Ammonia sets limit to life and alters physiology independently of pH in Halomonas meridiana

doi:10.21203/rs.3.rs-6178140/v1

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Ammonia sets limit to life and alters physiology independently of pH in Halomonas meridiana

https://doi.org/10.21203/rs.3.rs-6178140/v1

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published 04 Jun, 2025

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Significant amounts of ammonia are released anthropogenically, while ammonia is also retained within the water bodies of icy moons that are of astrobiological interest. Ammonia is toxic to many forms of life at high concentrations, and thus it is necessary to understand the habitability impact of ammonia on these environments. The survival limits and physiological response of aerobic bacteria in ammonia, and whether ammonia toxicity is distinct from toxicity by high pH, is poorly understood. Here, we investigate the survival thresholds, growth kinetics, and metabolomic response of Halomonas meridiana in ammonia-water solutions and pH-matched sodium hydroxide solutions. Using closed- and open-air systems to mimic environments with ammonia retention or dispersion, we found complete and partial cell death above 0.05 M ammonia, respectively. In open-air systems, a sub-set of cells survived up to 0.25 M ammonia; metabolomics revealed unique physiological responses to ammonia, including elevation of cyclic compounds and Coenzyme A metabolites, suggesting mechanisms of ammonia toxicity and adaptation. Ammonia and high pH toxicity were found to be distinct. These findings show that ammonia can impose a distinct geobiological limit, potentially constraining habitability of ammonia-rich terrestrial and extraterrestrial environments.

Earth and environmental sciences/Planetary science/Astrobiology

Biological sciences/Microbiology/Bacteria/Bacterial physiology

Biological sciences/Microbiology/Bacteriology

Biological sciences/Physiology/Metabolism/Metabolomics

Ammonia (hereafter considered as total ammonia, ammonia (NH₃) + ammonium (NH₄⁺)) is a prebiotic component of terrestrial environments. As such, ammonia is the preferred nitrogen source for many bacteria and plays an integral role in the global nitrogen cycle. Yet, despite biochemical significance, the toxicity of NH₃ has been well-documented across the domains of life ^1–4. NH₃ is gaseous under temperate conditions. The small size and non-polar properties of NH₃ facilitates passive permeation through membranes ^5–8, causing significant intracellular disruption ^9–11. The equilibrium between NH₃ and NH₄⁺ is influenced by factors such as pH (with NH₃ predominating at pH > 9.25), salinity and temperature ^12–14. Mono Lake, California, underscores the role of NH₃ in shaping microbial ecosystems; depth-dependent shifts in biodiversity are driven by the spatial distribution of abiotic factors including ammonia, which is predominately in the NH₃ form ^15,16.

Globally, several dozen tetragrams of ammonia are released annually from agriculture ^17,18. Ammonia deposition into soil, water and vegetation occurs as a consequence ^19,20. In alkaline soils (pH > 7), application of ammonium fertilizers leads to NH₃ volatilization; NH₄⁺ is transitioned to gaseous NH₃ and released into the atmosphere. Indeed, applied nitrogen losses of up to 66% have been recorded in alkaline soils as a result of ammonium fertilizer application ^21–23. Although toxicological effects are presumed, the downstream impact of NH₃ volatilization on microbial diversity and community structures in alkaline environments, where the relative abundance of NH₃ exceeds NH₄⁺, is not well understood. Likewise, the habitability impact of ammonia within extraterrestrial environments is also poorly considered. Ammonia is a primordial constituent of the solar system and has been detected at a volume mixing ratio of 0.61–1.3% on the icy moon Enceladus, orbiting Saturn ^29,30. The evidence of a saline, liquid water subsurface ocean ^24,25, and other physicochemical conditions suitable for life ^26–28, has encouraged strong astrobiological interest in this satellite. With a pH predicted at or above 9 ^27,28, the ocean of Enceladus would bear appreciable amounts of toxic NH₃. Due to the limited research on NH₃ tolerance in bacteria, coupled with the presence of carbon dioxide and hydrogen in the ocean, habitability assessments of Enceladus are currently limited to methanogenic organisms ^31–33.

To accurately assess the habitability impacts of ammonia, fundamental questions remain. Few studies have systematically investigated the survival limits or physiology of microorganisms under high NH₃ stress. Bacteria such as Bacillus pasteurii can grow in up to 0.5 M NH₃ ³⁴, however this species can utilise ammonia for ATP generation ³⁵, substrate permeability ³⁶ and oxidation of substrates ³⁷. Ammonia-oxidising bacteria (AOB) can metabolise NH₃ but are primarily adapted to NH₄^{+ 38,39}. Without specific adaptations, toxicity at 0.1 M ammonia (pH 9.5) has been observed in B. subtilis and Escherichia coli. However, similar toxicity was noted in sodium chloride solutions of matching pH ⁴⁰. The use of neutrophilic organisms has thus far limited our understanding of whether ammonia toxicity is intrinsically pH-based. There is compelling need for research examining the survival limits and physiological response of an alkaline adapted bacteria in ammonia solutions exceeding pH 9.25 (NH₃ > 50%), and comparing with a pH matched counterpart.

