Activity-based labelling of ammonia- and alkane-oxidizing microorganisms including ammonia-oxidizing archaea

doi:10.21203/rs.3.rs-3237955/v1

Download PDF

Article

Activity-based labelling of ammonia- and alkane-oxidizing microorganisms including ammonia-oxidizing archaea

https://doi.org/10.21203/rs.3.rs-3237955/v1

This work is licensed under a CC BY 4.0 License

Version 1

posted

You are reading this latest preprint version

Recently, an activity-based labelling protocol for the in situ detection of ammonia- and alkane-oxidizing bacteria became available. This functional tagging technique enabled targeted studies of these environmentally widespread functional groups, but it failed to capture ammonia-oxidizing archaea (AOA). Since their first discovery, AOA have emerged as key players within the biogeochemical nitrogen cycle, but our knowledge regarding their distribution and abundance in natural and engineered ecosystems is mainly derived from PCR-based and metagenomic studies. Furthermore, the archaeal ammonia monooxygenase is distinctly different from its bacterial counterparts and remains poorly understood. Here, we report on the development of a universal activity-based labelling protocol for the fluorescent detection of all ammonia- and alkane-oxidizing prokaryotes, including AOA. In this protocol, 1,5-hexadiyne is used as inhibitor of ammonia and alkane oxidation and as bifunctional enzyme probe for the fluorescent labelling of cells via the Cu(I)-catalyzed alkyne-azide cycloaddition reaction. Besides efficient activity-based labelling of ammonia- and alkane-oxidizing microorganisms, this method can also be employed in combination with deconvolution microscopy for determining the subcellular localization of their ammonia- and alkane-oxidizing enzyme systems. Labelling of these enzymes in diverse ammonia- and alkane-oxidizing microorganisms allowed their visualization on the cytoplasmic membranes, the intracytoplasmic membrane stacks of ammonia- and methane-oxidizing bacteria, and, fascinatingly, on vesicle-like structures in one AOA species. The development of this universal activity-based labelling method for ammonia- and alkane-oxidizers will be a valuable addition to the expanding molecular toolbox available for research of nitrifying and alkane-oxidizing microorganisms.

Biological sciences/Microbiology/Environmental microbiology

Biological sciences/Microbiology/Cellular microbiology

The oxidation of ammonia, methane, or short-chain alkanes is an environmentally widespread process catalyzed by a diverse range of microorganisms, which besides their fundamental role in the environment also are of large interest for biotechnological applications, e.g., for the treatment of wastewater and drinking water. Aerobic ammonia oxidation, the initial and rate-limiting step of the nitrification process, plays a key role in the biogeochemical nitrogen cycle and is catalyzed by chemolithoautotrophic ammonia-oxidizing prokaryotes. For decades, the metabolic potential to perform aerobic ammonia oxidation to nitrite had been considered a trait restricted to a few bacterial genera (ammonia-oxidizing bacteria; AOB). However, the discovery that archaea affiliated with the Thaumarchaeota phylum can also grow by oxidizing ammonia has fundamentally changed this perception¹ and the presence of ammonia-oxidizing archaea (AOA) has been verified in many ecologically diverse environments^2–6.

AOA are important microbial drivers in aquatic ecosystems, where they can account for up to 40% of all prokaryotes. AOA are also major players in nitrogen flux in terrestrial habitats, particularly acidic and natural soils^3,7,8. So far, the environmental identification of AOA has been largely based on the utilization of cultivation-independent sequencing-based techniques, like PCR and metagenomics⁴. These approaches have provided invaluable data regarding the distribution and abundance of AOA in a variety of environments but cannot directly detect their ammonia-oxidizing activity. The key enzyme of ammonia oxidation is ammonia monooxygenase (AMO), which catalyzes the oxygen-dependent conversion of ammonia to hydroxylamine. AMO enzymes belong to the copper-containing membrane monooxygenase (CuMMO) family, members of which catalyze a wide array of reactions, such as ammonia, methane, and short-chain hydrocarbon oxidation, and exhibit a high degree of genetic, structural, and catalytic similarities^9–11. Also, the archaeal and bacterial AMO enzymes exhibit genetic and structural similarities that indicate a common evolutionary history⁵. However, it has been demonstrated that there are substantial dissimilarities regarding substrate range and catalytic properties¹². For example, there are distinct differences in their sensitivity to the ammonia oxidation inhibitors allylthiourea (ATU)^13–15 and terminal n-alkynes¹². Furthermore, the subunit composition of the archaeal ammonia monooxygenase is notably different from the bacterial members of the CuMMO superfamily. For instance, the AMO from the AOA Nitrososphaera viennensis contains six subunits instead of the three typically found in bacterial CuMMOs¹⁶. In addition, the archaeal AMO shares a relatively low sequence similarity (40%) to bacterial CuMMOs, and the bacterial CuMMOs, despite performing different primary functions like methane and ammonia oxidation, share a greater sequence similarity to each other than with the archaeal enzyme. AOA also lack intracellular membrane stacks that harbor the AMO enzyme in bacterial ammonia oxidizers. Furthermore, co-oxidation of a wide range of substrates other than ammonia is known in bacterial AMOs, but this topic remains underexplored in AOA¹⁷. Having an activity-based probe for the archaeal AMO would be extremely useful for addressing the knowledge gaps in the function, cellular localization, and environmental activity of this important enzyme and exploring its role in the success and adaptation of AOA in the environment.

