Magnitude and controls of N2 fixation in the subtropical Northeast Atlantic

! ! ! ! ! ! ! ! ! ! ! ! ! D/Dª!Juan!Luis!Gómez!Pinchetti,!secretario!del!Departamento!de!Biología! de!la!Universidad!de!Las!Palmas!de!Gran!Canaria,!certifica:!! ! !que!el!Consejo!de!Doctores!del!Departamento!en!sesión!extraordinaria! tomó! el! acuerdo! de! dar! el! consentimiento! para!su! tramitación,! a! la! tesis! doctoral! titulada! “Magnitude! and! controls! of! N2! fixation!in! the!subtropical! Northeast! Atlantic”! presentada! por! el/la! doctorando/a! D/Dª! Mar! Benavides! Gorostegui! y! dirigida! por! el! Dr.! Javier! Arístegui! Ruiz! y! la! Dra.! Nona! Sheila! Agawin!Romualdo.! ! !Y! para! que! así! conste,! y! a! efectos! de! lo! previsto! en! el! Artº! 73.2! del! Reglamento!de!Estudios!de!Doctorado!de!esta!Universidad,!firmo!la!presente! en!Las!Palmas!de!Gran!Canaria,!a!3!de!Diciembre!de!2012.! ! ! ! ! ! ! ! Programa$de$doctorado$en$Oceanografía$ Bienio$2008G2010.$Con$Mención$de$Calidad$de$la$ANECA.$ !! !! Título!de!la!Tesis:! ! Magnitude$and$controls$of$N2$fixation$in$the$ subtropical$Northeast$Atlantic$ Magnitud!y!controles!de!la!fijación!de!nitrógeno!en!el!Nordeste!Atlántico! subtropical! !! Tesis! doctoral! presentada! por! Dª! Mar! Benavides! Gorostegui! para! optar! al! grado!de!Doctor!por!la!Universidad!de!Las!Palmas!de!Gran!Canaria.!! !! Dirigida!por:!! Dr.!D.!Javier!Arístegui!Ruiz!!!! !!!Dra.!Dª.!Nona!Sheila!Agawin!Romualdo! ! ! El/la$Director/a$$$$$$$$El/la$CoGDirector/a$$$$$$$$$$$$$$El/la$Doctorando! !! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! 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P,'!Q'2/$!3&$&./!*Q&0+&)./!.'!+&!4(.&G!+&!*(')*(&G!+&!3/+K%(*&G!-!%/./!+/!P,'!$'! )/$!Q&! (./! 3&$&)./!3/0! +&! *&X'1&I! jO0&*(&$! 3/0! %/./! +/! P,'! )/$! Q'2/$! 0'K./! R,)%&$k![-0:?0G1:%a]2O0&*(&$!3/0!%/./!%,!&3/-/!'$%/$!&d/$G!3/0!%/.&$!+&$!0($&$G! 3/0!%/.&!%,!&-,.&!*/)!M,*(Y'0!<-!*/)!%/./!+/!.'2W$FG!%'!4/-!&!'*Q&0!.'!2')/$I!8! 6()/%23/0!'$/$!0&%(%/$!%&)!&50&.&X+'$!')!+&!')%0&.&!.'!+&!Z&*,+%&.!/!%/2&)./! General%introduction% % ! ! ! 20! component! of! our! atmosphere!(78%),! it! is! mainly! unavailable! to! primary! producers.!This!is!caused!by!the!high!amount!of!energy!required!to!break!the! triple!bond!linking!both!nitrogen!atoms!of!the!N2!molecule.!Only!a!restricted! group!of!organisms!are!able!to!carry!out!biological!N2!fixation:!the!N2!fixers!or! diazotrophs!(from!the!greek!dis%GbisG,%azōos%GinertG,!and%trophikos%GnutritionG).! These! organisms! contain!a! nitrogenase! enzyme! complex,! composed! of! two! proteins:! dinitrogenase! (an! ironGmolybdenum! protein! encoded! by! the! nifDK! gene),!and!dinitrogenaseGreductase!(an!iron!protein!encoded!by!the!nifH!gene)! (Postgate,!1982).!Molybdenum!is!sometimes!replaced!by!vanadium!or!iron!in! some! alternative!nitrogenases.! The! nitrogenase! enzyme! complex! allows! diazotrophs!to!reduce!N2!to!ammonia!(NH3)!under!the!following!stoichiometry:!! ! N2!+!8H+!+!8eG!+!16ATP!!2NH3!+!H2!+!16ADP!+!16Pi!(1)! ! !As! previously! mentioned,! the! biological! fixation! of! N2! requires! a! high! amount! of! energy! (16! ATP).! Where! it! occurs,! this! process! enriches! the! food! web!through!the!leakage!of!ammonium!(NH4+)!and!amino!acids!that!serve!as!a! source!of!nitrogen!for!autotrophic!nonGdiazotrophic!phytoplankton!(Karl!et!al.,! 2002).! !Studying! the! nitrogen! cycle! is! of! major! interest.! Microbes! control! the! oceanic!nitrogen!cycle!through! a! set!of!gain!and!loss!processes,!and! sustains! the! marine! trophic! web! (Fig.! 2).! Inorganic! nitrogen! is! fixed! (or! ‘gained’)! through! N2!fixation! and!the!assimilation! of!NO3G,!nitrite!(NO2G)! and!NH4+,! and! remineralized!(or!‘lost’)!through!ammonification,!denitrification!and!anaerobic! ammonium!oxidation!(anammox).! !The!study!of!N2!fixation!has!become!of!great!interest!not!only!because!it! fuels!primary!production!in!large!oceanic!areas!(Capone!et!al.,!2005),!but!also! because!of!its!critical!role!in!balancing!the!oceanic!nitrogen!cycle.!Indeed,!N2! fixation! versus! denitrification! and! anammox! are! the! processes! thought! to! maintain!the!fixed!nitrogen!reservoir!balance!in!the!ocean! (Codispoti,! 2007).! Present!estimates!indicate!that!nitrogen!loss!via!denitrification!and!anammox! in! suboxic! and! anoxic!zones! far! exceed! rates! of! N2!fixation! in! surface! waters! (Table! 1).!However,! current! improvements! in! the! methodology! used! to! estimate! N2! fixation!(Mohr! et! al.,! 2010)!points! toward! a! smaller! difference! between! fixed! nitrogen! gains! and! losses,! which! could! eventually! balance! the! oceanic!fixed!nitrogen!budget!(Großkopf!et!al.,!2012).! ! ! PART%I:%Introduction% % ! ! 21! ! Fig.!2.!The!marine!nitrogen!cycle.!Modified!from!http://bit.ly/wC3xZo! ! !Recent! studies! have! shown! that! planktonic! diazotrophs! are! more! diverse!(Fig.!3)!and!more!widely!distributed!than!previously!thought!(Zehr!et! al.,! 2001,! 2008;! Moisander! et! al.,! 2010;! Riemann! et! al.,! 2010;! Farnelid! et! al.,! 2011).! Filamentous' heterocystous' cyanobacteria,! like!Anabaena,% Aphanizomenon%and!Nodularia,!are!frequently!present!in!estuaries!as!well!as!in! semiGenclosed!seas,!like!the!Baltic!Sea,!but!are!rare!in!open!ocean!waters!(Zehr,! 2011).! !Non5heterocystous'filamentous'cyanobacteria!such!as!Katagnymene! and!Trichodesmium!have!been!observed,!however,!in!open!ocean!regions!(e.g.! Langlois!et!al.,!2005),!although!the!species!of!Trichodesmium!are!undoubtedly! the! most! abundant! forms.! This! cyanobacterium! is! ubiquitous! in! the! tropical! and! subtropical! oceans,! where! it! often! forms! massive! blooms.! The! high! abundances!found!in!these!regions!lead!to!the!thought!that!it!was!the!principal! N2!fixer!in!the!oceans.!Its!diazotrophic!activity!was!discovered!in!the!early!60s! (Dugdale! et! al.,! 1961)! and,! ever! since,! Trichodesmium!has! been! the! focus! of! intensive! research.! As! a! result,! we! now! have! fairly! good! knowledge! of! its! nutritional! (e.g.! BermanGFrank! et! al.,! 2001a;! Dyhrman! et! al.,! 2002),! temperature!(e.g.!Breitbarth!et!al.,!2007)!and!physical!(Davis!and!McGillicuddy,! 2006)!controls,!global!distribution!(Luo!et!al.,!2012),!and!potential!responses! to!increased!atmospheric!CO2!levels!(e.g.!Hutchins!et!al.,!2007).!A!recent!review! by!Bergman!et!al.!(2012)!covers!all!of!these!aspects.! !Another!group!frequently!targeted!in!N2!fixation!studies!are!symbiotic' diazotrophs,! such! as! the! cyanobacteria! Richelia!and! Calothrix!which! are! symbionts!of!the!diatoms!Rhizosolenia,%Hemiaulus!or!Chaetoceros!(e.g.!Foster!et! al.,!2011).! General%introduction% % ! ! ! 22! Table!1.!Global!estimates!of!nitrogen!sinks!and!sources.!Modified!from!Gruber!(2008).!All!rates!are! in!Tg!N!yG1.! Process' Codispoti'et'al.' (2001)' Galloway'et'al.'' (2004)' Gruber'' (2004)' % Sources% % % % Pelagic!N2!fixation! 117! 106! 120! Benthic!N2!fixation! 15! 15! 15! River!input! 76! 48! 80! Atmospheric!deposition! 86! 33! 50! Total! 294! 202! 265! ! ! ! ! Sinks% % % % Organic!nitrogen!export! 1! ! 1! Benthic!denitrification! 300! 206! 180! Pelagic!denitrification! 150! 116! 65! Sediment!burial! 25! 16! 25! N2O!loss!to!atmosphere! 6! 4! 4! Total! 482! 342! 275! ! ! ! ! Sources%I%sinks% I188% I140% I10% ! ! !These! symbiotic! relationships! may! contribute! significantly! to! the! oceanic!carbon!‘biological!pump’.!Indeed,!diatomGdiazotroph!associations!have! been!recently!identified!as!the!predominant!cause!for!particulate!matter!export! peaks!during!the!summer!in!the!North!Pacific!Ocean!(Karl!et!al.,!2012).!Other! less! studied! symbiosis! of! the! pelagic! habitat! are!those! of!anaerobic! diazotrophic!bacteria!and!copepods!(Proctor,!1997).! !Recently,! the!application! of! molecular! biology! techniques! has! allowed! the!identification!of!a!new!wide!set!of!diazotrophs!through!the!detection!of!the! nifH!gene.! Zehr! et! al.! (1998;! 2001)! were! the! first! to! report! the! presence! of! unicellular' diazotrophic' cyanobacteria' (UCYN)!in!the!North!Pacific!Ocean.! In!the!following!years,!UCYN!of!groups!A,!B!and!C!(UCYNGA,!UCYNGB!and!UCYNG C)!were!detected!throughout!the!Atlantic!(Langlois!et!al.,!2005,!2008)!and!the! Pacific! Oceans! (Needoba! et! al.,! 2007),! where! they! occasionally!exceeded! Trichodesmium! N2! fixation! rates! (Falcón! et! al.,! 2004;! Montoya! et! al.,! 2004;! Benavides!et!al.,!in!press).!The!discovery!of!UCYNGA!was!especially!remarkable.! This!yetGuncultivated!small!organism!(<1!µm)!is!now!recognized!to!be!the!most! abundant! marine! diazotrophic! cyanobacterium!(Luo! et! al.,! 2012).!In! comparison!to!Trichodesmium!and!other!diazotrophs!which!need!warm!waters! to!grow!(generally!>20ºC),!UCYNGA!have!been!detected!at!higher!latitudes!and! depths! (Moisander! et! al.,! 2010),! and! are! even! present! in! upwelling! systems! where! the! temperatures! are! often! below! 17ºC! and! fixed! nitrogen! concentrations!are!high!(Sohm!et!al.,!2011b;!N.S.!Agawin,!unpublished!results).! PART%I:%Introduction% % ! ! 23! A! remarkable!peculiarity! of! UCYNGA! is! their! photoheterotrophic! metabolism.! These!organisms!lack!genes!for!the!oxygenGevolving!photosystem!II!and!carbon! fixation! (Zehr! et! al.,! 2008),! hence! they! are! believed!to! rely! on! compounds! produced! by! other! organisms! (Tripp! et! al.,! 2010).!Indeed,! Thompson! et! al.! (2012)! in! a! very! recent! study! found! UCYNGA! in! symbiosis! with! a! prymnesiophyte,! confirming! earlier! suspicions!of! their! mode! of! life.!These! authors!have!proposed!the!name!Candidatus%atelocyanobacterium!thalassa!for! the!UCYNGA.! ! ! Fig.!3.!Phylogenetic!diversity!of!the!nifH!gene!from!marine!planktonic!diazotrophs.!From!Riemann! et!al.!(2010).! ! ! !Besides! UCYN,! the! global! diazotrophic!domain! is! further! extended! to! non5cyanobacterial' diazotrophs,!which!are!found!in!all!four!clusters!of!the! nifH!gene!(Chien! and! Zinder,! 1996).! They! belong! mainly! to!αG,! βG,! γG! and! δG proteobacteria,! and! less! frequently! to! Archaea! and! anaerobic! bacteria! (Riemann!et!al.,!2010).!Recent!evidence!indicates!that!nonGcyanobacterial!nifH! is! more! abundant! and! diverse! than!UCYN! in! global! ocean! surface! waters! Riemann et al.: Non-cyanobacterial diazotrophs in marine waters gent non-functional archaeal nifH homologues, have been reported from the deep sea (Mehta et al. 2003). Based on clone libraries, nifH sequences from nonCyanobacteria appear to be not only diverse, but also abundant relative to those of Cyanobacteria. This is particularly evident from a recent compilation of published marine nifH gene clone libraries (Farnelid & Riemann 2008) showing that 73 to 91% (average 83%) of the total number of nifH sequences obtained were related to nonCyanobacteria. The data compiled here (Fig. 1) similarly show that 80% of the sequences from coastal marine and estuarine plankton samples derive from nonCyanobacteria. In the open ocean the proportion is lower, with ca. 36% of the sequences deriving from nonCyanobacteria, but this likely underestimates their numerical contribution, since many oceanographic studies sample most intensively in the euphotic zone, and clone libraries from below ~200 m consist almost exclusively of non-cyanobacterial nifH genes (e.g. Hewson et al. 2007a). While nifH clone libraries provide convincing evidence for a near-ubiquitous distribution of non237 Fig. 1. Phylogenetic tree illustrating the diversity among 2570 nifH genes amplified from microorganisms in plankton samples from marine and estuarine environments. Branches derived from sequences (n = 809) from open ocean samples are marked with a “s”. Green and blue branches belong to a loosely affiliated group of sequences designated as Cluster I (Chien & Zinder 1996), orange branches are affiliated with Clusters II and III. Purple branches are not assigned to the traditionally defined clusters. The tree (neighbour-joining, with no distance corrections) was generated using Arb (Ludwig et al. 2004) and is based on amino acid residues 46 to 151 (Azotobacter vinelandii numbering) translated from PCR-amplified fragments of nifH genes or transcripts. Sequences were downloaded as an Arb database that was last updated on 30 March 2009 (http://pmc.ucsc.edu/~wwwzehr/ research/database/) and aligned by a hidden Markov model for nifH available at the Pfam web site (Finn et al. 2010). Sequences from marine and estuarine samples (oceans, seas, bays, gulfs, harbours, lagoons) were retrieved from the database, and those from sediments, hydrothermal vents or associated with sessile plants and animals were excluded. Open ocean samples include those from the Atlantic and Pacific Oceans and the Arabian Sea, excluding nearshore environments (gulfs, bays, harbours, fjords) and inland seas (Baltic, Mediterranean). Clusters are labelled to indicate the phylogenetic affiliations of cultivated microorganisms whose nifH sequences most closely match those from the uncultivated microorganisms shown in the tree. A total of 66% of all sequences shown are non-Cyanobacteria, and 36% of those from the open ocean are non-Cyanobacteria General%introduction% % ! ! ! 24! (Farnelid!et!al.,!2011),!while!heterotrophic!N2!fixation!dominates!in!the!South! Pacific!Ocean!Gyre!(Halm!et!al.,!2011).!NonGcyanobacterial!nifH!is!also!the!most! abundant!below!200!m!depth!in!the!Sargasso!Sea!(Hewson!et!al.,!2007),!and!N2! fixation! rates! up! to! 0.3! nmol! LG1! dG1!associated! with!heterotrophs! have! been! measured!in!hypoxic!basins!in!the!Southern!California!Bight!(Hamersley!et!al.,! 2011).! All! this! suggests!that! typical!nifH!measurements! in! the! upper! water! column!underestimate!the!real!importance!of!nonGcyanobacterial!diazotrophy! (Riemann! et! al.,! 2010).!Nevertheless,! although! measurements!of! nonG cyanobacterial! nifH!abundance! and! distribution! are! increasing,! the! contribution!of!these!organisms!to!global!N2!fixation!remains!unknown.!! ! Controls%and%limitations% !There!are!several!environmental!factors!that!control!oceanic!N2!fixation,! but!the!extent!to!which!each!one!of!these!limits!or!enhances!N2!fixation!varies! among! diazotrophic! species.!Overall,! the! most!obvious!limiting! factor! is! oxygen'(which! is! a! byproduct! of! photosynthesis).! Oxygen! deactivates! the! nitrogenase!enzyme!complex,!inhibiting!diazotrophic!activity!(Postgate,!1982).! Photosynthetic!diazotrophic!organisms!have!developed!different!strategies!to! overcome! this! issue.! Some! filamentous! cyanobacteria! bear!heterocysts! that! create!a!microanaerobic!environment!and!thus!are!well!adapted!to!N2!fixation! in! oxygenated! waters.! However,! most! oceanic! photosynthetic! diazotrophic! cyanobacteria! are! nonGheterocystous! (Zehr,! 2011),! and! must! find! other! solutions!to!fix!N2!in!the!presence!of!oxygen.!SingleGcelled!diazotrophs!such!as! Crocosphaera!or! Cyanothece!confine! N2! fixation! to! the! night,! when! photosynthesis!and!subsequent!oxygen!production!are!absent.!Trichodesmium! lacks!heterocysts!but!surprisingly!fixes!CO2!and!N2!simultaneously!during!the! day!(Capone!et!al.,!1997;!BermanGFrank!et!al.,!2001b).!This!cyanobacterium!is! able!to!fix!N2!in! the! presence!of! oxygen!by!a!combined!spatial!and! temporal! segregation! strategy.! Temporal! segregation! strategies! such! as!transient! modifications!of!the!nitrogenase! enzyme! complex! or! its! rapid! turnover! have! been!discussed!(Capone!et!al.,!1997).!Another!possible!strategy!is!the!reduction! of!cellular! oxygen!by! ‘overfixation’! of! carbon,! which! likely! explains! the! high! carbonGtoGnitrogen!(C:N)!ratios!found!in!these!diazotrophs!(Mulholland,!2007).! Moreover,!BermanGFrank!et!al.!(2001b)!found!that!N2!fixation!takes!place!in!a! different!time!frame!than!photosynthesis!during!the!day.!Spatial!segregation!is! provided! by! structural!differences! along! the! trichomes:! the! filaments! have! zones!where!cells!are!light!and!less!granulated.!These!cells!termed!diazocytes! are!the!place!where!the!nitrogenase!is!found!