AEM Accepts, published online ahead of print on 8 February 2013 Appl. Environ. Microbiol. doi:10.1128/AEM.03705-12 Copyright © 2013, American Society for Microbiology. All Rights Reserved. 1 2 3 Denitrifying Alphaproteobacteria from the Arabian Sea that Express the Gene (nosZ) 4 Encoding Nitrous Oxide Reductase in Oxic and Sub-Oxic Waters 5 D o w 6 Michael Wyman#, Sylvia Hodgson, and Clare Bird1 n lo 7 ad e d 8 Biological and Environmental Sciences f r o m 9 School of Natural Sciences h t 10 University of Stirling tp : / / a 11 Stirling FK9 4LA e m 12 United Kingdom. .a s m 13 . o r g 14 Running title: Denitrifying alphaproteobacteria from the Arabian Sea / o n 15 D e c 16 Subject Category: Microbial ecology e m 17 b e r 18 # Corresponding author 20 , 2 19 0 1 8 20 1 Present address: School of Geosciences, University of Edinburgh, Grant Institute, b y g 21 The King's Buildings, West Mains Road, Edinburgh EH9 3JW u e s t 22 Abstract 23 Marine ecosystems are significant sources of the powerful greenhouse gas, nitrous 24 oxide (N O). A by-product of nitrification and an intermediate in the denitrification 2 25 pathway, N O is formed primarily in oxygen-deficient waters and sediments. We 2 26 describe the isolation of a group of alphaproteobacteria from the suboxic waters of the D o w 27 Arabian Sea that are phylogenetically affiliated with Labrenzia spp. and other n lo a 28 denitrifiers. Quantitative PCR assays revealed that these organisms were very broadly d e d 29 distributed in this semi-enclosed ocean basin. Their biogeographical range extended f r o m 30 from the productive, upwelling region off the Omani shelf to the clear, oligotrophic h t 31 waters that are found much further south and also included the mesotrophic waters tp : / / a 32 overlying the oxygen minimum zone (OMZ) in the north-eastern sector of the Arabian e m 33 Sea. These organisms actively expressed NosZ (N2O reductase, the terminal step in .a s m 34 the denitrification pathway) within the OMZ, an established region of pelagic . o r g 35 denitrification. They were found in greatest numbers outside of the OMZ, however, / o n 36 and nosZ mRNAs were also readily detected near the base of the upper mixed layer in D e c 37 nutrient-poor, oxic regions. Our findings provide firm molecular evidence of a e m 38 potential sink for N O within well-ventilated, oceanic surface waters in this b 2 e r 2 39 biogeochemically important region. We show that the Labrenzia-like denitrifiers and 0 , 2 40 their close relatives are habitual colonizers of the pseudobenthic environment 0 1 8 41 provided by Trichodesmium spp. We develop the conjecture that the O2-depleted b y g 42 microzones that occur within the colonies of these filamentous, diazotrophic, u e s 43 cyanobacteria might provide unexpected niches for the reduction of nitrogen oxides in t 44 tropical and sub-tropical surface waters. 45 46 47 Introduction 48 49 Emissions of the greenhouse gas, nitrous oxide (N O), have increased steadily since 2 50 the early part of the nineteenth century. Atmospheric N O concentrations are higher 2 51 today than at any time during the past 650,000 years (1, 2). Apart from its significant D o w 52 warming potential (~300-fold that of CO2 over a 100 year period), rising N2O is of n lo a 53 further environmental concern because it is presently the single-most destructive d e d 54 source of emissions contributing to stratospheric ozone depletion (3). The growing f r o m 55 inventory of atmospheric N O has occurred primarily as a result of an increase in 2 h t 56 emissions from the terrestrial environment owing to changes in agricultural practices, tp : / / a 57 the combustion of fossil fuels, and other anthropogenically-driven perturbations of the e m 58 nitrogen cycle (2). The marine environment is also an important net source of N2O to .a s m 59 the atmosphere, however, and the unperturbed (non-anthropogenic) rates of emissions . o r g 60 from coastal margins, shelf, and open waters are of the same order as those from land / o n 61 (1, 4, 5). D e c 62 e m 63 Significant feedbacks on marine N O emissions are anticipated over the coming b 2 e r 2 64 decades as a result of increasing ocean acidification (5) and the expansion of hypoxic 0 , 2 65 waters owing to surface warming and an acceleration in the prevailing rates of 0 1 8 66 eutrophication from anthropogenic nutrient loading (1, 6). In oxic waters, N2O is b y g 67 produced primarily as a by-product of nitrification, the oxidation of ammonium u e 68 (NH4+) to nitrite (NO2-) and nitrate (NO3-) carried out by ammonium-oxidizing st 69 Bacteria (AOB) and Archaea (AOA). Recent work has shown that ammonium 70 oxidation rates (and hence N O production by AOB/AOA) are exquisitely sensitive to 2 71 comparatively modest (~0.1 units) declines in pH (5) and show a mean reduction of 3 72 ~20 % in response to near future (20–30 years hence) ocean acidification. The pH of 73 oceanic surface waters is set to decline by 0.3 – 0.4 units by the end of the present 74 century (7) with the potential corollary of large negative feedbacks on marine 75 nitrification rates and N O emissions in the coming decades (5). 