An activity-based labelling method enabling the in situ detection of CuMMO-containing bacteria in complex microbial communities has lately been developed¹⁸. In this method, fluorescent labelling of CuMMO-containing bacteria, including the recently discovered complete ammonia-oxidizing (comammox) Nitrospira^19,20, was achieved by the use of the irreversible CuMMO inactivator 1,7-octadiyne (1,7OD) in combination with the highly specific copper-catalyzed alkyne-azide cycloaddition (CuAAC) reaction. Despite the structural similarities between the bacterial and archaeal AMOs, however, ammonia oxidation in AOA has been shown to be insensitive to long-chain alkynes, such as 1-octyne²¹ and, consequently, 1,7OD was unable to label AOA. Still, since the archaeal AMO is known to be sensitive to inactivation by shorter-chain terminal n-alkynes (< C₆)^21–23, their corresponding diynes are promising candidates for bifunctional enzyme probes to also label AOA.

In this study, we therefore set out to investigate the suitability of 1,5-Hexadiyne (1,5HD), the shortest commercially available diyne, to be employed as an enzyme inactivator for the activity-based fluorescent labelling of all ammonia, methane, and short-chain alkane oxidizers, including AOA. In addition, we use this activity-based fluorescent labelling method to investigate the subcellular localization of the respective CuMMO enzymes in phylogenetically diverse ammonia and methane oxidizers.

Cultivation

Pure cultures of Nitrosarchaeum koreense MY1 (Group 1.1a), Nitrosocosmicus franklandus (Group 1.1b), Nitrosomonas europaea (ATCC 25978) and Methylotetracoccus oryzae were grown as described before^24–27. Highly enriched cultures of Ca. Nitrosotenuis chungbukensis MY2²⁸ and Ca. Nitrospira kreftii were cultivated as described elsewhere²⁹. The pure cultures of the propane-oxidizing Rhodococcus sp. strain ZPP and the butane-degrading Thauera butanivorans strain Bu-B1211 (DSM 2080) were maintained as described in Sakoula et al¹⁸. Escherichia coli (DSMZ 498) cells were grown in LB medium (10 g L^− 1 NaCl, 5 g L^− 1 yeast extract, 10 g L^− 1 peptone), at 37 ^oC, 150 rpm. Unless stated otherwise, all cultures were harvested in the late exponential phase. AOA were harvested by filtration using Amicon Ultra-15 filters (10 kDa cutoff; Millipore, Burlington, MA, USA). The filters were equilibrated by washing twice with sterile medium prior to use. Biomass from the remaining strains was harvested by gentle centrifugation (2000 × g, 15 min), washed twice with the respective sterile medium, and resuspended in sterile medium to a final density of approximately 100 µg total cell protein ml^− 1.

