Table of Contents

• Coverage
• Dataset Description
  • Methods & Sampling
  • Data Processing Description
  • BCO-DMO Processing Description
• Data Files
• Supplemental Files
• Related Publications
• Parameters
• Instruments
• Deployments
• Project Information
• Funding

Sample collection

For all rate experiments, water was sampled through gas-tight Viton tubing into vials or glass bottles filling from the bottom with three times overflow. Vials or bottles were closed bubble-free with deoxygenated butyl rubber stoppers and kept cool/dark until return to the laboratory. Collection vessels included 20-mL glass vials (2021) or 250-mL glass serum bottles (2022) for N₂O production experiments, 500-mL glass bottles for N₂O consumption, anaerobic ammonia oxidation, denitrification, dissimilatory nitrite reduction to ammonium, and nitrite oxidation experiments, and 250-mL glass serum bottles for dark carbon fixation experiments.

Experimental set-up

Laboratory set up for N₂O production: In 2021, a 2-mL N₂ headspace was introduced in each 20-mL vial before it underwent 2 cycles composed of 30 seconds of vigorous shaking followed by 30 seconds of flushing with N₂. Each vial was then amended with one of three ¹⁵N-labeled tracer solutions (¹⁵NH₄⁺/¹⁴NO₂^-, ¹⁵NO₂^-/¹⁴NH₄⁺, ¹⁵NO₃^-/¹⁴NH₄) to quantify rates of N₂O production from NH₄⁺ oxidation and NO₃^- and NO₂^- reduction. Each tracer solution was added to six vials for a 2 µmol·L^-1 increase in the concentration of both the ¹⁵N and ¹⁴N  substrates. In 2022, a helium headspace was added to 250-mL bottles followed immediately by purging of the water for 30 minutes with helium. Each bottle was then amended with one of the three tracer solutions and the liquid dispensed into replicate 12-mL exetainer vials for incubation. Each vial was injected with 1µM unlabeled ¹⁴N₂O to stem the loss of ¹⁵N-labeled N₂O to other processes (2022 only). In both years, vials were incubated in the dark at in situ temperature and were terminated at 0 and 12 hours (2021) or 0, 9, and 18 hours (2022) by injection of saturated mercuric chloride.

Laboratory set up for N₂O consumption: A headspace was introduced followed immediately by purging of the water for 30 minutes with helium. Water was dispensed with > 2 volumes overflow into 12-mL exetainer vials that were immediately closed with deoxygenated chlorobutyl rubber septa. Each vial received a 2-mL helium headspace before undergoing 2 cycles composed of 30 seconds of vigorous shaking followed by 30 seconds of flushing with helium.  Each vial was then injected with ¹⁵N₂O (1 µM) and incubated in the dark at in situ temperature, and were terminated at four time points, 0, 6, 12 and 24 hours, by injection of saturated mercuric chloride.

Laboratory set up for anaerobic ammonia oxidation, denitrification, dissimilatory nitrite reduction to ammonium, and nitrite oxidation: A headspace was introduced and ¹⁵NO₂ added (5 µM) followed immediately by purging of the water for 30 minutes with helium. Water was dispensed with > 2 volumes overflow into 12-mL exetainer vials that were immediately closed with deoxygenated chlorobutyl rubber septa. Each vial received a 2-mL helium headspace before undergoing 2 cycles composed of 30 seconds of vigorous shaking followed by 30 seconds of flushing with helium.  Vials were then incubated in the dark at in situ temperature, and were terminated at four time points, 0, 6, 12 and 24 hours, by injection of saturated mercuric chloride.

Laboratory set up for dark carbon fixation: ¹³C-DIC was introduced into each serum bottle and a 20-mL helium headspace inserted, before undergoing 2 cycles composed of 30 seconds of vigorous shaking followed by 30 seconds of flushing with helium.  A sample of the headspace was placed into a 3 mL glass vial, and preserved with mercuric chloride, headspace free for labeling percentage determination (averaged 20 %). Serum bottles were then incubated in the dark at in situ temperature, and duplicate bottles were terminated at two time points 0 and 24 hours by filtration onto pre-combusted glass fiber filters (GF-75). Filters were subsequently oven dried and stored at room temperature until analysis.

N2O production: In 2021, rates of 15N2O production were calculated using equations in Ji et al (2020) and the bulk d15N-N2O measured by CRDS in each experimental vial. The 15N enrichment of each substrate pool (F) was calculated from stock tracer solution concentrations, confirmed using an Astoria Nutrient Autoanalyzer following the methodology of Barwell-Clarke and Whitney 1996, and a tracer addition volume of 0.1 mL. For 2022 experiments, the production of 15N15N and 15N15N18O were measured on a coupled gas chromatograph-isotope ratio mass spectrometer (GC-IRMS) and corrected for the labelling in the initial substrate pool (Bourbonnais et al 2021, Dalsgaard et al 2012). The 15N enrichment of each substrate pool (F) was determined by the concentrations of substrate before and after tracer addition as in (Dalsgaard et al 2012).