(Bergman!et!al.,!2012).!Once!N2!is! fixed,!it!is!rapidly!distributed!along!the!whole!trichome!(FinziGHart!et!al.,!2009).! PART%I:%Introduction% % ! ! 25! !There! is! a! wide! body! of! literature! addressing! N2!fixation! limitation! by! temperature.! Because! high! water! temperatures! prevent!oxygen! dissolution! and! enhance! cell! respiration,! temperature! is! thought! to! constrain! the! global! distribution!of!diazotrophic!cyanobacteria!(Staal!et!al.,!2003;!Stal,!2009).!For! example,!the!optimum!temperature!for!the!growth!of!Trichodesmium!has!been! estimated! at!27ºC! (Breitbarth! et! al.,! 2007),! which! confines! its! growth! to! the! tropical!and!subtropical!oceans.!Although!UCYN!are!also!most!abundant!in!the! tropical!and!subtropical!oceans!(Luo!et!al.,!2012),!the!presence!of!UCYNGA!and! heterotrophic! diazotrophs! at! higher! latitudes!and! depths!(Moisander! et! al.,! 2010),!in!coldGwater!upwelling!systems!(Sohm!et!al.,!2011b),!and!even!in!the! Arctic! (Blais! et! al.,! 2012)!indicates! that! the! diazotrophic! latitudinal! range! is! wider.! ! N2! fixation!may! also! be! limited! by! the! in! situ! availability! of! fixed' nitrogen!(e.g.! Krishnamurthy! et! al.,! 2007).!Additions! of! NO3G!suppress! the! expression! of! nif!genes,! and! the! production! of! diazocytes! in! Trichodesmium! (Mulholland! et! al.,! 2001;! Holl! and! Montoya,! 2005).! In! the! case! of! UCYN,! Dekaezemacker! and! Bonnet! (2011)! studied! the! effect! of! NO3G!and! NH4+! additions!on!the!diazotrophic!activity!of!two!cultured!strains!of!Crocosphaera.! They! found! that! N2!fixation! rates! decrease! with! increasing! concentrations! of! NH4+,! while! the! rates! were!not! affected! by! any! additions! of! NO3G.! This! may! explain! why! UCYN! are! usually! found! at! greater! depths! in! the! water! column! than! Trichodesmium,! and! even! in! coastal! upwelling! systems!(Sohm! et! al.,! 2011b).!! !UCYNGA! may! rely! on! dissolved! organic! matter! (DOM)! as! a! nutrient! source! (Tripp! et! al.,! 2010),! and! therefore! their! nitrogenase! activity! is! likely! unaffected!by!high!inorganic!nitrogen!in!situ!concentrations.!Indeed,!UCYNGA! are!dominant!in!the!northwest!African!upwelling!system!off!Cape!Ghir!(~30G 31ºN)! (N.S.R.! Agawin,! unpublished!results),! where! NO3G!concentrations! are! usually!>2!µM!(Benavides!et!al.,!2011).! !On!the!other!hand,!the!availability!of!other!nutrients!such!as!iron' and' phosphorus!influences! N2!fixation.! Diazotrophs! have! very! high! iron! cellular! quotas!(e.g.!BermanGFrank!et!al.,!2001a).!More!importantly,!iron!is!the!major! cofactor! of! the! nitrogenase! reductase,! which! makes! N2! fixation! directly! dependent!on!iron!availability.!Iron!reaches!the!open!ocean!mainly!through!the! deposition! of! atmospheric! dust,! although! this! flux! can! be! overpassed! by! vertical!diffusive!mixing!(Rijkenberg!et!al.,!2012).! !Due!to!the!proximity!of!the!Sahara!desert,!the!eastern!Atlantic!receives! the!greatest!inputs!of!desert!dust!in!the!world’s!ocean!(Prospero,!1981;!Fig.!4).! The! distribution! of! Saharan! dust! deposition! over! the! North! Atlantic! (Fig.! 4)! General%introduction% % ! ! ! 26! closely!matches!the!distribution!of!diazotrophic!activity!(Fig.!5),!suggesting!a! connection!between!iron!availability!and!N2!fixation.! ! ! Fig.!4.!Global!distribution!of!!average!dust!deposition!(g!mG2!yG1).!From!Jickells!et!al.!(2005).! ! This! has! been! further! confirmed! experimentally!by! adding! dust! to!seawater! samples! and!by! correlations! between! N2! fixation! rates! and! in! situ! dissolved! iron!concentrations!(e.g.!Mills!et!al.,!2004;!Moore!et!al.,!2009;!Fernández!et!al.,! 2010).! ! ! Fig.! 5.! Geometric! mean! of! depthGintegrated! N2! fixation! rates! (µmol! N! mG2! dG1).! From! Luo! et! al.! (2012).! ! macronutrients (42,43), and trace nutrients (such as Co and Zn) (2,44). Furthermore atmospheric inputs supply not only iron but also other nutrients and carbonate, which may influence ocean biogeochemistry (45,46). Luxury iron uptake has been demonstrated for some phytoplankton, allowing them to better adapt to episodic atmospheric supply (47). Iron availability influences algal community structure as well as overall productivity. Open ocean phytoplankton generally need less iron than coastal species, which have evolved in a more iron-rich environment, although iron-limited coastal systems are known (36). A reduced iron requirement can be achieved by reducing cell size or minimizing the number of ironcontaining enzymes (39). The success of Prochlorococus in HNLC areas depends on both strategies. Relief of iron stress results in the growth of phytoplankton taxa characterized by larger cells, particularly diatoms with less dense opal skeletons (36). A similar process may arise for coccolithophores as a result of Fe/Zn co-limitation (44). Changes in skeleton density should influence sinking rates and hence carbon export to depth, although this has not been seen in field experiments (48). Changes in coccolithophore abundance directly affect atmospheric partial pressure of CO 2 (pCO 2 ), because their calcification produces CO 2 (36,42). In addition to direct limitation of primary production in the HNLC regions, iron may limit (or co-limit with P) nitrogen fixation by photosynthetic diazotrophs in tropical oceans, where stratification creates high temperature and irradiance and low nitrate concentrations in surface waters, which favor this process (1,43). The best characterized photosynthetic diazotroph, Trichodesmium, requires 5 to 10 times more iron for growth based on nitrogen fixation, as compared to ammonium (47). The supply of dust to the oceans is very important in maintaining oceanic primary production and CO 2 uptake but is sensitive to climate change, although the overall effect will vary between ocean biogeochemical provinces (Table 2). In HNLC regions, changes in iron supply will directly affect primary production and species composition, whereas in subtropical/tropical oligotrophic regions, the impact will be mainly via changes in nitrogen fixation. The dust supply from the great North African and Asian deserts directly affects the tropical North Atlantic and temperate North Pacific, respectively, and effects in the two regions can be expected to be different. The largest HNLC region, the Southern Ocean (36), has the biggest potential to influence atmospheric CO 2 . Here atmospheric dust supply is low (Fig. 2), originating from small dust sources in Argentina, Australia, and South Africa (6). Changes in these small and littlestudied desert regions may have a disproportionately large global impact. Because the solubility of iron from dust is low, it follows that there is a large flux of particulate iron through the deep ocean, particularly beneath the major dust plumes. If some of this dust dissolves at depth, it will increase abyssal dissolved iron concentrations and, over the long term, productivity in upwelling regions such as the Southern Ocean. Deep-water dust dissolution will depend on organic ligand concentrations and possibly sediment redox (33). Martin (49)proposedthatincreaseddust transport during the last glaciation reduced iron limitation in HNLC regions, increasing primary production and CO 2 uptake. The complexity of iron biogeochemistry and nutrient co-limitation means that higher glacial dust loadings need not necessarily cause increased productivity. Current models and ice core data yield very different results, predicting that glacial/interglacial changes in dust fluxes will change atmospheric pCO 2 by 5 to 45 parts per million (ppm) as a contribution to the total change of 80 to 100 ppm (19,50). Bopp et al.(50) reviewed much of the existing marine sediment core data on glacial/interglacial ocean productivity changes and found no simple global pattern of change. However, there are regional patterns (51)with increases in productivity in the northwest Pacific, South Atlantic, and Indian Oceans north of the polar front, with decreases south of it. South Pacific productivity appears to be little changed. Some of these patterns can be reproduced in ocean models (50). Effect on Climate of Iron Inputs to the Oceans The oceans clearly exert a major influence on climate via heat transport and related physical processes (20). Large-scale reorganization of oceanic circulation will also affect the transport of iron, effects driven predominantly from within the ocean. Climate change will induce a variety of physicochemical changes in the open ocean, particularly by changing stratification and nutrient supply ratios (42), with unpredictable effects. We acknowledge these important issues but focus on the dust cycle, considering now ways in which this can affect the oceans and climate, beside the direct iron limitation of primary production and nitrogen fixation discussed above (Table 2). Changes in iron fluxes can result in species shifts and changes in phytoplankton size distribution, changing oceanic CO 2 uptake by altering the efficiency of organic carbon export to deep water. Dust may also play a direct role in regulating export via the ballast effect (52). In most areas, dust is a minor ballast component compared to opal and calcite, but their production is also influenced by dust/iron supply. Changes in ocean productivity and organic carbon export to deep water will influence subsurface oxygen levels and thereby denitrification in oxygen minima zones, oceanic nitrate inventories and productivity, and nitrous oxide emissions (53). Changes in sediment H 2 Sinsuchareascouldaffectdeepocean iron concentrations and productivity. Up to eightfold changes in dimethyl sulfide (DMS) concentrations are seen in iron addition experiments (54). DMS oxidizes in the atmosphere to form acidic sulfate aerosol, a highly effective scatterer of solar radiation. Modeling suggests that a twofold global rise in DMS fluxes produces a global temperature decrease of 1-C, proving a climate feedback and linking Fig. 2. Dust fluxes to the world oceans based on a composite of three published modeling studies that match satellite optical depth, in situ concentration, and deposition observations (11,14,15). The models have been extensively compared to observations, and although individual models show strengths and weaknesses, this composite appears to match observations well. Total atmospheric dust inputs to the oceans 0450 Tg year –1 . Percentage inputs to ocean basins based on this figure are as follows: North Atlantic, 43%; South Atlantic, 4%; North Pacific, 15%; South Pacific, 6%; Indian, 25%; and Southern Ocean, 6%. REVIEW 1 APRIL 2005 VOL 308 SCIENCE www.sciencemag.org 70 on February 1, 2010 www.sciencemag.orgDownloaded from ESSDD 5, 47–106, 2012 Database of diazotrophs in global ocean Y.-W. Luo et al. Title Page Abstract Instruments Data Provenance & Structure Tables Figures � � � � Back Close Full Screen / Esc Printer-friendly Version Interactive Discussion Discussion Paper | Discussion Paper | Discussion Paper | Discussion Paper | (a) (b) (c) (d) (e) (f) Fig. 6. (a) Geometric mean depth-integrated N2fixation rates binned on 3 ×3◦grid. Also geometric mean N2fixation rates on 3×3◦grid for 5 vertical layers of (b) 0–5 m, (c) 5–25 m, (d) 25–62.5 m, (e) 62.5–137.5 m and (f) 137.5–250 m. The color bar is in logarithmic scale. Zero values are marked with black triangles. 103 PART%I:%Introduction% % ! ! 27! !Phosphorus! is! needed!for! energyGcarrying! compounds! such! as! ATP! or! NADPH.!Phosphorus!is!available!at!low!concentrations!in!the!open!ocean!and! frequently!limits! N2! fixation! in! Trichodesmium!(e.g.! Hynes! et! al.,! 2009)! and! UCYN! such! as%Crocosphaera!(Dyhrman! and! Haley,! 2006).!To! overcome! this! limitation,! Trichodesmium!bears! enzymatic! systems! capable! of! uptaking! organic! phosphorus! (phosphonates;! Dyhrman! et! al.,! 2006)!or! inorganic! phosphate!(PO43G)!(Dyhrman!et!al.,!2002).!Crocosphaera%cannot!uptake!organic! phosphorus! forms! but! is! capable! of! inducing!a! high! affinity! PO43G!binding! system!when!phosphorus!supply!is!low!(Dyhrman!and!Haley,!2006).! 'Physical' forcing!controls!the! availability! of!inorganic! nutrients!in! the! upper! ocean!through!mixing,! upwelling! and! vertical! diffusion.!In! contrast! to! autotrophic! phytoplankton! species,! whose! growth!depends! on! NO3G! transported! from! deep! waters!or! NH4+!regenerated! in! situ,! diazotrophic! organisms! are! capable! of! growing! on! N2!as! the! only! source! of! nitrogen.! Theoretically,! this! sets! out! a! different! (with! less! competition)! scenario! for! these!organisms,!which!would!grow!better!in!the!core!of!warm!and!nutrientG poor!stratified!waters,!than!in!cold!nutrientGrich!waters.! !Although! the! role! of! physical! forcing! on! N2! fixation! has! been! seldom! studied,! previous! research! indicates! that! mesoscale! features! and! associated! density!fronts!influence!N2!fixation!rates!and!diazotrophic!organism!diversity! to! some! extent! (Sohm! et! al.,! 2011c).! Holl! et! al.!(2007)!measured! greater! N2! fixation! rates! in! the! core! of! warmGcore! eddies! than! in! the! core! of! coldGcore! eddies! off! the! west! coast! of! Australia.! In! an! anticyclonic! eddy! in! the! North! Pacific! Ocean! close! to! station! ALOHA! (22°45’! N,! 158°W),! Fong! et! al.!(2008)! found! relatively! high! concentrations! of! filamentous! cyanobacteria! and! UCYN! across!the!eddy,!and!enhanced!N2!fixation!rates!towards!its!edge.!Church!et!al.! (2009)!gathered!monthly!measurements!at!station!ALOHA!from!2004!to!2007! and!detected!an!increase!in!N2!fixation!rates!coinciding!with!periods!of!positive! sea!surface!height!anomaly!(SSHA).!Davis!and!McGillicuddy!(2006)!found!high! abundances!of!Trichodesmium!associated!with!anticyclonic!eddies!in!a! crossG Atlantic! cruise.! Similarly,! Benavides! et! al.! (2011)! found! N2! fixation! peaks! associated! with! density! fronts! and! high! concentrations! of! the! filamentous! diazotroph%Trichodesmium!at! the! edge! of! an! upwelling! filament!in! the! Northeast! Atlantic.! Also,! because%Trichodesmium!is! a! buoyant! organism! (Villareal! and! Carpenter,! 2003)! adapted! to! stratified! warm! waters,! high! turbulence!is!known!to!prevent!its!proliferation!(Carpenter!and!Price,!1976).! All! these! studies! suggest!that! physical! forcing! exerts! a! clear! influence!on! N2! fixation!activity!and!diazotrophic!organisms’!distribution.! ! General%introduction% % ! ! ! 28! Measuring%N2%fixation! !There!are!two!main!approaches!to!estimate!N2!fixation:!the!‘geochemical’! methods,!and!the!‘biological’!(or!‘direct’)!methods.! !Geochemical'methods!rely!on!the!study!of!the!horizontal!and!vertical! distribution!of! particulate!and!dissolved!nitrogen!pools! that! have! a!chemical! signature!indicative! of! N2! fixation.! There! are! two! principal! geochemical! methods!based!on!the!estimation!of!the!N*!and!δ15N!parameters.! !The! N*! parameter!is! based! on! the!relative! concentrations!of! NO3G!and! PO43G!in!seawater.!Redfield!et!al.!(1963)!established!that!carbon,!nitrogen!and! phosphorus! are! taken! up!and! remineralized! at!a! constant! ratio! of! 106:16:1! (C:N:P).!N2!fixation!and!denitrification!are!not!paralleled!by!equivalent!inputs! or! losses! of! phosphorus.! Therefore,! N2! fixation! raises! N:P! ratios,! while! denitrification!lowers!them,!i.e.:!nitrogen!production!in!excess!of!the!Redfield! stoichiometry! (N:P>16)! is! indicative! of! N2! fixation,! and! nitrogen! removal! in! deficit!of! the! Redfield! stoichiometry!(N:P<16)! is! indicative! of! denitrification.! These!processes!can!be!estimated!using!the!N*!parameter,!which!measures!the! concentration! of! NO3G!in! excess! (or! deficit)! of! that! expected! from! the! remineralization! of! PO43G! at! Redfield! stoichiometries,! following! the!general! formulation!below:! ! N*!