Here, we use a combination of growth kinetics and cell viability assays to identify habitability limits of Halomonas meridiana Slfth1 in ammonia solutions with high NH₃ content. This organism lacks specific adaptations to ammonia ⁴¹, but possesses alkalitolerant traits relevant for this investigation ⁴². We utilise growth systems permitting (“open-air system”) or preventing (“closed-air system”) gaseous escape to mimic environments where ammonia is environmentally dispersed and retained, respectively. By using an alkaline adapted organism, we resolve that ammonia toxicity is independent from pH toxicity and, and, using microscopy and metabolomics analysis, define the specific toxicity effects of ammonia on bacterial physiology independent of pH. Together, these findings advance the understanding of ammonia toxicity and should foster improved habitability assessments of NH₃-rich environments.

Physicochemical properties of ammonia and pH-matched solutions

Ammonia solutions were prepared from liquid ammonia to molar concentrations of 0.01, 0.025, 0.05, 0.1, 0.25, 0.5 and 1 M, equating to pH values between 8 and 11 (Fig. 1a). Positive control (PC) was unamended yeast media. The percentage abundance of NH₃ to NH₄⁺ at these concentrations and pH values is given in Fig. 1b. From concentrations of 0.05 M ammonia and higher, NH₃ accounts for over 50% of the total NH₃/NH₄⁺in solution. Solutions of increasing ammonia concentration showed decreased oxygen concentrations; however, this decrease was non-significant from pH-matched counterparts (Fig. 1c). Alterations to water activity with increasing ammonia and pH were also statistically non-significant (Fig. 1d).

Ammonia sets a distinct molarity threshold for viability and growth

Bacteria were grown in ammonia solutions utilising two system types: a ‘closed-air’ system – to determine growth limits in a perpetually ammonia exposed environments, and an ‘open-air’ system – to determine growth limits in an environment permitting ammonia dispersal. Fig. 2a shows the final colony forming units of H. meridiana after incubation in the closed-air system for 72 h. Colonies were evident following incubation in 0.01 M, 0.025 M and 0.05 M ammonia, but colony number decreased as NH₃ increased relative to NH₄⁺ (Fig. 1b). A lower number of viable colonies compared to positive control was observed in 0.01 M ammonia(p < 0.01). No significant difference in colony number was observed when H. meridiana was incubated in 0.025 M compared to positive control (p = 0.056) or 0.01 M ammonia (p = 0.323). This concentration of ammonia has a pH within the optimum range for H. meridiana (pH 9) (Fig. 1a). Lower viable cell numbers were observed when incubated in 0.05 M compared to positive control (p < 0.0001) and 0.025 M ammonia (p < 0.01). No viable colonies were present following incubation in 0.1 M ammonia or ammonia solutions of higher concentration (Fig. 2a).

Fig. 2b. shows the growth dynamics of H. meridiana Slthf1 over 48 h in increasing concentrations of ammonia in an open-air system. Lag phase duration, doubling time (T_d) and final OD₆₀₀ are presented in Fig. 2c-e, respectively. At concentrations exceeding 0.01 M ammonia, lag phase was greater with higher ammonia concentrations; incubation in 0.1 M and 0.25 M ammonia prolongs lag phase time by 6-fold and 25-fold compared to the positive control, respectively (Fig. 2c). The lag phase time in 0.1 M ammonia was higher than that in 0.05 M ammonia (p < 0.05). Likewise, the lag phase time in 0.25 M ammonia was higher than that in 0.1 M ammonia (p < 0.0001) (Fig. 2c). T_d was also higher with increasing ammonia concentration, with higher T_d compared to the positive control observed in 0.05 M (p < 0.05), 0.1 M (p ≤ 0.0001) and 0.25 M (p < 0.0001) solutions (Fig. 2d). The T_d was not significantly altered between positive control and 0.01 M (p = 0.965) or 0.1 M and 0.25 M solutions (p = 0.9972). Final OD₆₀₀after 48 h growth showed no statistically significant difference from positive control in 0.01 M (p = 0.932), 0.025 M (p = 0.330), 0.1 M (p = 0.248) and 0.25 M solutions (p = 0.135), but higher OD₆₀₀ values were observed in 0.05 M compared to the positive control (p < 0.01) (Fig. 2e). Cell density was lower in 0.5 and 1 M solutions compared to the positive control (0.5 M, p < 0.01; 1 M, p < 0.001) (Fig. 2e), where no distinguishable growth occurred within 48 hours (Fig. 2b).

The prolonged lag phase in cells inoculated into 0.1 M and 0.25 M ammonia indicate bactericidal reduction to the cell population. Alternatively, ammonia may act bacteriostatically, the effects of which cease once ammonia has dispersed. To assess this, temporal cell viability was compared with ammonia concentration over time in 0.1 M (Fig. 2f) and 0.25 M ammonia solutions (Fig. 2g). After 4 h exposure, viable cells in 0.1 M ammonia show a 1000-fold decrease from 0 h (t=28.28, df=4, p < 0.0001) (Fig. 2f). In 0.25 M ammonia, viable cells at 4 h are reduced 10-fold from 0 h (t=6.499, df=4, p < 0.01) (Fig. 2g). Bactericidal reduction to cell populations within 0.1 M ammoniasolutions cease between 4 h and 8 h, where a small but non-significant increase in cell viability is observed t=2.421, df=4, p = 0.0727) (Fig. 2f). Likewise, bactericidal effects are absent between 4 h and 8 h in 0.25 M ammoniasolutions where cell numbers stabilise (t=0.8934, df=4, p = 0.442) and increase from 8 h to 16 h (t=11.76, df=4, p < 0.001) (Fig. 2g). The increase in cell viability within 0.1 and 0.25 M ammoniasolutions at 8 h and 16 h, respectively, aligns with diminished ammonia levels to ≤ 0.05 M at 4 h and 13 hr, respectively (Fig. 2f, 2g). Thus, lag phase extension in the open-air system reflects two events: (1) an immediate bactericidal effect and (2) growth initiation after ammonia levels drop to sub-bactericidal levels (≤ 0.05 M) in surviving populations.

