Water Stress-Driven Changes in Bacterial Cell Surface Properties

ABSTRACT

Increased drought intensity and frequency exposes soil bacteria to prolonged water stress. While numerous studies reported on behavioral and physiological mechanisms of bacterial adaptation to water stress, changes in bacterial cell surface properties during adaptation are not well researched. We studied adaptive changes in cell surface hydrophobicity (CSH) after exposure to osmotic (NaCl) and matric stress (polyethylene glycol 8000 [PEG 8000]) for six typical soil bacteria (Bacillus subtilis, Arthrobacter chlorophenolicus, Pseudomonas fluorescens, Novosphingobium aromaticivorans, Rhodococcus erythropolis, and Mycobacterium pallens) covering a wide range of cell surface properties. Additional physicochemical parameters (surface chemical composition, surface charge, cell size and stiffness) of B. subtilis and P. fluorescens were analyzed to understand their possible contribution to CSH development. Changes in CSH caused by osmotic and matric stress depend on strain and stress type. CSH of B. subtilis and P. fluorescens increased with stress intensity, R. erythropolis and M. pallens exhibited a generally high but constant contact angle, while the response of A. chlorophenolicus and N. aromaticivorans depended on growth conditions and stress type. Osmotically driven changes in CSH of B. subtilis and P. fluorescens are accompanied by increasing surface N/C ratio, suggesting an increase in protein concentration within the cell wall. Cell envelope proteins thus presumably control bacterial CSH in two ways: (i) by increases in the relative density of surface proteins due to efflux of cytoplasmic water and subsequent cell shrinkage, and (ii) by destabilization of cell wall proteins, resulting in conformational changes which render the surface more hydrophobic.

IMPORTANCE Changes in precipitation frequency, intensity, and temporal distribution are projected to result in increased frequency and intensity of droughts and heavy rainfall events. Prolonged droughts can promote the development of soil water repellency (SWR); this impacts the infiltration and distribution of water in the soil profile, exposing soil microorganisms to water stress. Exposure to water stress has recently been reported to result in increased cell surface hydrophobicity. However, the mechanism of this development is poorly understood. This study investigates the changes in the physicochemical properties of bacterial cell surfaces under water stress as a possible mechanism of increased surface hydrophobicity. Our results improve understanding of the microbial response to water stress in terms of surface properties, the variations in stress response depending on cell wall composition, and its contribution to the development of SWR.

INTRODUCTION

In porous media such as soil, bacteria usually grow attached to solid-liquid interfaces rather than becoming suspended in the soil solution (1). Adhesion to solid surfaces, as well as to other microbes, by forming microcolonies enables bacteria to better cope with and survive environmental stress (2). Bacterial adhesion is determined by the surface properties of the substratum, as well as by bacterial cell wall physicochemical properties such as cell surface hydrophobicity (CSH), surface free energy, surface charge, and chemical composition (3–7). In the case of CSH, it has been observed that more hydrophobic cells show stronger adhesion to solid surfaces (6). Bacteria may exhibit a wide range of surface hydrophobicity degrees depending on the species, growth phase, and growth substrate (6, 8, 9). In addition, an increase in CSH has also been reported as a general stress response mechanism related to water stress, i.e., limited water availability. Water stress can be experienced by the cells in form of matric stress (caused by increasing proportion of capillary-bound water at low-soil water contents) or salt stress (osmotic stress resulting from increased salt concentration [10, 11]).

Adhesion of bacteria to surfaces has important implications within the environment. For example, living and dead bacterial biomass are important constituents of soil organic matter (SOM). Thus, bacteria imprint their properties to organic coatings on soil mineral surfaces (12) and therefore are thought to significantly contribute to the formation of soil water repellency (SWR) (3), i.e., a reduced wettability, as has been found in several soils (13–15) and identified as an important control of water distribution in soil. As a consequence, reduced wettability of soil particles also limits water availability for soil microorganisms (16), exposing them to water stress (osmotic and matric) and thereby potentially increasing their CSH. Despite its relevance to the ecological and hydrological functions of soils (17, 18), this feedback loop has been largely neglected. Therefore, we investigated how surface properties of soil bacteria change upon exposure to water stress.

Bacterial cell surface properties may be determined by the presence and conformation of various amphiphilic compounds carrying a range of functional groups (19). Proteins have been reported to play important roles in controlling the wettability of bacterial cell surfaces, with increasing protein content resulting in increasing hydrophobicity. Environmental factors, such as water stress, can provoke changes in the chemical properties of bacterial cell surfaces (11, 20), which can, for example, be determined by X-ray photoelectron spectroscopy (XPS) (21). The chemical composition of the cell envelope also strongly affects surface free energy, which determines bacterial cell wettability and can be quantified by contact angle (θ_w) measurements (22). Together with the electrostatic cell surface properties, which can be calculated from the zeta potential, the cell surface free energy determines the total interaction free energy and is crucial for the adhesion of bacterial cells to other particles such as mineral grains or other cells. Under physiological conditions, the bacterial cell surface is typically negatively charged due to the presence of carboxyl and phosphate groups on the cell envelope surface (19). Exposure to water stress, alongside the increasing ionic strength of the soil solution, can promote bacterial adhesion to solid surfaces by reducing the range of electrostatic repulsive forces of the electric double layer (5, 23–25).