2 76 D o w 77 The production of N2O by nitrifiers is oxygen-sensitive, however, and its emissions n lo a 78 increase very substantially under hypoxic conditions (8). Over the past fifty years the d e d 79 areal extent of hypoxic waters in coastal regions and at intermediate depths in the f r o m 80 North Pacific and tropical oceans has expanded and shoaled significantly (9, 10): a h t 81 trend that will only intensify as the global ocean continues to warm and lose oxygen tp : / / a 82 over the next few decades and beyond (11). Approximately ten percent of the e m 83 contemporary ocean is either hypoxic (O2 < 30 % saturation) or suboxic (O2 < 1 % .a s m 84 saturation) and even a modest expansion in the present volume of deoxygenated . o r g 85 waters is likely to increase N2O production by nitrifiers significantly (1, 6). The net / o n 86 impacts of increasing atmospheric CO2 on marine N2O emissions in the years to D e c 87 come, therefore, will depend on the overall balance of both positive and negative e m 88 feedbacks on ammonium oxidation rates and other climate sensitive sources of N O. b 2 e r 2 89 0 , 2 90 The predominant source of N2O emissions under suboxic conditions is not from 0 1 8 91 ammonium oxidisers but from a taxonomically diverse group of mainly heterotrophic b y 92 microbes known as denitrifiers (6). Denitrifiers use NO - as an alternative respiratory g 3 u e s 93 electron acceptor to oxidise an electron donor (usually organic carbon) to generate t 94 metabolic energy during anaerobic growth. Denitrification leads ultimately to the 95 liberation of dinitrogen gas (N ) in a four step reductive pathway from NO - that 2 3 96 proceeds via NO - and two obligate gaseous intermediates, nitric oxide (NO) and N O. 2 2 4 97 Not all denitrifiers are capable of the final reduction of N O to N , however, while for 2 2 98 others a sustained lag occurs during the induction of the necessary cellular machinery 99 to carry out this terminal step in the process (12). Some denitrifiers produce N only 2 100 in the complete absence of free oxygen but, nonetheless, are capable of partial 101 denitrification and liberate N O under hypoxic conditions (13). Denitrification is both D 2 o w 102 a sink and a potential source of emissions, therefore, and another key process in the n lo a 103 marine nitrogen cycle known to be sensitive to the future expansion in the volume of d e d 104 deoxygenated waters in the ocean (1, 6). f r o m 105 h t 106 The only known metabolic pathway for the consumption of N2O is that which requires tp : / / a 107 the copper containing enzyme, nitrous oxide reductase (NosZ). NosZ is found in all e m 108 denitrifiers that are capable of reducing NO3- as far as N2 and also in a few non- .a s m 109 denitrifying bacteria, such as Wolinella (Vibrio) succinogenes that can use N2O as a .o r g 110 terminal electron acceptor (14). These latter organisms are not themselves N2O / o n 111 producers, however, because they lack the NO-producing nitrite reductase (NirK or D e c 112 NirS) that is the definitive biochemical feature of denitrifiers (15), the primary sink e m 113 for this nitrogen oxide in the oceans (16). b e r 2 114 0 , 2 115 Uncertainties over the future scale and climatic impact of marine N2O emissions and, 0 1 8 116 in particular, their sensitivities to acidification and deoxygenation (17), has led to calls b y g 117 for a better understanding of the biological sources and sinks of this trace gas in the u e s 118 oceans (1, 18). Key to constraining the marine N2O budget is the development of a t 119 robust model of the distributions and activities of the organisms that produce and/or 120 consume N O and their responses to stress induced by global environmental change. 