Ammonia toxicity is independent from pH toxicity

Ammonia is a weak base that raises solution pH with increasing concentration. To delimitate ammonia toxicity from pH toxicity, growth experiments were repeated in NaOH solutions pH-matched to ammonia solutions. Fig. 3a presents the final CFU/mL of H. meridiana in solutions following 72 h closed-air system incubation. No significant differences were observed between cells grown in positive control and pH-matched solutions up to pH 10.78 (equivalent to 1 M ammonia). Fig. 3b presents the growth curve of H. meridiana grown under pH-matched NaOH solutions in an open-air system, with extrapolated parameters of lag phase (Fig. 3c), T_d (Fig. 3d) and final OD₆₀₀ (Fig. 3e) presented in log₂ fold changes (log₂FC) from growth in ammonia solutions of the same pH. Significantly lower lag phases were observed in cells grown in pH 8.96, 9.38, 9.73 and 10.18 compared to those grown at 0.025 M (t=4.399, df=3, p < 0.05), 0.05 M (t=11.26, df=3, p < 0.01), 0.1 M (t=7.357, df=3, p < 0.001) and 0.25 M ammonia (t=31.72, df=3, p < 0.0001) (Fig. 3c). The T_d compared between cells grown in ammonia and pH-matched counterparts was non-significantly different except for those grown in pH 9.38 (t=3.542, df=3, p < 0.05), which showed a lower T_d (Fig. 3d). Final OD₆₀₀ at 48 h was lower in pH 8.96 (t=4.882, df=3, p < 0.05), pH 9.38 (t=6.810, df=3, p < 0.01), pH 9.73 (t=4.123, df=3, p < 0.05) and pH 10.18 (t=7.206, df=3, p < 0.01) NaOH-solutions compared to growth in ammonia counterparts, while higher OD₆₀₀ was observed in pH 10.49 (t=16.09, df=3, p < 0.001) and pH 10.78 solutions (t=16.74, df=3, p < 0.001) compared to ammonia counterparts (Fig. 3e). These findings confirm that lag phase extension and absent growth in 0.5 and 1 M ammonia cannot be attributed to pH increases.

To broaden the pH comparison beyond NaOH, lag phase analysis was used to explore whether the prolonged lag phase in 0.25 M ammonia (Fig. 2c) could be replicated in KOH, Na2SiO3, K2CO3, and Na2CO3 at pH 10.18 (Fig. 3f). No differences in lag phase duration were observed between H. meridiana grown in Na2SiO3 (p = 0.779) or Na2CO3 (p = 0.996) compared to NaOH. Longer lag phases were seen in KOH (p < 0.05) and K2CO3 (p < 0.0001) compared to NaOH, possibly reflecting reduced adaptation to potassium in H. meridiana. All high-pH solutions showed significantly shorter lag phases than 0.25 M ammonia (p < 0.001), confirming the pH-independent toxicity of ammonia.

Ammonia toxicity exerts distinct changes to bacterial morphology

Specific bactericidal effects of ammonia on cells can be observed in Fig. 4. Under control growth conditions (unaltered yeast media) (Fig. 4a-b), H. meridiana exhibited ribosome-rich cytoplasm, visible nucleoids and clear division of outermembrane, periplasm and inner membrane. Cells exhibited irregular, undulating outermembrane, enlarged periplasmic spaces and PHA-like granules. Upon 2 h treatment of 1 M ammonia(Fig. 4c-d), cells showed intracellular aggregation and loss of ribosomes with some showing cytoplasmic loss and lysis. The periplasmic space volume decreased, and detachment of the inner membrane from the outermembrane was observed. Cells showed expansion of electrolucent cavities, possibly surrounding nucleoids. Condensed material within the electrolucent cavities exhibited splayed morphologies, possibly indicating disruption to DNA supercoiling. Cells exposed to a NaOH solutions pH-matched to 1 M ammonia(10.78 pH) also showed some cell lysis events (Fig. 4e-f). However, there were few morphological differences from cells grown in control media. Differences include uniform outermembrane, and fewer PHA-like granules.

Metabolic pathways altered in response to ammonia and high pH exposure

The survival of cells in up to 0.25 M ammonia in an open-air system suggests adaptations enabling tolerance to ammonia. Thus, untargeted metabolomics was performed on H. meridiana in unamended yeast media (hereafter denoted ‘control’) and yeast media with 0.25 M ammonia at pH 10.18 (hereafter denoted ‘0.25 M ammonia_’) to determine variations in metabolism that may account for the differences observed. To delimitate high pH adaptations from ammonia adaptations, metabolomics was also performed following exposure to yeast media adjusted to pH 10.18 with NaOH (hereafter denoted ‘NaOH pH 10.18’). Growth kinetics and sampling points in these conditions are indicated in Fig. 5a. Principal component analysis (PCA) (Fig. 5b) separated 0.25 M ammonia and NaOH pH 10.18 conditions from the control, with overlap between 0.25 M ammonia and NaOH pH 10.18 conditions indicating metabolic similarity. Univariate volcano analysis identified 23 features significantly altered in 0.25 M ammonia/control (Fig. 5c), 10 features significantly altered in 0.25 M ammonia/NaOH pH 10.18 (Fig. 5d), and 28 features significantly altered in NaOH pH 10.18/control (Fig. 5e). The volcano analysis dataset for each comparison group is shown in Supplementary Table 2-4.