Activity assays

Biomass was resuspended in the respective HEPES buffered (10 ml l^− 1 HEPES solution, consisting of 1 M HEPES and 0.6 M NaOH; pH 7.6) mineral salts medium and incubated in glass vials sealed with butyl rubber stoppers and aluminum crimp seals. Cells were incubated in the presence of 100 µΜ 1,5HD (aqueous concentration; 50% in pentane; Alfa Aesar, Haverhill, MA, USA) with either ammonium, hydroxylamine, or nitrite, and methane or methanol as substrates for ammonia- and alkane-oxidizing microorganisms, respectively. 1,5HD is reactive and unstable in the presence of oxygen and the commercial preparations contain an argon atmosphere. To avoid spontaneous breakdown, which could potentially interfere with the activity assays and labelling, the commercial stock of 1,5HD was aliquoted in an anaerobic cabinet after opening, and aliquots were stored under N₂ atmosphere until further use. Incubations with the same substrates with and without pentane addition (100 µΜ final concentration) but in the absence of 1,5HD were performed as controls. Abiotic hydroxylamine conversion was tested using sterile HEPES buffered pH 7.6 medium supplemented with 200 µΜ hydroxylamine (Figure S1). Bottles were incubated at the optimal growth temperature of each microorganism, in the dark without shaking. All incubations were performed in three biological replicates. Liquid samples (0.5 mL) were taken for the determination of ammonium, hydroxylamine, nitrite, nitrate, and methanol concentrations, as well as headspace gas samples (50 µl) for measuring methane, propane, and butane.

Analytical methods

Ammonium was measured fluorometrically using a modified orthophatal-dialdehyde assay³⁰. Nitrite and nitrate were measured colorimetrically by the Griess reaction³¹ in combination with nitrate reduction by vanadium(III)³², and hydroxylamine with 8-quinolinol³³. Methane, propane, and butane concentrations in the headspace were measured using gas chromatography (HP 5890a, equipped with a flame ionization detector and a Porapak Q column at 80°C). Methanol was measured colorimetrically according to Mangos and Haas³⁴. For the determination of protein content, cells were lysed with the Bacterial Protein Extraction Reagent (B-PER; Thermo Fisher Scientific, Waltham, MA, USA) according to manufacturer’s instructions in combination with mild sonication (1 minute, 30 Hz). Protein concentrations were measured using the Pierce bicinchoninic acid protein assay kit (Thermo Fisher Scientific, Waltham, MA, USA) according to the ‘Enhanced Test Tube’ protocol. For all fluorometric and colorimetric measurements a Spark M10 plate reader (Tecan Trading AG, Männedorf, Switzerland) was used.

In situ activity-based labelling

Active biomass was harvested as described above, washed twice, and resuspended in 50 ml medium without substrates. Subsequently, the biomass was incubated with 100 µM 1,5HD for 30 minutes in the dark without shaking, at the optimal temperature of each culture. Following inactivation, cells were pelleted by centrifugation (2000 × g, 10 minutes), washed twice in sterile PBS, pH 7.4, and fixed using a 50% (v/v) ethanol in PBS solution for 10 min at ambient temperature (RT). Subsequently, the CuAAC reaction was performed in plastic microcentrifuge tubes (1.5 mL) in a final volume of 250 µL to label 1,5HD-inhibited cells. For this, fixed biomass was washed with PBS, resuspended in 221 µl PBS and mixed with 12.5 µl of a freshly prepared 100 mΜ sodium ascorbate solution (99% purity, Merck KGaA, Darmstadt, Germany) and 12.5 µl of a freshly prepared 100 mM aminoguanidine hydrochloride solution (98% purity, Merck KGaA, Darmstadt, Germany). A dye mixture composed of 1.25 µl of a 20 mM CuSO₄ solution (99.99% purity, Merck KGaA, Darmstadt, Germany), 1.25 µl of 100 mM Tris(3-hydroxypropyltriazolylmethyl)amine in ddH₂O (THPTA; 95% purity, Merck KGaA, Darmstadt, Germany), and 0.3 µl of 5 mM Azide-Fluor 488 in DMSO (≥ 99% purity, Merck KGaA, Darmstadt, Germany) was prepared and left to react in the dark for 3 minutes. Subsequently, 2.8 µl dye mixture were added to the cell suspensions. The tubes were gently mixed and incubated for 60 min (RT, in the dark). CuAAC reactions were terminated by harvesting the cells by centrifugation (2000 × g, 10 minutes). Cell pellets were washed three times with PBS, resuspended in 1:1 (v/v) ethanol:PBS and stored at -20°C.

Fluorescence and deconvolution microscopy

Prior to microscopic analysis, 10 µl activity-based labelled samples were dried onto microscope slides, dehydrated using an increasing ethanol series (50, 80, and 100%; 3 min each) and stained with 0.1 µg/ml DAPI in PBS (4′,6-diamidino-2-phenylindole dihydrochloride; Merck KGaA, Darmstadt, Germany) for 15 minutes. Subsequently, the slides were washed in MQ water, air-dried and embedded in Vectashield mounting solution (Vector Laboratories Inc., Burlingame, CA, USA). CuAAC (activity-based) and DAPI-conferred fluorescent signals were recorded using the HyD hybrid detectors of a Leica TCS Sp8x confocal laser microscope (CLSM; Leica Microsystems B.V., Amsterdam, the Netherlands) equipped with a 405 nm UV diode and a pulsed white-light laser. Images were recorded using a 100× oil immersion objective at a resolution of 1024 × 1024 pixels and 8-bit depth.

