Skip to main content
  • ASM
    • Antimicrobial Agents and Chemotherapy
    • Applied and Environmental Microbiology
    • Clinical Microbiology Reviews
    • Clinical and Vaccine Immunology
    • EcoSal Plus
    • Eukaryotic Cell
    • Infection and Immunity
    • Journal of Bacteriology
    • Journal of Clinical Microbiology
    • Journal of Microbiology & Biology Education
    • Journal of Virology
    • mBio
    • Microbiology and Molecular Biology Reviews
    • Microbiology Resource Announcements
    • Microbiology Spectrum
    • Molecular and Cellular Biology
    • mSphere
    • mSystems
  • Log in
  • My alerts
  • My Cart

Main menu

  • Home
  • Articles
    • Current Issue
    • Accepted Manuscripts
    • COVID-19 Special Collection
    • Archive
    • Minireviews
  • For Authors
    • Submit a Manuscript
    • Scope
    • Editorial Policy
    • Submission, Review, & Publication Processes
    • Organization and Format
    • Errata, Author Corrections, Retractions
    • Illustrations and Tables
    • Nomenclature
    • Abbreviations and Conventions
    • Publication Fees
    • Ethics Resources and Policies
  • About the Journal
    • About AEM
    • Editor in Chief
    • Editorial Board
    • For Reviewers
    • For the Media
    • For Librarians
    • For Advertisers
    • Alerts
    • RSS
    • FAQ
  • Subscribe
    • Members
    • Institutions
  • ASM
    • Antimicrobial Agents and Chemotherapy
    • Applied and Environmental Microbiology
    • Clinical Microbiology Reviews
    • Clinical and Vaccine Immunology
    • EcoSal Plus
    • Eukaryotic Cell
    • Infection and Immunity
    • Journal of Bacteriology
    • Journal of Clinical Microbiology
    • Journal of Microbiology & Biology Education
    • Journal of Virology
    • mBio
    • Microbiology and Molecular Biology Reviews
    • Microbiology Resource Announcements
    • Microbiology Spectrum
    • Molecular and Cellular Biology
    • mSphere
    • mSystems

User menu

  • Log in
  • My alerts
  • My Cart

Search

  • Advanced search
Applied and Environmental Microbiology
publisher-logosite-logo

Advanced Search

  • Home
  • Articles
    • Current Issue
    • Accepted Manuscripts
    • COVID-19 Special Collection
    • Archive
    • Minireviews
  • For Authors
    • Submit a Manuscript
    • Scope
    • Editorial Policy
    • Submission, Review, & Publication Processes
    • Organization and Format
    • Errata, Author Corrections, Retractions
    • Illustrations and Tables
    • Nomenclature
    • Abbreviations and Conventions
    • Publication Fees
    • Ethics Resources and Policies
  • About the Journal
    • About AEM
    • Editor in Chief
    • Editorial Board
    • For Reviewers
    • For the Media
    • For Librarians
    • For Advertisers
    • Alerts
    • RSS
    • FAQ
  • Subscribe
    • Members
    • Institutions
Physiology

Reduction of the Temperature Sensitivity of Halomonas hydrothermalis by Iron Starvation Combined with Microaerobic Conditions

Jesse P. Harrison, John E. Hallsworth, Charles S. Cockell
K. E. Wommack, Editor
Jesse P. Harrison
aUK Centre for Astrobiology, School of Physics and Astronomy, The University of Edinburgh, Edinburgh, United Kingdom
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
  • ORCID record for Jesse P. Harrison
John E. Hallsworth
bInstitute for Global Food Security, School of Biological Sciences, Medical Biology Centre, Queen's University Belfast, Belfast, United Kingdom
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Charles S. Cockell
aUK Centre for Astrobiology, School of Physics and Astronomy, The University of Edinburgh, Edinburgh, United Kingdom
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
K. E. Wommack
Roles: Editor
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
DOI: 10.1128/AEM.03639-14
  • Article
  • Figures & Data
  • Info & Metrics
  • PDF
Loading

ABSTRACT

The limits to biological processes on Earth are determined by physicochemical parameters, such as extremes of temperature and low water availability. Research into microbial extremophiles has enhanced our understanding of the biophysical boundaries which define the biosphere. However, there remains a paucity of information on the degree to which rates of microbial multiplication within extreme environments are determined by the availability of specific chemical elements. Here, we show that iron availability and the composition of the gaseous phase (aerobic versus microaerobic) determine the susceptibility of a marine bacterium, Halomonas hydrothermalis, to suboptimal and elevated temperature and salinity by impacting rates of cell division (but not viability). In particular, iron starvation combined with microaerobic conditions (5% [vol/vol] O2, 10% [vol/vol] CO2, reduced pH) reduced sensitivity to temperature across the 13°C range tested. These data demonstrate that nutrient limitation interacts with physicochemical parameters to determine biological permissiveness for extreme environments. The interplay between resource availability and stress tolerance, therefore, may shape the distribution and ecology of microorganisms within Earth's biosphere.

INTRODUCTION

Knowledge of the physical and/or chemical parameters that can limit cell division and metabolic activity is of critical importance in fields such as ecology, agriculture, food preservation, biotechnology, and astrobiology (1–7). The physicochemical boundaries beyond which multiplication of all microorganisms is prevented are imposed by low water activity, extremes of temperature (approximately −20 to 120°C), and other situations, including high pH (>12), chaotropicity, and oxidative damage (e.g., due to UV radiation) (8–10). The stress mechanism for a number of these parameters involves changes in the entropic status of lipid bilayers and other macromolecular systems (see reference 9 and citations therein). Others, including the reactive oxygen species generated by UV radiation, act by inducing alterations in the primary structure of cellular macromolecules (11). One of the primary effects of extreme pH is the breakdown of electrochemical (and other) gradients across the plasma membrane (12).

An increasing body of evidence suggests that it is frequently the net effect of diverse stress parameters that defines the boundary space for life (8, 13–18). The sea ice bacterium Shewanella gelidimarina, for example, has been shown to exhibit an increased temperature range for cell division when cultured at high (NaCl) salinity, with increases in membrane lipid packing and fatty acid content conferring tolerance of both conditions (13). Other solutes (including MgCl2) have also been found to influence the temperature limits for microbial multiplication (15, 17). Moreover, adaptation to high hydrostatic pressure has been proposed as a mechanism that enables bacterial multiplication within hypersaline deep-sea environments (14). Such interactions between stress parameters are frequently likely to be multifactorial. Indeed, since no microbial isolates to date have been found to exhibit cell division under a combination of extremely high salinity, temperature (>60°C), and pH (>8), these conditions may be collectively prohibitive to life (8, 16). Other combinations of stress parameters may also determine the limits for cell division within extreme environments. For example, there is an absence of prokaryotic strains that are able to multiply under combinations of elevated (>10 MPa) hydrostatic pressure and extremely low (<4) or high (>9) pH (8).