The most significantly altered metabolites between these conditions were identified by ANOVA (Supplementary Table 5), depicted as a heatmap (Fig. 5f). Overall, high similarity was found between 0.25 M ammonia and NaOH pH 10.18 exposed samples; both conditions generally exhibited higher levels of unsaturated phospholipid and lower levels of linoleic acid and derivatives. Intermediates that feed glycerophospholipid biosynthesis, CMP-sialic acid (F = 2021.9, p < 0.0001) and glycerol-3-phosphate (F = 2022.2, p < 0.0001), were more abundant compared to the control. Amino acid pathway intermediates indole-3-ethanol (F = 2327.8, p < 0.0001), N, N-dimethylglycine (F = 30.626, p < 0.001) and N-acetylglutamate (F = 25.653, p < 0.01) were also altered compared to control. There were reduced levels of N-acetylserotonin (F = 18876, p < 0.0001) and N-acetyl-L-aspartic acid (F = 24.091, p < 0.01) that could suggest reactions with acetyl donors were less favourable at high pH in both conditions.

Ammonia exposure elicits a unique metabolomic response

Despite general similarities between cells exposed to 0.25 M ammonia and NaOH pH 10.18, ANOVA indicated five metabolites significantly altered only in 0.25 M ammonia exposed H. meridiana. Box plots of these metabolites are presented in Fig. 6a-e. Samples exposed to 0.25 M ammoniashowed significantly higher levels of unidentified metabolites, metabolite i at m/z=215.084 (Fig. 6a) and metabolite ii at m/z=157.0973 (Fig. 6b), compared to NaOH pH 10.18 exposed samples (metabolite i, p < 0.0001; metabolite ii, p < 0.0001) and control samples (metabolite i, F = 2112.3, p < 0.0001; metabolite ii, F = 4865.6, p < 0.0001). Library annotations matched metabolite i to nitrogen-containing heterocyclic compound atrazine (89.6% Q score) and metabolite ii to hydrocarbon 2,7-dimethylnaphthalene (96.5% Q score). Structures of atrazine and 2,7-dimethylnaphthalene are depicted in Fig. 6a and Fig. 6b, respectively. Multivariate analysis also revealed exposure to 0.25 M ammonia increased the relative abundance of pantothenate (F = 28.839, p < 0.001) (Fig. 6c), and amino acids D-allo-isoleucine (F = 28.686, p < 0.001) (Fig. 6d), and alanine (F = 25.391, p < 0.01) (Fig. 6e), compared to those in NaOH pH 10.18 and control conditions.

The influence of ammonia on habitability, particularly to microbes, is underrepresented in scientific literature. Yet, the pollution of ammonia into the environment could shape the underlying microbial community structures and biodiversity of alkaline environments. The presence of ammonia within the subsurface ocean of Enceladus, predicted at an alkaline pH, may also shape the habitability expectations of this satellite. Although the presence of life beyond Earth is speculative, terrestrial organisms can be utilised to explore the boundaries of known life under similar conditions to assess potential for habitability. In this study, we utilise alkalitolerant and halophilic H. meridiana as an analogue organism to investigate the molar concentration thresholds for survival, and physiological changes, that might be present in aerobic bacteria within extreme NH₃ environments.

The exposure of H. meridiana to increasing ammonia concentrations revealed that growth at 0.05 M ammonia, where the relative abundance of NH₃ is 62%, sets a consistent habitability limit regardless of whether in a closed-air or open-air system. The tolerance of microbes to ammonia varies and cannot be predicted by genus or species alone; strain-specific responses to stress have been well documented ^43–48. However, the molarity limit established in this study aligns with data for other bacteria, including B. subtilis strains T1, T2, T19, T22, T33, T39 ³⁴ and Enterobacter cloaecae HNR ⁹. The absolute limit for ammonia survival in the literature extends to 0.716 M (NH₄)₂SO₄ at pH 9, survived by strains of B. subtilis and Proteus morgani ³⁴. However, the relative abundance of NH₃ in these studies is below 40%. Our results characterise a higher habitability limit in NH₃ than previously established for an aerobic bacterium.

Independent ammonia and pH toxicity has been characterised in E. coli ⁴⁰, yet indistinguishable toxicity has been recorded with B. subtilis, Sporosarcina, Paenibacillus, Staphylococcus, Brevibacillus, Streptomyces, Pseudomonas and Arthrobacter ^2,40. Using an alkalitolerant organism, we establish ammonia toxicity is distinct from pH toxicity in H. meridiana, and does not exert toxicity via oxygen displacement or water activity changes. However, the degree of similarly between the metabolomic profiles of cells exposed to 0.25 M ammonia and a pH-matched NaOH solution indicates H. meridiana primarily responds to ammonia using high pH adaptations. Unique physiological responses to ammonia in H. meridiana included the accumulation of two cyclic compounds, one of which bore amine functional groups. The presence of these compounds may be indicative of intracellular NH₃-driven reactions ^49,50. The disruption of such structures to cellular components could be a mechanism of pH-independent toxicity. Indeed, H. meridiana treated with ammonia exhibited aggregation to intracellular components that would suggest significant disruption to cell contents.