For detecting the subcellular localization of the AMO- and particulate methane monooxygenase (pMMO)-derived fluorescent signal, the HyVolution deconvolution module of the Huygens Essential Suite (Scientific Volume Imaging B.V., Hilversum, The Netherlands) was used. Fluorescent images were acquired using the HyD hybrid detectors of a Leica Sp8x CLSM, a 100× oil immersion objective, a 0.5 AU pinhole size, at 1024 × 1024 pixels resolution. Deconvolution was performed using the resolution-optimized algorithm of the HyVolution module.

Inhibition of the archaeal ammonia oxidation by 1,5HD

AOA are strongly inhibited by low concentrations (10 µM) of short-chain alkynes (≤ C₅), but resistant to inhibition by longer-chain-length alkynes (≥ C₆), even at concentrations that can completely inhibit ammonia oxidation activity in AOB^12,22. Due to these compounds’ contrasting inhibition profiles of AOA and AOB, long-chain alkynes have been used in many ecological studies as differential inhibitors to distinguish bacterial and archaeal contributions to ammonia oxidation in soil^21,35,36.

In this study, we investigated whether the diyne counterparts to these alkynes displayed the same differential inhibition pattern in AOA and AOB. An earlier study¹⁸ found that 1,7OD effectively inhibited ammonia and alkane oxidation in all tested bacterial strains, but was inefficient in binding to the archaeal AMO, which is consistent with previous reports that AOA were resistant to 1-octyne²¹. As short-chain alkynes are effective inhibitors of ammonia oxidation in AOA, we hypothesized that short-chain diynes may be better suited for activity-based probing in AOA. 1,5HD is the shortest chain-length diyne commercially available and was therefore chosen for characterization on a pure culture of N. franklandus. Previous studies found 1-hexyne to be less inhibitory to AOA than < C₅ alkynes, but surprisingly, ammonia consumption by the AOA N. franklandus was completely inhibited by the addition of 100 µΜ 1,5HD, while 1 mM ammonium was stoichiometrically oxidized to nitrite in the absence of the inhibitor (Fig. 1A, B). No effect of 1,5HD on hydroxylamine oxidation was observed (Fig. 1C, D), confirming the specific interaction of 1,5HD with the AMO enzyme. These results indicated that 1,5HD, at least at 100 µΜ concentration, is able to efficiently inhibit ammonia oxidation in archaea, and thus could potentially be used for activity-based labelling of AOA.

Inhibition of bacterial ammonia and alkane oxidation by 1,5HD

To investigate the effect of 1,5HD on bacterial ammonia oxidation, we performed inhibition assays with a pure culture of N. europaea. Similar to N. franklandus, ammonia oxidation in N. europaea was completely suppressed by 100 µΜ 1,5HD (Fig. 2A, B). Again, hydroxylamine oxidation was not affected (Fig. 2C), demonstrating that 1,5HD specifically inhibits the AMO also in canonical AOB.

Comammox Nitrospira possess a phylogenetically distinct AMO^9,10,29 and thus might react differently to ammonia oxidation inhibitors. To test the effect of 1,5HD on comammox bacteria, a highly enriched culture of Ca. N. kreftii was used. In the absence of the inhibitor, the culture stoichiometrically oxidized 120 µM ammonium to nitrate, whereas addition of 100 µΜ 1,5HD resulted in complete inhibition of ammonia oxidation (Fig. 3A, B). While the influence on hydroxylamine oxidation was not tested in this enrichment culture, no influence of 1,5HD on nitrite oxidation was observed (Fig. 3C, D), again showcasing the specific reaction of 1,5HD with the AMO.