Despite our knowledge of the physicochemical boundaries for microbial cell division having been greatly advanced over the past three decades, we are only beginning to understand how interactions between different stressors define biological permissiveness for natural ecosystems (8, 13–18). In particular, while several studies have identified minimal and maximal limits for microbial life in response to the surrounding environment (1, 5, 19–21), there is a paucity of information on how rates of cell division are impacted by multiple stress parameters within extreme habitats. Detailed investigations of this topic are required to determine the limits of the biosphere under conditions that account for the physicochemical complexity of extreme environments on Earth, not only with reference to the minimal and maximal boundaries for life but also within the entire range for cell division (8). Moreover, we know relatively little about the impacts of small fluctuations in environmental conditions on rates of microbial multiplication and on the level of cellular sensitivity to water activity and other fundamental parameters (10).

Frequently, those chemical elements that can strongly affect rates of cell division are also known to play essential roles in cellular metabolism and microbially mediated redox processes. For example, iron is required for protein structure and function, and oxygen acts as the final electron acceptor during aerobic respiration. These and other elements can be scarce in both biologically hostile and biologically permissive environments. Yet, despite the fact that the majority of microorganisms on Earth are located in oligotrophic and/or oxygen-poor habitats (22–27), research into the physicochemical limits for life has most commonly relied on data obtained using resource-rich, aerobic media. Here, we sought to determine how iron (Fe3+) starvation and microaerobic conditions impact temperature and salt tolerance of the marine bacterium Halomonas hydrothermalis (28). This strain was employed as a model organism due to its ability to proliferate under a broad range of temperatures and salinities (14, 28). Rates of cell division and levels of viability were determined for 36 distinct culture conditions created from permutations of incubation temperature, NaCl concentration, Fe3+ availability, and (aerobic or microaerobic) atmospheric composition. This approach enabled us to obtain detailed insights into the growth phenotype of H. hydrothermalis under multiple-stress conditions that are permissible to cell division.

MATERIALS AND METHODS

Bacterial strain.Halomonas hydrothermalis DSM 15725T (28) was obtained from the German Collection of Microorganisms (DSMZ, Braunschweig, Germany). This strain exhibits cell division across temperatures of 2 to 40°C (optimal growth reported at 30°C), total salt concentrations of 0.5% to 22% (wt/vol) (optimal range of 4% to 7% [wt/vol]), and pH values of 5 to 12 (optimal range of 7 to 8) (28). An exponential-phase culture was obtained by culturing the bacterium in marine broth (DSMZ medium no. 514) with 3.5% (wt/vol) NaCl (pH 7.5; incubation at 30°C), and aliquots were prepared (25% [vol/vol] glycerol) for storage at −80°C. The stored cultures were used to inoculate marine agar (3.5% [wt/vol] NaCl), which was incubated at 30°C for 48 h and stored at 4°C until use.

Growth rate assays.Starter cultures were prepared by transferring cells to 6 ml of marine broth (3.5% [wt/vol] NaCl and pH 7.5) and to a loosely capped 10-ml Falcon tube. The cultures were kept overnight in a shaking incubator (30°C, 100 rpm) and diluted using fresh marine broth to give a cell density equivalent to an optical density at 600 nm (OD600) of 0.3. Optical density measurements for the starter cultures were performed for volumes of 2.5 ml, using a Helios Gamma spectrophotometer (Thermo Spectronic, Cambridge, United Kingdom).

Growth assays were initiated by adding starter culture to fresh marine broth at a ratio of 1:100 (vol/vol). Cultures (each 100 μl; n = 5) were incubated in 96-well microplates sealed with gas-permeable optical seals (4titude Ltd., Surrey, United Kingdom). Six variations of culture media based on marine broth were initially used, with 1%, 3.5%, or 11.8% (wt/vol) NaCl prepared in the presence or absence of Fe(III) citrate. The iron-depleted media were spiked with 100 μM diethylenetriaminepentaacetic acid (DETAPAC), an extracellular ferric chelator that has previously been employed to induce bacterial iron starvation (29–31). The media with and without Fe(III) citrate contained ∼0.8 mg liter−1 and ∼0.1 mg liter−1 of Fe, respectively. Complex media spiked with an iron chelator have occasionally been used for studies of iron starvation in bacteria (31, 32). Marine broth is an undefined medium that has been successfully employed as a model system and a proxy for seawater for in vitro studies of bacteria under conditions of various salinity and nutrient regimens (33–35).

All culture types were incubated at 22, 30, or 35°C. As with the salinities employed in our study, these temperatures have been previously reported to represent suboptimal, optimal, and supraoptimal values for cell division by H. hydrothermalis under growth conditions that were otherwise optimal (28). Although this strain can actively grow at 2°C, its rates of cell division at this temperature are very low (28), and therefore a higher minimum temperature was selected. The cells were kept under conditions of either aerobic or microaerobic (5% [vol/vol] O2, 10% [vol/vol] CO2, and 85% [vol/vol] N2) atmospheres by placing the microplates within a Synergy 2 microplate reader (BioTek Instruments Inc., Vermont, USA) or inside sealed 2.5-liter jars containing an Oxoid CampyGen envelope (36). For microaerobic treatments, all media types were equilibrated to microaerobic conditions prior to inoculation. CampyGen envelopes have previously been used to compare bacterial survival mechanisms in complex media incubated under either aerobic or microaerobic atmospheres (37). Cultures within both the microplate reader and the jars were shaken continuously (at 1,080 rpm and 100 rpm, respectively). Cell density was monitored for up to 7 days by OD600 measurements using the microplate reader, with medium-only controls (n = 3) included for each culture condition. In order to determine whether cells incubated under a combination of iron-deprived and microaerobic conditions remained metabolically active, a separate growth assay was performed for iron-depleted media with 18% (wt/vol) NaCl under a microaerobic atmosphere, with OD600 values monitored over 9 days. With the exception of this medium type, pH values of both aerobic and microaerobic media (incubated at 30°C) were monitored by duplicate daily measurements.

Determination of cell viability.To assess cell viability, all culture types (see “Growth rate assays” above) were sampled on two consecutive days during the stationary-growth phase (measurements performed on days 3 and 4 following the onset of the growth assays at 22 and 30°C and on days 2 and 3 for 35°C). Earlier sampling times were employed for cultures incubated at 35°C due to cells frequently entering the death phase following incubation at this temperature for 4 days. Stationary-phase cultures were selected for analyses of cell viability to account for the net effect of different stress parameters on bacterial survival during earlier growth stages (i.e., both the lag and log phases).

