. chlororaphis accumulates glutamate (Jones, 1988). Cultures for experiments involving 14C labeling of cellular constituents were grown in -3.0 MPa 21-C medium containing 0.33 g L-1 of yeast extract (Difco) and 12.5 mM of either glutamate (P. chlororaphis) or proline (S. griseus). Exponential phase cultures
were 14C-labeled by adding 1.63 x 105 Bq [l-14C] glutamate or 1.41 x 105 Bq [l-14C] proline
, (Amersham Co., Arlington Heights, IL). A base trap containing 0.5 mL of 1M NaOH was added to each flask to measure respired 14C-CO2. Cultures were incubated for 30 min after labeling and then harvested as described below. Base traps were removed at harvest and assayed for 14C.
Cultures were harvested by centrifugation at 20000 g and 5°C for 10 min, resuspended in a -3.0 MPa isotonic solution and recentrifuged; a 1-mL sample of the supernatant was saved to determine the amount of 14C-labeled substrate remaining. The cell pellets were resuspended and pooled in a -3.0 MPa isotonic solution and exposed to a 2 MPa dilution stress as described above. Each culture flask was fitted with a base trap immediately after dilution and allowed to equilibrate at 25°C for 15 min after which the base traps were removed to determine the amount of 14C-CO2 that was respired. The cultures were harvested by centrifugation, and a 1-mL sample of the supernatant was used to determine the amount of 14C-labeled cellular constituents released into the extracellular environment. Intracellular solutes were extracted from cell pellets with TCA as described earlier and the resulting supernatant was analyzed to determine the amount of 14C-labeled solutes. The remaining cell pellet was resuspended in 1 mL of an isotonic NaCl solution to determine the amount of 14C-labeled macromolecules. Samples for 14C analysis were placed in glass scintillation vials, mixed with 10 mL of Scintiverse 2 scintillation cocktail (Fisher Scientific, Pittsburgh, PA), capped, shaken vigorously, and assayed in a liquid scintillation counter.
Data Presentation and Analysis
In all experiments, the proportion of cellular constituents released was derived by dividing extracellular contents by the sum of the intra- and extracellular contents. The data are presented as mean net percentage release ± standard error of the mean (SEM) of three to four replications. These values were obtained by subtracting control values from treatments in which there was a water potential increase. For the experiment comparing the effects of lysozyme–EDTA treatments and toluene treatments on amino acid and protein release, the data were arcsin-transformed for the ANOVA, and treatment means were compared by Duncan's new multiple range test (Snedecor and Cochran, 1980). Culturability was expressed as the percentage culturable relative to the undiluted control.
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Results |
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TOPABSTRACTINTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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Effects of Dilution on Culturability
In general, dilution did not decrease S. griseus and B. pumulis culturability on TSA (data not shown). Culturability of P. chlororaphis was affected more by the medium onto which it was plated, whereas P. fluorescens was affected more by the magnitude of the dilution (Fig. 4)
. Reduction in culturability (70–80%) following a 2-MPa dilution for the two Pseudomonas spp. was greater than the level of cell lysis (1%) at this dilution (Fig. 1). This suggests that following large increases in water potential the Pseudomonas spp. became physiologically impaired and nonculturable.
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Discussion |
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TOPABSTRACTINTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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Previous studies of two California soils showed that sudden wetting of dry soil resulting in water potential increases of 2.8 MPa caused the release of 17 to 27% of the soil microbial biomass C (Kieft et al., 1987). The responses of the four bacterial species tested here suggests that some of the soil microbial biomass C released upon wetting of dry soils may result from export of organic compatible solutes. In this study, we examined only bacterial responses to changes in solute water potential. Yet, microbes in dry soils would experience sudden changes in both matric and solute water potentials when dry soils are wetted. Although bacterial response mechanisms to sudden changes in matric potential could be different than those reported here, it is generally accepted that the response mechanisms for adjusting intracellular water potentials would be similar for sudden increases in matric and solute potential (Brown, 1990). While we report only on response mechanisms used by soil bacteria, soil fungi appear to respond to water potential increases using similar physiological strategies (Brown, 1990; Carlile and Watkinson, 1994).