Besides the ammonia monooxygenases, the CuMMO family includes enzymes that catalyze diverse reactions like the oxidation of alkanes such as methane and other short-chain (C₂–C₄) hydrocarbons, but still exhibit a high degree of structural similarity^9–11. Thus, the ability of 1,5HD to inhibit methane-oxidizing bacteria was tested. Indeed, methane oxidation by the type Ib methanotroph M. oryzae was fully inhibited in the presence of 100 µΜ 1,5HD, while no effect on subsequent methanol oxidation was observed (Fig. 4). Thus, together with the successful labelling of additional ammonia- and alkane-oxidizing microorganisms (see below), our results indicate that 1,5HD can be employed as a universal inhibitor of CuMMO-containing microorganisms.

In situ activity-based fluorescent labelling of ammonia- and alkane-oxidizers

The diyne 1,7OD irreversibly inactivates bacterial CuMMOs via a suicide inactivation mechanism²³ and has successfully been employed as a bifunctional enzyme probe that, in combination with a subsequent CuAAC reaction, allows the activity-based fluorescent staining of ammonia- and alkane-oxidizing bacteria¹⁸. However, AOA are only partly inhibited by octyne (≤ 40 µM) and this inhibition is fully reversible, indicating a different, non-covalent interaction with the archaeal AMO¹². Consequently, activity-based staining of AOA cells was not possible using 1,7OD¹⁸.

In this study, we were able to demonstrate that the short-chain diyne 1,5HD efficiently inhibited ammonia oxidation in the AOA N. franklandus. Thus, its ability to be used for in situ activity-based labelling of AOA was further investigated. For this, active cultures of N. koreense, N. franklandus and Ca. N. chungbukensis were used. Following incubation with 1,5HD, ethanol fixation, and CuAAC reactions, all cultures were efficiently fluorescently stained (Fig. 5), indicating that 1,5HD can function as a bifunctional enzyme probe for the fluorescent labelling of phylogenetically diverse AOA. As the inhibition by 1,5HD was AMO-specific and did not interfere with hydroxylamine oxidation (Fig. 1), the observed fluorescence can be attributed to specific binding of 1,5HD to the AMO enzyme, rather than alternative targets in the cell. Additionally, the fact that its addition is required for fluorescence suggests the binding of 1,5HD to be covalent and irreversible. This makes 1,5HD only the third known irreversible inhibitor of the archaeal AMO besides acetylene and phenylacetylene²². Being able to fluorescently label active AOA will be a useful tool in environmental studies, as it will enable the identification of active members of mixed microbial communities based on monooxygenase enzyme activity.

The successful inhibition of ammonia- and alkane-oxidizing bacteria by 1,5HD (Figs. 2–4) also confirmed interaction of 1,5HD with the bacterial CuMMO enzymes. Consequently, when 1,5HD-treated biomass was subjected to the CuAAC protocol, strong fluorescent labelling was achieved for ammonia-oxidizing N. europaea, methane-oxidizing M. oryzae, propane-oxidizing Rhodococcus sp. ZPP and butane-oxidizing Thauera butanivorans cells (Fig. 6). In contrast, no background or unspecific labelling was observed when E. coli cells were subjected to the same protocol (Figure S2). This efficient and specific labelling indicated that 1,5HD can be used as a universal probe for the function-based detection of CuMMO-containing microorganisms. For instance, coupling activity-based probing with 1,5HD to fluorescence-activated cell sorting and sequencing-based approaches could yield novel insights into the ecology and metabolism of ammonia- and alkane-oxidizing microorganisms.

Localization of the AMO enzyme in AOA

Like its bacterial homolog, the archaeal AMO enzyme is believed to reside on and be strongly associated with the cytoplasmic membrane⁵ and fluorescent signals derived from AMO labelling are thus expected to colocalize with the cytoplasmic membrane. This assumption is supported by all characterized members of the CuMMO superfamily being membrane-bound¹⁰, by the archaeal AMOs containing alpha helices predicted to span the membrane¹⁶, by the native archaeal AMO complex being recovered from the membrane fraction of N. viennensis¹⁶, and by the fact that AOA lack the membrane stacks found in AOB. However, until now the archaeal AMO had not been visualized in its native environment. To visualize the subcellular AMO localization, three AOA cultures were subjected to the activity-based labelling protocol in combination with DAPI counter-staining. As expected, the localization of the AMO-derived signal was observed to coincide with the cytoplasmic membrane in high-resolution deconvolution micrographs in all cells (Fig. 7). Surprisingly, in addition to the plasma membrane in the periphery of the cells, some fluorescent labelling was evident inside the cells of N. franklandus (Fig. 7A). This distribution of fluorescent signal in N. franklandus was more akin with bacterial ammonia oxidizers (Fig. 8) than with the other AOA tested in this study. While most AOA do not contain intracellular compartments or structures, N. franklandus has been described to have some intracellular compartmentalization, including vesicle-like structures of unknown function³⁷. While fluorescence microscopy cannot conclusively proof that these vesicle-like structures harbor AMO, it is tempting to speculate based on the signal distribution we observed that they serve a similar function in increasing available membrane space available for ammonia oxidation as has been proposed for AOB. This should be further explored in future studies using, e.g., electron microscopy approaches.