Viability was measured using a LIVE/DEAD BacLight staining kit (Life Technologies, Paisley, United Kingdom) (38, 39) in combination with flow cytometry. Aliquots of both the cultures (n = 3) and negative controls (n = 1) were transferred at a ratio of 1:100 (vol/vol) to sterile 1.5% (wt/vol) NaCl solution and stained in a total volume of 100 μl according to the manufacturer's instructions. The stained samples were analyzed using a FACSCanto II flow cytometer (BD Biosciences, California, USA) equipped with a 488-nm laser and FACSDiva software (Version 6.1.3; BD Biosciences). Syto 9 fluorescence and propidium iodide fluorescence were detected using a 525/30-nm-wavelength band-pass filter (fluorescein isothiocyanate [FITC] channel) and a 670-nm-wavelength long-pass filter (peridinin chlorophyll protein [PerCP] channel), respectively. A minimum of 35,000 events were measured for each replicate.

Prior to the analysis of experimental samples, gates corresponding to intact cells, compromised cells, and nonspecific fluorescence signals were defined, using negative-control samples and overnight cultures stained with either a single dye or both dyes. Both untreated (“live”) and isopropanol-treated (“dead”) cultures were stained to facilitate the adjustment of gating settings (see Fig. S1 in the supplemental material). Additionally, mixtures of these live and dead cultures were analyzed to confirm that a 1:1 (vol/vol) ratio of the two dye components produced reliable staining results for H. hydrothermalis (linear regression, r2 = 0.99), following the manufacturer's instructions.

Data analysis and statistics.Specific growth rates (h−1) were calculated using OD600 measurements corresponding to either the entire exponential portion of the growth curve or a region of maximal slope on the curve when exponential growth was absent (14). Growth rates were calculated using the following formula (40): μ=ln 2td where μ = growth rate (h−1) and td = population doubling time (h−1).

Proportions of viable bacteria (percentages of total fluorescence events) were calculated using the FlowJo software package (version X.0.6; TreeStar Inc., California, USA), following gating (see “Determination of cell viability” above).

For comparisons of growth rates, 2-way analyses of variance (ANOVA) were performed with salinity (1% to 11.8% [wt/vol] NaCl) and temperature (22 to 35°C) as the factors. Analyses were performed for each combination of iron availability and type of atmosphere (see “Growth rate assays” above), followed by post hoc Tukey's honestly significant difference (HSD) tests. Measurements obtained for cells incubated under aerobic and iron-rich conditions were Box-Cox transformed (λ = −0.05) in order to alleviate heteroscedasticity (41). This transformation has been previously employed in several microbiological studies (see, e.g., references 42, 43, and 44). For comparisons of the proportions of viable cells across each combination of iron availability and atmospheric composition, a one-way ANOVA and Box-Cox-transformed data (λ = 2.8) were used in conjunction with a Tukey's HSD test. All statistical analyses were performed using R v3.0.2 (45).

RESULTS AND DISCUSSION

Under aerobic conditions with plentiful iron, bacterial growth rates were susceptible to moderate changes in temperature and salinity (Table 1 and Fig. 1A). For example, the mean growth rates of cultures incubated at a temperature-salinity combination of 35°C and 11.8% (wt/vol) NaCl were 43% and 44% lower than those at 35°C or 30°C and 3.5% (wt/vol) NaCl, respectively (Fig. 1A). Moreover, mean growth rates at 22°C and 11.8% (wt/vol) NaCl were 67% lower than those at 30°C and 3.5% (wt/vol) NaCl (Fig. 1A). Under these aerobic and iron-rich conditions, there was no evidence of an interaction between temperature and salinity in terms of their impacts on bacterial growth rates (Table 1). However, some Halomonas species incubated in aerobic high-salinity media have exhibited enhanced growth rates at temperatures below 22°C (14).

View this table:
  • View inline
  • View popup
TABLE 1

Two-way ANOVA results for bacterial growth ratesa

FIG 1
  • Open in new tab
  • Download powerpoint
FIG 1

Growth rates of Halomonas hydrothermalis under different culture conditions. (A) Cells grown under conditions of plentiful iron and oxygen. (B) Cells grown in iron-deprived media under aerobic conditions. (C) Cells grown in the presence of freely available iron under microaerobic conditions. (D) Cells grown in iron-deprived media under microaerobic conditions. Low, Mid, and High, 1%, 3.5%, and 11.8% (wt/vol) NaCl, respectively. Growth rates were calculated from growth curves as described in Materials and Methods. Data are presented as untransformed means ± standard errors of the means (SE) (n = 5). Different letters above the bars indicate significant differences between samples within a given combination of conditions of iron availability and type of atmosphere (aerobic or microaerobic) (P < 0.05 [Tukey's HSD test following 2-way ANOVA; see Table 1 for ANOVA results]).

Under iron starvation and aerobic conditions, H. hydrothermalis cultures generally exhibited reduced growth rates (Fig. 1B) in comparison with nonstarved aerobic cultures at equivalent temperatures and salinities (Fig. 1A). Both temperature and salinity significantly impacted these rates (Table 1), with iron starvation suppressing bacterial sensitivity to sub- and supraoptimal salinities at 22°C but not at higher temperatures, as shown by a Tukey's HSD test (Fig. 1B). The combined impacts of low oxygen availability and high CO2 concentrations on bacterial growth rates differed, with high rates maintained at most salinities at 30 or 35°C, despite an overall reduction in growth rates at 22°C (Fig. 1C). Under these microaerobic conditions, temperature and salinity, both as individual parameters and in combination, remained key determinants for rates of cell division (Table 1). Interestingly, the most pronounced impact of microaerobic culture conditions involved the growth rate limitation of cultures at suboptimal salinities and/or temperatures (Fig. 1A to C).

When H. hydrothermalis was cultured under conditions of both iron starvation and a microaerobic atmosphere (Fig. 1D), mean growth rates were reduced by as much as 83% relative to those of nonstarved cells (Fig. 1A). In fact, cell division was suppressed across all culture conditions to the extent that bacterial growth rates no longer varied as a function of temperature (Table 1). As maximal OD600 values of >0.2 were observed under all the examined incubation conditions (corresponding to mean CFU counts ranging from 2.2 × 108 ml−1 to 5.6 × 108 ml−1 in aerobic cultures incubated at 30°C [n = 3]; data not shown), this result was not attributable to low biomass or to optical density values residing below the detection limit for turbidimetry. Therefore, to assess whether this phenomenon was mediated by a decline in cell viability (46), we calculated the proportions of surviving stationary-phase cells under each set of culture conditions (see Materials and Methods; see also Fig. S1 in the supplemental material). The greatest difference between the different conditions was minimal (8.4%; Fig. 2) and did not correlate with growth rates (Fig. 1). Moreover, no significant differences (P > 0.05) in viability were observed across the majority of the culture conditions (Fig. 2), indicating that the growth rate values were accurate (and were not an artifact occurring due to differential rates of cell death). In addition, bacteria were able to proliferate under a combination of iron starvation and microaerobic conditions and a NaCl concentration of 18% (wt/vol) (see Fig. S2 in the supplemental material), which is close to the upper limit of salinity for cell division in H. hydrothermalis (28). Collectively, these data evidence the continued metabolic activity of the cultures despite their reduced sensitivity to temperature and suggest that the incubation conditions used in our study did not influence the upper limit of salinity for cell division in H. hydrothermalis (47).

