Microbial nitrogen removal versus recycling in the redox transition zone of a meromictic lake and its coupling to sulfur

Microbial nitrogen removal versus recycling in the redox transition zone of a meromictic lake and its coupling to sulfur | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Microbial nitrogen removal versus recycling in the redox transition zone of a meromictic lake and its coupling to sulfur View ORCID Profile Jana Tischer , View ORCID Profile Moritz F. Lehmann , View ORCID Profile Guangyi Su , View ORCID Profile Fabio Lepori , View ORCID Profile Jakob Zopfi doi: https://doi.org/10.1101/2025.01.03.627289 Jana Tischer 1 Department of Environmental Sciences, University of Basel , Basel, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jana Tischer For correspondence: jana.tischer{at}unibas.ch jakob.zopfi{at}unibas.ch Moritz F. Lehmann 1 Department of Environmental Sciences, University of Basel , Basel, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Moritz F. Lehmann Guangyi Su 1 Department of Environmental Sciences, University of Basel , Basel, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Guangyi Su Fabio Lepori 2 Department for Environment, Constructions and Design, University of Applied Sciences and Arts of Southern Switzerland , Canobbio, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Fabio Lepori Jakob Zopfi 1 Department of Environmental Sciences, University of Basel , Basel, Switzerland Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jakob Zopfi For correspondence: jana.tischer{at}unibas.ch jakob.zopfi{at}unibas.ch Abstract Full Text Info/History Metrics Supplementary material Data/Code Preview PDF Abstract Organotrophic denitrification is an important nitrogen (N) removal process in lakes, but alternative N reduction processes such as lithotrophic sulfur (S)-oxidizing denitrification may be greatly underappreciated. We studied the redox transition zone (RTZ) in the meromictic water column of the North Basin of Lake Lugano (Switzerland) to characterize N transformation pathways coupled to the S and carbon (C) cycles. Incubations with 15 N-labeled and unlabeled nitrate showed low denitrification rates and a general limitation of organic electron donors. The most accessible fractions of exported primary production biomass may have been largely consumed in the oxic water column during sedimentation, and did not reach the RTZ at ∼100 m. Conversely, sulfide (H 2 S) and methane (CH 4 ), major end products of anaerobic degradation of the more recalcitrant organic matter fractions in the sediment, represent a continuous source of energy to the RTZ, fostering the establishment of a community of S- and CH 4 -dependent nitrate reducers, dominated by Sulfuritalea and Candidatus Methylomirabilis over several years of observation. Anoxic incubation experiments with H 2 S amendments revealed a strong stimulation of dissimilatory nitrate reduction to ammonium (DNRA), but not denitrification. High relative abundances of the archaeal ammonia oxidizer Candidatus Nitrosopumilus and bacterial nitrifiers indicate intense nitrate regeneration by nitrification in the upper RTZ. The potential interaction between nitrification and S-driven DNRA is unclear. However, their importance in close proximity suggests that, at least under conditions of carbon limitation, N recycling between the nitrate and ammonium pools, predominates over N removal via complete denitrification in the Lake Lugano North Basin. Introduction Due to the widespread use of fertilizers in agriculture and waste water release, large amounts of reactive (i.e., fixed) nitrogen (N) have entered and altered natural ecosystems ( Gruber and Galloway 2008 ). It is estimated that up to 75% of the anthropogenic N introduced into inland waters are removed along the freshwater/seawater continuum before reaching coastal marine ecosystems. In this regard, lakes serve as efficient N sinks, playing a particularly important role in mitigating anthropogenic nutrient loads (Howarth et al. 1996). Canonical organotrophic denitrification, the microbial reduction of nitrate (NO 3 - ) to dinitrogen gas (N 2 ) with organic compounds as substrate, is considered to be the most important N removal process ( Seitzinger 1988 ). However, there is increasing evidence for alternative substrates such as methane (CH 4 ) (Raghoebarsing et al. 2006) and inorganic electron donors like reduced sulfur (S) compounds ( Hulth et al. 2005 ; Burgin and Hamilton 2007 ). Anaerobic ammonium oxidation to N 2 (anammox) and the dissimilatory nitrate reduction to