Elevated levels of alanine, D-allo-isoleucine, and pantothenate were also exclusively observed in ammonia exposed H. meridiana. The presence of an isoleucine derivative and pantothenate suggested modulation to the Coenzyme A pathway that affects the cell membrane ^51,52, and amino acid metabolism ^52–54. Indeed, D-amino acids have been implicated in cell wall remodelling in Vibrio cholerae and B. subtilis ⁵⁵. Alanine dehydrogenase catalyses the production of alanine by a reaction between pyruvate and ammonium ^56–58. Thus, alanine could be a by-product of elevated ammonia levels. Notably, D-amino acid aminotransferase catalyses the production of pyruvate and D-glutamate from D-alanine and 2-oxoglutarate, for which D-allo-isoleucine can act as an amino donor in Thermotoga maritima ⁵⁹. The presence of D-allo-isoleucine may therefore lead to enrichment of pyruvate and, in turn, enrichment of D-alanine or L-alanine via the alanine dehydrogenase pathway, or both. D-alanine may act to impede ammonia permeation; D-alanine can increase rigidity, and decrease permeability, of the peptidoglycan layer ^60–62. L-alanine may act to prevent excessive ammonia metabolism; in some bacteria, L-alanine is an allosteric inhibitor of the nitrogen assimilation enzyme glutamine synthetase ^63–65.

Interpreted from the point of view of planetary habitability, the ammonia toxicity limit established in this study is higher than the predicted 0.01–0.1 M ammonia content of the Enceladus ocean ^28,30. The closed-air system is the most accurate representation of icy moon subsurface oceans as ammonia concentrations are presumed to remain constant. With oceanic temperatures estimated at 0 ºC ²⁶ and salinity at 4% ²⁷, the Enceladus ocean would require a pH of < 9.96 to satisfy the NH₃ < 62% habitability boundary suggested in this study, which is in range of current estimations ^27,28. In the terrestrial environment, up to 78 tetragrams of ammonia per annum are expelled globally from agricultural processes ^17,18. A water volume of 1 litre would require just 1 gram of ammonia to exceed an ammonia concentration of 0.05 M. In alkaline environments, this may alter bacterial community structures and biodiversity once the relative abundance of NH₃ exceeds thresholds for survival, as has been observed in Mono Lake ¹⁵. However, unless NH₃ deposition is continuous, bactericidal effects may be transient due to the open-air dispersal of NH₃ – repopulation of H. meridiana in an open-air ammonia system was observed at concentrations above 0.05 M ammonia.

Without field-based environmental surveys, the effects of NH₃ on terrestrial and extraterrestrial habitability are only estimated. Additionally, without isolation and biological characterisation of metabolites i and ii, we can only speculate the contribution of these molecules in ammonia toxicity. However, we have shown that bacterial growth and viability may become impacted when molar concentrations of ammonia are above 0.05 M, where NH₃ exceeds NH₄⁺. Moreover, we deduce that ammonia and pH toxicity are distinct, suggesting a need to modify the current understanding of NH₃ toxicity in bacteria. Together, these findings can be integrated into the current knowledge regarding the impact of ammonia on terrestrial geomicrobiology and may inform future habitability assessments of NH₃-rich extraterrestrial environments.

Solution preparation

Solutions of liquid ammonia were made to the required molar concentrations—0.01, 0.025, 0.05, 0.1, 0.25, 0.5, 1 M ammonia_—in yeast media using 35% NH₃ (Fisher Scientific, CAS Number: 1336-21-6). The pH of the solutions was determined with a Jenway 3510 benchtop pH meter. pH-matched sodium hydroxide solutions (NaOH; Fisher Scientific, CAS Number: 1310-73-2) were prepared by gradual addition of 1 M NaOH into yeast media. To compare growth in different basic solutions at a pH equivalent to that of 0.25 M ammonia (pH 10.18), several bases were selected that have a natural pH at or greater than pH 10.18 and high solubility in water. Thus, yeast media was made to pH 10.18 ± 0.01 using gradual addition of 1 M NaOH, 1 M potassium hydroxide (KOH), 1 M sodium metasilicate (Na₂SiO₃), 1 M potassium carbonate (K₂CO₃) or 1 M sodium carbonate (Na₂CO₃). Growth experiments proceeded as outlined below. pH was matched to within ± 0.01 pH units. All solutions were filter-sterilised through a 0.22-micron pore before use.

NH₃/NH₄⁺ percentage abundance

The percentage concentration of ammonia was determined by indirect calculation based on commonly utilised equations developed by Hampson ¹². Firstly, the ionic strength of the prepared solutions based on known salinity (0.2 M NaCl) was determined by Eq. 1.0

\(\:I=19.9273\left(S\right)/(1000-1.005109\left(S\right))\)

Eq. 1.0

Where I = molar ionic strength; S = salinity (parts per thousand, ppt).

The stoichiometric acid hydrolysis constant of ammonium ions, pKa^s, based on I, was then calculated. pKa^s values for given ionic strengths were provided by Hampson ¹². These values were plotted to obtain a linear regression for calculating pKa (Supplementary Fig. 1). pKa^s was calculated from Eq. 2.0.