FIG 2
  • Open in new tab
  • Download powerpoint
FIG 2

Viability of stationary-phase Halomonas hydrothermalis cells under different culture conditions. Cultures were incubated under iron-rich (Fe+) or iron-starved (Fe−) conditions, in the presence of an aerobic atmosphere (Aer+) or a microaerobic atmosphere (Aer−). Data are presented as untransformed means ± SE (n = 54). Different letters above the bars indicate significant differences between samples within a given combination of conditions of iron availability and type of atmosphere (aerobic or microaerobic) (P < 0.05 [Tukey's HSD test]), determined following a one-way ANOVA using Box-Cox-transformed data (P < 0.001) (F3,212 = 6.13). The measurements are based on LIVE/DEAD staining of cells followed by flow cytometry, with data pooled from two consecutive days. For details on the experimental protocol and instrumental calibration, see Materials and Methods and Fig. S1 in the supplemental material.

Compounded impacts of iron availability and atmospheric gas composition on microbial temperature and salt sensitivities have not been previously reported, despite observations that nutrient-starved cells frequently exhibit increased tolerance of heat, UV-B radiation, antibiotics, and oxidative stress (48–53). Interactions between starvation conditions, and the ability of abiotic parameters to influence bacterial cell division, have implications for the habitability and productivity of natural systems, including marine waters (14, 22, 24, 27, 54) and nutrient-poor rocks (55, 56), and are also likely to modulate the growth and persistence of microorganisms within nutrient-limited artificial environments (57). Such interactions may additionally influence the distribution and composition of microbial communities within and between habitats. Rates of microbial cell division within biologically hostile habitats (2, 8–10, 16, 18), therefore, can potentially be impacted by effects of resource limitation on microbial sensitivity to extremes, which in turn are likely to depend on other environmental conditions.

Microbial responses to individual stress parameters such as supraoptimal salinities and temperatures are well characterized, involving mechanisms that include the regulation of intracellular ion (e.g., Na+ and K+) concentrations, solute accumulation, and alterations in membrane fluidity (58, 59). However, the biophysical and/or metabolic mechanism(s) by which responses to iron starvation and microaerobic conditions can determine tolerance of specific stresses has yet to be fully characterized. It is known that several general-stress proteins can be triggered in response to nutrient starvation and are additionally induced by oxygen limitation as well as other physicochemical challenges, such as heat shock, acid stress, hydrophobic stressors, reactive oxygen species, and osmotic stress (60–62). It is plausible that, in the present study, iron starvation and microaerobic conditions exerted a compound influence on H. hydrothermalis tolerance of differences in temperature and salinities either directly or as a consequence of secondary stress responses (such as those induced by a reduction in pH under CO2-enriched conditions; see Table S1 in the supplemental material). Indeed, exposure to a given extreme can frequently confer cross-protection against other environmental stressors in both prokaryotes and eukaryotes, due to overlapping physiological adaptations and stress-response networks (8, 60, 62–65).

In summary, the findings of the current study suggest that both the cellular stress mechanisms and stress responses of microbial extremophiles can be fully understood only when related to the wider metabolic energy budget of a given habitat. Moreover, the specific impacts of nutrient availability on rates of microbial cell division are likely to depend on other environmental factors, including the ambient gas composition. Since the majority of microbial taxa within the biosphere are adapted to nutrient-poor conditions, the ability of microorganisms to exhibit differential responses to nutrient limitation is likely to impact population sizes within specific habitats (66, 67) and the biological permissiveness of extreme environments on Earth. The current study focused on characterizing the phenotype-level impacts of multiple stress parameters on the growth rates and viability of H. hydrothermalis under conditions that enable cell division. However, it is possible that interactions between nutrient availability and stress sensitivity can affect the physicochemical growth limits of diverse microbial taxa (1, 5, 13, 19–21, 47). The findings of our study, therefore, suggest that the habitability of extreme environments (including hypersaline habitats) is often defined by the interplay between several physicochemical and biological factors rather than by one parameter alone.

ACKNOWLEDGMENTS

We thank Martin Waterfall for assistance with flow cytometry.

This work was supported by the Science and Technology Facilities Council (STFC Consolidated Grant no. ST/1001964/1) and The University of Edinburgh Development Trust (Innovation Initiative Grant no. GR000859).

FOOTNOTES

    • Received 4 November 2014.
    • Accepted 7 January 2015.
    • Accepted manuscript posted online 16 January 2015.
  • Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.03639-14.

  • Copyright © 2015, American Society for Microbiology. All Rights Reserved.