ammonium (DNRA) represent additional, possibly underappreciated, N turnover processes in lakes ( Schubert et al. 2006 ; Roland et al. 2018 ). Yet, while these major N-transformations are well known, their relative contributions to N cycling, their different trophic modes (i.e., organotrophic versus lithotrophic), as well as the responsible microbial actors in lakes are not well understood. This is particularly important in the context of lacustrine N budgets: in contrast to denitrification and anammox, DNRA promotes N recycling rather than fixed-N removal. Denitrification and DNRA have in common, besides the initial reduction step from NO 3 - to NO 2 - , that both processes can be performed by organotrophic as well as lithotrophic microorganisms ( Pandey et al. 2020 ). It is assumed that DNRA dominates under conditions with a high availability of organic carbon (C org ) relative to NO 3 - , as e.g., in lake sediments, while denitrification is favored in high-NO 3 - environments ( Kelso et al. 1997 ; Dong et al. 2011 ). Furthermore, the type and availability of alternative inorganic substrates may also affect the mode of nitrate reduction ( Brunet and Garcia-Gil 1996 ; Cojean et al. 2020 ). Organotrophic N reduction is performed by a wide variety of ubiquitous facultative or strict anaerobic microorganisms, including many Proteobacteria, as well as some Firmicutes and Actinobacteria ( Shapleigh 2013 ; Pandey et al. 2020 ). Moreover, a range of obligate and facultative chemolithotrophic bacteria (e.g., species of the families Rhodocyclaceae , Sulfurimonadaceae , or Hydrogenophilaceae ) can couple, for example, the oxidation of sulfide (H 2 S), elemental sulfur (S 0 ), or thiosulfate (S 2 O 3 2- ) with the reduction of NO 3 - to NO 2 - , and from there to N 2 gas or to NH 4 + ( Zumft 1997 ; Shao et al. 2010 ; Pandey et al. 2020 ). To date, only few specialized microorganisms are known (e.g., Candidatus Methylomirabilis) that oxidize CH 4 anaerobically via NO 3 - or NO 2 - reduction to N 2 (Raghoebarsing et al. 2006; Yao et al. 2024) The absolute and relative importance of these different N-turnover modes in the natural environment can be highly variable. In lacustrine water columns, reported N-transformation rates vary by several orders of magnitude (i.e., between 10 µmol N L - 1 d -1 ; Table 1 ). Alternative modes of suboxic N reaction were shown to contribute substantially to total N 2 production (sometimes even exceeding rates of canonical organotrophic denitrification), such as S-dependent denitrification in Wintergreen Lake ( Burgin et al. 2012 ), methane-dependent denitrification in several Indian reservoirs (Naqvi et al. 2018), or anammox in Lake Tanganyika ( Schubert et al. 2006 ). Studies that concurrently investigated the rates of denitrification, DNRA, and anammox within the same lake water column are rare. Lake Kivu stands out as an exception, with all three processes confirmed to co-occur there ( Roland et al. 2018 ). Obviously, the ecosystem functioning of lakes in general, e.g., the mitigation of excessive N loading versus internal recycling, will strongly depend on the relative importance of denitrification and anammox on the one hand versus DNRA on the other. View this table: View inline View popup Download powerpoint Table 1. Potential rates of denitrification, with and without additional electron donor (ambient or added), dissimilatory nitrate reduction to ammonium (DNRA), and anammox in the water column of different lakes. From each study, the highest measured rates are reported. Rates of anammox include values determined with different 15 N-substrates: 15 NH 4 + , 15 NO 2 - , or 15 NO 3 - . nd = not detectable, na = not analyzed. Lake Lugano has been heavily impacted by increased inputs of phosphorous (P) and N from municipal wastewater, particularly between the 1960s and the 1980s ( Barbieri and Mosello 1992 ; Lepori et al. 2018 ; Studer et al. 2024 ). The fluxes of fixed N, and N cycling, in Lake Lugano have been the focus of research for many years (e.g., Barbieri and Mosello 1992 ; Lehmann et al. 2004 ). More recently, Wenk et al. (2013 ; 2014 ) investigated the modes of N loss in the water column of the northern basin, and reported that canonical organotrophic denitrification in the deep, meromictic northern basin of Lake Lugano plays only a minor role as N sink ( Wenk et al. 2013 ). Instead, S-driven chemolithotrophic denitrification, and to a much lesser extent anammox, were proposed as the main N-removing processes in the North Basin’s redox transition zone (RTZ). However, the