\(\:pK{a}^{s}=0.1179\left(I\right)+9.246\)

Eq. 2.0

Percentage abundance of NH₃ at given salinity, pH and temperature was then calculated from Eq. 3.0. As salinity, temperature, pressure and pKa^s are constant in the experiment, only pH was modified to give percentage abundance of NH₃ at each pH value (Supplementary Table 1).

\(\:N{H}_{3}=N{H}_{3}+N{H}_{4}^{+}/(1+{10}^{\left({pKa}^{s}+0.0324\left(298-T\right)+0.0415\left(P\right)/T-1\right)}\)

Eq. 3.0

Where P = 1 atm pressure; T = temperature (K)

Oxygen measurements

Oxygen concentrations (µmol/L) of unamended yeast media, ammonia solutions and pH-matched solutions, prepared as described above, were recorded using an oxygen microsenser with tip diameter of < 2 mm (OX-500, Unisense, Denmark). Solutions were equilibrated for 0.5 h before reading at 28 ºC in 96-well plates sealed with aluminium foil to prevent gas loss.

Water activity

Unamended yeast media, ammonia solutions and pH-matched solutions were prepared as described above. Solutions were equilibrated for 1.5 h and water activities measured at 28 ºC with a Rotronic HP23-AW water activity meter (Rotoronic AG, Bassersdorf, Switzerland).

Bacterial strain selection

Halomonas meridiana Slthf1 (DSM 15724; Gram negative) was obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ). The organism was originally isolated at a depth of 2000 m from low temperature hydrothermal fluid in the East Pacific Rise. The strain exhibits halophilic growth up to 22% NaCl, and shows tolerance up to pH 12 ⁴². The strain is therefore representative of the physiological attributes that could be necessary to grow in saline, alkaline fluids. A complete genome sequence is available for this strain ⁴¹. This organism is not known to have specific adaptations to ammonia; this was a deliberate choice. The purpose of this study was not to study microbial biochemical adaptation to high ammonia, but rather to study whether NH₃ can act as an abiotic factor influencing bacterial growth, survival and physiology.

Bacterial culture

An aerobic culture of H. meridiana was grown in glass conical Erlenmeyer flasks at 28 ºC in an orbital benchtop shaking incubator set to rotate at 150 RPM. We assume ammonia-rich environments present optimal nutrient conditions (i.e., available organic debris) with a salinity < 2% for the purpose of this study. Thus, H. meridiana Slthf1 was grown in a yeast media consisting of 1g/100 mL Bacto™ yeast extract (Becton, Dickinson and Company), 0.2 M NaCl (1.17% salinity) (Thermo Fisher Scientific, CAS Number: 7647-14-5) and deionised water (dH₂O).

Growth experiments

Two growth systems were implemented: ‘closed’ growth experiments were conducted in tightly sealed 15 mL falcon tubes with sufficient headspace (14 mL) to allow aerobic metabolism but prevent ammonia evaporation; ‘open-air’ growth experiments were conducted in 96-well plates to permit ammonia evaporation and to monitor growth dynamics. Growth in closed-air system experiments were assessed by colony forming units (CFU) after 72 h incubation at 28 ºC. Growth in open-air systems were assessed in 96-well plates by optical density (OD) readings in a BMG SPECTROstar Nano Microplate Reader at 600 nm with reading taken every half an hour for 48 h at 28 ºC. Plate wells were prepared with 190 µL of a selected solution and seeded with 10 µL overnight H. meridiana culture to OD₆₀₀ = 0.05. Positive control wells were 190 µL unamended yeast media inoculated with 10 µL overnight H. meridiana culture to OD₆₀₀ = 0.05. Negative controls had no inoculation. To avoid condensation, plate lids were coated with Triton X-100 (0.05%) in 20% ethanol. Plates were shaken at 200 RPM before each reading.

Growth parameters

Experiments were conducted to yield three standard microbiological growth parameters: lag phase duration, doubling time (T_d), and final OD₆₀₀. The lag phase was calculated with the web-based Microbial Lag Phase Duration Calculator⁶⁶, with the following parameters: algorithm = parameter fitting to a model; pre-processing applied: cut data at some time = yes, max time = 24 or 48 hours; smooth data = no; initial biomass = first observation; model to fit = logistic; NLS fitting algorithm = auto; max number of iterations = 1000. T_d was calculated by first determining the growth rate in the exponential growth phase (µ) as per Eq. 4.0, followed by calculation of T_d as per Eq. 5.0.

\(\:\mu\:=\left({Log}_{10}\left(N\right)-{Log}_{10}\left({N}_{0}\right)\right)2.303/(t-{t}_{0})\)

Eq. 4.0

Where N₀ = OD₆₀₀ at the beginning of a selected time interval (t₀) in the exponential growth phase; N = OD₆₀₀ at the end of a selected time interval (t) in the exponential growth phase. t and t₀ are recorded in minutes.

\(\:{T}_{\text{d}}=ln2/\mu\:\)

Eq. 5.0

The final OD₆₀₀ reached after 48 hours was recorded and used to represent the final cell concentration reached. Cell viability versus OD₆₀₀ over 24 h was assessed by PrestoBlue^™ cell viability reagent (Thermo Fisher Scientific, Massachusetts, USA). Calibration curves confirm changes to observed OD₆₀₀ corresponds to increasing number of viable cells (Supplementary Fig. 2). Growth or lack of growth was determined by the presence or absence of a defined lag phase, exponential phase, and stationary phase within a 48 h growth period.