The cell surface may exhibit specific physical characteristics and topographic features, such as surface roughness. Atomic force microscopy (AFM) is one of the most widespread techniques for examining the physical properties of biological substances at the nanometer scale (26). The ability to image cells and cell wall ultrastructure under physiological conditions has given new insights into various processes, e.g., cell growth (27), cell division (28), cell wall assembly (29), and membrane damage (30). AFM is also suitable for studying the nanomechanical properties of cells (31). Thereby, AFM maps and force spectroscopy may be used to obtain information on the physical properties of bacteria, such as surface roughness and elasticity, which are important for their adhesion to substrates such as soil minerals, but also to study their wettability (32).

While numerous studies have reported on the impact of water stress (mostly in the form of osmotic stress) on different bacterial properties and functions (33), there are no comprehensive studies on its impact on the surface properties of typical soil bacteria. Likewise, little is known about the effect of water stress induced by matric potential, the main component of total water potential in nonsaline soils. Moreover, most studies are performed in liquid cultures, although these conditions are rarely present in non-water-saturated soil, where only relatively thin water films are present on particle surfaces.

Therefore, this study aims to gain more insights into the impact of (i) NaCl-induced osmotic stress and (ii) polyethylen glycol 8000 (PEG 8000)-induced matric stress on an entire set of physicochemical properties of microbial cell surfaces. Water stress-driven changes in microbial CSH, a potential contributor to SWR development, were analyzed for six typical soil bacteria. For the two strains with the most pronounced reaction to osmotic and matric stress, additional properties were analyzed to reveal the mechanisms of the water-stress response. These included important factors controlling bacterial adhesion to solid phases, i.e., bacterial cell surface zeta potential, solid surface free energy, and surface chemical composition, as well as nanoscale form and elasticity of the cells. We hypothesized that exposure to osmotic and matric stress will result in increased CSH promoted by changes in cell wall physicochemical properties.

RESULTS

Bacterial growth inhibition and cell surface hydrophobicity.

Bacterial growth was inhibited with decreasing water potential for all strains under both osmotic and matric stress (Fig. S1 in the supplemental material). Osmotically driven growth inhibition was also observed during submerged and surface growth. Because matric stress was not tested for surface growth, no information on the roles of growth conditions at different levels of matric stress was obtained from our study. In summary, the extent of the bacterial response to water stress varied depending on strain, growth condition, and stress type.

Strain-dependent variations in the θ_w of unstressed bacterial cells were observed to range between 40° and 110°. With exception of A. chlorophenolicus, which exhibited higher levels of surface hydrophobicity during surface growth, no differences in the θ_w of unstressed cells were observed between surface and submersed growth. An increase in CSH was observed for most of the strains in response to reduced water potential for one or both stress conditions (Fig. 1). Increased contact angles were observed for P. fluorescens and B. subtilis for both stress types. Under osmotic stress, cell surface θ_w had already increased at water potential values of −0.6 MPa (P < 0.0001). Similarly, −0.85 MPa matric potential resulted in significant increases in θ_w for both strains (P < 0.05). The responses of A. chlorophenolicus and N. aromaticivorans differed depending on stress type (osmotic versus matric stress). R. erythropolis (109 ± 1°) and M. pallens (82 ± 1°) already exhibited high hydrophobicity in cells grown without water stress and no significant changes in θ_w with increasing water potential. Furthermore, response to osmotic stress varied depending on growth conditions. Osmotic stress affected cell surface θ_w during surface growth only for P. fluorescens and N. aromaticivorans. Both strains already showed a significant increase in θ_w at an osmotic potential of −0.5 MPa (P < 0.0001).

FIG 1.

Bacterial cell surface hydrophobicity, expressed as contact angle, under different growth conditions. Green, osmotic stress in submersed growth; blue, osmotic stress in surface growth; orange, matric stress in submersed growth. Values are arithmetic means of 8 measurements for each of n = 3 biological replications, with error bars indicating standard error.

Solid surface free energy.

Solid surface free energy and its components showed pronounced differences between the two strains studied, B. subtilis and P. fluorescens, and were affected by the intensity and type of stress (Fig. 2). Total solid surface free energy (γ_s ^TOT) ranged between 26 and 45 mJ m⁻² and was dominated by the nonpolar Lifshitz-van der Waals component (γ_s^LW), except for osmotically stressed cells of B. subtilis, with an additional considerable contribution of the acid/base component (γ_s^AB). Regarding the effect of stress and the type of stress treatment, no clear and consistent pattern was found for total solid surface free energy (γ_s ^TOT) as well as for the acid/base (γ_s^AB) and Lifshitz-van der Waals (γ_s^LW) components. However, while the electron acceptor (γ_s⁺) component was generally low (<2.2 mJ m⁻²), the electron donor (γ_s⁻) component of both, B. subtilis and P. fluorescens, was considerably lower for both osmotic and matric stress-affected cells than for unstressed cells.

FIG 2.