conditions that promote fixed-N elimination by lithotrophic (i.e., S-dependent) rather than organotrophic N 2 production, and whether the controls on the modes of denitrification change seasonally (e.g., due to fluctuations in C org export/substrate availability), remained largely unresolved. Furthermore, it is still uncertain to what extent N recycling occurs within the RTZ of the Lake Lugano North Basin, as possible modes of DNRA have not been determined. Wenk et al. (2014) found, based on their natural abundance N and O isotope measurements, that there is little scope for nitrate regeneration by microaerobic nitrification within the RTZ. Here we combine 15 N isotope-label incubation experiments and 16S rRNA gene amplicon sequence data covering several years of observation to i) shed light on the controls on, and importance of, microbial fixed N removal versus fixed N recycling (e.g., by DNRA and nitrifying organisms) within/along the RTZ in the meromictic Lake Lugano North Basin, and ii) identify the main S-oxidizing nitrate reducers, as well as other N- and S-transforming microorganisms involved. We examine whether the previously proposed predominance of S-driven denitrification in the basin’s RTZ is a permanent or a seasonal feature, and we aim to understand why a thermodynamically less favorable inorganic substrate, such as H 2 S, is preferred over organic matter (OM) as driver for N cycling. Methods Study site Lake Lugano is a south-alpine lake on the Swiss-Italian border at an altitude of 271 m above sea level. A causeway built on a moraine separates the lake into the southern basin (93 m) and the deep and narrow (288 m) northern basin. Increasing eutrophication led to biogenic meromixis from ∼1960 onward. This state was only interrupted by two mixing events in 2005 and 2006, when cold and windy winters caused complete overturning and the transient oxygenation of the entire water column ( Holzner et al. 2009 ). In situ profiling and sample collection We studied the water column of the northern basin of Lake Lugano at the deepest spot (46°00’37.7”N, 9°01’14.9”E) off the village of Gandria in April, June, and October 2015, March, September, and November 2016, February and October 2017, and April 2018. A conductivity, temperature, depth (CTD) probe (Idronaut Ocean Seven 316Plus) was used to determine oxygen (O 2 ) concentrations, temperature, conductivity, and, from July 2016 onward, chlorophyll a (Chl a ). Monthly primary production and nitrate concentration data were obtained through a monitoring program conducted by the University of Applied Sciences and Arts of Southern Switzerland ( www.cipais.org ). A RTZ separates the fully oxic upper water column from the anoxic monimolimnion. In this study, the upper RTZ boundary is set at the depth where O 2 falls below 5 µM, and its lower boundary at the depth, where oxygen-sensitive reduced chemical compounds like Fe 2+ or H 2 S rise above background levels. Samples were collected across the RTZ starting at around 80 m depth to a maximum depth of 155 to 165 m using 5 L Niskin bottles. Sample water was filled into sterile 1 L plastic or borosilicate bottles for DNA analysis and into 1 L borosilicate bottles for enrichment cultures, each without leaving any headspace. Water samples were kept cold and in the dark until further processing. For DNA analysis water samples (∼1.1-1.2 L) were filtered through 0.2 µm polycarbonate membrane filters (Cyclopore, Whatman) within 24 h of sample collection. For 15 N-label incubations, water from the Niskin bottle was filled directly into sterile 160 mL serum vials (bubble free with ∼1-2 volumes overflow), and the vials were closed with grey rubber stoppers (VWR). The bottles were kept cold and in the dark at all times, until the incubation experiments were started in the home laboratory within 10 h after sampling. From each depth, additional samples were collected and prepared for various chemical analyses (see below). Chemical analyses Nitrogen species Water samples for the analysis of dissolved inorganic N concentrations were filtered (0.45 µm pore size) right after collection. NH 4 + concentrations were determined using the colorimetrical indophenol reaction ( Hansen and Koroleff 1999 ), and NO 2 - using sulphanilamide and N-(1-Naphthyl)ethylenediamine ( Hansen and Koroleff 1999 ). NO x (i.e., NO 2 - + NO 3 - ) was determined using a NO x -Analyzer (Antek Model 745) involving the reduction of NO x in a hot acidic V 3+ solution to NO gas, and subsequent chemiluminescence detection ( Braman and Hendrix 1989 ). We determined