=![NO3G]!G!16[PO43G]! (2)! ! where![NO3G]!and![PO43G]!are!the!concentrations!of!NO3G!and!PO43G,!respectively.! This!parameter!was!firstly!introduced!in!the!90s!by!Michaels!et!al.!(1996).!The! equation!was!amended!by!adding!constants!that!bring!the!global!N:P!ratio!to! 16!and! the! intercept! to! zero!(Michaels! et! al.,! 1996;! Gruber! and! Sarmiento,! 1997).!Extensive!nutrient!databases!from!projects!such!as!WOCE!or!GEOSECS! have!been!used!to!estimate!the!distribution!and!magnitude!of!N*!in!the!oceans.! !The!second!geochemical!method!most!commonly!used!is!based!on!the! δ15N! parameter.! This! parameter!measures! the! relative! abundance! of! the! nitrogen!isotopes!14N!and!15N!in!a!sample!with!respect!to!standard!atmospheric! N2!(Equation!3):! ! δ15N!=![(15N/14N)sample/(15N/14N)standard!–!1]!x!1000! ! (3)! ! ! PART%I:%Introduction% % ! ! 29! !Atmospheric!N2!has!a!δ15N~0.6‰,!while!NO3G!has!a!δ15N~5‰!(Karl!et! al.,!2002).!The!δ15N!of!a!given!dissolved!or!particulate!nitrogen!pool!is!given!by! the! isotopic! composition! of! its! source! and! the! isotopic! fractionation! experienced!along!its!assimilation.!Therefore,!organic!nitrogen!produced!by!N2! fixation!is!depleted!in!δ15N,!while!higher!δ15N!values!are!expected!in!organic! nitrogen!pools!produced!by!the!assimilation!of!NO3G.! !In!general,!geochemical!methods!have!the!disadvantage!that!N2!fixation! signatures! (N*>2.5!µmol!kgG1!or!low!δ15N!values)!can!be!caused!by!processes! other!than!N2!fixation.!For!example,!high!N:P!fixed!nitrogen!reaches!the!ocean! via!atmospheric!deposition!of!anthropogenic!materials!derived!from!fossil!fuel! combustion! and! agriculture! fertilizers! (Zamora! et! al.,! 2010).! Moreover,! this! anthropogenic!atmospheric!nitrogen!has!low!δ15N!values!too!(Hastings!et!al.,! 2003).! Other!misinterpretations! stem! from! isotopic! fractionation! during! nitrogen! cycling! (Altabet,! 1988).!Despite! these! drawbacks,! the! geochemical! methods!have!the!advantage!that!they!can!be!used!in!large!oceanic!regions!at! higher!spatial!resolutions!than!it!is!possible!using!biological!methods.! The!biological'methods!measure!the!actual!amount!of!N2!fixed!by!in! situ! diazotrophic! organisms! during! a! given! period! of! incubation.! In! oceanographic!cruises,!incubations!are!generally!performed!on!deck!on!lightG adapted!incubators!cooled!with!surface!seawater,!or!alternatively!using!in!situ! freeGfloating!arrays.!There!are!two!principal!biological!methods:!the!acetylene! reduction!assay!(ARA)!and!the!assimilation!of!the!stable!isotope!15N2.!! The! ARA!measures! N2!fixation! indirectly! relying! on! the! fact! that! the! nitrogenase!enzyme!is!capable!of!reducing!acetylene,!a!tripleGbonded!molecule! structurally!comparable!to!N2.! The! reduction!of!acetylene!to!ethylene!can! be! directly! compared! with! the! reduction! of! N2!to! NH4+.! To! convert! ethylene! produced!to!N2!fixation!fixed,!a!conversion!ratio!must!be!applied.!The!common! theoretical!ratios!used!are!3:1!or!4:1!(C2H4:N2),!but!empirically!obtained!ratios! are!usually!higher!(Mulholland!et!al.,!2004,!2006;!Benavides!et!al.,!2011).!The! use! of! the! ratio! 3:1! or! 4:1! depends! on! whether!the! recycling! of! hydrogen! is! considered! or! not.! Hydrogen! is! an! obligate! byproduct! of! N2! fixation! (see! Equation! 1).! Two! electrons! are! used! when! reducing! acetylene! to! ethylene,! while! eight! electrons! are! used! to! reduce! N2!to! 2NH3,! yielding!a! conversion! factor!of!4:1!(Equation!5).!The!byGproduced!hydrogen!can!be!recycled!through! an! uptake! hydrogenase! enzyme,! using! two! electrons! less! and! yielding! a! 3:1! ratio!(Equation!6).!! ! ! $365/5%+;N6.)/B65%4(,%+-):/(6% % ! ! ! "U! (-*./0353!0/!&'+!3)2&-(<01.%!,(-&'!$&%./&01!(9+-!&'+!7T=#V,!<.-.%%+%=!W/!(-4+-! &(! 04+/&06?! :'01'! 6.1&(-3! :+-+! 1(/&-(%%0/*!&'+! 40.A(&-(<'01! .1&090&?;! :+! 1(5<.-+4!6-.1&0(/.&+4!,7!608.&0(/!./4!OP,!-+%+.3+!-.&+3!:0&'!6.1&(-3!3)1'!.3! &+5<+-.&)-+;! 3.%0/0&?;! 3+.! 3)-6.1+! '+0*'&! ./(5.%0+3;! /)&-0+/&! 1(/1+/&-.&0(/3! ./4!.&5(3<'+-01!4)3&!%(.43=! ! 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Chapter 1: N2 fixation in the subtropical Northeast Atlantic 40 Abstract We measured gross and net nitrogen fixation in fractionated samples (organisms >10 μm and <10 μm), and the density of Trichodesmium, during a cruise along the northeast Atlantic boundary current system and during 2 mesoscale experiments in the upwelling systems of Cape Silleiro (northwest Iberia) and Cape Ghir (northwest Africa). The density of Trichodesmium (<0.5 trichomes L−1) and its associated rates of nitrogen fixation (<0.1 μmol N m−2 d−1) were low. Trichomes appeared to accumulate at frontal sites — such as upwelling filaments and the Azores Front. Gross and net rates of nitrogen fixation were always <0.4 nmol N L−1d−1 except off the northwest African coast where a gross nitrogen fixation peak of 0.98 nmol N L−1 d−1 was measured. The <10 μm fraction contributed more to both gross and net nitrogen fixation than did the >10 μm fraction in most of the areas studied. The <10 μm fraction was responsible for 70 to 92% of the total nitrogen fixation in cold nutrient-rich areas. The contribution of small diazotrophs to nitrogen fixation in the upwelling sites suggests that the distribution and activity of these organisms are more widespread than previously thought. Introduction Nitrogen is the main nutrient that limits primary production in the open ocean oligotrophic environment where the quasi-permanent stratification of the upper water column prevents mixing with the denser and nutrient-rich deep waters (Falkowski, 1997). Dinitrogen (N2) fixation in the ocean is predominantly attributed to the photoautotrophic cyanobacteria. This process enriches the food web with combined nitrogen through leakage of ammonium and amino acids which serve as a source of nitrogen for autotrophic non-diazotrophic phytoplankton (Mahaffey et al., 2005). Indeed, N2 fixation is thought to fuel 50% of the primary production in these ‘oceanic deserts’ (Capone et al., 2005). Nitrogen is removed from the oxygen minimum zones (OMZ) and from sediments through denitrification. Based on present knowledge, the rates of denitrification exceed those of N2 fixation (Codispoti, 2007), suggesting imbalances in the cycling of nitrogen in the ocean. However, some models suggest that areas of N2 fixation and denitrification are coupled and could result in a homeostatic nitrogen cycle (Deutsch et al., 2007). Among marine diazotrophic microorganisms, Trichodesmium has been considered as the principal N2 fixer in the ocean. However, recent discoveries recognized unicellular diazotrophic cyanobacteria as important contributors to the PART II: Results 41 oceanic nitrogen budget (Zehr et al., 2001). Although the presence of noncyanobacterial nifH genes has been reported in the oligotrophic oceans (Falcón et al., 2004; Langlois et al., 2005; 2008), their contribution to overall N2 fixation requires further study. While Trichodesmium seems to be restricted to tropical oligotrophic areas (Capone et al., 1997), the unicellular cyanobacteria of groups A (UCYN-A), B and C may be more widely distributed. In the Atlantic Ocean, Langlois et al. (2008) found unicellular diazotrophs from the equator to ~35°N. Many of the group A and group B sequences were highly similar to those found previously in the Pacific Ocean, suggesting that these unicellular diazotrophs have a cosmopolitan distribution (Langlois et al., 2005). Unicellular oceanic diazotrophs have been documented in the North Pacific (Zehr et al., 2001), South Pacific (Moisander et al., 2010), the Mediterranean Sea (Le Moal and Biegala, 2009) and the Arabian Sea (Mazard et al., 2004). Organisms of group A (UCYN-A) are the most abundant and widely distributed unicellular diazotrophs in the open ocean (Langlois et al., 2005; Church et al., 2008; Langlois et al., 2008; Moisander et al., 2010). Although present mainly in tropical latitudes, UCYN-A occur in waters with a wide range of temperatures (Moisander et al., 2010); they are found in surface waters but also occur at greater depths where temperature and light intensity are lower and the concentration of inorganic nitrogen is higher (Montoya et al., 2004; Moisander et al. 2010). Organisms of groups B and C seem to occupy more narrow ranges of temperature (Langlois et al., 2005; 2008). An increasing number of studies have reported on the distribution and abundance of diazotrophs, but only few have also measured N2 fixation. Diazotrophy in the ocean is thought to be limited by iron and/or phosphorus (Mills et al., 2004; Moore et al., 2009). The Northeast Atlantic is expected to have high rates of N2 fixation because of the recurrent input of iron and phosphorus through aeolian transport from the nearby Sahara desert (Prospero, 1981). However, N2 fixation in this area could also be restricted as a result of the cold, nutrient-rich upwelling waters from the Iberian−Canary Current system, which extend hundreds of kilometers offshore through upwelling filaments (Álvarez-Salgado et al., 2007). Both the availability of combined nitrogen and the low temperatures could prevent N2 fixation, limiting its relevance in upwelling regions—as happens in the cold, high-latitude seas (Gruber and Sarmiento, 1997). The majority of the N2 fixation studies conducted in the Northeast Atlantic used a geochemical approach (e.g. Mahaffey et al., 2003; Álvarez and Álvarez-Salgado, 2007; Bourbonnais et al., 2009). The role of diazotrophy in nitrogen cycling in the Northeast Atlantic Ocean remains unclear and more direct measurements of N2 Chapter 1: N2 fixation in the subtropical Northeast Atlantic 42 fixation are needed. We measured N2 fixation by size-fractionated plankton from the Northeast Atlantic in samples from oligotrophic open ocean waters and from two active coastal upwelling regions of the Canary Current Large Marine Ecosystem (Arístegui et al., 2009): Cape Silleiro (northwest Iberia) and Cape Ghir (northwest Africa). The aims of this study were (1) to provide a first estimate of the diazotrophic activity in two active upwelling areas of the Northeast Atlantic, and (2) to compare the relative contribution of Trichodesmium with that of smaller diazotrophs. We combined the two most commonly used methods to measure (1) gross N2 fixation, using the acetylene reduction assay (ARA) (Stal, 1988), and (2) net N2 fixation, using the 15N2 tracer technique as employed by Montoya et al. (1996) (Gallon et al., 2002; Mulholland et al., 2004). The difference between the rates obtained by these two approaches is thought to be a proxy for the release of dissolved organic nitrogen (DON) (Gallon et al., 2002; Mulholland et al., 2004). DON fuels an important amount of autotrophic production (Berman and Bronk, 2003; Bronk et al., 2007), and some diazotrophs may release up to 50% of their recently fixed N2 as DON (Mulholland and Bernhardt, 2005); therefore, the potential influence of DON in supporting primary production and global nitrogen cycling should not be overlooked (Bronk et al., 1994; Mulholland, 2007). Materials and methods Sampling and hydrographic measurements The sampling was carried out within the framework of the project ‘Shelf−Ocean Exchanges in the Canaries−Iberian Large Marine Ecosystem’ (CAIBEX) on board the R/V Sarmiento de Gamboa during the summer of 2009. The cruise was divided into three legs: 2 mesoscale experiments off Cape Silleiro, northwest Iberia (6 to 24 July), and off Cape Ghir, northwest Africa (16 August to 5 September), and a large-scale open-ocean grid (CAIBOX, 25 July to 14 August) connecting the two cape regions (Fig. 1). Cape Silleiro and Cape Ghir are sites where upwelling filaments typically develop (Arístegui et al., 2009). However, the filaments of Cape Silleiro did not develop at the time of the cruise, so the sampling stations corresponded only to sites of coastal upwelling (stations S05 to S08) and to the open ocean (stations S01 to S03). A strong upwelling filament developed during the Cape Ghir cruise, allowing tracking and sampling of the structure during six consecutive days. Samples were collected along the path of the filament at stations G12, G17, G22, and G40, G44, G48, as well as at two stations outside the filament (stations GM2 and GT2). Finally, PART II: Results 43 17 additional stations were sampled along the box-grid of the CAIBOX cruise, connecting the northwest Iberia upwelling with the Canary Islands waters through a meridional section at 20°N (Fig.1). Fig. 1. Sampling stations superimposed on sea surface temperature (SST) maps obtained from AquaMODIS. Cape Silleiro SST data were averaged from 11 to 20 July 2009, CAIBOX from 2 to 4 July (dates when the ship sailed over the Azores Front), and Cape Ghir represents day 26 August (when station GT2 was sampled). More detailed descriptions of the hydrographic features found during these cruises may be found elsewhere (Carracedo et al., 2012; Troupin et al., 2012). Temperature, salinity, fluorescence and photosynthetically active radiance (PAR) data were recorded by a CTD SeaBird 911 plus, a Sea-Tech fluorometer and a LiCor PAR sensor, all mounted on a General Oceanics 24 bottle rosette sampler. Chapter 1: N2 fixation in the subtropical Northeast Atlantic 44 Plankton net casts and counts of Trichodesmium At each station, a plankton net of 50 μm mesh size was deployed two or three times from the deep chlorophyll maximum (DCM) and hauled vertically to the surface at a mean speed of 20 m min−1. Typically, 20 to 60 m3 were filtered by the net, depending on the depth of the DCM and the number of tows. The sample was concentrated to 240 mL, of which 60 mL were fixed with 10% formaldehyde to a final concentration of 4%. The formaldehyde was buffered with phosphatebuffered saline (PBS, Sigma Aldrich) and adjusted to pH 8.5. The samples were stored in the dark at ambient temperature until used for counting Trichodesmium in the laboratory with an inverted microscope. The number of free trichomes ranged from 1 to 1200 per sample. The colonial tufts or puffs were present in low numbers (typically 1 to 4 per sample). The other 180 mL of the plankton net sample were used for measuring N2 fixation in the > 50 μm fraction (see next section). Measurements of N2 fixation Water from the near-surface (5 m) was collected at each station between 08:00 h and 12:00 h (UTC). N2 fixation was measured by the ARA technique (Stal, 1988) and the 15N2 stable isotope technique (Montoya et al., 1996). The incubations for both techniques were done in on-deck incubators cooled with surface seawater. Neutral density screens (Lee Filters) were used to reproduce the incident PAR light measured at 5 m depth. ARA incubations were 3 to 4 h, and 15N2 incubations were for 24 h. For the ARA, triplicate samples of bulk seawater, each of 2 L, were filtered through 25 mm Whatman GF/F filters to obtain total N2 fixation rates. The <10 μm fraction was obtained by pre-filtering whole seawater through 47 mm white polycarbonate filters of pore size 10 μm (GE-Osmonics Poretics) and subsequently filtering onto GF/F filters. The activity of the >10 μm fraction was computed by subtracting the rates of the <10 μm fraction from the total N2 fixation rates. All GF/F filters were placed in 10 mL crimp-top vials (Varian/Chrompack), humidified with 0.5 mL of filtered (GF/F) seawater, and sealed with a rubber stopper and an aluminium cap using a seal crimper. After sealing, 2 mL of acetylene were injected into each of the samples using gas-tight syringes (Hamilton). Acetylene was generated from calcium carbide (CaC2, Sigma Aldrich) by adding Milli-Q water in a reaction flask (Stal, 1988). The gas was recovered in tedlar gas bags (volume 1 L) with polypropylene valves (SKC). The ethylene contamination of the generated acetylene was <12 