Contact angle and solid surface free energy (γ_s) components of unstressed (control) and stressed (osmotic stress: −2.9 MPa; matric stress: −3.8 MPa) cells of Bacillus subtilis and Pseudomonas fluorescens. γs⁺, γs^–, γ_s^AB, and γ_s^LW indicate the electron acceptor, electron donor, acid/base, and Lifshitz-van der Waals components, respectively, and γ_s^TOT indicates the total solid surface free energy. Error bars indicate standard errors (n = 9, for each of the three testing liquids) and different lowercase letters indicate significant (P < 0.05) differences in contact angle between treatments of the respective strain.

Zeta potential.

To measure the surface charge of the bacterial cell wall, the zeta potentials of P. fluorescens and B. subtilis were measured, and pronounced differences in zeta potential were found between the two strains. B. subtilis had a significantly (P < 0.001) more negative zeta potential (−37 ± 1 mV) than P. fluorescens (−9 ± 1 mV; Fig. 3).

FIG 3.

Zeta potentials of unstressed (blue) and stressed (green, osmotic stress: −2.9 MPa; orange, matric stress: −3.8 MPa) cells of B. subtilis and P. fluorescens. Error bars indicate standard error (n = 10) and letters indicate significant (P < 0.05) differences between treatments.

The strains showed contrasting behavior with respect to stress, resulting in slightly more negative zeta potentials for B. subtilis (P < 0.05) and P. fluorescens (P = 0.001) cells for osmotically stressed compared to unstressed control cells. Exposure to matric stress caused a significantly more negative zeta potential for P. fluorescens (P < 0.001) compared to unstressed and osmotically stressed cells, while no difference was observed for B. subtilis (P = 0.27) between cells exposed to matric stress and unstressed cells.

X-ray photoelectron spectroscopy.

XPS measurements revealed some pronounced differences in surface elemental composition between the strains (Fig. 4). Surface C concentration was found to be significantly lower (P < 0.001), and surface O and P concentration were significantly higher (P < 0.01 and P < 0.001, respectively) for B. subtilis than for P. fluorescens.

FIG 4.

Surface chemical composition of unstressed (control) and stressed (osmotic stress: −2.9 MPa; matric stress: −3.8 MPa) cells of B. subtilis (dark blue) and P. fluorescens (light blue). Error bars indicate standard error (n = 3) and different lowercase (B. subtilis) or uppercase (P. fluorescens) letters indicate significant (P < 0.05) differences in surface chemical composition between treatments of the respective strains.

Regarding the effect of stress on surface chemical composition, the XPS measurements revealed an inconsistent picture for both B. subtilis and P. fluorescens. Specifically, we could not find a statistically significant change that pointed in the same direction for both strains and both stress types for any of the surface elements. Moreover, we found that both types of stress tended to result in an increase in surface N and a decrease in surface C. This combination resulted in a significant increase in the surface N/C ratio (P < 0.05, except for matric stress-affected cells of B. subtilis), being directly related to the contact angle (Fig. 5a). Estimation of the mass percentage of the main cell surface compounds revealed a stress-induced increase in proteins and a decrease in hydrocarbon-like compounds (Fig. 5b) for both strains and both stress types. For both strains, osmotic stress decreased, and matric stress increased the amount of surface polysaccharides.

FIG 5.

Relationship between (a) contact angle and the atomic N/C ratio and (b) mass percentage of surface polysaccharides, proteins, and hydrocarbons for unstressed (blue) and stressed (green, osmotic stress [Ψ_O]: −2.9 MPa; orange, matric stress [Ψ_M]: −3.8 MPa) cells of B. subtilis (circles) and P. fluorescens (triangles). Error bars indicate standard error (contact angle: n = 9; X-ray photoelectron spectroscopy: n = 3).

Atomic force microscopy.

The nanoscale physical and mechanical properties of B. subtilis and P. fluorescens do not significantly differ between strains (Fig. 6). Exposure to osmotic stress resulted in a significant decrease in the average height for both strains and an increase in the average log DMT modulus (Derjaguin-Muller-Toporov model of the Young’s modulus). However, the latter was significant only for P. fluorescens. The increased log DMT modulus indicates a reduced elasticity and increased stiffness of the cell surface upon osmotic stress that was more strongly expressed in P. fluorescens. In contrast, there was a clear but insignificant trend that the shrinkage in height was, on average, more strongly expressed in B. subtilis.

FIG 6.

(a) Exemplary AFM maps of unstressed (control) and stressed (osmotic stress: −2.9 MPa) cells of B. subtilis and P. fluorescens of height sensor and log DMT channels. (b and c) Boxplots of (b) height and (c) log DMT modulus of 18 to 30 unstressed and osmotically stressed cells of B. subtilis and P. fluorescens, with different letters indicating significant (P < 0.05) differences.

DISCUSSION

Our results demonstrate reduced growth and changes in cell surface properties for all bacterial strains studied upon exposure to both osmotic and matric stress. We found characteristic shifts in bacterial growth rates, cell surface contact angles, cell surface chemical properties, and charge, as well as cell height and stiffness, after exposure to increased external osmotic and matric potentials.

Growth rate reduction by water stress.