NO 3 − concentrations by subtracting NO 2 − from NO x . Iron Aliquots of unfiltered and filtered (0.2 µm) water samples were fixed with ∼150 mM (final concentration) nitric acid (HNO 3 ) for the analysis of total and dissolved iron concentrations, respectively. Concentrations were quantified by inductively coupled plasma optical emission spectrometry (ICP-OES, Agilent Technologies) with a detection limit of ∼1.8 µM and 5-10% measurement uncertainty. We calculated concentrations of particulate Fe from the difference between total and dissolved Fe. Sulfur compounds We analyzed concentrations of dissolved H 2 S, S 2 O 3 2- , and SO 3 2- according to Zopfi et al. (2008) . Briefly, 450 µL unfiltered water samples were fixed in a mixture of 25 µM of HEPES-EDTA buffer (pH 8, 500 mM, 50 mM) and 25 µL of ∼45 mM monobromobimane in the dark. After 30 min, we stopped the derivatization reaction by adding 50 µL of 312 mM methanesulfonic acid. The samples were stored at -20 °C until analysis with reversed-phase high-performance liquid chromatography (RP-HPLC, Dionex) using a LiChrosphere 60RP select B column (125×4 mm, 5 μm; Merck) and a Waters 470 scanning fluorescence detector (excitation at 380 nm; detection at 480 nm). In certain cases, H 2 S concentrations were determined photometrically through the methylene blue reaction in unfiltered samples, immediately fixed with zinc acetate (0.5% final concentration, w/v) upon collection ( Cline 1969 ). For the analysis of suspended S 0 , we filtered 60 mL of water sample through a glass microfiber (GF/F) filter (Whatman), and subsequently stabilized the S 0 on the filter with 2-3 mL 5% (w/v) zinc acetate solution. The filters were stored at -20 °C until analysis. Using 2 mL of HPLC grade methanol, S 0 was extracted overnight from the filters and subsequently quantified by RP-HPLC using a Knauer C18-column (Eurospher II, 100-5 C18 H, 125x4 mm) and UV-detection at 265 nm ( Zopfi et al. 2008 ). SO 4 2- concentrations were analyzed by ion chromatography and UV detection (940 Professional IC Vario, Metrohm). Methane We analyzed methane concentrations in November 2016 and October 2017, as described in Su et al. (2023) . Briefly, water samples were collected in 120 mL serum bottles, crimp-sealed with buthyl rubber stoppers, and a 20 mL air headspace was created before fixing the sample with 5 mL of 12.5 M NaOH. Methane concentrations were measured using a gas chromatograph (SRI 8610C, SRI Instruments) with a flame ionization detector ( Lehmann et al. 2004 ). Water column stability and turbulent flux calculations The static stability of the water column was calculated as the Brunt-Väisälä frequency N 2 ( Wüest et al. 1992 ): where g represents the gravitational acceleration (9.81 m/s 2 ), ρ is the density of water, and δρ/δz denotes the density gradient over a specific depth increment. We calculated the pure water density as function of the measured temperature (T) with the following equation: In a second step, we calculated the conductivity-dependent density as function of conductivity according to Wüest et al. (1992) : where the constant β 20 is the relative change of density per unit electrical conductivity (7.05 x 10 -7 [µS x cm -1 ] -1 ) and κ 20 is the conductivity at 20°C. Turbulent diffusive fluxes of NO 3 - , NH 4 + , H 2 S, and CH 4 towards the RTZ were determined according to Wenk et al. (2013) : where F z represents the vertical solute flux, K z the vertical eddy diffusivity, and dC/dz the concentration gradient of the respective solute. Vertical eddy diffusivity was calculated as: where a 0 is a system-specific constant, which we adapted from Wenk et al. (2013) for the Lake Lugano North Basin (0.00014 cm 2 s -2 ). The N 2 and the K z values were calculated using the average CTD data integrated over 5 m intervals. To estimate NO 3 - fluxes towards the RTZ from above, we used the average K z value from all samplings between 2015 and 2018 of 0.85 ± 0.10 m 2 d -1 (standard deviation, SD) for the interval 80-110 m. Similarly, to calculate fluxes of NH 4 + , H 2 S, and CH 4 from the deeper hypolimnion towards the RTZ, the average K z value for the depth interval 95-150 m (1.31 ± 0.22 SD m 2 d -1 ) was used. These values for K z are within the range of previously reported vertical eddy diffusivities for the North Basin of Lake Lugano ( Wüest et al. 1992 ; Wenk et al. 2013 ). DNA extraction, PCR amplification, Illumina sequencing, and data analysis Filters with collected DNA were stored at -70 °C until extraction of DNA using the Fast DNA Spin Kit for Soil (MP Biomedicals). In