ppm. PART II: Results 45 Blank GF/F filters were incubated with the same volume of filtered seawater and acetylene. A set of 3 blanks was made once every 5 stations. As we corrected ARA rates with their respective blanks every 5 stations, we obtained specific detection limits for each set of stations. The detection limit of the ARA technique ranged between 0.0185 and 0.1268 nmol ethylene, defined as three times the standard deviation of the difference between the ethylene produced after the incubation period and the ethylene content of the blanks. We divided the 180 mL of sample left over from the plankton net casts (see above) into three parts, each filtered through a GF/F filter and processed for the ARA as described above. It was assumed that this activity corresponds to the N2 fixation attributed to Trichodesmium or to endosymbiont cyanobacteria, such as Richelia intracellularis, which is the typical endosymbiont of the diatoms Rhizosolenia spp. or Hemiaulus spp., which range in size from 15 to 100 μm and from 15 to 35 μm, respectively. After incubation, 10 mL of headspace were sampled and transferred to preevacuated Hungate tubes, which were finally sealed with thermofusible glue and stored in the dark at ambient temperature until analysis. In previous tests, the retention of gas in Hungate tubes was found to be greatly enhanced after sealing with thermofusible glue, so this approach was used in the present study. Acetylene and ethylene were measured using a gas chromatograph (Agilent Technologies, model HP 5890) equipped with a flame ionization detector (FID), fitted with a Varian (Middelburg) wide-bore column (ref. CP7584) packed with CP-PoraPLOT U (27.5 m length, 0.53 mm inner diameter, 0.70 mm outer diameter, 20 μm film thickness). The column flow rate was 4 mL min−1 at a pressure of 5 psi. Helium was used as a carrier gas at a flow rate of 30 mL min−1. Hydrogen and airflow rates were set at 30 mL min−1 and 365 ml min−1, respectively. Helium and hydrogen were obtained from Carburos Metálicos (Air Products Group). Oven, injection and detector temperatures were set at 52, 120 and 170°C, respectively. Acetylene reduction rates were calculated using acetylene as an internal standard (Stal, 1988) and converted to N2 fixation rates using a factor of 4:1. Daily rates were computed from hourly rates multiplied by the number of light hours in each specific date and geographic position. For the 15N2 technique, unfiltered surface and <10 μm seawater (pre-filtered with polycarbonate filters as detailed above) was transferred to transparent polycarbonate bottles (volume 1.24 L, Nalgene). The bottles were completely filled using silicone tubing to prevent the introduction of air bubbles. They were then sealed with septum screw-caps before trace additions of 15N2 (2 ml 99 atom %; Tracer Tec) were injected through the septum using a gas-tight syringe. Enrichments varied from 9.8 to 11.2%. The pressure across the septum was Chapter 1: N2 fixation in the subtropical Northeast Atlantic 52 Fig. 4. Box plot summary of the (a) >10 μm fraction and (b) <10 μm fraction contribution to total net N2 fixation rates (%). Associated measured C2H4:15N2 ratios are presented as black circles. Error bars represent the range of values recorded. The upper and lower limits of the boxes represent the upper and lower quartiles, respectively. Bands crossing the boxes represent the median. Indeed, the densities of Trichodesmium reported in the literature for the Northeast Atlantic typically range from 0 to 100 trichomes L−1 (Tyrrell et al., 2003; Moore et al., 2009; Fernández et al., 2010; González-Taboada et al., 2010; present study), while densities in the Northwest Atlantic easily reached 100 to 1000 trichomes L−1 (Capone et al., 1997; Mahaffey et al., 2003). Few studies have addressed N2 fixation in upwelling areas. Staal et al. (2007) measured N2 fixation from 14°N to 13°S along the west African coast (1 to 20°N). They did not find N2 fixation in the areas affected by the coastal upwelling PART II: Results 53 (roughly 20 to 15°N). These authors found average N2 fixation rates ranging from 2.2 to 3.7 μmol N m−2 d−1, which is in the same range as reported here. Integrating our volumetric ARA values to the depth of the mixed layer (MLD), we calculate an average total ARA-derived N2 fixation rate of 2.14 ± 0.99 and 6.08 ± 7.73 μmol m−2 d−1 in the upwelling areas of Cape Silleiro and Cape Ghir, respectively. Staal et al. (2003) argued that a high temperature is needed to prevent oxygen deactivation of nitrogenase. This would be even more crucial for unicellular cyanobacteria because of their higher surface-to-volume ratio. In contrast to this paradigm, Raimbault and Garcia (2008) found N2 fixation rates as high as 3.6 nmol N L−1 d−1 off the Chilean upwelling system, where the water temperature was ~15°C. These authors did not detect the presence of Trichodesmium and therefore they attributed the measured N2 fixation rates to nanoplanktonic and picoplanktonic diazotrophs. In our study, N2 fixation in the small size fraction (< 10 μm) was always < 0.4 nmol N L−1 d−1 except for a peak of 0.98 nmol N L−1 d−1 at one station off Cape Ghir (Fig. 3c). The contribution of the small size fraction was generally higher in these colder and nutrient-richer waters of the northwestern Iberian and northwestern African upwelling systems than it was in the open-ocean (CAIBOX). Molecular biological studies performed during the CAIBEX cruises have confirmed that 66% of the < 3 μm clones sequenced belonged to the UCYN-A group (N. S. R. Agawin et al. unpublished results). Moreover, we cannot exclude the possibility that heterotrophic diazotrophs were present in the <10 μm fraction, although their importance for oceanic N2 fixation is debated (Riemann et al., 2010). The contribution of >10 μm and <10 μm diazotrophs to gross and net N2 fixation appeared to be highly variable across the studied areas (Table 2). The percentage contribution of each fraction to total release of DON did not correlate to either water temperature or concentrations of inorganic nutrients. DON release by diazotrophs may be influenced by physical and hydrographic factors (temperature, nutrients, light, turbulence), and biological factors such as the actual physiological state of the cells, interactions with the surrounding planktonic community (virus, bacteria, grazers), or varying direct uptake of DON by other phytoplankton. Studies performed to date indicate that the DON released by Trichodesmium may vary greatly among environments (see summary tables in Mulholland et al., 2006). Also, Trichodesmium might release DON when experiencing abrupt changes in temperature and/or light (Mulholland, 2007), which might have been the case in our study area, where hydrographic features such as upwelling filaments and fronts were found. Bronk (1999) found a higher release of DON in cultures of non- Chapter 1: N2 fixation in the subtropical Northeast Atlantic 54 diazotrophic unicellular cyanobacteria in nitrogen-replete conditions, in contrast to the release of dissolved organic carbon (DOC), which usually increases in nutrient-depleted situations. However, studies on the dynamics of DON release by unicellular diazotrophic cyanobacteria are lacking in the literature (Mulholland, 2007), and the mechanism of release of DON may be different in diazotrophic and non-diazotrophic cyanobacteria. The difference between N2 fixation derived from ARA and from 15N2 assimilation has been interpreted as the release of DON (Gallon et al., 2002; Mulholland et al., 2004; 2006). Notwithstanding this, new insight into N2 fixation techniques has demonstrated that the use of 15N2 bubbles potentially underestimates N2 fixation rates (especially in short incubations) due to slow dissolution of the 15N2 into the sample water (Mohr et al., 2010). The use of 15N2saturated water, instead of bubbles, might decrease the difference between the ARA and 15N2 approaches to some extent, but it will probably not invalidate the subtraction of ARA and 15N2 fixation as a proxy for DON release (see ‘Materials and Methods’). Nevertheless, this aspect deserves attention in future studies. We have demonstrated that organisms in the <10 μm fraction are more important than Trichodesmium for the fixation of N2 in the subtropical Northeast Atlantic Ocean. Gross rates of N2 fixation found in the upwelling zones were similar to those in the open ocean. This finding increases the latitudinal and habitat range of diazotrophic organisms, even when the rates were invariably low (<0.4 nmol N L−1 d−1). Acknowledgements This work was supported by project CAIBEX (CTM2007-66408-CO2-O2) to J.A. and N.S.R.A., a Ramón station Cajal grant and project CYFOD (CTM2008-00915E) to N.S.R.A., and an FPI fellowship (BES-2008-006985) to M.B., all from the Spanish Ministry of Science and Innovation (MICINN). We acknowledge V. Vieitez, M. Espino, Y. Santana and the analytical laboratory of CEME (NIOO-KNAW, Yerseke) for sample analysis, and C. Troupin for providing the Aqua-MODIS SST maps used in this work. We also thank the captain and crew of R/V Sarmiento de Gamboa, as well as the UTM for their kindness and professionalism. The authors thank three anonymous reviewers for their constructive comments and suggestions that helped us to improve the manuscript. This is publication 5121 of NIOO-KNAW. Table 2. Summary of average of gross and net N2 fixation, percentage of total N2 fixation lost as dissolved organic nitrogen (DON), and percentage contribution to total gross and total net N2 fixation by each fraction (>10 μm and <10 μm). Average rates or contributions by the >50 μm fractions have been included only where applicable (as no net N2 fixation by the > 50 µm fraction was assayed). Average gross N2 fixation (nmol N L-1 d-1) Average net N2 fixation (nmol N L-1 d-1) Fixed N2 lost as DON (% of total N2 fixation) Contribution to total gross N2 fixation (%) Contribution to total net N2 fixation (%) >10 <10 >50 (x10-4) Total >10 <10 Total >10 <10 >10 <10 >50 >10 <10 CAIBOX North of AF 0.03 0.04 1.47 0.07 0.002 0.04 41.1 99.7 24.2 40.25 59.8 0.2 5.7 94.3 CAIBOX South of AF 0.05 0.06 3.87 0.11 0.01 0.02 76.4 28.5 58.12 42.28 57.8 0.4 35.7 64.3 Cape Silleiro 0.04 0.10 3.11 0.09 0.05 0.08 44.1 42.7 64.07 29.65 70.4 0.5 26.3 73.7 Cape Ghir 0.01 0.24 4.03 0.25 0.03 0.08 54.1 14.9 89.73 7.62 92.4 0.2 12.8 87.2 !"#$%&'()*(#+%,-,.-#($#(/'0(1'23$'.4( ! "#$%&!'()(*(++%! !"#$%&'()( ,%--&./(*!&$0#)%1!)%+$&0()!#)*!1#$2&)!$(.(#-(!23!#! 4#$%)(!5)%1(..5.#$!*%#6&+$&78%1!13#)&2#1+($%54! ! '()#/%*(-!"9!:0#;%)!<=>9!:$?-+(05%!@9!A(()(!@9!=+#.!B@! :C5#+%1!"%1$&2%#.!D1&.&03E!F)!$(/%-%&)E! Chapter 2: DON and DOC release by unicellular diazotrophic cyanobacteria 58 Abstract Dinitrogen (N2) fixation rates may be underestimated when recently fixed N2 is released as dissolved organic nitrogen (DON). DON release (DONr) is substantial in the filamentous cyanobacterium Trichodesmium but has never been reported in unicellular diazotrophic cyanobacteria. In this study we used axenic cultures of the marine unicellular diazotroph Cyanothece sp. Miami BG 043511 to measure dissolved organic matter release under N2-fixing conditions. DONr was measured as the transfer of 15N2 from the culture medium to the extracellular DON pool. On average, the DON released represented ~1% of the total N2 fixed. The average release of dissolved organic carbon (DOC) as determined by 14C, represented ~2% of the total carbon fixed. These results suggest that cultured populations of unicellular diazotrophs do not release much dissolved organic matter, but it cannot be excluded that DONr is important in the field when grazers and bacteria are present, or when the organism is exposed to environmental stresses such as turbulence, excess light, temperature changes or nutrient limitation. Introduction The release of fixed dissolved organic carbon (DOC) by healthy marine plankton has been studied since the 1960s (e.g. Fogg, 1962). Release of organic matter in an oligotrophic environment might seem disadvantageous for phytoplankton as it represents a loss of energy and promotes growth of heterotrophic bacteria that compete for nutrients. Hence, these organisms may be outcompeted by those that do not release DOC. The organic carbon may serve as a substrate for autotrophic and heterotrophic plankton (Wood and Van Valen, 1990). Therefore, the release of organic compounds by healthy cells has often been regarded as a paradox in biological oceanography, although it may also have advantages. For example, organic compounds help chelating elements with low availability and solubility such as trace metals (Barbeau, 2006). The release of dissolved organic compounds also helps to protect the photosynthetic apparatus in high-light regimes by dissipating excess energy, and shortens the phytoplankton’s lag-phase after nutrient-impoverished periods (Wood and Van Valen, 1990; Wannicke et al., 2009). DOC release (DOCr) by phytoplankton has been widely studied during the past decades (Fogg, 1962; Sharp, 1977; Wood and Van Valen, 1990). Diazotrophic plankton such as blooming filamentous cyanobacteria are also known to release high amounts of DOC when the bloom decays, feeding grazers PART II: Results 59 and stimulating bacterial production (Sellner, 1997). DOCr has also been observed in unicellular cyanobacteria (Bertilsson et al., 2005). However, little is known about the magnitude and ecological significance of the release of dissolved organic nitrogen (DONr) by oceanic diazotrophic cyanobacteria. Most marine dinitrogen (N2) fixation field-studies have been conducted by measuring the 15N enrichment of particulate nitrogen (PN) after a certain time of incubation. The filtrates are usually discarded and therefore the N2 fixed and subsequently released as dissolved organic or inorganic nitrogen (DON or DIN) is not taken into account. Thus, the amount of N2 that has been fixed is potentially underestimated (Bronk et al., 1994; Gallon et al., 2002; Mulholland, 2007). Global oceanic denitrification exceeds N2 fixation by ~200 Tg N y-1 (Mahaffey et al., 2005; Codispoti, 2007). Recent research indicates that this disequilibrium could be less severe when N2 fixation measurements are expanded to higher latitudes and greater depths (e.g. Moisander et al., 2010), and when these measurements are indeed accurate, which has been questioned recently (Mohr et al., 2010). However, less attention has been paid to the potential underestimation of global oceanic N2 fixation rates caused by not taking into account DONr. Estimating DONr is also important because it is a source of new nitrogen for autotrophic and heterotrophic plankton (Berman and Bronk, 2003; Bronk et al., 2007). For instance, transfer of fixed N2 from large diazotrophic cyanobacteria to picoplankton has been described for the Baltic Sea (Ohlendieck et al., 2000) and for the Southwest Pacific (Garcia et al., 2007). N2 fixed by the filamentous cyanobacterium Trichodesmium can also sustain the growth of larger autotrophic plankton, such as diatoms (Lee Chen et al., 2010), or even promote harmful algae blooms, as for example those of Karenia brevis off the Florida shelf (Bronk et al., 2004). Agawin et al. (2007) studied interactions between N2-fixing and non-N2-fixing cyanobacteria in a coupled modeling-chemostat culture experiment and found that the DON released by the N2-fixing unicellular cyanobacterium Cyanothece induced a four-fold increase of the abundance of the non-diazotrophic Synechococcus than it would have achieved in monoculture. This suggests that organic compounds exuded by diazotrophs can fuel primary production and may exert considerable control on the composition of the plankton community in the oligotrophic ocean. Among marine diazotrophs, Trichodesmium was long thought to be the principal N2 fixer in the oceans (e.g. Capone et al., 