Bacterial growth rates were reduced by both osmotic and matric stress for all bacterial strains. However, the maximum stress level without growth reduction and the slope of growth reduction varied depending on the strain and the type of stress imposed (osmotic versus matric). Nevertheless, our results agree with those of a number of earlier studies reporting osmotically driven growth-rate reduction in pure culture experiments (10, 34, 35) and in soil (36–39). In general, exposure to stress results in increased energy requirements for cell maintenance, reducing the energy available for new biomass production (40, 41), resulting in reduced growth rates.

Surface physicochemical properties of the strains and their implications for soil particle surface properties.

Bacteria can exhibit a wide range of degrees of surface hydrophobicity, and the degree of hydrophobicity can vary between different species of the same genus. Furthermore, the degree of CSH was found to be positively correlated with the ability to adhere to various surfaces (6), such as soil minerals. In a previous study using differently sizes of quartz minerals as models for soil particles, attachment of P. putida cells to mineral surfaces was demonstrated, and cell attachment induced mineral surface hydrophobicity (3). The surface hydrophobicity of the formed cell-mineral associations is more pronounced if the cells are exposed to osmotic stress prior to the attachment. We performed additional experiments to test how the attachment of Gram-positive (B. subtilis) and Gram-negative (P. fluorescens) bacteria onto mineral surfaces would impact the surface hydrophobicity of the resulting cell-mineral associations. In agreement with the results reported by Achtenhagen et al. (3), the surface hydrophobicity of cell-mineral associations increased, with B. subtilis having a stronger impact than P. fluorescens (42). These results are in line with our those of our further investigation of the bacterial community compositions of two soils with different degrees of hydrophobicity, showing that a higher degree of SWR was associated with a higher relative abundance of Gram-positive bacteria, in particular, representatives of Actinobacteria (42).

Our results also show a wide range of θ_w in unstressed cells, with most strains having rather hydrophilic surface properties. Only M. pallens and R. erythropolis exhibited high levels of hydrophobicity for the unstressed cells. This can be attributed to the presence of mycolic acids in the cell walls of bacteria of the Mycolata taxon (including Mycobacterium and Rhodococcus [43]). These compounds render the surfaces of these cells hydrophobic (44).

Further investigation of cell surface properties of B. subtilis and P. fluorescens revealed a significantly more negative zeta potential of B. subtilis cells at the selected pH of 7. This can be attributed to the higher abundance of anionic phosphate groups within the teichoic acids present in the peptidoglycan layer of Gram-positive strains (45). However, studies assessing the impact of various cationic agents (46) and antimicrobial peptides (47) on bacterial cell wall zeta potential have reported contradicting results, showing slightly higher zeta potentials for Gram-negative Escherichia coli compared to the Gram-positive Staphylococcus aureus. It must be kept in mind, however, that the zeta potential may vary with strain, growth conditions, and media composition, as well as with the conditions of zeta potential measurements, including the ionic strength and pH (48).

XPS measurements of B. subtilis and P. fluorescens cell surface chemistry also showed pronounced differences in surface elemental composition between the strains occurring due to the differences between the cell wall structures of Gram-positive and Gram-negative bacteria (49, 50). The cell surface of B. subtilis showed a lower protein and a higher polysaccharide concentration than P. fluorescens. This, again, can be explained by the presence of teichoic acids. Similarly, the significantly larger amounts of P found in the cell wall of Gram-positive B. subtilis is also due to the presence of teichoic acids, which are composed largely of polyol phosphates (45, 51, 52).

Water stress-induced changes in cell surface hydrophobicity.

Using our experimental approach, we showed that reduction in the growth rates of typical soil bacteria due to water stress (osmotic and matric) is accompanied by an increase in bacterial CSH during surface and submerged growth conditions. The response to water stress varies depending on stress type and level, growth condition, and strain. Variations in changes in CSH can be best attributed to the different cell wall compositions of different strains, their susceptibility to environmental conditions, and their initial physiological state (53, 54). Stress-induced increase in surface hydrophobicity was more pronounced for strains with lower initial degrees of surface hydrophobicity in unstressed cultures. Increases in cell surface contact angle have been already reported for different osmotically stressed P. putida strains (3, 10, 11), associated with the release of outer membrane vesicles (11). In contrast, the CSH of initially hydrophobic cells such as M. pallens and R. erythropolis were not affected by water stress. Exposure to osmotic stress has been reported to result in changes to the mycolic acid composition of R. opacus PWD4 accompanied by increasing cell surface θ_w (44). This contradicts our observation because neither Mycobacterium nor Rhodococcus strains exhibited responses to water stress in our study. However, the cell surface contact angles of five different mycolic acid-containing Rhodococcus (including two R. erythropolis, 34 to 56 C atoms) and three Mycobacterium (60 to 90 C atoms) were reported to range from 70° to 100° and 85° to 98°, respectively, which is in accordance with the value attained in this study.