REFERENCES

  1. 1.↵
    1. Ratkowsky DA,
    2. Lowry RK,
    3. McMeekin TA,
    4. Stokes AN
    . 1983. Model for bacterial culture growth rate throughout the entire biokinetic temperature range. J Bacteriol 154:1222–1226.
    OpenUrlAbstract/FREE Full Text
  2. 2.↵
    1. Pikuta EV,
    2. Hoover RB,
    3. Tang J
    . 2007. Microbial extremophiles at the limits of life. Crit Rev Microbiol 33:183–209. doi:10.1080/10408410701451948.
    OpenUrlCrossRefPubMedWeb of Science
  3. 3.↵
    1. Lammer H,
    2. Bredehöft JH,
    3. Coustenis A,
    4. Khodachenko ML,
    5. Kaltenegger L,
    6. Grasset O,
    7. Prieur D,
    8. Raulin F,
    9. Ehrenfreund P,
    10. Yamauchi M,
    11. Wahlund J-E,
    12. Grießmeier J-M,
    13. Stangl G,
    14. Cockell CS,
    15. Kulikov YN,
    16. Grenfell JL,
    17. Rauer H
    . 2009. What makes a planet habitable? Astron Astrophys Rev 17:181–249. doi:10.1007/s00159-009-0019-z.
    OpenUrlCrossRefWeb of Science
  4. 4.↵
    1. Oren A
    . 2011. Thermodynamic limits to microbial life at high salt concentrations. Environ Microbiol 13:1908–1923. doi:10.1111/j.1462-2920.2010.02365.x.
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    1. Corkrey R,
    2. Olley J,
    3. Ratkowsky D,
    4. McMeekin T,
    5. Ross T
    . 2012. Universality of thermodynamic constants governing biological growth rates. PLoS One 7:e32003. doi:10.1371/journal.pone.0032003.
    OpenUrlCrossRefPubMed
  6. 6.↵
    1. Lineweaver CH,
    2. Chopra A
    . 2012. The habitability of our Earth and other Earths: astrophysical, geochemical, geophysical, and biological limits on planet habitability. Annu Rev Earth Planet Sci 40:597–623. doi:10.1146/annurev-earth-042711-105531.
    OpenUrlCrossRefWeb of Science
  7. 7.↵
    1. Lievens B,
    2. Hallsworth JE,
    3. Belgacem ZB,
    4. Pozo MI,
    5. Stevenson A,
    6. Willems KA,
    7. Jacquemyn H
    . 3 September 2014, posting date. Microbiology of sugar-rich environments: diversity, ecology, and system constraints. Environ Microbiol doi:10.1111/1462-2920.12570.
    OpenUrlCrossRef
  8. 8.↵
    1. Harrison JP,
    2. Gheeraert N,
    3. Tsigelnitskiy D,
    4. Cockell CS
    . 2013. The limits for life under multiple extremes. Trends Microbiol 21:204–212. doi:10.1016/j.tim.2013.01.006.
    OpenUrlCrossRefPubMed
  9. 9.↵
    1. Rummel JD,
    2. Beaty DW,
    3. Jones MA,
    4. Bakermans C,
    5. Barlow NG,
    6. Boston PJ,
    7. Chevrier VF,
    8. Clark BC,
    9. de Vera J-PP,
    10. Gough RV,
    11. Hallsworth JE,
    12. Head JW,
    13. Hipkin VJ,
    14. Kieft TL,
    15. McEwen AS,
    16. Mellon MT,
    17. Mikucki JA,
    18. Nicholson WL,
    19. Omelon CR,
    20. Peterson R,
    21. Roden EE,
    22. Sherwood Lollar B,
    23. Tanaka KL,
    24. Viola D,
    25. Wray JJ
    . 2014. A new analysis of Mars “special regions”: findings of the second MEPAG Special Regions Science Analysis Group (SR-SAG2). Astrobiology 14:887–968. doi:10.1089/ast.2014.1227.
    OpenUrlCrossRefPubMed
  10. 10.↵
    1. Stevenson A,
    2. Burkhardt J,
    3. Cockell CS,
    4. Cray JA,
    5. Dijksterhuis J,
    6. Fox-Powell M,
    7. Kee TP,
    8. Kminek G,
    9. McGenity TJ,
    10. Timmis KN,
    11. Timson DJ,
    12. Voytek MA,
    13. Westall F,
    14. Yakimov MM,
    15. Hallsworth JE
    . 28 September 2014, posting date. Multiplication of microbes below 0.690 water activity: implications for terrestrial and extraterrestrial life. Environ Microbiol doi:10.1111/1462-2920.12598.
    OpenUrlCrossRef
  11. 11.↵
    1. Imlay JA
    . 2013. The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium. Nat Rev Microbiol 11:443–454. doi:10.1038/nrmicro3032.
    OpenUrlCrossRefPubMed
  12. 12.↵
    1. Padan E,
    2. Bibi E,
    3. Ito M,
    4. Krulwich TA
    . 2005. Alkaline pH homeostasis in bacteria: new insights. BBA Biomembranes 1717:67–88. doi:10.1016/j.bbamem.2005.09.010.
    OpenUrlCrossRefPubMedWeb of Science
  13. 13.↵
    1. Nichols DS,
    2. Olley J,
    3. Garda H,
    4. Brenner RR,
    5. McMeekin TA
    . 2000. Effect of temperature and salinity stress on growth and lipid composition of Shewanella gelidimarina. Appl Environ Microbiol 66:2422–2429. doi:10.1128/AEM.66.6.2422-2429.2000.
    OpenUrlAbstract/FREE Full Text
  14. 14.↵
    1. Kaye JZ,
    2. Baross JA
    . 2004. Synchronous effects of temperature, hydrostatic pressure, and salinity on growth, phospholipid profiles, and protein patterns of four Halomonas species isolated from deep-sea hydrothermal vent and sea surface environments. Appl Environ Microbiol 70:6220–6229. doi:10.1128/AEM.70.10.6220-6229.2004.
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    1. Hallsworth JE,
    2. Yakimov MM,
    3. Golyshin PN,
    4. Gillion JLM,
    5. D'Auria G,
    6. de Lima Alves F,
    7. La Cono V,
    8. Genovese M,
    9. McKew BA,
    10. Hayes SL,
    11. Harris G,
    12. Giuliano L,
    13. Timmis KN,
    14. McGenity TJ
    . 2007. Limits of life in MgCl2-containing environments: chaotropicity defines the window. Environ Microbiol 9:801–813. doi:10.1111/j.1462-2920.2006.01212.x.
    OpenUrlCrossRefPubMedWeb of Science
  16. 16.↵
    1. Bowers KJ,
    2. Mesbah NM,
    3. Wiegel J
    . 2009. Biodiversity of poly-extremophilic Bacteria: does combining the extremes of high salt, alkaline pH and elevated temperature approach a physico-chemical boundary for life? Saline Systems 5:9. doi:10.1186/1746-1448-5-9.
    OpenUrlCrossRefPubMed
  17. 17.↵
    1. Chin JP,
    2. Megaw J,
    3. Magill CL,
    4. Nowotarski K,
    5. Williams JP,
    6. Bhaganna P,
    7. Linton M,
    8. Patterson MF,
    9. Underwood GJC,
    10. Mswaka AY,
    11. Hallsworth JE
    . 2010. Solutes determine the temperature windows for microbial survival and growth. Proc Natl Acad Sci U S A 107:7835–7840. doi:10.1073/pnas.1000557107.
    OpenUrlAbstract/FREE Full Text
  18. 18.↵
    1. Yakimov MM,
    2. Lo Cono V,
    3. La Spada G,
    4. Bortoluzzi G,
    5. Messina E,
    6. Smedile F,
    7. Arcadi E,
    8. Borghini M,
    9. Ferrer M,
    10. Schmitt-Kopplin P,
    11. Hertkorn N,
    12. Cray JA,
    13. Hallsworth JE,
    14. Golyshin PN,
    15. Giuliano L
    . 6 August 2014, posting date. Microbial community of the deep-sea brine Lake Kryos seawater-brine interface is active below the chaotropicity limit of life as revealed by recovery of mRNA. Environ Microbiol doi:10.1111/1462-2920.12587.
    OpenUrlCrossRef
  19. 19.↵
    1. Cuppers HGAM,
    2. Oomes S,
    3. Brul S
    . 1997. A model for the combined effects of temperature and salt concentration on growth rate of food spoilage molds. Appl Environ Microbiol 63:3764–3769.
    OpenUrlAbstract/FREE Full Text
  20. 20.↵
    1. Presser KA,
    2. Ross T,
    3. Ratkowsky DA
    . 1998. Modelling the growth limits (growth/no growth interface) of Escherichia coli as a function of temperature, pH, lactic acid concentration, and water activity. Appl Environ Microbiol 64:1773–1779.