addition to the 2015-2018 samples, we used samples already collected in 2009 and 2010 (see Su et al. 2023 ). The 16S rRNA gene library preparation, amplicon sequencing, and bioinformatic treatment of raw sequences are described in detail in Su et al. (2023) . Briefly, the updated Earth Microbiome PCR primers for bacteria and archaea 515F-Y and 926R, targeting the V4 and V5 regions of the 16S rRNA gene, were used for the first PCR (Parada et al. 2015). Sample indices and Illumina adaptors were added by the second PCR before sequencing the purified amplicons on an Illumina MiSeq platform at the Genomics Facility Basel. Amplicon Sequence Variants (ASVs) were identified by denoising amplicons to zero-radius OTUs using the UNOISE algorithm in USEARCH v10.0.240 ( Edgar 2010 , 2013 ). Finally, we used SINTAX (Edgar 2016) and the SILVA 16S rRNA reference database v138 ( Quast et al. 2013 ) for the taxonomic assignment of ASVs. We performed downstream sequence analysis and visualization in R v4.1.1 using the packages phyloseq v1.36.0 ( McMurdie and Holmes 2013 ), vegan v2.5-7 (Oksanen et al. 2020), ggplot2 v3.3.5 ( Wickham 2016 ), and dplyr v1.0.7 ( Wickham et al. 2021 ). The sequence data were cleaned by removing mitochondrial and chloroplast sequences, as well as ASVs with unknown phylum-level taxonomy. We examined alpha diversity measures, specifically the observed ASV richness and Shannon diversity, using rarefied data ( Su et al. 2023 ). In addition, we analyzed the temporal evolution of alpha diversity in each redox zone by determining Pearson correlation coefficients of the alpha diversity measure Observed using months as measure of time between sampling timepoints. Beta diversity was assessed using weighted UniFrac distances and principal coordinate analysis (PCoA). Based on the environmental conditions, we divided the samples into three groups: the oxic water column above the RTZ (“oxic”), the RTZ (“RTZ”), and the anoxic water layer below the RTZ (“anoxic”). Pairwise t-tests were performed by using the t.test() function in R to test for significance between the three different redox zones of the alpha diversity measures and of the weighted UniFrac distances by comparing the beta diversity distance pairs within the oxic zone samples, within the RTZ samples and within the anoxic samples. Permutational Multivariate Analysis of Variance (PERMANOVA) was performed using the adonis2() function in vegan to test for significant differences between the different redox zones and/or between timepoints, as well as well their interaction. We visualized key phyla and classes based on relative abundances of 16S rRNA gene amplicons (taxa with a relative abundance of ≥1% in at least one of the samples). In addition, the temporal change of the microbial community at a given redox zone was monitored by selecting one sample per timepoint, respectively, from the oxic water column (5-10 m above the upper boundary of the RTZ), one from the RTZ (middle of the RTZ), and one from the anoxic water column (40-45 m below the lower boundary of the RTZ). We screened for S- and N-cycling taxonomic groups (Table S1) including ammonia-oxidizing archaea (Yang et al. 2021), ammonia-oxidizing bacteria ( Kowalchuk and Stephen 2001 ), nitrite-oxidizing bacteria ( Daims et al. 2016 ), anammox bacteria ( Jetten et al. 2009 ), organotrophic nitrate/nitrite reducers ( Zumft 1997 ; Pandey et al. 2020 ), S-oxidizing N reducers ( Shao et al. 2010 ; Kojima and Fukui 2011 ), CH 4 -oxidizing denitrifiers ( Kits et al. 2015 ), and S-reducers ( Rosenberg et al. 2014 ). Additionally, we selected taxa with names that designate one of the specific before-mentioned metabolisms, e.g., “denitrificans”, “sulfuri”, or “desulf”. In our screening efforts, we also included genera that appeared to thrive and enrich in our incubation experiments with added NO 3 - , i.e., that showed a higher (≥2%) relative abundance (ΔRA = RA end – RA initial ) after incubation (limiting ourselves to microorganisms known for N reduction, denitrification, or DNRA genes; identified using www.kegg.jp ) (Tischer et al. 2024/in prep). We only considered taxa with an abundance of ≥0.005% in at least one of the in-situ water-column samples. Batch incubation experiments and microbial enrichments We performed batch incubations with lake water sampled between 2016 and 2018 in order to investigate NO 3 - reduction with different electron donors. In 1-L borosilicate bottles with water samples, we introduced a N 2 headspace (∼130 mL), and purged for 30 minutes with N 2 to ensure anoxic incubation