1997). However, in the last decade molecular techniques revealed that unicellular diazotrophic Chapter 2: DON and DOC release by unicellular diazotrophic cyanobacteria 60 cyanobacteria are more abundant and more widely distributed than Trichodesmium (Luo et al., 2012), and their N2 fixation rates are often higher (Falcón et al., 2004; Garcia et al., 2007; Moisander et al., 2010). Field populations of Trichodesmium are known to release up to ~50% of the recently fixed N2 as DON (Glibert and Bronk, 1994), but in cultures the release is much less (Mulholland et al., 2004). Due to the wider geographical distribution of unicellular diazotrophs, it is necessary to study their DONr activity in order to estimate the potential underestimation of N2 fixation rates. However, direct estimates of DON release by unicellular diazotrophs have not been reported (Mulholland, 2007). DONr rates can be estimated as the difference between gross and net N2 fixation rates, measured by the acetylene reduction assay (ARA) and 15N2 assimilation into biomass, respectively (Gallon et al., 2002; Mulholland et al., 2004). Alternatively, one can measure the 15N atom % enrichment of the DON pool and use common tracer equations to estimate DON release rates (e.g. Glibert and Bronk, 1994; Mulholland et al., 2004). Both approaches have been used to indirectly estimate DONr by unicellular diazotrophs. Benavides et al. (2011) performed size-fractionated paired ARA and 15N2 assimilation measurements in the eastern North Atlantic and estimated that the <10 µm diazotrophs potentially released up to 67% of their recently fixed N2. These facts suggest that unicellular diazotrophs in the field release DON similarly as Trichodesmium. In this paper we used cultures of Cyanothece sp. Miami BG 043511 (hereafter Cyanothece) as a model organism to test whether the release of dissolved organic matter in unicellular diazotrophic cyanobacteria may be an important process. Materials and methods Culture conditions and experimental setup N2 fixation and DONr (experiment #1, 10 days), carbon fixation and DOCr (experiment #2, 8 days) were examined in this study. Cultures of Cyanothece sp. Miami BG 043511 (formerly classified as Synechococcus) were obtained from the Culture Collection Yerseke (strain CCY 0408). Cultures were grown in 250 mL transparent polypropylene tissue flasks with filter screw caps (VWR) in an illuminated incubator (Snijders ECD01E) at 27ºC under a 12-12 h light-dark cycle and a light irradiance of ~50 µmol photons m-2 s-2. Standard YBCII medium devoid of a source of combined nitrogen (Chen et al., 1996) was used in experiments #1 and #2. In both experiments two replicate cultures PART II: Results 61 were inoculated with 1% of an exponentially growing stock culture. Experiment #1 lasted 10 days and experiment #2 lasted 8 days. Therefore, 20 and 16 culture replicates were prepared for experiment #1 and #2, respectively. Every experiment day, two replicates were sacrificed for the assays and analyses described below. Chlorophyll, biomass, cell abundance and cell size During both experiments, chlorophyll a (Chl a) concentrations, cell abundance and cell size were monitored daily. For Chl a analysis, culture aliquots were filtered onto 25 mm GF/F filters. Chl a was extracted with 90% cold acetone for 24 h and subsequently analyzed by means of a Cary Eclipse fluorescence spectrophotometer, previously calibrated with pure Chl a (Sigma), and concentrations calculated using the equations in Ritchie (2006). Cell counts and cell size measurements were performed daily in triplicate vortexed fresh samples using a Multisizer 3 Coulter Counter (Becton Dickinson). Bacterial contaminant numbers were checked daily by phase contrast microscopy and did not exceed 3% of the cell number of the cyanobacteria throughout the experiments. N2 fixation Net N2 fixation was assayed once a day in the middle of the dark period using the stable isotope tracer method by Montoya et al. (1996). For this technique, 125 mL glass vials were filled with culture to overflow, closed with rubber stoppers (allowing the excess culture to escape by a sterile syringe tip piercing the septum) and crimp-sealed with aluminum caps. Trace additions of 15N2 (500 µL; 98.3 at.% 15N, Euriso-top, Saint-Aubin, France) were made using a gas-tight syringe (Hamilton) and the samples were incubated for 3 h. The 15N2 incubated samples were filtered onto pre-combusted GF/F filters (6 h, 450ºC) to obtain the 15N enrichment and PN concentration. DONr and intracellular DON production (inDONp) In experiment #1, DONr was estimated as the transfer of 15N from the culture medium to the extracellular DON pool, using the ammonium diffusion technique by Slawyk and Raimbault (1995) to isolate the labeled DON, and equations in Slawyk et al. (1998). In this experiment, we also investigated the inDONp (i.e. N2 fixed as DON but not released during the incubation period), Table 2: Average concentrations of 15N in the particulate nitrogen (PN) and extracellular and intracellular dissolved organic nitrogen (exDON and inDON) pools, at the start and at the end of the incubations performed with additions of 15N2 to Cyanothece cultures (experiment #1). The sum of 15N in all pools at the start and at the end of the incubations is given. All values are given in µmol N L-1. For initial values, the concentration of 15N2 added to the samples is included in the sum as well (0.16 µmol N L-1). A percentage of 15N recovery is given based on the amount of 15N in the samples at the start of the incubations with the amount of 15N recovered in at their end. Standard deviation values are given in brackets. Initial Final Day PN exDON inDON Total (+15N2) PN exDON inDON Total % recovery 1 0.53 (0.09) 0.04 (0.00) 0.02 0.75 1.06 (0.02) 0.04 (0.0) 0.05 (0.01) 1.15 153.33 2 0.82 (0.1) 0.04 0.02 (2.59) 1.04 2.02 (0.63) 0.17 (0.19) 0.06 (0.07) 2.25 216.35 3 1.15 (0.19) 0.05 (0.01) 0.03 (0.01) 1.39 1.73 (0.03) 0.08 (0.02) 0.12 (0.04) 1.93 138.85 4 1.65 (0.34) 0.15 (0.06) 0.08 (0.02) 2.04 2.06 (0.06) 0.16 (0.04) 0.13 (0.02) 2.35 115.19 5 1.95 (0.04) 0.12 (0.03) 0.1 (0.01) 2.33 1.95 (0.32) 0.14 (0.02) 0.1 (0.09) 2.19 93.99 6 2.74 (0.38) 0.29 (0.06) 0.09 (0.00) 3.28 3.17 (0.01) 0.2 (0.01) 0.12 (0.04) 3.49 106.40 7 3.38 (0.21) 0.25 (0.12) 0.09 (0.04) 3.88 3.42 (0.55) 0.25 (0.02) 0.15 (0.04) 3.82 98.45 8 4.08 (0.22) 0.29 (0.03) 0.12 4.65 3.68 (0.38) 0.24 (0.0) 0.15 4.07 87.53 9 3.87 (0.15) 0.2 (0.03) 0.08 4.31 4.22 (0.19) 0.24 (0.0) 0.06 (0.01) 4.52 104.87 10 3.96 (0.79) 0.26 (0.07) 0.04 4.42 3.51 (0.05) 0.31 (0.14) 0.03 (0.01) 3.85 87.10 PART II: Results 69 The inDONp rates (transfer of 15N2 to the intracellular DON pool) were generally one order of magnitude higher than DONr, with values ranging from 0.003 to 0.2 µmol N L-1 h-1 when 100% dissolution of the 15N2 bubble was considered, and 0.008 to ~0.5 µmol N L-1 h-1 when only 28.8% dissolution is considered. The overall tendency of inDONp rates was to decrease along the experiment, and the lowest rates were observed during the last three days. Carbon pools, carbon fixation, DOCr and inDOCp Rates of net carbon fixation, DOCr and inDOCp, and the concentration of carbon in the particulate and dissolved pools of Cyanothece cultures during experiment #2 are given in Table 4. Carbon fixation rates are given either as net carbon fixed (carbon fixed into biomass), or as ‘total carbon fixed’ (net carbon fixed + DOC released + intracellular DOC produced). Total carbon fixation ranged from 54 to 151 µmol C L-1 h-1, close to net carbon fixation, which ranged from 37 to 101 µmol C L-1 h-1. The rate of DOCr rates ranged from <1 to 3 µmol C L-1 h-1, while the rate of inDOCp was one to two orders of magnitude higher, ranging from 15 to 55 µmol C L-1 h-1. DOCr rates represented only 1 – 4% of the total carbon fixed (average ~2%). Particulate carbon (PC) ranged from 950 to 5600 µmol C L-1. Extracellular DOC concentrations ranged from ~70 to 600 µmol C L-1. Intracellular DOC concentrations were generally higher than extracellular DOC concentrations, except on the last three days of experiment #2 (days 6 – 8). C:N fixation ratios The ratios provided here come from two different experiments (#1 and #2, see Materials and methods). C:N fixation ratios would be more accurate if both carbon and N2 fixation rates had been measured simultaneously in the same culture. However, experiment #2 was done independently because samples treated with radioactivity (14C) need to be manipulated in a separate laboratory and cannot be further processed by an elemental analyzer or IRMS. Carbon was fixed at higher rates than nitrogen, as reflected by the high molar ratios of net carbon fixation: net N2 fixation (mol C: mol N) (Table 5). C:N ratios increased along the experiment and exceeded Redfield in almost all experiment days. When net N2 fixation rates were calculated considering 100% dissolution of the 15N2 bubble, C:N ranged from 8 to ~40. Table 3: Average rates of net N2 fixation (A), dissolved organic nitrogen (DON) released (B), intracellular DON produced (C), and total nitrogen fixed (A+B+C) measured considering 100% dissolution of the 15N2 bubble, and, alternatively, considering 28.8% dissolution of the 15N2 bubble after a 3 h incubation period. All rates are given in µmol N L-1 h-1. The last column includes the percentage of DON released compared to the total N2 fixed (A+B+C), which is equal for both 15N2 bubble % dissolutions. Note that the total N2 fixed column does not include any N2 fixed and released as dissolved inorganic nitrogen (DIN). Standard deviation values are given in brackets. 100% dissolution of 15N2 bubble 28.8% dissolution of 15N2 bubble Day Net N2 fixation (A) DONr (B) inDONp (C) Total N2 fixed (A+B+C) Net N2 fixation (A) DONr (B) inDONp (C) Total N2 fixed (A+B+C) % DONr 1 3.66 (0.27) 0.00 (0.00) 0.06 (0.04) 3.72 9.46 (0.68) 0.01 (0.002) 0.16 (0.11) 9.62 0.08 2 7.36 (3.06) 0.07 (0.10) 0.15 (0.10) 7.59 18.9 (7.80) 0.19 (0.2) 0.39 (0.25) 19.25 0.98 3 4.85 (0.52) 0.04 (0.02) 0.10 (0.02) 4.99 12.3 (1.36) 0.09 (0.04) 0.26 (0.04) 12.51 0.72 4 4.28 (0.11) 0.03 (0.03) 0.20 (0.03) 4.51 10.8 (0.31) 0.08 (0.07) 0.49 (0.07) 10.95 0.69 5 3.44 (1.18) 0.03 (0.01) 0.18 (0.01) 3.65 8.62 (2.99) 0.07 (0.02) 0.45 (0.02) 8.75 0.76 6 3.47 (0.38) 0.03 (0.01) 0.06 (0.01) 3.56 8.60 (0.97) 0.08 (0.03) 0.15 (0.03) 8.77 0.95 7 2.76 (1.01) 0.04 (0.01) 0.13 (0.01) 2.93 6.78 (2.47) 0.09 (0.01) 0.33 (0.01) 6.97 1.35 8 3.08 (0.19) 0.06 (0.02) 0.01 (0.02) 3.15 7.51 (0.48) 0.14 (0.05) 0.02 (0.05) 7.79 1.78 9 2.90 (0.08) 0.03 (0.01) 0.00 (0.01) 2.94 7.02 (0.16) 0.08 (0.03) 0.01 (0.03) 7.18 1.09 10 1.83 (0.21) 0.02 (0.01) 0.00 (0.01) 1.85 4.4 (0.52) 0.05 (0.02) 0.01 (0.01) 4.49 1.01 Table 4: Average rates of net carbon fixation (A), rate of release of dissolved organic carbon (DOCr) (B), and rate of production of intracellular DOC (inDOCp) (C), as well as the rate of the total carbon fixed (sum of A, B and C) measured using 14C. All rates are given in µmol C L-1 h-1. The percentage of carbon fixed which is released as DOC is given. The three final columns represent the concentration of carbon in the particulate carbon (PC), extracellular and intracellular DOC pools (exDOC and inDOC) at the end of the incubation period. Concentrations are expressed in µmol C L-1. Standard deviation values are given in brackets. Day Net carbon fixed (A) DOCr (B) inDOCp (C) Total carbon fixed (A+B+C) % DOCr PC exDOC inDOC 1 37.47 (4.1) 2.41 (0.12) 14.55 (3.6) 54.43 (5.8) 4.43 948.92 (0.4) 71.39 (17.7) 299.97 (62.9) 2 61.38 (7.7) 0.70 (0.4) 37.44 (7.6) 99.52 (12.9) 0.70 1119.28 (0.03) 75.42 (1.7) 108.84 (39.3) 3 89.31 (7) 0.96 (0.3) 54.80 (6.32) 145.07 (11.8) 0.66 1651.62 (0.17) 111.60 (6.7) 172.30 (1.9) 4 82.62 (10.3) 1.88 (0.2) 52.50 (1.2) 137 (16.9) 1.37 2117.82 (0.13) 160 (2) 306.50 (8.7) 5 97.6 (4.5) 3.11 (0.8) 46.10 (2) 146.81 (7.8) 2.11 2735.87 (1.41) 274.77 (11.3) 559.24 (203.7) 6 84.7 (1.9) 2.54 (0.2) 46.04 (2.8) 133.28 (3.4) 1.91 4373.13 (0.24) 324 (28.2) 282.50 (172.6) 7 101 (2.7) 2.70 (0.1) 47.52 (12.2) 151.22 (4.1) 1.79 5615.35 (0.1) 459.10 (45) 360.90 (40.4) 8 98.59 (9.01) 2.43 (0.2) 49.34 (14.5) 150.36 (13.8) 1.62 5543 (0.9) 599.24 (36.9) 297.75 (17.1) Chapter 2: DON and DOC release by unicellular diazotrophic cyanobacteria 72 When 28.8% dissolution of the 15N2 bubble was assumed, net N2 fixation rates were ~2.5 fold higher. Consequently, when 28.8% dissolution of the 15N2 bubble was applied C:N ratios were lower and did not exceed Redfield, except on days 1 and 2. From day 3 on, the C:N ratios ranged from 3 to ~15. DOCr : DONr ratios give an estimation of the C:N ratio of the dissolved organic matter released. These ratios were also high, ranging from ~10 to 116.3 when 100% dissolution of the 15N2 bubble was considered for the calculation of N2 fixation rates, and from ~4 to 46.5 when 28.8% dissolution was assumed. On day 1, the DONr rates were close to zero (Table 3), inflating DOCr : DONr ratios (Table 5). Discussion Measuring N2 fixation and nitrogen release: methodological issues There are a number of methodological issues which affect the estimation of N2 fixation and DONr rates: (1) potential underestimation of N2 fixation rates when the 15N2 bubble method is used, (2) unsuccessful recovery of all 15N added at the end of an incubation period, (3) methodological problems related to DON isolation for IRMS analysis, (4) artifacts associated to sample manipulation, and (5) intracellular isotopic dilution. Each will be discussed in detail below. Table 5: Net carbon fixation : net N2 fixation, and dissolved organic carbon release (DOCr) : dissolved organic nitrogen release (DONr) ratios (mol C:mol N). N2 fixation rates were calculated assuming 100% dissolution of the 15N2 bubble or, alternatively, assuming 28.8% dissolution of the 15N2 bubble after a 3 h incubation period. 