Furthermore, high-resolution deconvolution microscopy of the AOB N. europaea and the type Ib methanotroph M. oryzae revealed that the respective CuMMO-derived signals were localized along the intracytoplasmic membrane stacks and cytoplasmic membranes present in these organisms (Fig. 8), in agreement with previous studies showing the cytoplasmic membrane-associated localization of the AMO and pMMO enzymes^38,39. Thus, in addition to its usability for the targeted detection of ammonia- and alkane-oxidizing microorganisms in complex environmental samples, the activity-based CuMMO labelling method presented here also has great potential to study their cellular organization using high-resolution microscopy.

The results presented here demonstrate that the diyne 1,5HD can be used to efficiently inhibit the activity of ammonia- and alkane-oxidizing microorganisms. Furthermore, we demonstrate that 1,5HD can be employed as a universal bifunctional enzyme probe for the activity-based fluorescent labelling of CuMMO-containing cells. The development of this universal labelling method for CuMMO-containing bacteria and archaea is an important advance as the existing 1,7OD-based method to label these organisms excluded AOA, which are very abundant ammonia-oxidizers in different environments and crucial for nitrogen cycling. This imposed a significant limitation when studying active AOA populations. When combined with 16S rRNA-targeted fluorescence in situ hybridization (FISH), the activity-based labelling can be used for the phylogenetic identification of catalytically active ammonia- and alkane-oxidizing microorganisms present in complex microbial communities. In addition, this method is a promising tool for characterization of the cellular localization of the archaeal AMO, which to date remains elusive. In conclusion, we are convinced that the development of this universal activity-based labelling method for ammonia- and alkane-oxidizers is a valuable addition to the molecular toolbox available to study these ubiquitous organisms with great environmental and biotechnological significance.

Acknowledgements

The authors would like to thank M.Y. Jung and M. Ghashgavi for providing biomass. D.S. acknowledges the Federation of European Microbiological Societies for a FEMS Travel and Research grant. Funding was provided by the European Research Council (ERC Advanced Grant Ecomom 339880 and Synergy Grant MARIX 854088) and the Netherlands Organization for Scientific Research (NWO; grants 016.Veni.192.062, 016.Vidi.189.050, SIAM 024.002.002). L.L.M was supported by a Royal Society Dorothy Hodgkin Research Fellowship (DH150187) and an ERC Starting Grant (UNITY 852993). A.S. was supported by a Royal Society Fellowship Enhancement Award (RGF\EA\180300).

Author Contributions

D.S., M.A.H.J.v.K., and S.L. conceived research. D.S., L.L.M., M.A.H.J.v.K, and S.L. planned the project. D.S., A.S., and P.B. executed experiments. D.S. analyzed data. D.S., L.L.M., M.A.H.J.v.K., and S.L. wrote the manuscript. All authors discussed results, and commented and agreed on the final manuscript.

Competing interests

The authors declare no competing interests.

Data availability

The datasets generated and/or analyzed during the current study are available from the corresponding authors on reasonable request.