Analysis of solid surface free energy and its components showed that the electron donor (γ_s⁻) component of both B. subtilis and P. fluorescens was considerably lower for both osmotic and matric stress-affected cells than for unstressed cells. According to van Oss (22), the electron donor component is associated with the interaction between colloidal particles and collector surfaces, allowing the derivation of general trends for the adhesion potential of bacterial cells to mineral particles. Increasing cell adhesion with an increasing γ_s^LW/γ_s^– ratio has been reported in a study of biofilm adhesion behavior (55). This suggests that stressed cells of both B. subtilis and P. fluorescens have a higher tendency toward adhesion and the formation of more stable cell-mineral associations compared to unstressed cells. This is in line with the observed stress-induced increase in hydrophobicity, which is generally known to promote bacterial adhesion (56) as an important strategy to cope with environmental stress (2).

Changes in surface elemental composition upon stress.

Although the effect of stress on single surface elements was inconsistent for B. subtilis and P. fluorescens, both types of stress resulted in an increased surface N/C ratio, which was positively correlated with CSH. This suggests a stress-induced increase in N-rich compounds, presumably proteins, at the cell surface. These results support findings by Cowan et al. (57) for Gram-positive cocci, suggesting that particularly N-containing compounds (i.e., proteins) may control bacterial CSH (20). The change in the abundance of N-containing compounds on the cell surface can thus be related to the changes in physical cell properties upon stress.

Surface properties determined by physical stress response.

Our AFM analyses revealed that water stress induced cell-size reduction and increased cell stiffness, whereas XPS indicated an increased surface N/C ratio. Based on these results, we propose that changes in surface hydrophobicity of bacterial cells occur as a result of changes in both the density and conformation of cell membrane surface proteins. If not exposed to osmotic stress, bacterial cells keep the amount of biomass components (in terms of dry mass per cell volume) approximately constant, independent of nutrient conditions, during their cell cycle (58–60). However, exposure to osmotic stress may result in efflux of cytoplasmatic water from the cell (61), while the dry weight stays unchanged (62). This loss of water and consequently of cell volume from bacterial cells can result in membrane molecular crowding at the cell surface (63, 64). As a result, the protein/lipid ratio of the cell envelope increases, accompanied by an increase in the overall CSH. In addition, osmotically driven membrane macromolecular crowding can lead to the destabilization of cell envelope proteins (65), affecting the structure and conformational stability of cell wall proteins (63, 66). An increase in protein density on the cell wall can lead to increased protein-protein interaction. This has been shown to affect the native structure of proteins, causing conformational shifts (67), and may be an explanation for the increased stiffness observed in our study. Changes in protein conformation (e.g., partial unfolding) would presumably expose the hydrophobic regions to the outer membrane surface, potentially leading to increased CSH. An increase in the relative content of hydrophobic domains on bacterial cell walls is also supported by the reduction of the electron donor (γ_s^–) component of cell surface free energy. The different responses of the two strains to osmotic stress, in terms of stiffness (increase in P. fluorescens and no significant impact in B. subtilis), can be attributed to the thick and initially more rigid cell wall of the Gram-positive B. subtilis, compared to the Gram-negative P. fluorescens cells with their thinner and softer peptidoglycan layer (50).

Conclusions.

We analyzed water stress-driven changes in the cell surface wettability of various Gram-negative and Gram-positive soil bacteria, combined with nanoscale physical properties (e.g., size and stiffness), surface chemical composition, and charge. Our results contributed to the understanding of probable mechanisms responsible for changes in bacterial cell hydrophobicity by providing insights into the changes in surface physicochemical properties which occur simultaneously with an increase in CSH and their potential contribution to the development of SWR.

Increased CSH due to stress exposure was observed for most strains with hydrophilic cell surfaces. In addition, we found that water stress-driven increases in CSH were correlated with increases in the N/C ratio, reductions in cell size, and strain-dependent increases in cell stiffness.

Surface hydrophobicity of soil bacteria may significantly alter upon drought stress, but the reactions of different strains are highly variable. The degree and type of changes depend on the type of drought stress, growth conditions, and cell wall architecture (i.e., Gram-positive versus Gram-negative). Nevertheless, the results reported here suggest that cell wall hydrophobicity can aid bacterial survival under water stress, because water stress resulted in increased surface hydrophobicity for all the strains with initially more hydrophilic cell walls. Presumably, a hydrophobic cell surface protects the cell from further excessive water loss and increases attachment of cells to surfaces and biofilm formation. This allows protection of the cell interior from hostile environmental conditions.

The efflux of cytoplasmatic water under osmotic stress resulted in cell shrinkage, driving structural and conformational changes in bacterial cell walls. A significant increase in content and possible osmotically driven destabilization of the conformation of surface proteins suggests an increased protein density and an increase in hydrophobic domains on the cell surface due to conformational shifts.

Surface hydrophobicity of bacterial cells plays a crucial role in their adhesion to surfaces. Therefore, one possible consequence of water stress-driven increases in bacterial CSH may be an increase in the propensity of microorganisms to adhere to soil particles, thereby reducing the wettability of these particles. Furthermore, bacterial living biomass and necromass affect the quality of SOM, a determining factor for the occurrence of SWR, indicating a possible involvement of soil bacteria in the development of soil hydrophobicity. The general mechanism we observed may be modulated by the texture or other physicochemical properties of the soil as well as by the architecture of the soil microbial community present.

MATERIALS AND METHODS

Microorganisms and growth conditions.