100% dissolution of 15N2 bubble 28.8% dissolution of 15N2 bubble Day Net carbon fixation : net N2 fixation DOCr : DONr Net carbon fixation : net N2 fixation DOCr : DONr 1 10.24 3.89 830.1 321.3 2 8.33 3.19 9.5 3.7 3 18.42 7.11 27.0 10.6 4 19.29 7.50 62.7 24.8 5 28.34 11.11 116.3 46.5 6 24.44 9.66 75.6 30.5 7 36.62 14.61 70.6 28.7 8 32.02 12.87 42.6 17.5 PART II: Results 73 (1) Recently, Mohr et al. (2010) demonstrated that the routinely used 15N2 bubble tracer method underestimates N2 fixation rates due to the slow dissolution of the gas into the medium or the sample seawater. These authors recommended adding 15N2 already dissolved in the water instead of as a bubble, as previously done by Glibert and Bronk (1994), in order to provide a constant 15N enrichment of the source N2 pool. Mohr et al.’s (2010) study was published while the experiments reported in this paper were being carried out. As the experiments had already started, we chose not to change the methodology to maintain consistency. In order to give an approximation of what the rates would have been like using more realistic dissolution of 15N2 after 3 h incubation, we recalculated net N2 fixation, DONr and inDONp rates assuming 28.8% dissolution of the 15N2 (Table 3), which is the dissolution of 15N2 observed after 3 h of incubation in YBCII medium at 27ºC (W. Mohr, personal communication). When 28.8% dissolution is applied, net N2 fixation rates explain better the daily PN increase observed (Tables 1 and 3), corroborating that the 15N2 bubble method underestimates N2 fixation rates. (2) Another problem often encountered during 15N experiments is that the label is not fully recovered. Bronk and Glibert (1994) demonstrated that the fate of this ‘missing’ 15N differs between marine systems. In particular, 15N transfer from the source pool to the extracellular DON pool seems to be more important in oligotrophic systems, which supports the need to include DONr measurements in routine nitrogen uptake experiments in order to avoid underestimations (Bronk et al., 1994). (3) Measuring the 15N enrichment of the DON pool is prone with difficulties. Extracting DON for isotope analysis is difficult and has prompted intensive discussion in the past (Slawyk et al., 1998; Bronk and Ward, 2000; Slawyk et al., 2000). The protocol for extracting DON used here (Slawyk and Raimbault, 1995) (see Materials and methods) could underestimate DONr because the alkaline hot experimental conditions (60ºC) of the extraction process may hydrolyze some fraction of the DON as ammonium (McCarthy and Bronk, 2008). The alternative DON isolation method is the ion retardation column (Bronk and Glibert, 1991), which is no longer commercially available in its previous quality and now retains variable amounts of DON, and therefore is not reliable (McCarthy and Bronk, 2008). Hence, consistent DON extraction protocols await development. (4) Other methodological problems may affect the measurement of DONr, such as the breakage of cells during vacuum filtration and exposure to light and temperature changes during sample manipulation (Feuillade et al., 1990; Wannicke et al., 2009). Vacuum pressure was kept low throughout our Chapter 2: DON and DOC release by unicellular diazotrophic cyanobacteria 74 experiments (<100 mm Hg) and care was taken to avoid temperature and light changes. Light was kept constant in our experiments (~50 µmol photons m-2 s-2), and was well below natural intensities in tropical and subtropical waters where Cyanothece occurs (Langlois et al., 2008). We therefore think that culture settings or sample manipulation did not affect DONr analysis. (5) The intracellular isotopic dilution in cells with high intracellular DON pools might have affected the measurements of the DONr rates. When the intracellular DON pool and its turnover time are high, isotopic equilibrium between the intracellular pool and the extracellular medium is slow. This causes artificially low DON release rates, as the DON recovered in the extracellular pool is 14N-rich material, which was inside the cell previously to 15N2 addition. (Mulholland et al., 2004). Nitrogen versus carbon metabolism in Cyanothece Diazotrophs are known to fix carbon at rates exceeding the C:N Redfield ratio, (Mulholland et al., 2006; Mulholland, 2007). Reasons why this phenomenon occurs in natural populations include (1) substantial nitrogen and/or carbon release due to methodological problems associated to cell manipulation, (2) excess carbon fixation (this ballast may cause sinking of the cells), and (3) excess carbon fixation to decrease cellular oxygen concentrations which may inhibit the nitrogenase (Mehler reactions), or for ATP production (Mulholland, 2007). Errors in global estimates of N2 fixation would be lessened if accurate C:N fixation ratios of marine diazotrophs were used instead of theoretical elemental stoichiometries (Mulholland, 2007). The DON released during our experiments was similar to the DOC released when expressed as a percentage of the total nitrogen or carbon fixed. On average, Cyanothece released ~1% of the recently fixed gross N2 as DON, while ~2% of the gross carbon fixed was released as DOC. In a series of culture experiments, Wannicke et al. (2009) found that cultured Trichodesmium released 71% of the fixed N2 as DON and ammonium and 50% of the carbon fixed as DOC. In our study DOCr/DONr C:N ratios were high and always exceeding Redfield stoichiometry, with the only exception of day 2 when N2 fixation rates were calculated using 28.8% dissolution of the 15N2 bubble (Table 5). These high ratios suggest that the dissolved organic matter released by Cyanothece is enriched in carbon. It should be noted that DOC and DON were treated as independent pools, but the exudate material of natural populations of diazotrophic organisms is normally composed of both DOC and DON. For example, this is the case for PART II: Results 75 amino acids (Gallon et al., 2002). Whereas eukaryotic phytoplankton organisms are known to exude dissolved organic compounds lacking nitrogen such as carbohydrates (Newell et al., 1972), diazotrophic cyanobacterial exudates are commonly rich in nitrogen (Capone et al., 1994; Mulholland, 2007). DONr by unicellular diazotrophic cyanobacteria Trichodesmium is thought to release fixed N2 to provide other trichomes in the colony with nitrogen (Mulholland and Capone, 2000), because not every cell in a trichome and not all trichomes express nitrogenase (Bergman and Carpenter, 1991), or are in the same phase of the cell cycle (Wannicke et al., 2009). Some unicellular cyanobacteria release organic compounds to feed neighboring cells. For example, Gloeothece cells live embedded in a mucilaginous sheath that acts as an ‘extracellular vacuole’ (Flynn and Gallon, 1990). The advantage of releasing DON by free-living unicellular diazotrophs is not clear, if there is any. Possibly free-living unicellular diazotrophs release DON as a response to abiotic factors such as light stress (Wood and Van Valen, 1990). In our study, the DON released represented on average ~1% of the total N2 fixed by Cyanothece during experiment #1 (Table 3). Given that optimum culture conditions were used, the growth of the cells was balanced and DONr was negligible. Cultured nitrogen-starved eukaryotic phytoplankton have been reported to release dissolved organic compounds when reaching stationary phase (Newell et al., 1972). Contrarily, in our study similar levels of DONr and DOCr were observed throughout the experiment (Tables 3, 4). A decrease in DONr rates was observed during the last three days of experiment #1, when the cells were reaching the stationary phase (Fig. 1). We hypothesize that this decrease of DONr rates might have been caused by the increasing availability of DIN and DON in the medium during the last days of the experiment #1, and/or the simultaneous uptake of DON by Cyanothece during the experiment (Bronk and Glibert, 1993). Similarly, Bronk (1999) observed a decrease in DONr rates in Synechococcus cultures when the cells became nutrient limited. Cyanothece cells were incubated under a 12-12h light-dark regime and DONr rates were measured during the dark phase. However, we were unable to quantify how much of the DON released during the dark phase was taken up during the next light period. The percentages of fixed N2 released as DON obtained in this study are low (~1% of total N2 fixation), even lower than those obtained with cultures of Chapter 2: DON and DOC release by unicellular diazotrophic cyanobacteria 76 Trichodesmium IMS101 (~8%; Mulholland et al., 2004), and much lower than in field studies of Trichodesmium (~50%; Glibert and Bronk, 1994). To our knowledge, there are no reports on DONr by unicellular diazotrophs (Mulholland, 2007). Recently, we measured DONr by the <10 µm planktonic fraction in surface waters of the North Atlantic Ocean and estimated that on average ~20% of the recently fixed N2 was released extracellularly as DON (M. Benavides, submitted). The <10 µm fraction in natural waters may contain a variety of different diazotrophs, probably mostly UCYN-A and Crocosphaera which are dominant in this area of the Atlantic (Langlois et al., 2008). Unfortunately, we cannot discern which diazotrophs release DON and which do not from a <10 µm planktonic sample. However, this difference in DONr dynamics between natural and cultured of unicellular diazotrophs is comparable to that observed by Glibert and Bronk (1994) and Mulholland et al. (2004) in natural and cultured populations of Trichodesmium, respectively. The constantly growing evidence of the diversity and wide distribution of oceanic unicellular diazotrophs emphasize the need for accurate N2 fixation rates and to provide better estimates of DONr by these organisms. Among the oceanic unicellular diazotrophs, UCYN-A are the most abundant (Luo et al., 2012). These cyanobacteria lack the genetic material to fix carbon, which makes them depend on compounds produced by other organisms (Tripp et al., 2010). Therefore, it is unlikely that they release any of their recently fixed N2 unless being under some sort of environmental or predatory stress. However, recent research on extracellular polysaccharides (EPS) production by Crocosphaera (Sohm et al., 2011a) suggests that the DONr dynamics in oceanic UCYN-B diazotrophs could be important. We chose Cyanothece as a model organism because it is easier to grow in cultures, facilitating experiments. This diazotroph is similar to Crocosphaera in size (~2-6 µm) and in their nitrogenase activity patterns. Thus, their organic matter excretion metabolism could be indeed comparable. However, UCYN-A and Crocosphaera are much more abundant in the open ocean than Cyanothece (Luo et al., 2012). Therefore, DONr should be measured in these other species before we draw any conclusion of the importance of DONr in underestimating global N2 fixation rates. Acknowledgements We are indebted to M. Houtekamer, C. Kleppe van Zettenand, Lennart van IJzerloo and the rest of the analytic laboratory team at NIOZ-Yerseke. We also thank J. Ly, J. Kromkamp, H. Bolhuis, H.T.S. Boschker and S. Malkin for PART II: Results 77 helpful advice on the setup and development of the experiments. We thank D.A. Bronk for many valuable comments on earlier versions of this manuscript. We also thank A. Santana for his help with data analysis. This work was supported by project CAIBEX (CTM2007-66408-CO2-O2) to J.A., a Ramón y Cajal grant and project CYFOD (CTM2008-00915-E) to N.S.R.A., and a FPI fellowship (BES2008-006985) to M.B., all from the Spanish Ministry of Science and Innovation (MICINN). Chapter 3: Longitudinal variability of size-fractionated N2 fixation and DON release 84 Therefore, pre-fractionation is necessary to study DONr dynamics by each size fraction separately. We sampled one station per day but alternated between day (0900-1200) and night samplings (1900-2100, local time) (Figure 1a). This approach allowed us to study the diazotrophic activity of organisms that fix N2 in the light (e.g. Trichodesmium, heterotrophic diazotrophs), and organisms that only fix N2 in the dark to avoid oxygen deactivation of the nitrogenase enzyme system (e.g. Crocosphaera). N2 fixation and DON release rates were measured using 15N-labeled N2 gas during 3 to 4 h incubations, which is the average incubation time frequently used in other DONr studies (e.g. Glibert and Bronk, 1994; Bronk, 1999). In this kind of experiments, incubation times must be long enough to allow sufficient 15N label to reach the DON pool (according to the detection limit of the isotope ratio mass spectrometer -IRMSequipment used), and short enough to protect the cells from usual bottle-effects. There are two general approaches used to add the labeled gas to a sample: the addition of water that was saturated with 15N2 (e.g. Glibert and Bronk, 1994; Mohr et al., 2010), and the addition of a bubble (Montoya et al., 1996). In this study we used the addition of saturated water in fractionated seawater samples and the bubble method in whole (unfiltered) seawater samples (see below). With the addition of a bubble, the N2 fixation rate is potentially underestimated due to slow dissolution of the gas bubble in water (Mohr et al., 2010). This is particularly true in short-term incubations where the gas bubble may not have sufficient time to equilibrate with the sample. If the gas bubble does not equilibrate, the actual atom percent (atom %) enrichment of the available N2 pool will be lower than calculated resulting in an underestimation of the fixation rate. To prepare the 15N2-saturated water, surface seawater (~5 m) was recovered from the flow-through system of the ship and filtered through a 47 mm GF/F filter. This was done during the upcast of the CTD to ensure that the 15N2saturated water was the same as the sample water. This filtered seawater was decanted into 0.5 L transparent polycarbonate bottles (Nalgene) and degassed as outlined in Mohr et al. (2010). Each bottle was filled to overflow, closed with a septum screwcap and 5 mL of 15N2 (99 at.% 15N; Tracetec) were injected using a Hamilton gas-tight syringe. The bottles were vigorously shaken for 10 - 20 min and then kept in the on-deck incubators until the rosette was back onboard (0.5 - 2.5 h depending on the station depth) to ensure that the 15N-enriched seawater had the same temperature as the sample at the time of mixing, in order to protect the organisms from thermal shocks. To check the real 15N enrichment of the 15N2-amended seawater added to PART II: Results 85 the samples, replicate samples of the 15N2-amended seawater prepared onboard were taken in 10 mL crimp vials (Chrompack), filled to overflow, and sealed with teflon-lined stoppers and aluminum caps using a seal crimper. These were stored at room temperature in the dark until being analyzed by membrane inlet mass spectrometry (MIMS) in S. Joye’s lab. MIMS analysis showed that the real enrichment of the 15N2-amended seawater was ~100% (99 ± 2%) of that expected from complete dissolution of the 15N2 bubble. To collect water used to measure fractionated rates, near surface (~5 m) seawater was collected with a 30 L Niskin bottle at each station. The samples were pre-fractionated by filtering two 2 L replicates through a 10 µm Nitex custommade sieve. The <10 µm fraction was recovered in 2.4 L transparent polycarbonate bottles (Nalgene). The >10 µm fraction was recovered from the sieve by gently washing and concentrating the biological material using a water sprayer filled with filtered (GF/F) seawater from the same station. Then 200 mL of 15N2-enriched filtered seawater was added to all bottles, they were filled to their neck with filtered seawater from the same station and placed in the on-deck incubators for 3 - 4 hr. The incubators were connected to the ship’s flow-through system and covered with neutral density screens (Lee Filters). After the incubation, the samples were filtered through precombusted 25 mm GF/F filters, which were subsequently stored in sterile cryovials (VWR) and frozen. The GF/F filtrates were then filtered through 0.2 µm polycarbonate filters (GE-Osmonics Poretics) using gentle vacuum pressure (<100 mm Hg) to remove bacteria prior to isolating the dissolved N pools. Rigorous care was taken to avoid light stress in the samples. The filtrates stored frozen in triplicate 50 mL polypropylene tubes (VWR) and used to measure the concentrations of NO3-, NO2-, NH4+ and total dissolved N (TDN) as described below. Whole seawater net N2 fixation rates Parallel to seawater collection for fractionation, whole seawater was transferred to 2.4 L transparent polycarbonate bottles. The bottles were completely filled using silicone tubing to prevent the introduction of air bubbles. They were then sealed with septum screw-caps before 2 mL of 15N-labeled N2 gas were injected through the septum. The pressure across the septum was equilibrated by allowing the excess water to escape through a sterile syringe tip piercing the septum. The bottles were placed in the on-deck incubator for the same incubation period. After the incubation, samples were also filtered through precombusted GF/F filters, stored in cryovials, and frozen until IRMS analysis ashore. N2 fixation rates measured with the bubble method were calculated as outlined in Montoya et al. (1996). Chapter 3: Longitudinal variability of size-fractionated N2 fixation and DON release 86 Chemical analyses and rate calculations Nutrient pools in filtrates were analyzed in D. Bronk’s lab after the cruise. Concentrations of NH4+ were analyzed in duplicate with the manual phenol hypochlorite method (Hansen and Koroleff, 1983) using a Shimadzu UV-1601 spectrophotometer (detection limit of ~0.03 μmol L−1). Concentrations of NO3and NO2were determined with a Lachat QuikChem 8500 autoanalyzer (detection limits of 0.1 μmol L−1 and 0.03 μmol L−1, respectively). TDN concentrations