Könneke, M. et al. Isolation of an autotrophic ammonia-oxidizing marine archaeon. Nature 437, 543–546 (2005).
Herbold, C. W. et al. Ammonia-oxidising archaea living at low pH: Insights from comparative genomics. Environ. Microbiol. 19, 4939–4952 (2017).
Lehtovirta-Morley, L. E., Stoecker, K., Vilcinskas, A., Prosser, J. I. & Nicol, G. W. Cultivation of an obligate acidophilic ammonia oxidizer from a nitrifying acid soil. Proc. Natl. Acad. Sci. 108, 15892 (2011).
Prosser, J. I. & Nicol, G. W. Relative contributions of archaea and bacteria to aerobic ammonia oxidation in the environment. Environ. Microbiol. 10, 2931–2941 (2008).
Schleper, C. & Nicol, G. W. Ammonia-oxidising archaea--physiology, ecology and evolution. Adv. Microb. Physiol. 57, 1–41 (2010).
Daebeler, A. et al. Cultivation and Genomic Analysis of “Candidatus Nitrosocaldus islandicus,” an Obligately Thermophilic, Ammonia-Oxidizing Thaumarchaeon from a Hot Spring Biofilm in Graendalur Valley, Iceland. Front. Microbiol. 9, (2018).
Karner, M. B., DeLong, E. F. & Karl, D. M. Archaeal dominance in the mesopelagic zone of the Pacific Ocean. Nature 409, 507–510 (2001).
Verhamme, D. T., Prosser, J. I. & Nicol, G. W. Ammonia concentration determines differential growth of ammonia-oxidising archaea and bacteria in soil microcosms. ISME J. 5, 1067–1071 (2011).
Holmes, A. J., Costello, A., Lidstrom, M. E. & Murrell, J. C. Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132, 203–208 (1995).
Khadka, R. et al. Evolutionary history of copper membrane monooxygenases. Front. Microbiol. 9, (2018).
Rochman, F. F. et al. Novel copper-containing membrane monooxygenases (CuMMOs) encoded by alkane-utilizing Betaproteobacteria. ISME J. 14, 714–726 (2020).
Taylor, A. E. et al. Inhibitory effects of C2 to C10 1-alkynes on ammonia oxidation in two Nitrososphaera species. Appl. Environ. Microbiol. 81, 1942–1948 (2015).
Hatzenpichler, R. et al. A moderately thermophilic ammonia-oxidizing crenarchaeote from a hot spring. Proc. Natl. Acad. Sci. U. S. A. 105, 2134–2139 (2008).
Lehtovirta-Morley, L. E., Verhamme, D. T., Nicol, G. W. & Prosser, J. I. Effect of nitrification inhibitors on the growth and activity of Nitrosotalea devanaterra in culture and soil. Soil Biol. Biochem. 62, 129–133 (2013).
Shen, T., Stieglmeier, M., Dai, J., Urich, T. & Schleper, C. Responses of the terrestrial ammonia-oxidizing archaeon Ca. Nitrososphaera viennensis and the ammonia-oxidizing bacterium Nitrosospira multiformis to nitrification inhibitors. FEMS Microbiol. Lett. 344, 121–129 (2013).
Hodgskiss, L. H. et al. Unexpected complexity of the ammonia monooxygenase in archaea. ISME J. 17, 588–599 (2023).
Rasche, M. E., Hyman, M. R. & Arp, D. J. Factors Limiting Aliphatic Chlorocarbon Degradation by Nitrosomonas europaea: Cometabolic Inactivation of Ammonia Monooxygenase and Substrate Specificity. Appl. Environ. Microbiol. 57, 2986–2994 (1991).
Sakoula, D. et al. Universal activity-based labeling method for ammonia- and alkane-oxidizing bacteria. ISME J. (2021) doi:10.1038/s41396-021-01144-0.
Daims, H. et al. Complete nitrification by Nitrospira bacteria. Nature 528, 504–509 (2015).
van Kessel, M. A. H. J. et al. Complete nitrification by a single microorganism. Nature 528, 555–559 (2015).
Taylor, A. E. et al. Use of aliphatic n-alkynes to discriminate soil nitrification activities of ammonia-oxidizing thaumarchaea and bacteria. Appl. Environ. Microbiol. 79, 6544–6551 (2013).
Wright, C. L., Schatteman, A., Crombie, A. T., Murrell, J. C. & Lehtovirta-Morley, L. E. Inhibition of Ammonia Monooxygenase from Ammonia-Oxidizing Archaea by Linear and Aromatic Alkynes. Appl. Environ. Microbiol. 86, (2020).
Bennett, K., Sadler, N. C., Wright, A. T., Yeager, C. & Hyman, M. R. Activity-Based Protein Profiling of Ammonia Monooxygenase in Nitrosomonas europaea. Appl. Environ. Microbiol. 82, 2270–2279 (2016).