Six strains of typical soil bacteria were selected for the experiments, Bacillus subtilis DSM 10, Arthrobacter chlorophenolicus DSM 12829, Pseudomonas fluorescens DSM 55090, Novosphingobium aromaticivorans DSM 12444, Rhodococcus erythropolis DSM 43066, and Mycobacterium pallens DSM 45404. All strains were obtained from the Leibniz Institute DSMZ German Collection of Microorganisms and Cell Cultures. All experiments described below were performed with pure cultures of these strains.

Pure cultures were grown in the respective media as recommended by the DSMZ. Prior to the experiments, precultures were prepared by inoculating 50 mL sterile mineral salt medium containing 7 g Na₂HPO₄, 2.8 g KH₂PO₄, 0.5 g NaCl, 1 g NH₄Cl, 0.1 g MgSO₄ · 7 H₂O, 0.01 g FeSO₄ ·7 H₂O, 5 mg MnSO₄ H₂O, 6.4 mg ZnCl₂, 1 mg CaCl₂ · 6 H₂O, 0.6 mg BaCl₂, 0.36 mg CuSO₄ · 7 H₂O, 0.36 mg CuSO₄ ·5 H₂O, 6.5 mg H₃BO₃, 0.01 g EDTA and 146 μL HCl (37%) per L of distilled water (68), supplemented with 4 g L⁻¹ sodium succinate and 1 g L⁻¹ yeast extract as carbon sources. The cultures were incubated overnight at 30°C on an orbital shaker (160 rpm) and used as inoculants for the growth inhibition experiments. Solid media were prepared with the same ingredients supplemented with 15 g agar per L of medium.

Adjustment of osmotic (ΔΨ_O) and matric (ΔΨ_M) potentials of growth media.

NaCl and PEG 8000 (Carl Roth, Karlsruhe, Germany) were used to adjust the osmotic and matric potentials, respectively, of the mineral medium to −0.5, −1.5, −2.5, and −3.5 MPa. Osmotic potential was lowered by addition of different amounts of NaCl (Table S1 in the supplemental material). High-molecular weight PEGs were used to lower the matric potential in the growth medium because they hold water in their structure, mimicking the conditions in dry soil (69). The amount of PEG 8000 to be added was calculated based on empirical regression model equation 1 (70):

$$[\text{PEG}]=4-{(5.16\Psi T-560\Psi +16)}^{0.5}/(2.58T-280)$$ (1)

where [PEG] is the amount (g L⁻¹), Ψ is the water potential (Pa), and T is the air temperature (K).

To verify the achieved water potentials of the samples, the water activities (A_w) of the liquid and solid media were measured with a LabMaster-aw instrument (Novasina AG, Lachen, Switzerland). For A_w measurements, 5 mL of liquid or solid medium was filled into a dry sample cup (Ø: 40 mm, 12 mm, polypropylene, ePW sample cups; Novasina). System parameters were set to 5 min stable observation time for temperature and water activity. The measurements were performed at 30°C. Measured A_w values were converted to osmotic and matric potentials using equation 2 (71):

$$\Psi =-1,065T{\text{A}}_{\text{w}}$$ (2)

where Ψ is the matric/osmotic potential (kPa), T is the air temperature (K), A_w is the water activity (–) and 1,065 is a derived constant (72).

Water potential values derived from A_w measurements (Table S1) were used for presentation and interpretation of the obtained results rather than the calculated values.

Quantification of growth in submerged culture.

For growth inhibition experiments of submerged cultures, 50 mL of sterile mineral salt medium supplemented with different amounts of NaCl or PEG 8000 (Table S1) and glucose and yeast extract as C sources, as described above, was inoculated with overnight cultures to obtain an initial optical density at 560 nm (OD₅₆₀) of 0.05 in 250-mL clear screw-cap vials. The samples were incubated at 30°C on an orbital shaker (160 rpm) for 6 to 22 h, depending on the growth rates in the respective growth medium.

Cell growth was monitored by OD₅₆₀ using a UV/VIS spectrophotometer (Lambda2S; PerkinElmer, Waltham, USA). For analysis of their physicochemical properties, cells were harvested during the late exponential growth phase by centrifugation at 11,000 × g for 15 min (Hermle Z383K), resuspended in 2 mL KNO₃ (10 mM [pH 7.0]) and transferred to 2-mL reaction tubes. Each sample was washed twice with 2 mL KNO₃ followed by 1 min centrifugation at 10,000 × g. The washed biomass was resuspended in 1 mL KNO₃ and stored at 5°C until contact angle analysis.

Quantification of growth in surface cultures.

Five milliliters of solid mineral salt medium supplemented with 4 g L⁻¹ sodium succinate and 1 g L⁻¹ yeast extract as carbon sources was inoculated with a 5-μL drop of the overnight culture in the center and incubated at 30°C in six-well microplates (Ø: 3.5 cm; Thermo Fisher Scientific), for 30 to 65 h depending on the growth rates of the strains.

Destructive sampling of biomass in individual wells was performed at the end of the incubation period (again depending on the growth rates of the strains) by washing the biomass off the agar surface at different times after inoculation with 2 mL KNO₃ to monitor the growth via OD₅₆₀ measurements. At the end of the incubation, the biomass was washed twice with 2 mL KNO₃ followed by 1 min centrifugation at 10,000 × g, and then stored overnight at 4°C for contact angle measurements.