conditions. Subsequently, we added NO 3 - , NO 3 - and H 2 S, or NO 3 - and sodium acetate (NaAc) from sterile anoxic stock solutions. Targeted initial concentrations for NO 3 - and NaAc were 25 µM, for H 2 S they ranged between 25 and 100 µM ( Table 2 ). One bottle was left as a live control treatment without any amendments, and we incubated a dead control with NO 3 - and H 2 S, killed with 15 mL 50% (w/v) ZnCl 2 , to ensure that no abiotic H 2 S oxidation was taking place (data not shown). Incubations were carried out at ∼8 °C in the dark with gentle agitation. We monitored the concentrations of dissolved N and S species by taking subsamples (∼10 mL) at specific time intervals. To prevent contamination with O 2 , we initially pressurized the incubation bottles to around 2 bars and used N 2 -flushed syringes for sampling. When H 2 S was consumed before the complete reduction of NO 3 - , we added additional H 2 S (∼25 µM final concentration). At the end of the experiment, we filtered the remaining water (approximately 300-800 mL) to collect biomass for DNA extraction and sequencing, as described above. This allowed us to identify microbial taxa that responded positively to the different substrate additions. We calculated NO 3 - turnover rates based on the change in NO 3 - concentration over time, using samples from two intervals, i) the first 2-3 days and ii) the first 4-5 days of incubation. We tested differences between treatments with a pairwise t-test and differences between intervals with an analysis of variance (ANOVA). Similarly, we calculated turnover rates for H 2 S, S 0 , SO 3 2- , S 2 O 3 2- , and SO 4 2- using samples collected during the first 3-4 days of incubation. A t -test was performed on the rates to test for significance. View this table: View inline View popup Download powerpoint Table 2. Incubation conditions and potential rates determined in 15 N-tracer incubation experiments. Values are presented with standard error of means for triplicate or quintuplicate incubations. Percentages refer to the relative contribution of a given process with respect to the combined production of 15 N-N 2 and 15 N-NH 4 + . 15 N-label incubation experiments The 15 N-label incubation experiments were performed in October 2017 and April 2018 to determine rates of denitrification, anammox, and DNRA at different depths in the water column. We used a modified version of the protocol described in Wenk et al. (2013) , adjusted for the additional quantification of 15 NH 4 + . We introduced a 10 mL helium (He) headspace to the 160 mL sample vials, and purged for 10 minutes with He to remove potential traces of O 2 . Subsequently, we added 15 N-NO 3 - tracer from a sterile and anoxic stock solution of 7.5 mM 15 N-KNO 3 (99% 15 N-KNO 3 , Cambridge Isotope Laboratories, Inc.), aiming for an initial 15 NO 3 - concentration of 25 µM. Additionally, H 2 S or Na-acetate (each 25 µM final concentration) was added in the respective treatments ( Table 2 ). All incubations were performed in triplicate in October 2017 and in quintuplicate in April 2018. Incubations were performed in the dark, at ∼8 °C with gentle agitation. Samples for N 2 isotope analysis were collected at five timepoints (∼12, 36, 60, 108, and 325 h) by sampling 2 mL of headspace, in exchange with He, using an airtight 3 mL Luer-lock plastic syringe (Braun) with a plastic valve flushed 3-times with He. The samples were stored in 3 mL exetainers after replacing sterile anoxic water with the gas sample. In addition to headspace samples, we took 5-mL liquid samples, filtered them using 0.2 µm membrane filters, and stored them frozen until analysis. The resulting negative pressure in the incubation vials was equilibrated by adding He. We determined 15 N-N 2 production (m/z 29/28 and 30/28 ratios) via isotope ratio mass spectrometry (IRMS; Flash-EA-ConfloIV-DELTA V Advanced, Thermo Scientific), and calculated the rates of denitrification and anammox according to the isotope pairing equations of Thamdrup and Dalsgaard (2002) and Thamdrup et al. (2006), where we used only the first three timepoints. The 15 N-NH 4 + samples were transformed to 15 N 2 by oxidation with hyprobromite ( Risgaard-Petersen et al. 1995 ), and analyzed as described above. NH 4 + standards of different concentrations and 15 N/ 14 N ratios were included in the analysis. We referenced the samples against air N 2 and used the quantified ammonium-derived 15 N 2 to assess 15 N-NH 4 + concentrations in the liquid sample according to Cojean et al. (2020) . DNRA rates were then calculated from the