were analyzed on a Shimadzu TOC-V equipped with a TNM module. DON concentrations were calculated by subtracting NO3-, NO2and NH4+ concentrations from TDN concentrations; propagation of error calculations were used to estimate errors associated with DON concentrations (Bronk et al., 2000). We also isolated the NO3-, NH4+ and TDN pools to determine their 15N atom % enrichment so that we could quantify rates of DON release. To isolate the NH4+ pool, we used solid phase extraction (SPE) columns (C18) (Dudek et al., 1986) and then spotted the isolated NH4+ on a filter prior to mass spectrometry analysis. The atom % of the particulate organic N (PON) and NH4+ pools were analyzed on a Europa GEO 20/20 IRMS with an automated nitrogen and carbon analyzer for solids and liquids (ANCA-SL). To isolate the TDN pool, 15 mL samples were oxidized to NO3via persulfate oxidation (Valderrama, 1981). The NO3was then converted to nitrous oxide (N2O) using denitrifying bacteria lacking N2O-reductase activity (Sigman et al., 2001). Isotope ratios of N2O were then measured using a ThermoFinnigan GasBench + PreCon trace gas concentration system interfaced to a ThermoScientific Delta V Plus IRMS at the University of California Davis Stable Isotope Facility (Davis, CA). The same approach without the persulfate oxidation was used to isolate the NO3pool. The 15N atom % enrichment of each of these pools was used to measure DON release rates as follows: atom%TDN x (TDN)=(atom%NO3x (NO3-))+(atom%NH4+ x (NH4+))+(atom%DON x (DON)) (1) where (TDN), (NO3-),(NH4+) and (DON) are the concentrations of TDN, NO3-, NH4+ and DON, respectively. Solving Equation (1), the 15N atom % enrichment of the DON pool was calculated as follows: PART II: Results 87 (2) N2 fixation and DON release rates were then calculated using standard tracer equations: (3) where atom% excess is the 15N% enrichment over natural abundance. N2 fixation rates obtained from 15N2 incorporation into biomass were considered ‘net’ N2 fixation rates. The sum of net N2 fixation rates and DON release rates were considered ‘gross’ N2 fixation rates (Mulholland et al., 2004). Results Hydrographic features and nutrient concentrations Low values of SST and SSS of ~19ºC and 36.8 respectively, were observed near the coastal upwelling off NW Africa (Figure 2). From east to west, SST increased up to 24.26ºC at 54.03ºW, and SSS up to 37.54 at 37.57ºW. Moving westwards, SST presented sharp decreases of 1 - 2ºC coincident with strong negative SSHA (<-12 cm). In contrast, SSS decreased until 62ºW (station 104), where it reached 36.28. From 62ºW to the west, SSS increased in two steps coincident with positive or low values of SSHA, and finally reached ~36.74. The surface concentrations of NO3- + NO2-, PO43and SiO2 were generally maximum at the eastern end of the transect (coinciding with the NW African coastal upwelling), and decreased towards the west. The lowest concentrations were observed west of ~45ºW, coinciding with the oligotrophic Sargasso Sea. Along with SST and SSS increments and nutrients high values, the first seven N2 fixation stations show positive SSHA values, which then decrease considerably until station 71 (44.5ºW). From this position to the west, high and predominantly negative SSHA dominate. The overall variability of SSHA along the transect can be observed in Figure 3. atom%DON = atom%TDN x [TDN] ( ) - (at%NO3 - x [NO3 -]) + (at%NH4 + x [NH4 +]) ( ) [DON] rate = atom% excess target pool atom% excess source pool x time x target pool concentration Chapter 3: Longitudinal variability of size-fractionated N2 fixation and DON release 88 Dissolved 15N2 versus 15N2 bubble net N2 fixation rates The sum of net N2 fixation rates in the >10 µm and <10 µm fractions measured using the dissolved method (Mohr et al., 2010) was compared to net N2 fixation rates (in whole seawater samples collected at the same stations) using the 15N2 bubble classic method (Montoya et al., 1996). The difference between both rates tended to increase towards the east (Figure 4a). Indeed, the percent underestimation of the bubble method was significantly correlated with longitude (rs = 0.518, p =0.001). The average percent underestimation of the bubble method was 49 ± 39%. Because dissolved organic matter (DOM) coating 15N2 bubbles could slow its dissolution in seawater (Mohr et al., 2010), we checked for any significant correlations between percentage underestimations and DON concentrations. The concentrations of DON in >10 µm and <10 µm samples did not correlate significantly with the percentage underestimation of N2 fixation rates, but followed a similar longitudinal distribution (Figure 4b). Fig. 2: Values of (a) sea surface height anomalies (SSHA), (b) sea surface temperature (SST) and sea surface salinity (SSS), and (c) surface concentrations of nitrate plus nitrite (NO3- + NO2-), phosphate (PO43-) and silicon (SiO2) (all in µmol L-1) at stations were N2 fixation experiments were performed. PART II: Results 89 N2 fixation and DON release estimated by the dissolved 15N2 method Both >10 and <10 µm organisms presented a similar range of N2 fixation rates (~0.01to ~0.44 nmol N L-1 h-1), although >10 µm rates were statistically different from <10 µm rates (Wilcoxon test, p<0.0001). On average, net N2 fixation rates of organisms >10 µm were slightly higher than those <10 µm (0.16 and 0.1 nmol N L-1 h-1, respectively). Both rates showed a tendency to decrease towards the west (Figure 5), showing significant relationships with longitude (Spearman’s correlation rank coefficient rs = -0.487, p = 0.002, and rs = -0.496, p = 0.001, for the >10 µm and <10 µm fractions, respectively). N2 fixation rates of organisms >10 µm measured during the day were not significantly different from those measured during the night (Wilcoxon test, p = 0.936), while the differences between rates of organisms <10 µm measured during the day and the night were almost statistically significant (Wilcoxon test, p = 0.059). DON release rates in organisms >10 µm and <10 µm ranged from 0.001 to ~0.09 nmol N L-1 h-1 (Figure 6a). Considering gross N2 fixation as net N2 fixation plus DON release (see Materials and methods), DON release represented ~14% of >10 µm gross N2 fixation and ~23% of <10 µm gross N2 fixation (Figure 6b). Significant differences in DON release were not found between >10 µm and <10 µm fractions, neither between day and night (Wilcoxon test, p>0.05). Also, DON release rates did not show a clear trend with longitude (rs = 0.203, p = 0.391, and rs = -0.046, p = 0.848, for the >10 µm and <10 µm fractions, respectively). Discussion Dissolved versus bubble methods Glibert and Bronk (1994) were the first to use dissolved 15N2 to measure N2 fixation in Trichodesmium colonies, although they did not perform 15N2 bubble versus dissolved 15N2 comparisons. Mohr et al. (2010) observed that N2 fixation rates in cultures of Crocosphaera watsonii were 40% higher when the dissolved method was used, compared to the bubble method. Recently, Wilson et al. (2012) compared both methods in natural waters of the North Pacific Ocean (station ALOHA), and obtained rates that were 2 – 3.5 fold higher with the dissolved method than they were with the bubble method. Großkopf et al. (2012) observed that both methods differ by 62% when the diazotrophic community is dominated by Trichodesmium, and up to 570% when dominated by diazotrophs other than Trichodesmium (symbionts of diatoms, UCYN, heterotrophic diazotrophs). Chapter 3: Longitudinal variability of size-fractionated N2 fixation and DON release 90 Fig. 3: West to east maps of sea surface height anomalies (SSHA) averaged every three N2 fixation stations Our N2 fixation rates estimated by the dissolved method were ~50% higher than those estimated with the bubble method. This underestimation is at the lower end of the few comparisons available in the literature. Adding to differences in the diazotrophic community, the factors influencing N2 fixation differences between the dissolved and bubble methods include (1) temperature -which affects gas dissolution into water-, (2) the agitation of the incubation bottles, (3) the volume of the incubation bottle, (4) the volume of 15N2 injected, (5) the duration of the incubation, (6) the time the incubation starts relative to the onset of nitrogenase activity -which differs between diazotroph species-, and (7) the DOM coating of the 15N2 bubble (Mohr et al., 2010). These factors vary widely among the previously published N2 fixation data measured using the bubble method, which makes the recalculation of rates difficult if not impossible (Großkopf et al., 2012). PART II: Results 91 Fig. 4: (a) N2 fixation rates obtained as the sum of >10 µm and <10 µm rates using the 15N2 dissolved method, and by incubation of whole seawater with a 15N2 bubble. (b) DON concentrations (in µmol L-1) in the >10 µm and <10 µm fractions. Error bars represent the standard deviation of the mean. With regards to DOM, we observed that the differences between the dissolved and bubble methods were greater in the eastern part of the transect, coinciding with higher DON concentrations (Figure 4b). Correlations between DON and the difference between the dissolved and bubble methods were not statistically significant (data not shown), but their longitudinal trend was similar. The difference between both methods would have probably correlated better with dissolved organic carbon (DOC), but unfortunately those data are not available. This study does not include diazotrophic organisms’ abundance and distribution, so we cannot analyze how this affected the percentage underestimation of the bubble method with regards to the dissolved method. However, the compilation of diazotroph abundance data shows that Trichodesmium clearly dominates in the tropical Northwest Atlantic, while UCYN are somewhat more abundant in the eastern than in the western side of the basin (Luo et al., 2012) and at higher latitudes (Moisander et al., 2010). Chapter 3: Longitudinal variability of size-fractionated N2 fixation and DON release 92 Fig. 5: >10 µm and <10 µm net N2 fixation rates in stations sampled during (a) the day, and (b) during the night. Error bars represent the standard deviation of the mean. This distribution of diazotrophs and percentage underestimation by the bubble method versus the dissolved method agrees with Großkopf et al. (2012), although our percentage underestimation values are much lower. Additionally, we must note that the sum of fractionated N2 fixation rates (>10 µm + <10 µm) may be not directly comparable to whole seawater N2 fixation rates. Sample fractionation is not always successful. For example, if <10 µm organisms are retained in the >10 µm fraction due to clogging of the mesh used for fractionation. However, due to the oligotrophic character of the area of study, seawater samples flowed easily through the mesh used for fractionation, and thus significant clogging or cell disruption may not be significant. DON release DON release rates were not significantly different between fractions, nor were they different between day and night, or significantly correlated with PART II: Results 93 longitude. This lack of spatial and daily variability in both size fractions suggests that the release of a percentage of recently fixed N2 is inherent to natural assemblages of marine diazotrophs, although we cannot discern if this release was active or passive, via predation, viral infection, or cell death. Axenic cultures of Cyanothece sp. Miami BG 043511 grown in optimum conditions released ~1% of their recently fixed N2 as DON (M. Benavides et al., in revision). This is much lower than the ~23% DON release obtained in this study for the <10 µm fraction However, Cyanothece or other UCYN belonging to group C (UCYN-C) diazotrophs are less frequent in the North Atlantic Ocean, and therefore it can be expected that the <10 µm diazotroph community in this transect was rather composed of Crocosphaera (UCYN group B - UCYN-B), UCYN of group A (UCYN-A), and heterotrophic diazotrophs (Langlois et al., 2008). 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(XSL73135) Ue"r":-.J-1:3 Uncultured marine bacterium clone TAI",U0515 (AY89638 4) L_____ Cyano!hece sp. PCC 8801 (CP001287) L____ Gloeof!tece sp. PCC 6909 (Ll5554) UnCllltllrcd bactenllm clone 5202A67 _t7 (DQ273169) Uncultured marine bacterium clone TAS1406 (AY896461) ,------Anabae na PCC7120 (AKANIFBH) Anabaena aphanizomenoides ~ ... f17 clone 7101A04 L_____ TolYPofhrix sp. PCC7601 (AY76842l) L______ Ch/orogfoell(lsi.\· sp. (CSU73138) L__________ (vngbya "ro/lei clone alexl (EF422067) .-'-'--- ...... l'riclwdesmiwn group L_______ .~~vmploca sp. PCC 8002 (AY221816) Phonnidium sp. ADl (AF227927) L------;9l9955 L ____ Phormidium sp. (PSU73132) L_____ Methylomonas rubra clone 23-4 (AF484673) f----------j 0.05 !"#$%&'()*(+!,-./(01234#45&(34(%"&(678%'1$35#9(-:(/%9#4%35( ( ( ! "N#! 1'B)&2!V<"L!/'M)+-,)+7!/)-*',('B%!-34+/-+1&2!8?%4(1%!&,!-.EL!:JJI^!U')2-+/&(! &,!-.EL!:J"J;E! ! $%&! /)2,()34,)'+-.! B-,,&(+! '5! 1&..2! )+! ,%&! >?@9:<5)=! .)+&-7&! -+/! ,%&)(! (-,&2! 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(a) The non upwelling-affected area, sampled from 25 July to 14 August 2009. The dashed line indicates the approximate position of the Azores Front (AF) during the cruise. White dots indicate stations where dissolved inorganic and organic nitrogen concentrations were measured. (b) The upwelling-affected area, sampled from 16 August to 5 September 2009. Nitrogen uptake, nitrogen regeneration and DON release NO3and NH4+ uptake, regeneration and associated release as DON was measured at the surface (5 m) and DCM depth of stations X2, X4, X6, X8, X10, X12, X14 and X16 in the non upwelling-affected area (Fig. 1a), and at all stations of the upwelling-affected area (Fig. 1b). For 15N experiments, water was directly transferred from the Niskin bottles to 2 L transparent polycarbonate bottles (Nalgene) in duplicate. 15N-labeled substrate was added AF a) CAIBOX b) Cape Ghir PART II: Results 149 to the bottles as 1 mL of 15KNO3 (200 μM; 99 at.%), or 1 mL of 15NH4Cl (20 μM; 99 at.%) (Sigma-Aldrich). For the non upwelling-affected stations, isotope enrichments were on average 22 and 27% for NO3– and NH4+, respectively. In the case of the upwelling-affected stations, average enrichments were 5 and 56% for NO3– and NH4+, respectively. Replicate bottles were immediately filtered after inoculation of the labeled substrates to obtain the initial 15N enrichment of samples. The other set of replicate bottles were equally inoculated and placed in on-deck incident light-calibrated containers cooled with surface seawater for 3–4 h. A hand light-meter (Biospherical Instruments) and neutral density filters (Lee filters) were used to mimic the light levels measured on PAR profiles at each station. After the incubation period, the samples were gently filtered through precombusted GF/F filters (vacuum pressure <100 mm Hg), wrapped in precombusted aluminium foil and stored at –20ºC. The uptake of NO3– was calculated according to the equations in Dugdale and Wilkerson (1986). NH4+ uptake rates were corrected for isotopic dilution as indicated by Glibert et al. (1982). Wherever under detection limit NH4+ concentrations were observed, values were substituted by the detection limit of the method (i.e. 5 nM), resulting in 50% enrichments, which can be considered as the threshold between real and potential uptake rates (Fernández et al., 2009). The filtrates were kept to analyze final NO3–, NO2–, NH4+ and DON concentrations (see above), and the 15N-enrichment of the final NO3–, NH4+ and DON pools using the protocol proposed by Slawyk and Raimbault (1995). For the latter, 80 mL of filtrate were stored in 100 mL borosilicate flasks (Duran Schott) and poisoned with 1% HgCl2 (Sigma-Aldrich) to interrupt any microbial activity. The extraction protocol consists of three steps: (1) NO3– and NO2– are reduced and stripped off together with initial NH4+ as ammonium sulfate, (2) oxidation of the remaining dissolved nitrogen (i.e. the DON pool) to NO3– by persulfate oxidation (Valderrama, 1981), and (3) repetition of step (1) to strip off DON as ammonium sulfate. From the samples incubated with 15NO3–, rates of NO3– uptake and DON release from NO3– are obtained. From the samples incubated with 15NH4+, rates of NH4+ uptake, NO3– regeneration, NH4+ regeneration and DON release from NH4+ are obtained. Nitrogen regeneration rates (NO3– and NH4+) were calculated by applying the equations in Fernández and Raimbault (2007). DON release rates from NO3– and NH4+ uptaken by microorganisms were calculated with the equations in Slawyk et al. (1998). N2 fixation was assayed only in the surface (~5 m) depth as outlined in Benavides et al. (2011), at stations X1 to X17 in the non upwelling-affected area, and at all stations in the upwelling-affected area. Briefly, seawater was transferred to 1 L transparent polycarbonate bottles (Nalgene). The bottles Chapter 6: N2 fixation contribution to new production and excess nitrogen 150 were filled to overflow using silicone tubing and preventing the introduction of air bubbles. Then they were sealed with septum screw caps before 2 mL of 15N2-gas (99 at.