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    1. Wijtzes T,
    2. Rombouts FM,
    3. Kant-Muermans MLT,
    4. van 't Riet K,
    5. Zwietering MH
    . 2001. Development and validation of a combined temperature, water activity, pH model for bacterial growth rate of Lactobacillus curvatus. Int J Food Microbiol 63:57–64. doi:10.1016/S0168-1605(00)00401-3.
    OpenUrlCrossRefPubMed
  22. 22.↵
    1. Church MJ,
    2. Hutchins DA,
    3. Ducklow HW
    . 2000. Limitation of bacterial growth by dissolved organic matter and iron in the Southern Ocean. Appl Environ Microbiol 66:455–466. doi:10.1128/AEM.66.2.455-466.2000.
    OpenUrlAbstract/FREE Full Text
  23. 23.↵
    1. Ley RE,
    2. Williams MW,
    3. Schmidt SK
    . 2004. Microbial population dynamics in an extreme environment: controlling factors in talus soils at 3750 m in the Colorado Rocky Mountains. Biogeochemistry 68:313–335. doi:10.1023/B:BIOG.0000031032.58611.d0.
    OpenUrlCrossRef
  24. 24.↵
    1. Harpole WS,
    2. Ngai JT,
    3. Cleland EE,
    4. Seabloom EW,
    5. Borer ET,
    6. Bracken MES,
    7. Elser JJ,
    8. Gruner DS,
    9. Hillebrand H,
    10. Shurin JB,
    11. Smith JE
    . 2011. Nutrient co-limitation of primary producer communities. Ecol Lett 14:852–862. doi:10.1111/j.1461-0248.2011.01651.x.
    OpenUrlCrossRefPubMed
  25. 25.↵
    1. Swanner ED,
    2. Templeton AS
    . 2011. Potential for nitrogen fixation and nitrification in the granite-hosted subsurface at Henderson Mine, CO. Front Microbiol 2:254. doi:10.3389/fmicb.2011.00254.
    OpenUrlCrossRefPubMed
  26. 26.↵
    1. Hobbie JE,
    2. Hobbie EA
    . 2013. Microbes in nature are limited by carbon and energy: the starving-survival lifestyle in soil and consequences for estimating microbial rates. Front Microbiol 4:324. doi:10.3389/fmicb.2013.00324.
    OpenUrlCrossRefPubMed
  27. 27.↵
    1. Sebastián M,
    2. Gasol JM
    . 2013. Heterogeneity in the nutrient limitation of different bacterioplankton groups in the Eastern Mediterranean Sea. ISME J 7:1665–1668. doi:10.1038/ismej.2013.42.
    OpenUrlCrossRefPubMed
  28. 28.↵
    1. Kaye JZ,
    2. Márquez MC,
    3. Ventosa A,
    4. Baross JA
    . 2004. Halomonas neptunia sp. nov., Halomonas sulfidaeris sp. nov., Halomonas axialensis sp. nov. and Halomonas hydrothermalis sp. nov.: halophilic bacteria isolated from deep-sea hydrothermal-vent environments. Int J Syst Evol Microbiol 54(Pt 2):499–511. doi:10.1099/ijs.0.02799-0.
    OpenUrlCrossRefPubMed
  29. 29.↵
    1. Pericone CD,
    2. Park S,
    3. Imlay JA,
    4. Weiser JN
    . 2003. Factors contributing to hydrogen peroxide resistance in Streptococcus pneumoniae include pyruvate oxidase (SpxB) and avoidance of the toxic effects of the Fenton reaction. J Bacteriol 185:6815–6825. doi:10.1128/JB.185.23.6815-6825.2003.
    OpenUrlAbstract/FREE Full Text
  30. 30.↵
    1. Varghese S,
    2. Tang Y,
    3. Imlay JA
    . 2003. Contrasting sensitivities of Escherichia coli aconitases A and B to oxidation and iron depletion. J Bacteriol 185:221–230. doi:10.1128/JB.185.1.221-230.2003.
    OpenUrlAbstract/FREE Full Text
  31. 31.↵
    1. Wu Y,
    2. Outten FW
    . 2009. lscR controls iron-dependent biofilm formation in Escherichia coli by regulating type I fimbria expression. J Bacteriol 191:1248–1257. doi:10.1128/JB.01086-08.
    OpenUrlAbstract/FREE Full Text
  32. 32.↵
    1. Ronpirin C,
    2. Jerse AE,
    3. Cornelissen CN
    . 2001. Gonococcal genes encoding transferrin-binding proteins A and B are arranged in a bicistronic operon but are subject to differential expression. Infect Immun 69:6336–6347. doi:10.1128/IAI.69.10.6336-6347.2001.
    OpenUrlAbstract/FREE Full Text
  33. 33.↵
    1. Fernández-Martínez J,
    2. Pujalte MJ,
    3. García-Martínez J,
    4. Mata M,
    5. Garay E,
    6. Rodríguez-Valera F
    . 2003. Description of Alcanivorax venustensis sp. nov. and reclassification of Fundibacter jadensis DSM 12178T (Bruns and Berthe-Corti 1999) as Alcanivorax jadensis comb. nov., members of the emended genus Alcanivorax. Int J Syst Evol Microbiol 53(Pt 1):331–338. doi:10.1099/ijs.0.01923-0.
    OpenUrlCrossRefPubMedWeb of Science
  34. 34.↵
    1. Sebastian M,
    2. Ammerman JW
    . 2009. The alkaline phosphatase PhoX is more widely distributed in marine bacteria than the classical PhoA. ISME J 3:563–572. doi:10.1038/ismej.2009.10.
    OpenUrlCrossRefPubMedWeb of Science
  35. 35.↵
    1. Feng S,
    2. Powell SM,
    3. Wilson R,
    4. Bowman JP
    . 2013. Light-stimulated growth of proteorhodopsin-bearing sea-ice psychrophile Psychroflexus torquis is salinity dependent. ISME J 7:2206–2213. doi:10.1038/ismej.2013.97.
    OpenUrlCrossRefPubMedWeb of Science
  36. 36.↵
    1. Bolton FJ,
    2. Wareing DRA,
    3. Sails AD
    . 1997. Comparison of a novel microaerobic system with three other gas-generating systems for the recovery of Campylobacter species from human faecal samples. Eur J Clin Microbiol Infect Dis 16:839–842. doi:10.1007/BF01700415.
    OpenUrlCrossRefPubMed
  37. 37.↵
    1. Moen B,
    2. Oust A,
    3. Langsrud Ø,
    4. Dorrell N,
    5. Marsden GL,
    6. Hinds J,
    7. Kohler A,
    8. Wren BW,
    9. Rudi K
    . 2005. Explorative multifactor approach for investigating global survival mechanisms of Campylobacter jejuni under environmental conditions. Appl Environ Microbiol 71:2086–2094. doi:10.1128/AEM.71.4.2086-2094.2005.
    OpenUrlAbstract/FREE Full Text
  38. 38.↵
    1. Boulos L,
    2. Prévost M,
    3. Barbeau B,
    4. Coallier J,
    5. Desjardins R
    . 1999. LIVE/DEAD® BacLight™: application of a new rapid staining method for direct enumeration of viable and total bacteria in drinking water. J Microbiol Methods 37:77–86. doi:10.1016/S0167-7012(99)00048-2.
    OpenUrlCrossRefPubMedWeb of Science
  39. 39.↵
    1. Leuko S,
    2. Legat A,
    3. Fendrihan S,
    4. Stan-Lotter H
    . 2004. Evaluation of the LIVE/DEAD BacLight kit for extremophilic archaea and environmental hypersaline samples. Appl Environ Microbiol 70:6884–6886. doi:10.1128/AEM.70.11.6884-6886.2004.
    OpenUrlAbstract/FREE Full Text
  40. 40.↵
    1. Jensen RA,
    2. Stenmark SL,
    3. Champney WS
    . 1972. Molecular basis for the differential anti-metabolite action of D-tyrosine in strains 23 and 168 of Bacillus subtilis. Arch Mikrobiol 87:173–180.
    OpenUrlCrossRefPubMed
  41. 41.↵
    1. Box GEP,
    2. Cox DR
    . 1964. An analysis of transformations. J R Stat Soc B 26:211–252.
    OpenUrl
  42. 42.↵
    1. Lahlali R,
    2. Serrhini MN,
    3. Friel D,
    4. Jijakli MH
    . 2006. In vitro effects of water activity, temperature and solutes on the growth rate of P. italicum Wehmer and P. digitatum Sacc. J Appl Microbiol 101:628–636. doi:10.1111/j.1365-2672.2006.02953.x.
    OpenUrlCrossRefPubMed
  43. 43.↵
    1. Vieira-Silva S,
    2. Rocha EPC
    . 2010. The systemic imprint of growth and its uses in ecological (meta)genomics. PLoS Genet 6:e1000808. doi:10.1371/journal.pgen.1000808.
    OpenUrlCrossRefPubMed
  44. 44.↵