2 121 Approximately half of the annual emissions of N O in the contemporary ocean come 2 5 122 from the three major oxygen minimum zones (OMZs) that are located in the eastern 123 tropical North Pacific (ETNP), the eastern tropical South Pacific (ETSP) and the 124 Arabian Sea (19, 20). The net contributions of different biological sources of N O in 2 125 these regions are debated (see 18) but there is general agreement that the OMZs are 126 “hotspots” within the ocean that are especially vulnerable to warming and D o w 127 deoxygenation over the coming decades (1, 17). n lo a 128 d e d 129 In the present study we describe the isolation into culture of a novel group of pelagic f r o m 130 denitrifying, alphaproteobacteria from the suboxic waters of the Arabian Sea, one of h t 131 the most intense regions of N2O production in the world ocean (20). Employing a tp : / / a 132 quantitative PCR protocol targeting the nosZ gene from these organisms, we show e m 133 that they have a broad biogeographical distribution in this ocean basin, ranging from .a s m 134 the upwelling region along the Omani shelf to the highly oligotrophic equatorial . o r g 135 waters to the south. We further show that these organisms were expressing the / o n 136 cognate mRNA for nosZ in the suboxic intermediate waters of the OMZ located in the D e c 137 northeastern sector of the Arabian Sea and, quite unexpectedly, were also active at e m 138 shallower depths within the upper mixed layer of waters well outside of the region of b e r 2 139 the OMZ. 0 , 2 140 0 1 8 141 b y g 142 Materials and Methods u e s 143 t 144 Study Site, Sample Collection and Nucleic Acid Purification 145 6 146 Observations were made in September 2001 aboard RRS Charles Darwin during the 147 NERC-funded AMBITION cruise (CD132) in the Arabian Sea. Eleven stations were 148 occupied along the length of a 5150 km transect between Victoria, Seychelles and 149 Muscat, Oman (21). Hydrographical data were collected at each station with a Sea- 150 Bird 911plus CTD profiler (Sea-Bird Electronics, Inc., Bellevue WA, USA) D o w 151 configured with a Chelsea Aquatracker III fluorometer (Chelsea Instruments, West n lo a 152 Molesey UK) to measure chlorophyll fluorescence and auxiliary sensors for dissolved d e d 153 oxygen and PAR (photosynthetically active radiation). f r o m 154 h t 155 Seawater samples were collected from discrete depths at selected stations (Figure 1) tp : / / a 156 with a rosette of twenty-four, 30-litre volume Niskin bottles mounted on the CTD e m 157 profiler. Plankton samples were obtained by filtering 4 – 5 litres of seawater .a s m 158 (unscreened) through 90 mm diameter, 0.2 μm pore size polycarbonate filters . o r g 159 (Osmonics Inc., Minnetonka MN USA) at a negative pressure of < 20 mm Hg. Cell / o n 160 material collected on the filters was taken up in DNA isolation buffer (250 mM NaCl, D e c 161 100 mM EGTA, 100 mM Tris-HCl [pH 8.0], and 1% [wt/vol] lithium dodecyl e m 162 sulphate) and stored frozen at -70 °C or, for RNA samples, preserved in RNAlater b e r 2 163 (Applied Biosystems, Warrington, UK) and refrigerated at 4 °C. At the end of the 0 , 2 164 cruise, the preserved nucleic acid samples were shipped by air to the UK on dry ice 0 1 8 165 and subsequently stored at -80 °C prior to the extraction and purification of DNA and b y g 166 RNA as described by Bird et al. (21). u e s 167 t 168 Plankton net hauls for the collection of Trichodesmium spp. colonies and extraction 169 of DNA 170 7 171 Plankton samples were also collected from depths of 5 or 10 m at each station during 172 short (10 – 15 minutes) horizontal hauls using a standard WP2 conical net (200 μm 173 mesh size) towed at a ship speed (through the water) of 1 knot. Trichodesmium 174 colonies were sorted into sterile, filtered (0.2 μm pore size polycarbonate membranes) 175 surface seawater using a sterile, disposable inoculating loop, rinsed in two changes of D o w 176 sterile seawater and transferred to 0.5 ml RNAlater prior to storage at 4 °C. DNA was n lo a 177 extracted from the sorted Trichodesmium colonies (~5 colonies per extraction) after d e d 178 rinsing twice in sterile ASW medium (22) to remove RNAlater. The washed colonies f r o m 179 were collected on 0.2 μm pore size polycarbonate membranes and then lysed in TE h t 180 buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA) amended to 0.1 % Triton X-100 and tp : / / 181 0.2 mg ml-1 lysozyme (Sigma cat. number L7651) at 37 °C for 15 minutes. The ae m 182 lysates were brought to 50 μg ml-1 proteinase K (Roche) and 0.2 % SDS and .a s m 183 incubated at 56 °C for a further 10 minutes before purification of DNA using the . o r g 184 Qiagen DNeasy kit using the supplier’s abridged protocol for the purification of DNA / o