Bacteria produce a variety of organic solutes that are compatible with cell physiology when they are grown in high osmolarity media. We only examined the release of amino acids and carbohydrates following dilution stress and other osmotically active organic or inorganic constituents (e.g., glycine betaine, ectoines, N-acetylglutaminylglutamine amide, nucleotides, or K+) could have been released (Britten and McClure, 1962; Christian, 1962; Malin and Lapidot, 1996; Reed et al., 1986; Schleyer et al., 1993; Tschicholz and Trüper, 1990). Quantification of the full complement of compatible solutes released following dilution of a greater variety of soil bacteria would provide for a better understanding of the importance of this response mechanism in causing the pulse of mineralization known to occur after dry soil is wetted.
The mechanism by which solutes are released and whether solute release is an active or passive process is not known. Pressure- or stretch-activated channels have been reported in both Gram-negative (Berrier et al., 1992; Martinac et al., 1987) and Gram-positive bacteria (Berrier et al., 1992); differences in the osmolarity across a membrane of as little as a few milliosmolar is sufficient to activate these channels. There is also evidence to suggest that different stretch-activated channels exist with different molecular mass exclusion limits (Berrier et al., 1992). When cells that are adapted to low water potentials are exposed to a dilution stress, solutes need to be released very rapidly, and stretch-activated channels may be sufficiently large to accommodate passage of a variety of compatible solutes.
Several investigators have demonstrated that within minutes after dilution, cells begin to preferentially reassimilate some of their previously released solutes (Gauthier et al., 1991; Schleyer et al., 1993; Tschicholz and Trüper, 1990). This period of reassimilation occurs before there are visible signs of growth. Since we allowed our postdilution cultures to equilibrate for 15 min prior to analysis for solute release, the values reported may reflect some reassimilation and may underestimate total solute release. Hyperosmotically grown Escherichia coli reassimilate solutes after dilution, and solute reassimilation enhances survival of E. coli in seawater (Gauthier et al., 1991). We observed that culturability of P. chlororaphis after dilution was substantially affected by the medium on which it was plated (Fig. 4). The better culturability of P. chlororaphis on TSA than GSA medium could be due to the greater abundance of osmotically active solutes in TSA than GMA. In soil, the release of solutes into the environment following a wetting event might yield a relatively rich source of solutes and nutrients for cells that have been impaired by dilution stress.
The responses of the two Gram-positive isolates and two Pseudomonas spp. to dilution varied considerably. In general, B. pumulis and S. griseus were more tolerant to dilution, since culturability was unaffected and the amount of solutes released was very small (Fig. 2 and 4). Pseudomonas fluorescens appeared less tolerant of dilution stress than P. chlororaphis. Pseudomonas fluorescens tolerated a 0.5 MPa water potential increase, but water potential increases of greater magnitude resulted in a dramatic reduction in culturability and a greater release of solutes (Fig. 2, 3, and 4). This decrease in culturability was apparently not due to cell lysis (Fig. 1) and possibly reflects physiological stress in response to dilution. Such a marked threshold response to dilution may be related to salt toxicity; P. fluorescens was isolated from a nonsaline soil (Gamble, 1977). Microorganisms from saline environments have been shown to be more salt tolerant and to more effectively remove Na+ once it has entered the cell (Imhoff et al., 1983; Kogut and Russell, 1984; Vreeland, 1987).
We have compiled data on the effect of rapid increases of water potential on the release of solutes from previous studies and present them with those from our study to identify patterns in solute release between different types of organisms and the habitats from which they originated (Table 3) . Since the solute release data reported in Table 3 reflect the time frame of analysis, the choice of organic solutes examined, the type of medium used for cultivating the organism, and the different osmotic activity of each solute, it is difficult to directly compare data from different studies. It is evident, however, that bacteria commonly respond to rapid and large increases in water potential by releasing a significant portion of their organic solute pool. In addition, Gram-positive bacteria release less of their internal solute pool per unit change in water potential than do Gram-negative bacteria. This is consistent with the theory proposed by Harris (1981) that Gram-positive bacteria are relatively tolerant of dilution stress due to their cell wall architecture and greater cell turgor pressure.
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ACKNOWLEDGMENTS |
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Received for publication June 28, 1999.
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TOPABSTRACTINTRODUCTIONMaterials and methodsResultsDiscussionREFERENCES |
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