For both submersed and surface cultures, the growth rate μ (h⁻¹) of each culture in the exponential growth phase was calculated based on the development of OD₅₆₀, and the growth rates of stressed cultures are presented as the percentage of the growth rate in the corresponding unstressed cultures.

Contact angle measurement.

To assess bacterial CSH, the θ_w of the samples were measured during growth inhibition experiments. Depending on the OD of the sample, 100 to 500 μL of the biomass was suspended in 20 mL KNO₃ and the suspension was filtered through a cellulose nitrate membrane filter (pore size 0.45 μm, Ø: 25 mm, NC 45; Whatman) to produce a homogenously covered filter surface. The filters were mounted to a microscope slide with double-sided adhesive tape and dried for 2 h. The θ_w was determined with the sessile drop method (73) using a drop-shape analysis system (DSA 100, Krüss GmbH, Hamburg, Germany). A single water droplet was placed on the filter surface and the solid-water θ_w was determined after 30 ms by image analysis with the DSA software. Two filters were prepared for each sample, four measurements were performed per filter at different spots, and the average θ_w was calculated. For surface free energy determination, additional θ measurements for unstressed and stressed B. subtilis and P. fluorescens cells were carried out with 1 μL deionized water, ethylene glycol, and α-bromonaphthalene as testing liquids, using a contact angle microscope equipped with a video camera (OCA 15; DataPhysics, Filderstadt, Germany).

In a few cases, bacterial growth was inhibited so strongly by the water stress applied that not enough biomass was available to cover the filters completely and homogeneously. This prevented the determination of θ_w for these samples.

Surface free energy determination.

Solid and liquid interfacial properties are related to each other through the solid-liquid θ as expressed by equation 3 (74):

$$\left(1+\text{cos}\theta \right){\gamma }_l=2\left(\sqrt{{\gamma }_{\text{s}}^{\text{LW}}{\gamma }_{\text{l}}^{\text{LW}}}\hspace{0.17em}+\hspace{0.17em}\sqrt{{\gamma }_{\text{s}}^{+}{\gamma }_{\text{l}}^{-}}\hspace{0.17em}+\hspace{0.17em}\sqrt{{\gamma }_{\text{s}}^{-}{\gamma }_{\text{l}}^{+}}\right)$$ (3)

where θ is the solid-liquid contact angle (°), γ_l is the liquid surface free energy (J m⁻²), and γ_s is the solid surface free energy (J m⁻²). The superscripts ‘LW’, ‘−’, and ‘+’denote the nonpolar Lifshitz-van der Waals component, electron-donor (base) component, and electron-acceptor (acid) component, respectively. The three unknown variables in equation 3, γ_s^LW, γ_s^–, and γ_s⁺, were determined by solving a system of three independent linear equations, using the contact angles determined with deionized water, ethylene glycol, and α-bromonaphthalene and the respective liquid surface free energy components (22). The polar acid/base component (γ_s^AB) was calculated as follows:

$${\gamma }_{\text{s}}^{\text{AB}}=2\sqrt{({\gamma }_{\text{s}}^{+}{\gamma }_{\text{s}}^{-})},$$

and the total solid surface free energy (γ_s^TOT) was calculated as the sum of γ_s^AB and γ_s^LW. (For more details on the contact angle determination and surface free energy calculations, see reference 75.)

Zeta potential analysis.

For the measurement of the zeta potential, cell pellets were suspended in 10 mM KNO₃ to obtain cell suspensions with concentrations of ~10⁹ cells L⁻¹. The pH (measured potentiometrically with a glass electrode) was adjusted with KOH to pH 7, which corresponds to the pH of the growth medium. The pH-dependent zeta potential of unstressed und stressed (osmotic stress: −3.1 MPa; matric stress: −3.8 MPa) B. subtilis and P. fluorescens cells was calculated from electrophoretic mobility measured by phase analysis light scattering (ZetaPALS; Brookhaven Instruments Corp., Holtsville, USA) using Smoluchowski’s equation (76). The mean zeta potential of each sample was determined based on 10 consecutive runs comprised of 10 cycles each.

X-ray photoelectron spectroscopy.

The surface elemental composition of unstressed and stressed B. subtilis and P. fluorescens cells was analyzed by XPS using an Axis Ultra DLD (Kratos Analytical, Manchester, United Kingdom) with monochromatic AlKα radiation (1,486.6 eV; emission current: 20 mA, voltage: 6 kV). For the measurement, the central portion (approximately 25 mm²) of an air-dried filter, prepared the same way as for the contact angle analysis, was fixed on a bar using carbon conductive tape (Agar Scientific Elektron Technology UK Ltd., Stansted, United Kingdom). Survey spectra in the binding energy range of 1,200 to 0 eV (1-eV resolution) was obtained at a pressure of 4 × 10⁷ Pa with a pass energy of 160 eV, a dwell time of 500 ms, and three sweeps per measurement cycle at a take-off angle of 0°. For each sample, three spectra at different locations (spot size: 300 × 700 μm) were recorded. After charge correction for the C 1-s peak at 284.8 eV (C to C, C to H), the spectra were evaluated using Vision 2 software (Kratos Analytical, Manchester, United Kingdom). Surface elemental composition was quantified in terms of atomic-% (at.-%) using the relative sensitivity factors implemented in the software.