%; Tracer Tec) were injected through the septum using a gastight syringe (Hamilton). The pressure across the septum was equilibrated by allowing the excess air to escape through a sterile syringe tip piercing the septum at the same time as the 15N2-gas was being injected. Finally, the bottles were placed in the on-deck incubator for 24 h. After the incubation, samples were filtered through precombusted GF/Fs (vacuum pressure <100 mm Hg), wrapped in precombusted aluminium foil and stored at –20ºC until analysis. N2 fixation rates were calculated after Montoya et al. (1996). The concentration of particulate organic nitrogen (PON) and the isotopic ratio (15N/14N) of samples was obtained by means of a Thermo Flash 1112 elemental analyzer interfaced by a Conflo III with a Thermo Delta V Advantage isotope ratio mass spectrometer (IRMS). Results Hydrography Sections of temperature and salinity in the upper 200 m of the open ocean transect (non upwelling-affected area, Fig. 1a) are shown in Fig. 2. Temperature increased southwards along the transect (Fig. 2a). A sharp thermocline was observed in the northern part of the transect (stations X2 to X6, Fig. 1a), where temperatures of 19–20ºC were measured in the upper 40 m, then decreasing to 15–17ºC at 50–60 m depth. The Azores Front was located slightly north of station X8 and can be clearly identified by the tilting in the isotherms. South of the Azores Front, the entire water column warmed up and temperatures > 19ºC were observed down to ~80 m depth. This pattern was only disrupted after station X16, when reaching the coastal transition zone (CTZ) between the NW African upwelling ecosystem and the Canary Islands (Fig. 1b). Salinity mimics the structure of temperature in the upper 200 m of the open ocean transect (Fig. 2b). In the northern part of this transect, salinity decreased sharply from 38 to ~36.5 in the upper ~60 m depth. The Azores Front was also easily depicted by the tilting isohalines observed near to station X8. South of the Azores Front, the isohalines deepened (38 at ~100 m depth) and a stratified area was observed from stations X8 to X16. Stratification disruption associated with the CTZ was also observed in the isohalines (Fig. 2b). PART II: Results 151 Fig.2: (a) Temperature and (b) salinity sections from the non upwelling-affected area. The position of the stations sampled is displayed over the upper x-axis of (a). Surface and 50 m temperature and salinity distributions from the upwellingaffected area are shown in Fig. 3. We chose to represent temperature and salinity at 50 m because it is representative of the DCM at most stations sampled in this area (Table 1). From the distribution of temperature and salinity at 5 m we see that an upwelling filament extended up to ~11.5ºN, which is >150 km offshore. A tongue of cold upwelled water (16–18ºC) and associated low salinity (~36.2) extended over the shelf from the northern part of the NW African coast and the sampled stations. The coldest SST values (<17ºC) were detected at ~31ºN (Fig. 3a). An area of warmer (20-21ºC) and saltier water (36.5–36.6) was observed leeward of the upwelling-affected stations, over the bay of Agadir (Morocco). Chapter 6: N2 fixation contribution to new production and excess nitrogen 152 Figure 3: Temperature and salinity at the surface (~5 m) and 50 m from the upwelling-affected area. Black dots represent stations where 15N experiments were preformed. Surface temperature (a), and salinity (b) were obtained from thermosalinograph data (white line). Temperature and salinity at 50 m (figures c and d, respectively) were obtained by the combination of CTD casts (white circles) and Seasoar measurements (white squares). At 50 m depth, the filament decreased in width. The coldest temperatures (15ºC) were observed near the coast leeward of the upwelling-affected stations. Stations G12, G17 and G22 were clearly situated over the path of the filament, were temperatures at 50 m depth ranged from 17 to 17.5ºC. Station GT2 was situated at the edge of the filament, and stations G40, G44 and G48 at its tipend, where higher temperatures (17.5 to 18ºC) were measured. NO3–, NH4+, DON and DOC distributions In situ surface NO3– concentrations were generally <1 µM at all the stations sampled in both study areas with few exceptions (Table 1). These surface concentrations were fairly stable along the non upwelling-affected stations (at ~0.1 µM), being somewhat more variable at the DCM. Peaks of NO3– were measured in the upwelling-affected area associated with the coastal upwelling (station GM2), and stations over the upwelling filament (G17 and G44). In the non upwelling-affected area, DCM NO3– concentrations equaled those of the surface at most stations, while in the upwelling-affected area PART II: Results 153 increases of up to 1 µM were observed with respect to surface values. NH4+ concentrations were under the detection limit at several stations (indicated as n/d in Table 1). Where detectable, NH4+ concentrations were <50 nM in most cases, being somewhat higher north of the Azores Front (stations X2 to X8). NH4+ concentrations at the DCM were generally higher than at the surface. DON concentrations were very stable along the non upwelling-affected stations’ transect, generally ranging from 5.0 to 6.5 µM, except at station X4, where anomalously high concentrations of 7–9 µM were recorded. Generally, DON concentrations were higher at the surface than at the DCM, although stations X14 and X16 were an exception. DOC concentrations ranged from ~50 to 80 µM. The influence of the Azores Front was again noted in surface DOC concentrations. The largest values were found in the area affected by the front (stations X8 - X10) and then decreased towards the NW African coast. This effect was not observed at the DCM. DOC concentrations at the DCM remained stable along the non upwelling-affected area with the exception of station X14. In the upwelling-affected area, DON concentrations were much more variable. Surface concentrations averaged ~7 µM, but a peak > 9 µM was observed at the most offshore station (G48). At the DCM, the concentrations were also variable and a peak >10 µM was observed at station G40. DOC concentrations were lower in the upwelling-affected area than in the non upwelling-affected area, ranging from ~50 to 60 µM. Surface DOC concentrations tended to increase from the coast to the open ocean, while DCM values remained ~54 µM, with the exception of two peaks of ~63 µM at stations GM2 and G17. Nitrogen uptake, regeneration and release Surface and DCM NO3– uptake, NH4+ uptake, NO3– regeneration, NH4+ regeneration, DON release from uptaken NO3– and NH4+, N2 fixation, and the associated new and regenerated production rates (Pnew and Preg, respectively) were averaged for the non upwelling-affected and upwellingaffected areas (Table 2). Note that N2 fixation rates are only available for the surface. NO3– uptake was two orders of magnitude lower than NH4+ uptake in the non upwelling-affected area, while these were similar in the upwellingaffected area (Table 2). The regeneration of NO3– was low in the non upwellingaffected area and not detectable in the upwelling-affected area. NH4+ regeneration was one order of magnitude lower than NH4+ uptake in the non upwelling-affected area, and two orders of magnitude lower in the upwellingaffected area. DON release from NO3– was similar in both areas, while DON release from NH4+ was one order of magnitude greater in the upwellingaffected area than in the non upwelling-affected area. N2 fixation was very low in both areas, being somewhat higher in the upwelling-affected area. Chapter 6: N2 fixation contribution to new production and excess nitrogen 154 Pnew was corrected by adding N2 fixation -as another source of new nitrogen-, and by subtracting NO3– regeneration -a source of regenerated nitrogen- (Raimbault and Garcia, 2008). Preg was corrected by adding NH4+ regeneration to NH4+ uptake. DON released from NO3or NH4+ uptake was added to Pnew and Preg, respectively (considering the ‘loss’ of inorganic nitrogen to the DON pool; Raimbault and Garcia, 2008). Thus, the f-ratio was calculated as follows: f-ratio = (NO3– uptake + DON release from NO3– + N2 fixation – NO3– regeneration) / (NO3– uptake + N2 fixation – NO3– regeneration + NH4+ uptake + DON release from NH4+ + NH4+ regeneration). The NO3– regeneration rates measured were low or undetectable in the two sampled areas, so its effect in reducing Pnew was minimal. Similarly, N2 fixation rates were several orders of magnitude lower than NO3– uptake, therefore N2 fixation usually contributed <1% to Pnew (Table 2). Pnew was much greater than Preg in the upwelling affected area, while they were similar in the non upwellingaffected area. Accordingly, the f-ratio was very low in the non upwellingaffected oligotrophic waters (0.08), and higher (0.48) in the upwelling area (Table 2). If we consider gross NO3– and gross NH4+ uptake as NO3– uptake plus DON release from NO3–, and NH4+ uptake released from NH4+ (Bronk et al., 1994), we find that DON release from NO3– is more important in the non upwelling-affected area, while DON release from NH4+ is more important in the upwelling-affected area (Table 3). Nevertheless, the variability was very high (note high standard deviation values in Table 3). Discussion Uptake and regeneration of NO3and NH4+ and associated DON release in open ocean and upwelling zones It is generally assumed that primary production in upwelling systems is mainly supported by NO3- (‘new nitrogen’), which is regenerated at the base of the euphotic zone and reaches the photic layer through upward diffusive fluxes. Instead, productivity in open ocean oligotrophic ecosystems is thought to rely on NH4+ (‘regenerated nitrogen’) due to the year-round persistence of a strong thermocline that prevents nutrient-rich deep waters to reach the photic layer (Dugdale and Goering, 1967). The f-ratio quantifies the contribution of Pnew to total production (Pnew + Preg) (Eppley and Peterson, 1979). Table 1: Position of the mixed layer depth (MLD) and the deep chlorophyll maximum (DCM) at the stations sampled in the non upwelling-affected and upwellingaffected areas. In situ concentrations (previous to incubations with added 15NH4+ or 15NO3–) of nitrate (NO3–), ammonium (NH4+), dissolved organic nitrogen (DON) and dissolved organic carbon (DOC) (all in µM) at the surface and at the DCM of all the stations sampled. Under detection limit NH4+ concentrations are depicted as n/d (not detectable). Area Station Latitude (ºN) Longitude (ºN) MLD (m) DCM (m) Surface NO3– (µM) DCM NO3– (µM) Surface NH4+ (µM) DCM NH4+ (µM) Surface DON (µM) DCM DON (µM) Surface DOC (µM) DCM DOC (µM) Non upwelling-affected X2 41.50 10.58 40 60 0.13 0.12 – 0.117 6.5 5.8 68.8 63.0 X4 41.50 15.29 30 67 0.09 0.09 0.064 0.070 8.6 6.9 73.6 65.4 X6 41.44 19.26 38 55 0.15 0.05 0.157 0.179 6.1 5.3 71.4 63.4 X8 37.98 20.01 25 80 0.14 0.14 0.013 0.087 – – 78.7 63.0 X10 34.99 19.97 40 80 0.08 0.08 n/d 0.028 6.6 5.6 79.1 67.3 X12 31.50 20.00 45 105 0.13 0.11 0.020 0.015 6.2 6.1 70.2 61.2 X14 29.85 19.19 45 90 0.17 0.18 n/d 0.019 5.6 6.2 64.9 74.5 X16 29.25 15.93 50 110 0.05 0.05 n/d 0.012 5.0 5.8 68.4 61.4 Upwelling-affected GM2 31.00 10.01 15 25 3.00 2.77 n/d n/d – – 62.56 59.34 G12 30.74 10.65 15 30 0.61 1.95 n/d 0.009 6.88 3.73 – – G17 30.90 10.80 20 40 1.35 2.20 0.002 0.072 6.91 6.53 54.14 62.75 G22 31.00 10.91 25 40 0.38 1.56 0.015 0.305 6.74 – 49.18 53.08 GT2 31.11 10.60 25 45 0.82 0.93 n/d n/d 5.47 – 56.99 62.68 G40 31.01 11.31 30 48 0.55 1.65 n/d n/d 7.22 10.44 55.08 53.91 G44 30.95 11.40 28 43 1.50 0.38 n/d 0.021 6.18 7.44 59.32 53.51 G48 30.89 11.45 27 50 0.76 1.46 0.007 n/d 9.46 3.45 60.05 53.95 Table 2: Average and standard deviation (± SD) values of NO3– and NH4+ uptake and regeneration, and dissolved organic nitrogen (DON) release derived from NO3– uptake (DON-NO3–) and from NH4+ uptake (DON-NH4+), and N2 fixation, all in µmol L–1 h–1. All values are average measurements made at the surface (5 m) and at the deep chlorophyll maximum (DCM), except for N2 fixation, which was only measured at the surface. NO3– regeneration was not detectable (n/d) in the upwellingaffected stations. NO3– uptake NH4+ uptake NO3– regeneration NH4+ regeneration DON-NO3– release DON-NH4+ release N2 fixation Pnew Preg f-ratio Non upwellingaffected 0.008 ± 0.007 0.143 ±0.187 0.002 ± 0.002 0.012 ± 0.013 0.008 ± 0.486 0.001 ± 0.002 1.3 ± 0.7 x10–6 0.014 0.157 0.084 Upwelling affected 0.122 ± 0.086 0.105 ± 0.078 n/d 0.008 ± 0.023 0.004 ± 0.006 0.020 ± 0.033 5 ± 5 x10–6 0.126 0.133 0.487 PART II: Results 157 Table 3: Percentage DON release contribution to gross NO3– and NH4+ uptake in non upwellingaffected and upwelling-affected areas. Area DON release from NO3– contribution to gross NO3– uptake (%) DON release from NH4+ contribution to gross NH4+ uptake (%) Non upwelling-affected 26.48 ± 30.41 2.21 ± 3.18 Upwelling-affected 2.59 ± 2.67 14.16 ± 13 Upwelling systems are expected to have high f-ratios, while oligotrophic systems usually present low f-ratios. Nevertheless, the recent inclusion of N2 fixation, NH4+ and NO3regeneration and DON release rates in the computation of f-ratios has shown that Pnew is generally overestimated when nitrification is not taken into account, while Preg is generally underestimated when NH4+ regeneration is not taken into account (e.g. Fernández and Raimbault, 2007), while gross NO3and NH4+ uptake rates are underestimated if the associated DON released is not accounted for (e.g. Bronk et al., 1994). In this study, all the above-mentioned fluxes have been included in the calculation of Pnew, Preg and f-ratios, enhancing the robustness of our results. The NO3uptake rates in the upwelling-affected zone were in the range of those observed in the upwelling systems off California (0.1 - 0.55 µmol N L-1 h-1; Dugdale et al., 2006), and off Peru (0.11 - 0.14 µmol N L-1 h-1; Blasco et al., 1984), although more recently, rates one to two orders of magnitude lower (0.003 - 0.02 µmol N L-1 h-1) have been reported for the Peruvian and Chilean upwelling systems (Raimbault and Garcia, 2008; Fernández et al., 2009). While these authors observed considerably high NO3regeneration (nitrification) rates in the surface waters of the South Pacific Ocean upwelling systems (0.125 × 10-3 - 0.001 µmol N L-1 h-1), this flux was not detectable during our study in the coastal upwelling off Cape Ghir. 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