Ammonia evaporation and cell viability

Overnight culture of H. meridiana was inoculated to OD₆₀₀ = 0.05 in 0 (positive control), 0.01, 0.025, 0.05, 0.1, 0.25, 0.5 and 1 M ammonia solutions into separate 96-well plates. Plates were incubated at 28 ºC on an orbital shaker for 4, 8, 16 and 24 hours. Ammonia content at these time intervals was recorded by the CHEMetrics High Range VACUette Ammonia test kit (K-1510C) using direct nesslerization. The parts per million (ppm) of ammonia in the sample was determined by colorimetric analysis of the Nessler reaction product at 420 nm ⁶⁷. Viability of cells was also assessed at these times by CFU on yeast media agar plates.

Transmission electron microscopy

An overnight culture of H. meridiana was pelleted, resuspended and exposed for 2 h to the following solutions: unamended yeast media (control), 1 M ammonia and a pH-matched solution (pH 10.78). The matched pH solution was created by gradual addition of 1 M NaOH to yeast media. Following incubation, cells were washed and resuspended in phosphate buffered saline (PBS). Samples were fixed in 3% glutaraldehyde in 0.1 M Sodium Cacodylate buffer, pH 7.3, for 2 hours then washed in three 10-minute changes of 0.1 M Sodium Cacodylate. Specimens were then post-fixed in 1% Osmium Tetroxide in 0.1 M Sodium Cacodylate for 45 minutes, then washed in three 10-minute changes of 0.1 M Sodium Cacodylate buffer. These samples were then dehydrated in 50%, 70%, 90% and 100% ethanol (X3) for 15 minutes each, then in two 10-minute changes in Propylene Oxide. Samples were then embedded in TAAB 812 resin. Sections, 1 µm thick were cut on a Leica Ultracut ultramicrotome, stained with Toluidine Blue, and viewed in a light microscope to select suitable areas for investigation. Ultrathin sections, 60 nm thick were cut from selected areas, stained in Uranyl Acetate and Lead Citrate then viewed in a JEOL JEM-1400 Plus TEM. Representative images were collected on a GATAN OneView camera at 4K resolution. Images were processed using ImageJ ⁵⁷

Metabolomics sampling and extraction

An overnight culture of H. meridiana was inoculated to OD₆₀₀ = 0.05 within 0.25 M ammonia, pH-matched yeast media (pH 10.18) or unamended yeast media (control) within a 24-well plate. The pH-matched solution was created by gradual addition of 1 M NaOH to yeast media. Growth occurred within plates at 28 ºC and was assessed by OD₆₀₀ reading every half an hour using a BMG SPECTROstar Nano Microplate Reader. Samples were harvested during the log phase of growth at OD₆₀₀ = 0.5. All the following procedures occurred at 4°C. Samples were aliquoted into microcentrifuge tubes and quenched by rapid cooling through submersion in a dry ice/70% (v/v) ethanol bath after brief incubation on ice. During cooling, samples were mixed vigorously to prevent freezing. Cells were separated from spent medium by centrifugation at 1,000 g for 10 minutes and supernatant discarded. Metabolite extraction occurred by addition of ice-cold chloroform/methanol/water (1:3:1 ratio). Cell lysis was encouraged by sonication with ice for 5 minutes at 37 kHz within an ultrasonication bath (Elmasonic S 60 H). Extraction mixtures were mixed vigorously at 1,200 rpm on an orbital shaker for 1 h. Following mixing, samples were centrifuged at 13,000 g for 3 minutes. The supernatant, containing metabolites, was collected into sterile microcentrifuge tubes. A pooled quality control sample was generated by combining equal volumes of metabolites from each sample. All samples were stored at − 80°C until analysis.

Untargeted metabolomics analysis

The untargeted metabolomics analysis was performed using liquid chromatography (LC) coupled to ion mobility (IM) quadrupole time-of-flight (qTOF) mass spectrometry (MS) as described previously (Ref 1). The instrumentation consisted of an Agilent 1290 Infinity II series UHPLC system hyphenated with an Agilent 6560 IM-qTOF with a Dual Agilent Jet Stream Electron Ionization source. LC separation was performed on an InfinityLab Poroshell 120 HILIC-Z, 2.1 mm × 50 mm, 2.7 µm UHPLC column (Agilent Technologies 689775–924) coupled to an InfinityLab Poroshell 120 HILIC-Z, 3.0 mm × 2.7 µm UHPLC guard column (Agilent Technologies 823750–948). A 3.5 min gradient was run using organic buffer (acetonitrile) combined with an aqueous buffer with low pH (10 mM ammonium formate, pH 3) or high pH (10 mM ammonium acetate, pH 9) for positive and negative ionization modes, respectively. Data was acquired using MassHunter Data Acquisition 10.0 software on 1 µL of sample separated on the column with a flow rate of 800 µL/min. The quality control sample was injected five times at the beginning of the experiment to condition the column and after every five test samples to monitor the instrument state throughout data acquisition. Data were acquired in the 50 to 1700 m/z range, with an MS acquisition rate of 0.8 scans/s. The metabolomics analyses were carried out by the EdinOmics research facility (RRID: SCR_021838) at the University of Edinburgh.