Jung, M.-Y. et al. A mesophilic, autotrophic, ammonia-oxidizing archaeon of thaumarchaeal group I.1a cultivated from a deep oligotrophic soil horizon. Appl. Environ. Microbiol. 80, 3645–3655 (2014).
Lehtovirta-Morley, L. E. et al. Isolation of ‘Candidatus Nitrosocosmicus franklandus’, a novel ureolytic soil archaeal ammonia oxidiser with tolerance to high ammonia concentration. FEMS Microbiol. Ecol. 92, fiw057 (2016).
Ghashghavi, M. et al. Methylotetracoccus oryzae Strain C50C1 Is a Novel Type Ib Gammaproteobacterial Methanotroph Adapted to Freshwater Environments. mSphere 4, e00631-18 (2019).
Koops, H. P., Bötther, B., Möller, U. C., Pommerening-Röser, A. & Stehr, G. Classification of eight new species of ammonia-oxidizing bacteria: Nitrosomonas communis sp. nov., Nitrosomonas ureae sp. nov., Nitrosomonas aestuarii sp. nov., Nitrosomonas marina sp. nov., Nitrosomonas nitrosa sp. nov., Nitrosomonas eutropha sp. nov., Nitrosomonas oligotropha sp. nov. and Nitrosomonas halophila sp. nov. Microbiology, vol. 137 1689–1699 (1991).
Jung, M.-Y. et al. Enrichment and characterization of an autotrophic ammonia-oxidizing archaeon of mesophilic crenarchaeal group I.1a from an agricultural soil. Appl. Environ. Microbiol. 77, 8635–8647 (2011).
Sakoula, D. et al. Enrichment and physiological characterization of a novel comammox Nitrospira indicates ammonium inhibition of complete nitrification. ISME J. (2020) doi:10.1038/s41396-020-00827-4.
Taylor, S., Ninjoor, V., Dowd, D. M. & Tappel, A. L. Cathepsin B2 measurement by sensitive fluorometric ammonia analysis. Anal. Biochem. 60, 153–162 (1974).
Griess, P. Bemerkungen zu der Abhandlung der HH. Weselky und Benedikt ueber einege Azoverbindungen. Berichte Dtsch. Chem. Ges. 12, 426–428 (1879).
Miranda, K. M., Espey, M. G. & Wink, D. A. A rapid, simple spectrophotometric method for simultaneous detection of nitrate and nitrite. Nitric Oxide Biol. Chem. 5, 62–71 (2001).
Frear, D. S. & Burrell, R. C. Spectrophotometric Method for Determining Hydroxylamine Reductase Activity in Higher Plants. Anal. Chem. 27, 1664–1665 (1955).
Mangos, T. J. & Haas, M. J. Enzymatic Determination of Methanol with Alcohol Oxidase, Peroxidase, and the Chromogen 2,2‘-Azinobis(3-ethylbenzthiazoline-6-sulfonic acid) and Its Application to the Determination of the Methyl Ester Content of Pectins. J. Agric. Food Chem. 44, 2977–2981 (1996).
Ouyang, Y., Norton, J. M., Stark, J. M., Reeve, J. R. & Habteselassie, M. Y. Ammonia-oxidizing bacteria are more responsive than archaea to nitrogen source in an agricultural soil. Soil Biol. Biochem. 96, 4–15 (2016).
Lu, X., Bottomley, P. J. & Myrold, D. D. Contributions of ammonia-oxidizing archaea and bacteria to nitrification in Oregon forest soils. Soil Biol. Biochem. 85, 54–62 (2015).
Klein, T. et al. Cultivation of ammonia-oxidising archaea on solid medium. FEMS Microbiol. Lett. 369, fnac029 (2022).
Kitmitto, A., Myronova, N., Basu, P. & Dalton, H. Characterization and Structural Analysis of an Active Particulate Methane Monooxygenase Trimer from Methylococcus capsulatus (Bath). Biochemistry 44, 10954–10965 (2005).
Fiencke, C. & Bock, E. Immunocytochemical localization of membrane-bound ammonia monooxygenase in cells of ammonia oxidizing bacteria. Arch. Microbiol. 185, 99–106 (2006).

There is no conflict of interest

03SakoulaetalSuplementarymaterial.pdf
Supplementary Material

Download PDF

Version 1

posted

You are reading this latest preprint version

Activity-based labelling of ammonia- and alkane-oxidizing microorganisms including ammonia-oxidizing archaea

Status:

Version 1

Abstract

Figures

Introduction

Materials and Methods

Cultivation

Activity assays

Analytical methods

Fluorescence and deconvolution microscopy

Results and Discussion

Inhibition of the archaeal ammonia oxidation by 1,5HD

Inhibition of bacterial ammonia and alkane oxidation by 1,5HD

Localization of the AMO enzyme in AOA

Concluding remarks

Declarations

References

Additional Declarations

Supplementary Files

Status:

Version 1