slope of the increase in 15 N-NH 4 + concentration over time. Given that anammox rates were low ( Table 2 ), we assumed that the consumption of 15 N-NH 4 + produced by DNRA was negligible. Results Water column characteristics and hydrochemistry Water column CTD profiles from the different sampling campaigns are shown in Fig. S1. Water column stability (Fig. S1e) was generally strongest in the subsurface waters, where temperature and density gradients were highest; no significant density gradient was observed at the RTZ (Fig. S1d). Water column O 2 profiles indicate that all O 2 was consumed approximately at a water depth of ∼80 and 90 m (four selected timepoints in Fig. 1 ), which was 35-45 m higher up in the water column compared to 2009 and 2010 ( Wenk et al. 2013 ). Download figure Open in new tab Figure 1. Concentration profiles at the water column redox transition zone (RTZ) of the Lake Lugano North Basin from four different sampling campaigns. The grey bars indicate the extension of the RTZ. The biweekly O 2 monitoring data from 2015 to 2018 showed highest concentrations between 0 and 20 m during spring and summer, while in late summer and autumn, the water column was generally less oxygenated with an O 2 depletion at ∼20 to 30 m (Fig. S2a). Among the sampling dates when we collected water samples for incubation experiments, April 2018 was the only date with a distinct O 2 peak (up to ∼500 µM at ∼15 m) in shallow waters (Fig. S1a). The Chl a concentrations in the surface waters were highest in spring and summer (Fig. S2b). The NO 3 - concentration profiles of the entire water column showed highest concentrations between 20 and 50 m depth, ranging from 24 to 38 µM (Fig. S2b). Detailed concentration profiles at the RTZ ( Fig. 1 , Fig. S3) indicate that NO 3 - decreased to <0.5 µM within the RTZ. For most timepoints, NO 2 - did not exceed 0.05 µM in the RTZ. NH 4 + concentrations started to increase around the depth of O 2 depletion and reached concentrations between 15 and 25 µM at ∼155 m. NH 4 + and NO 3 - profiles generally overlapped, with concentrations of ∼1-2 µM each. H 2 S, on the other hand, was first detected approximately 15 m below the depth where NO 3 - reached its lowest levels <0.5 µM ( Fig. 1 , Fig. 2a-c ), whereas S 0 , the most abundant product among the intermediates of H 2 S oxidation, was detectable up to the upper border of the RTZ at levels up to ∼0.25 µM ( Fig. 2d-f ). Low SO 3 2- and S 2 O 3 2- concentrations of <0.15 µM were detected in most of the anoxic water samples ( Fig. 2d-f ). In this regard, the October 2017 profiles stand out, as S 0 , S 2 O 3 2- , and SO 3 2- consistently showed two distinct concentration peaks in the anoxic water column. Dissolved Fe 2+ concentrations rose up to ∼3 µM around 160 m depth, while particulate Fe remained below 1 µM, reaching a maximum at 125-130 m in October 2017 and April 2018 (Fig. S3e). CH 4 concentration profiles (November 2016 data have been previously published in Su et al. 2023 ; Fig. 1 ) clearly overlapped with NO 3 - profiles. Below the RTZ, CH 4 concentrations increased to up to ∼50 µM (e.g., October 2017). Turbulent diffusive solute fluxes towards the RTZ between 2015 to 2018 were on average 828 ± 315 SD µmol for NO 3 - m -2 d -1 , 379 ± 122 SD µmol for NH 4 + m -2 d -1 , 199 ± 87 SD µmol for H 2 S m -2 d -1 , and 882 ± 597 SD µmol for CH 4 m -2 d -1 (Table S2). Download figure Open in new tab Figure 2. Depth profiles of sulfur compounds across and below the redox transition zone (RTZ) (grey bars), from February 2017 (a+d) , October 2017 ( b+e) , and April 2018 (c+f) . Microbial community structures The 16S rRNA gene sequencing revealed differences between the microbial communities in the oxic water column just above the RTZ, within the RTZ, and in the anoxic water below the RTZ with regard to their alpha and beta diversities ( Fig. 3 ). The observed ASV richness was significantly higher in the anoxic water layers below the RTZ compared to the oxic water (pairwise t-test; t = -6.1, df = 19, p < 0.001) layer and the RTZ (pairwise t-test; t = -14.2, df = 93, p < 0.001). Interestingly, ASV richness increased with time during the multiannual survey of microbial community structures in the water column (Fig. S4), especially in the anoxic water below the RTZ (Pearson correlation coefficient; t = 3.0, n= 68, r = 0.34). The Shannon diversity index, which quantifies both richness and evenness, was higher in the oxic water compared to the RTZ (pairwise t-test; t = 4.2, df = 26, p < 0.001) and the anoxic water layer (pairwise t-test; t = 