    1. Bru D,
    2. Ramette A,
    3. Saby NPA,
    4. Dequiedt S,
    5. Ranjard L,
    6. Jolivet C,
    7. Arrouays D,
    8. Philippot L
    . 2011. Determinants of the distribution of nitrogen-cycling microbial communities at the landscape scale. ISME J 5:532–542. doi:10.1038/ismej.2010.130.
    OpenUrlCrossRefPubMedWeb of Science
  45. 45.↵
    Development Core Team R. 2013. R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria.
  46. 46.↵
    1. Nyström T,
    2. Olsson RM,
    3. Kjelleberg S
    . 1992. Survival, stress resistance, and alterations in protein expression in the marine Vibrio sp. strain S14 during starvation for different individual nutrients. Appl Environ Microbiol 58:55–65.
    OpenUrlAbstract/FREE Full Text
  47. 47.↵
    1. Nichols DS,
    2. Greenhill AR,
    3. Shadbolt CT,
    4. Ross T,
    5. McMeekin TA
    . 1999. Physicochemical parameters for growth of the sea ice bacteria Glaciecola punicea ACAM 611T and Gelidibacter sp. strain IC158. Appl Environ Microbiol 65:3757–3760.
    OpenUrlAbstract/FREE Full Text
  48. 48.↵
    1. Matin A
    . 1991. The molecular basis of carbon-starvation-induced general resistance in Escherichia coli. Mol Microbiol 5:3–10. doi:10.1111/j.1365-2958.1991.tb01819.x.
    OpenUrlCrossRefPubMedWeb of Science
  49. 49.↵
    1. McDougald D,
    2. Gong L,
    3. Srinivasan S,
    4. Hild E,
    5. Thompson L,
    6. Takayama K,
    7. Rice SA,
    8. Kjelleberg S
    . 2002. Defences against oxidative stress during starvation in bacteria. Antonie Van Leeuwenhoek 81:3–13. doi:10.1023/A:1020540503200.
    OpenUrlCrossRefPubMedWeb of Science
  50. 50.↵
    1. Rangel DEN,
    2. Anderson AJ,
    3. Roberts DW
    . 2008. Evaluating physical and nutritional stress during mycelial growth as inducers of tolerance to heat and UV-B radiation in Metarhizium anisopliae conidia. Mycol Res 112:1362–1372. doi:10.1016/j.mycres.2008.04.013.
    OpenUrlCrossRefPubMed
  51. 51.↵
    1. Fung DKC,
    2. Chan EWC,
    3. Chin ML,
    4. Chan RCY
    . 2010. Delineation of a bacterial starvation stress response network which can mediate antibiotic tolerance development. Antimicrob Agents Chemother 54:1082–1093. doi:10.1128/AAC.01218-09.
    OpenUrlAbstract/FREE Full Text
  52. 52.↵
    1. Nguyen D,
    2. Joshi-Datar A,
    3. Lepine F,
    4. Bauerle E,
    5. Olakanmi O,
    6. Beer K,
    7. McKay G,
    8. Siehnel R,
    9. Schafhauser J,
    10. Wang Y,
    11. Britigan BE,
    12. Singh PK
    . 2011. Active starvation responses mediate antibiotic tolerance in biofilms and nutrient-limited bacteria. Science 334:982–986. doi:10.1126/science.1211037.
    OpenUrlAbstract/FREE Full Text
  53. 53.↵
    1. Poole K
    . 2012. Stress responses as determinants of antimicrobial resistance in Gram-negative bacteria. Trends Microbiol 20:227–234. doi:10.1016/j.tim.2012.02.004.
    OpenUrlCrossRefPubMedWeb of Science
  54. 54.↵
    1. Saito MA,
    2. Goepfert TJ,
    3. Ritt JT
    . 2008. Some thoughts on the concept of colimitation: three definitions and the importance of bioavailability. Limnol Oceanogr 53:276–290. doi:10.4319/lo.2008.53.1.0276.
    OpenUrlCrossRefWeb of Science
  55. 55.↵
    1. Cockell CS,
    2. Olsson K,
    3. Knowles F,
    4. Kelly L,
    5. Herrera A,
    6. Thorsteinsson T,
    7. Marteinsson V
    . 2009. Bacteria in weathered basaltic glass, Iceland. Geomicrobiol J 26:491–507. doi:10.1080/01490450903061101.
    OpenUrlCrossRefWeb of Science
  56. 56.↵
    1. Gorbushina AA,
    2. Broughton WJ
    . 2009. Microbiology of the atmosphere-rock interface: how biological interactions and physical stresses modulate a sophisticated microbial ecosystem. Annu Rev Microbiol 63:431–450. doi:10.1146/annurev.micro.091208.073349.
    OpenUrlCrossRefPubMedWeb of Science
  57. 57.↵
    1. La Duc MT,
    2. Dekas A,
    3. Osman S,
    4. Moissl C,
    5. Newcombe D,
    6. Venkateswaran K
    . 2007. Isolation and characterization of bacteria capable of tolerating the extreme conditions of clean room environments. Appl Environ Microbiol 73:2600–2611. doi:10.1128/AEM.03007-06.
    OpenUrlAbstract/FREE Full Text
  58. 58.↵
    1. Lamosa P,
    2. Martins LO,
    3. Da Costa MS,
    4. Santos H
    . 1998. Effects of temperature, salinity, and medium composition on compatible solute accumulation by Thermococcus spp. Appl Environ Microbiol 64:3591–3598.
    OpenUrlAbstract/FREE Full Text
  59. 59.↵
    1. Ventosa A,
    2. Nieto JJ,
    3. Oren A
    . 1998. Biology of moderately halophilic bacteria. Microbiol Mol Biol Rev 62:504–544.
    OpenUrlAbstract/FREE Full Text
  60. 60.↵
    1. Hecker M,
    2. Völker U
    . 2001. General stress response of Bacillus subtilis and other bacteria. Adv Microb Physiol 44:35–91. doi:10.1016/S0065-2911(01)44011-2.
    OpenUrlCrossRefPubMedWeb of Science
  61. 61.↵
    1. Petersohn A,
    2. Brigulla M,
    3. Haas S,
    4. Hoheisel JD,
    5. Völker U,
    6. Hecker M
    . 2001. Global analysis of the general stress response of Bacillus subtilis. J Bacteriol 183:5617–5631. doi:10.1128/JB.183.19.5617-5631.2001.
    OpenUrlAbstract/FREE Full Text
  62. 62.↵
    1. Bhaganna P,
    2. Volkers RJM,
    3. Bell ANW,
    4. Kluge K,
    5. Timson DJ,
    6. McGrath JW,
    7. Ruijssenaars HJ,
    8. Hallsworth JE
    . 2010. Hydrophobic substances induce water stress in microbial cells. Microb Biotechnol 3:701–716. doi:10.1111/j.1751-7915.2010.00203.x.
    OpenUrlCrossRefPubMed
  63. 63.↵
    1. Hallsworth JE,
    2. Heim S,
    3. Timmis KN
    . 2003. Chaotropic solutes cause water stress in Pseudomonas putida. Environ Microbiol 5:1270–1280. doi:10.1111/j.1462-2920.2003.00478.x.
    OpenUrlCrossRefPubMedWeb of Science
  64. 64.↵
    1. Rangel DEN
    . 2011. Stress induced cross-protection against environmental challenges on prokaryotic and eukaryotic microbes. World J Microbiol Biotechnol 27:1281–1296. doi:10.1007/s11274-010-0584-3.
    OpenUrlCrossRefPubMed
  65. 65.↵
    1. Spector MP,
    2. Kenyon WJ
    . 2012. Resistance and survival strategies of Salmonella enterica to environmental stresses. Food Res Int 45:455–481. doi:10.1016/j.foodres.2011.06.056.
    OpenUrlCrossRef
  66. 66.↵
    1. Cray JA,
    2. Bell ANW,
    3. Bhaganna P,
    4. Mswaka AY,
    5. Timson DJ,
    6. Hallsworth JE
    . 2013. The biology of habitat dominance; can microbes behave as weeds? Microb Biotechnol 6:453–492. doi:10.1111/1751-7915.12027.
    OpenUrlCrossRefPubMed
  67. 67.↵
    1. Oren A,
    2. Hallsworth JE
    . 2014. Microbial weeds in hypersaline habitats: the enigma of the weed-like Haloferax mediterranei. FEMS Microbiol Lett 359:134–142. doi:10.1111/1574-6968.12571.
    OpenUrlCrossRefPubMed
PreviousNext
Back to top
Download PDF
Citation Tools
Reduction of the Temperature Sensitivity of Halomonas hydrothermalis by Iron Starvation Combined with Microaerobic Conditions
Jesse P. Harrison, John E. Hallsworth, Charles S. Cockell
Applied and Environmental Microbiology Feb 2015, 81 (6) 2156-2162; DOI: 10.1128/AEM.03639-14