n 185 lysates. D e c 186 e m 187 Enrichment Culture for the Isolation of Nitrate-Respiring (Denitrifying) Bacteria b e r 2 188 from the Arabian Sea 0 , 2 189 0 1 8 190 Seawater was collected from a depth of 120 m in suboxic waters at station 10 (24° 19' b y 191 N, 58° 10' E; depth 2863 m; Figure 1) and amended with 1.5 g l-1 Tryptic Soy Broth g u e s 192 (Thermo Fisher Scientific, Basingstoke, UK). The amended seawater was further t 193 supplemented with 1 x Guillard’s (F/2) nutrient and vitamins solution (Sigma, Poole, 194 UK) and incubated in filled, air-tight, sterile bottles at 25 °C. Following the visible 195 establishment of bacterial growth after 3–5 days, sub-cultures were transferred 8 196 aseptically to degassed (boiled), filter-sterilized seawater from the same station 197 amended with 0.5 g l-1 NaNO , 1.0 g l-1 NH Cl, 0.006 g l-1 FeCl , 0.001 g l-1 EDTA, 3 4 2 198 0.003 g l-1 K HPO , 1 ml l-1 A5 trace metal mixture (23) and 1 g l-1 of sodium acetate. 2 4 199 The sub-cultures were incubated in filled, air-tight, sterile bottles at 25 °C until the 200 medium turbidity increased (7–14 days). After two further rounds of subculture, the D o w 201 bacterial suspensions were transferred to streak plates of the same medium solidified n lo 202 with 10 g l-1 Difco Bactoagar (Becton, Dickinson and Co., Oxford, UK). The ad e d 203 inoculated plates were incubated at 25 °C in sealed, anaerobic jars in an oxygen-free f r o m 204 nitrogen atmosphere. Individual colonies were isolated by repeated subculture on agar h t 205 streak plates and maintained thereafter at 25 °C under these same incubation tp : / / a 206 conditions. e m 207 .a s m 208 PCR Amplification of nosZ, nirS and 16S rRNA Genes from Bacteria Isolated from . o r g 209 the Arabian Sea / o n 210 D e c 211 Independent isolates of putative, nitrate-respiring bacteria were grown aerobically at e m 212 25 °C for 24–48 h in 3 ml ASW medium (22) amended with 10% (vol/vol) Luria b e r 213 Bertani broth (10 g l-1 tryptone, 5 g l-1 yeast extract, 10 g l-1 NaCl) in an orbital 20 , 2 214 incubator at 180 r.p.m. The cultures were harvested by centrifugation at 16,000 x g for 0 1 8 215 2 minutes and DNA was isolated from the pelleted cell material using a DNeasy b y g 216 Tissue kit following the suppliers (Qiagen Ltd., Crawley UK) recommended protocol u e s 217 for bacteria. The DNA samples were screened for the presence of nosZ by the PCR t 218 using Thermoprime DNA polymerase master mix (Fisher Scientific, Loughborough, 219 UK) in reaction volumes of 25 µL containing 2mM MgCl , 10 ng DNA and 50 pmol 2 220 each of the primers nosZF1 and nosZR (Table 1). 9 221 222 The degenerate primers nosZF1 and nosZR were designed to target the conserved 223 motifs DV(H/Q)YQPGH and CHA(M/I/L)H(L/M)EM identified in the majority of 224 complete NosZ sequences available from the GenBank database at the start of this 225 study and correspond to the nucleotide positions 1276 – 1298 and 1864 – 1887 of D o w 226 nosZ from Pseudomonas stutzeri (ZoBell strain) ATCC 14405 (GenBank accession n lo a 227 number CAA37714.2) respectively. After denaturation at 95 °C for 2 minutes, the d e d 228 cycling conditions were 94 °C for 30 seconds, 55 °C for 30 seconds and 72 °C for 1 f r o m 229 minute for 25 cycles followed by a final extension at 72 °C for 10 minutes. h t 230 tp : / / a 231 The PCR products obtained were resolved in 1% (wt/vol) agarose gels and purified e m 232 using the Wizard SV Gel and PCR Clean-Up System (Promega Ltd., Southampton, .a s m 233 UK) before TA cloning in the plasmid vector pCR-TOPO as recommended by the . o r g 234 supplier (Invitrogen, Paisley UK). The cloned PCR products from four independent / o n 235 isolates (designated 1N, 4N, 5N and 8N) were DNA sequenced on both strands using D e c 236 M13 forward and reverse primers in parallel reactions performed with a DYEnamic e m 237 ET Terminator cycle sequencing kit (GE Healthcare Life Sciences, Little Chalfont b e r 2 238 UK) and an ABI Prism 377 automated sequencer. 0 , 2 239 0 1 8 240 An ~890 bp fragment of the gene encoding NirS (nitrite reductase) was amplified b y g 241 from genomic DNA purified from the Arabian Sea isolate designated 4N using the u e s 242 primer pair nirS1F and nirS6R (24; Table 1). The reactions were carried out as t 243 described above for nosZ except that the annealing temperature was increased to 58 244 °C. Genomic DNA from this isolate was also used to amplify a fragment of the 16S 245 rRNA gene with the universal primer pair 27F and 1492R (25) under similar reaction 10