For P. fluorescens, the proportions of surface carbon associated with proteins (C_Pr/C), polysaccharides (C_PS/C), and hydrocarbon-like compounds (C_HC/C) were estimated from the observed elemental concentration ratios [N/C]_obs and [O/C]_obs using the following set of equations (77):

$${\left[\text{N/C}\right]}_{\text{obs}}=0.279({\text{C}}_{\mathrm{Pr}}/\text{C})$$ (4)

$${\left[\text{O}/\text{C}\right]}_{\text{obs}}=0.325({\text{C}}_{\text{Pr}}\text{/C})\hspace{0.17em}+\hspace{0.17em}0.833({\text{C}}_{\text{PS}}\text{/C})$$ (5)

$${\left[\text{C}/\text{C}\right]}_{\text{obs}}\left(\frac{{\text{C}}_{\text{Pr}}}{\text{C}}\right)+\left(\frac{{\text{C}}_{\text{PS}}}{\text{C}}\right)+\left(\frac{{\text{C}}_{\text{HC}}}{\text{C}}\right)=1$$ (6)

In the case of B. subtilis, the previous set of equations is not applicable because phosphate oxygen is not negligible with respect to total oxygen (51). To account for this, equation 5 was replaced by

$${\left[(\text{O}-4\text{P})/\text{C}\right]}_{\text{obs}}\text{=\hspace{0.17em}0}{\text{.325(C}}_{\text{Pr}}/\text{C})+0{\text{.833(C}}_{\text{PS}}\text{/C)}$$ (7)

to estimate the respective proportions of surface carbon for B. subtilis (21). The coefficients used for the protein constituent are based on the composition of the major outer protein of P. fluorescens. However, according to Genet et al. (77), the choice of the model protein composition is not decisive because bacterial protein compositions are very similar.

The mass percentage of the respective component i (i.e., Pr, PS, HC) was then calculated according to the formula

$$\text{Constituent}i\left(\text{mass\%}\right)=\frac{({\text{C}}_i/\text{C})}{M_{\text{i}}}\cdot \frac{1}{\frac{({\text{C}}_{\text{PS}}\text{/C})}{M_{\text{Pr}}}+\frac{({\text{C}}_{\text{Pr}}\text{/C})}{M_{\text{PS}}}+\frac{({\text{C}}_{\text{HC}}\text{/C})}{M_{\text{HC}}}}$$ (8)

where M_i is the carbon concentration of each constituent (43.5, 37.0, and 71.4 mmol C g⁻¹ for Pr, PS, and HC, respectively) (21).

Atomic force microscopy.

AFM mapping was used to obtain information on the impact of water stress on the elasticity and height of bacterial cells. B. subtilis and P. fluorescens cells (OD₅₆₀ = 0.9) exposed to different levels of osmotic stress were fixed on a smooth surface of adhesive resin glue (tempfix; PLANO GmbH, Wetzlar, Germany). Selected spots of appropriate cell density were mapped in 10 mM KNO₃ using an atomic force microscope (AFM, Dimension Icon; Bruker Corporation, Billerica, MA, USA) in the Peak Force Quantitative Nanomechanical Mapping (PFQNM) mode. Before analysis, chemically functionalized probes (SmartTips; NanoAndMore GmbH, Wetzlar, Germany) were calibrated for their deflection sensitivity (Sv) on mica and their spring constant, k, was determined by the thermal noise method (78). By post blind tip reconstruction (79) on a standard titanium roughness sample, the tip radii (30 to 40 nm) were obtained using the NanoScope Analysis software (version 8.15, Bruker, USA). From the height sensor and the log DMT modulus channel, the maximum height and the respective elastic Derjaguin-Muller-Toporov (DMT) modulus at the highest points of 18 to 30 cells were determined.

Statistics.

The bacterial surface chemical composition results were tested for statistically significant differences between control and stressed cells by one-way repeated measures analysis of variance (ANOVA) followed by the Holm-Sidak post hoc test when normality (tested by Shapiro-Wilk test) and equal variance (Brown-Forsythe test) were met. In all other cases, repeated measures ANOVA on ranks followed by Tukey’s post hoc test was performed, using SigmaPlot 13.0 (Systat Software, Inc., San Jose, USA).

Normal distribution and variance homogeneity of all data sets were tested by Shapiro-Wilk (shapiro.test) and Levene’s test (t test) using the statistical software R (R Core Team, 2020). The statistically significant differences in bacterial growth rate and contact angle were tested by pairwise t test (pairwise.t.test). For normally distributed data, statistical significance of differences in height and elastic modulus obtained from AFM measurements between stressed and unstressed cells of B. subtilis and P. fluorescens were tested by pairwise t test if variances of the data sets were equal and by Welch’s test if variances were not equal (pairwise.t.test). If the data sets were not normally distributed, Wilcoxon rank-sum test (pairwise.wilcox.test) was used for this purpose.

Data availability.

All data presented in this study are compiled in Table S2 in the supplemental material.