Citation Manager Formats

  • BibTeX
  • Bookends
  • EasyBib
  • EndNote (tagged)
  • EndNote 8 (xml)
  • Medlars
  • Mendeley
  • Papers
  • RefWorks Tagged
  • Ref Manager
  • RIS
  • Zotero
Print

Alerts
Sign In to Email Alerts with your Email Address
Email

Thank you for sharing this Applied and Environmental Microbiology article.

NOTE: We request your email address only to inform the recipient that it was you who recommended this article, and that it is not junk mail. We do not retain these email addresses.

Enter multiple addresses on separate lines or separate them with commas.
Reduction of the Temperature Sensitivity of Halomonas hydrothermalis by Iron Starvation Combined with Microaerobic Conditions
(Your Name) has forwarded a page to you from Applied and Environmental Microbiology
(Your Name) thought you would be interested in this article in Applied and Environmental Microbiology.
CAPTCHA
This question is for testing whether or not you are a human visitor and to prevent automated spam submissions.
Share
Reduction of the Temperature Sensitivity of Halomonas hydrothermalis by Iron Starvation Combined with Microaerobic Conditions
Jesse P. Harrison, John E. Hallsworth, Charles S. Cockell
Applied and Environmental Microbiology Feb 2015, 81 (6) 2156-2162; DOI: 10.1128/AEM.03639-14
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
  • Top
  • Article
    • ABSTRACT
    • INTRODUCTION
    • MATERIALS AND METHODS
    • RESULTS AND DISCUSSION
    • ACKNOWLEDGMENTS
    • FOOTNOTES
    • REFERENCES
  • Figures & Data
  • Info & Metrics
  • PDF

Related Articles

Cited By...

About

  • About AEM
  • Editor in Chief
  • Editorial Board
  • Policies
  • For Reviewers
  • For the Media
  • For Librarians
  • For Advertisers
  • Alerts
  • RSS
  • FAQ
  • Permissions
  • Journal Announcements

Authors

  • ASM Author Center
  • Submit a Manuscript
  • Article Types
  • Ethics
  • Contact Us

Follow #AppEnvMicro

@ASMicrobiology

       

ASM Journals

ASM journals are the most prominent publications in the field, delivering up-to-date and authoritative coverage of both basic and clinical microbiology.

About ASM | Contact Us | Press Room

 

ASM is a member of

Scientific Society Publisher Alliance

 

American Society for Microbiology
1752 N St. NW
Washington, DC 20036
Phone: (202) 737-3600

Copyright © 2021 American Society for Microbiology | Privacy Policy | Website feedback

 

Print ISSN: 0099-2240; Online ISSN: 1098-5336