N2O consumption: The production of 15N15N was analyzed on a gas chromatography isotope ratio mass spectrometer (Dalsgaard et al 2012).

Anaerobic ammonia oxidation and Denitrification: The production of 14N15N and 15N15N was analyzed on a GC-IRMS and corrected for the labelling in the initial substrate pool (Dalsgaard et al 2012).

Dissimilatory nitrite reduction to ammonium: After conversion of 15NH4 to N2 with alkaline hypobromite (Warembourg 1993) the production of 14N15N and 15N15N was analyzed on a GC-IRMS and corrected for the labelling in the initial substrate pool (Dalsgaard et al 2012).

Nitrite oxidation: After conversion of 15NO3 to N2 with cadmium/sulfamic acid (Bristow et al 2016), the production of 14N15N and 15N15N was analyzed on a GC-IRMS and corrected for the labelling in the initial substrate pool (Dalsgaard et al 2012).

Dark carbon fixation: After acidification, the 13C enrichment was analysed by an elemental analyzer - isotope ratio mass spectrometer.

* converted nd to blanks (no data, i.e., experiments were not run)
* converted dates and times to ISO formats

NSF Award Abstract:

Nitrous oxide (N2O) is a gas produced by microbes in both aquatic and terrestrial environments, and, like other greenhouse gases, it contributes to global warming. Furthermore, N2O can destroy ozone, a gas responsible for protecting the earth from dangerous ultraviolet radiation. In the ocean, N2O production is largely controlled by the amount of available dissolved oxygen, with more N2O being produced under low oxygen concentrations; however, when no oxygen is available, a scenario referred to as anoxia, microbes in the ocean switch from producing N2O to consuming N2O. In recent years, it has become evident that zones of low oxygen are expanding in some areas of the oceans, and this has raised concern that more N2O will be produced. If this occurs, more N2O will be emitted to the atmosphere, and will lead to further global warming and ozone destruction. Because of this, research has largely focused on understanding how much N2O is produced in the ocean under low oxygen conditions. If, however, anoxic zones also increase in size, this could act to balance out, at least to some degree, the predicted increase in N2O production caused by the expansion of zones where oxygen is present but in low concentrations. This study aims to simultaneously measure N2O production and consumption, in both low oxygen and anoxic zones and identify the microbes responsible for N2O production and consumption. Our results will: 1) lead to a much better understanding of how N2O consumption in anoxic zones could help to balance out an increase in N2O production if low oxygen zones in the ocean continue to expand, 2) help to inform models aimed at predicting oceanic N2O production and emissions to the atmosphere under future ocean conditions, and 3) allow us to better understand the microbes involved in N2O production and consumption. Our study will support a postdoc and undergraduate students who will work at the interface of marine chemistry and community genomics. The PIs plan to specifically consider applications from underrepresented minorities and students at institutions with limited opportunities. The PIs also plan a number of other educational/outreach programs ranging from teacher-training workshops, teacher internships, and academic and public lecture series.

The oceanic production of the potent greenhouse and ozone destroying gas nitrous oxide (N2O) increases as dissolved oxygen (DO) concentrations transition from oxic to hypoxic. Marine DO concentrations have decreased globally with climate change and oceanic hypoxic zones have expanded and predicted to continue expanding. This increase is cause for concern that N2O production in the ocean will increase in the future which would lead to higher emissions to the atmosphere. As a result, much research has focused on quantifying the oxygen thresholds that correspond to large increases in N2O production. In contrast, relatively few studies have aimed to quantify the capacity for net N2O consumption, resulting from microbial N2O reduction to N2 under anoxic conditions, to buffer against predicted N2O production increases if anoxic zones expand in conjunction with hypoxic zones. To this end, this study aims to simultaneously quantify N2O production and consumption from oxic-hypoxic-anoxic water column zones, in order to determine the potential for N2O consumption to counteract predicted increases in N2O production. Our field work be conducted in Saanich Inlet, a British Columbian fjord which is an ideal natural laboratory for our study, as it is characterized by a well-established oxycline and anoxic zone. Specifically, we aim to 1) measure bulk N2O concentrations, and, using 15N tracer techniques, quantify N2O production and consumption rates as DO concentrations decrease from oxic to anoxic conditions, 2) quantify the magnitude by which N2O consumption in the anoxic zone balances increased N2O production in the overlying hypoxic region, and 3) definitively link observed N2O production and consumption rates to the microorganisms mediating this process, focusing specifically on distinguishing N2O consumption via denitrifier (NO3- to N2) versus non-denitrifier (N2O to N2 only) taxa. Ultimately, our results will provide quantitative information on N2O consumption rates over fluctuating ocean conditions, thereby helping constrain models of oxygen effects on net N2O production and ocean-to-atmosphere greenhouse gas fluxes. Furthermore, this work will identify the taxonomic breadth of microbes capable of N2O reduction and their linkage to actual N2O reduction rates, thereby providing a quantitative understanding of whether or not the detection of specific bio-signatures is predictive of marine N2O dynamics.

This award reflects NSF's statutory mission and has been deemed worthy of support through evaluation using the Foundation's intellectual merit and broader impacts review criteria.