Metabolomics data processing and statistical analysis

The raw data files were processed using the Agilent MassHunter software suite. Briefly, ion multiplexed data files and calibration files were demultiplexed using the PNNL PreProcessor v2020.03.23 (the default settings for demultiplexing, moving average smoothing, saturation repair and spike removal were applied to the data). The data files were recalibrated for accurate mass and drift time using the AgtTofReprocessUi and the IM-MS Browser 10.0, respectively. Molecular features were extracted in Mass Profiler 10.0 with a retention time tolerance of ± 0.3 min, drift time tolerance of ± 1.5% and accurate mass tolerance of ± (5 ppm + 2 mDa). Features were annotated based on accurate mass and collision cross section (CCS) values using McLean CCS Compendium PCDL library ⁶⁸. Multivariate and univariate statistical analysis was performed using the MetaboAnalyst 6.0 web-based platform ⁶⁹. The input data were log-transformed and Pareto-scaled. The results were visualised using principal component analysis (PCA), box and whisker plots, volcano plots and heatmaps. Further aesthetic adjustments, including font changes and line thickness modifications, were made using Inkscape (version 1.0.1). Chemical structures were drawn using ChemDraw (version 23.1.2) and exported in SVG format for inclusion in the figures. The raw data associated with this dataset is available to view in the Supplementary Dataset. This study focuses on metabolomic changes in ammonia, pH and control samples, but the broader metabolomics profiling also included samples exposed to (NH₄)₂SO₄. For analysis, only ammonia, pH and control samples were included in this study. The metabolomics of (NH₄)₂SO₄ exposed samples are to be addressed in a separate analysis.

Statistics and reproducibility

All data were compiled from a minimum of three different cultures inoculated on different days with new media (n = 3–4 biological replicates). The normality of data was assessed with the Shapiro-Wilk test. For comparison of two groups, paired or unpaired two-tailed t-tests were used. The Wilcoxon matched-pairs signed-rank test was used for paired groups where the assumption of normality was violated. An unpaired t test with Welch's correction was applied for groups with unequal variance. For comparison of two or more groups, equal variance was assessed with the Brown-Forsythe test. Samples of equal variance were analysed by analysis of variance (ANOVA) followed by Tukey’s post-hoc test. For samples where variance was not equal, Welch’s ANOVA test with Games-Howell’s post-hoc test was used. Statistical tests are specified in figure legends. Significance was considered when p < 0.05. All statistical analyses were performed using GraphPad Prism version 8.0.2 (GraphPad Software Inc.).

Data Availability

All data generated or analysed during this study are included in this published article within the Supplementary Dataset, except when specified as part of the Supplementary Material.

Acknowledgements

Funding for this research was provided by the Natural Environmental Research Council (NERC) though an E4 Doctoral Training Partnership (DTP) studentship (NE/S007407/1). C.H. and C.S.C thanks the Science and Technology Facilities Council (STFC) for support under grant ST/V000586/1 and ST/Y001788/1. We also acknowledge the support of the Wellcome Trust Multiuser Equipment Grant (WT104915MA) for use of the JEOL JEM-1400 Plus TEM. We acknowledge the EdinOmics research facility at the University of Edinburgh where the metabolomics analyses were carried out.

Author information

Cassie M. Hopton & Charles S. Cockell - School of Physics and Astronomy, University of Edinburgh, James Clerk Maxwell Building, Peter Guthrie Tait Road, Edinburgh, EH9 3FD, United Kingdom

Peter Nienow - School of Geosciences, University of Edinburgh, Drummond St, Edinburgh, EH8 9XP, United Kingdom

Author contributions

C.H. conducted the experiments, data analysis and paper writing. C.S.C and P.N. jointly supervised this work, provided instruments and materials for experiments, and provided input in the writing of the paper.

Corresponding author

Cassie M. Hopton, [email protected], ORCID ID: 0000-0003-2830-7787

Competing interests

All authors confirm that this research was carried out without any commercial or financial affiliations that could be perceived as a potential competing interest.

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Ammonia sets limit to life and alters physiology independently of pH in Halomonas meridiana

Status:

Journal Publication

Version 1

Abstract

Figures

Introduction

Results

Discussion

Materials and Methods

Solution preparation

NH₃/NH₄⁺ percentage abundance

Oxygen measurements

Water activity

Bacterial strain selection

Bacterial culture

Growth experiments

Growth parameters

Ammonia evaporation and cell viability

Transmission electron microscopy

Metabolomics sampling and extraction

Untargeted metabolomics analysis

Metabolomics data processing and statistical analysis

Statistics and reproducibility

Declarations

References

Additional Declarations

Supplementary Files

Status:

Journal Publication

Version 1

Ammonia sets limit to life and alters physiology independently of pH in Halomonas meridiana

Status:

Journal Publication

Version 1

Abstract

Figures

Introduction

Results

Discussion

Materials and Methods

Solution preparation

NH3/NH4+ percentage abundance

Oxygen measurements

Water activity

Bacterial strain selection

Bacterial culture

Growth experiments

Growth parameters

Ammonia evaporation and cell viability

Transmission electron microscopy

Metabolomics sampling and extraction

Untargeted metabolomics analysis

Metabolomics data processing and statistical analysis

Statistics and reproducibility

Declarations

References

Additional Declarations

Supplementary Files

Status:

Journal Publication

Version 1

NH₃/NH₄⁺ percentage abundance