4.4, df = 19, p < 0.001). The community structure in the oxic water column seems to be more evenly distributed than in the RTZ and the anoxic zone. Particularly the communities in the anoxic layer show a high richness (i.e., a large number of different ASVs) but lower Shannon diversity, indicating a community with few high abundance and many very low abundance taxa. Download figure Open in new tab Figure 3. Characterization of the microbial communities in the water column of the Lake Lugano North Basin depending on redox regime (“oxic”, redox transition zone: “RTZ”, “anoxic”) and/or sampling timepoint using 16S rRNA gene amplicon sequencing variant (ASV) data from 2009 to 2018. a) Observed ASV richness and Shannon diversity, b) principal coordinate analysis (PCoA) of weighted UniFrac distances, and c) weighted UniFrac distances of microbial community structures within the three distinct redox zones. Stars indicate significance of pairwise t-tests ( p < 0.001). The PCoA showed that the microbial community structures followed the three redox zones, although the transitions were gradual and zones were overlapping ( Fig. 3b ). The community structures in the three defined redox zones were significantly different from each other (PERMANOVA; F 2,116 = 114.3, R 2 = 0.38, p < 0.001). Furthermore, significant differences were observed across timepoints (PERMANOVA; F 11,116 = 13.3, R 2 = 0.24, p < 0.001) and in the interaction between timepoints and redox zones (PERMANOVA; F 21,116 = 5.5, R 2 = 0.19, p < 0.001). The microbial community structure below the RTZ exhibited the least variability between different timepoints ( Fig. 3b ). This observation was confirmed by the pairwise Weighted UniFrac distances between samples within a specific redox zone, where the beta diversity below the RTZ was significantly lower than in the RTZ and in oxic-water samples ( Fig. 3c ; pairwise t-test; oxic vs. RTZ samples: t = 9.3, df = 196, p < 0.001; oxic vs. anoxic samples: t = 13.9, df = 138, p < 0.001; RTZ vs. anoxic samples: t = 8.0, df = 695, p < 0.001). Phylogenetic analysis of 16S rRNA gene amplicons revealed Proteobacteria, Bacteroidota, Actinobacteriota, Chloroflexi, and Crenarchaeota as the most common phyla (Fig. S5a) and Actinobacteria, Bacteroidia, and Gammaproteobacteria as the most common classes (Fig S5b). Below the RTZ, the relative abundance of Desulfobacterota and Nanoarchaeota increased with depth in the water column. The composition of the microbial community (Fig. S6; phyla and classes) in the different redox zones over time showed that in the oxic water column, changes in relative abundance are more pronounced compared to the RTZ and the anoxic water column. Identified ASVs in the dataset associated to N- and S-cycling are summarized in Table S1 and Fig. S7-S11, whereas vertical distribution profiles of prominent representatives in the water column are shown in Fig 4 . We detected the ammonia-oxidizing archaeon Candidatus Nitrosopumilus, the ammonia-oxidizing bacterium Nitrosospira and Nitrosomonas , and the nitrite-oxidizing bacterium Nitrospira with highest relative abundances at low O 2 conditions above the RTZ ( Fig. 4a-c ). Furthermore, we detected the anammox-associated family Brocadiaceae ( Fig. 4d ). We also identified several taxa capable of N reduction, e.g., in the classes of Alphaproteobacteria, Gammaproteobacteria, and Campylobacteria (former Epsilonproteobacteria), which showed maximum relative abundances within the RTZ (examples in Fig. 4e-j ). These taxa included organotrophic N reducers, such as Denitratisoma, Sterolibacterium , and Dechloromonas , S-oxidizing N reducers as Sulfuritalea, Sulfuricurvum , and Sulfurimonas , and NO 2 - -reducing, CH 4 -oxidizing Ca. Methylomirabilis. Some taxa showed two distinct maxima in their relative abundance, the first near the oxic-anoxic interface and another one further below within anoxic waters, e.g., Sulfuritalea and Sulfurimonas in March 2016. A diverse community of taxa capable to reduce SO 4 2- and other S-compounds was present, mostly belonging to the Desulfobacterota (former Deltaproteobacteria), e.g., Desulfobacca , Desulfocapsa ( Fig. 4k-l ), and Desulfovibrio. In addition, we identified sulfate-reducing Firmicutes, including Desulfosporosinu s and Desulfurispora . Relative abundances of S-reducers generally increased with depth, but some genera exhibited distinct peaks in and/or below the RTZ, including Desulfosporosinus (Fig. S11a), Desulfomonile (Fig. S11g), and the versatile NO 3 - /Fe(III)/S 0 -reducing Geobacter (Fig. S11j). Download figure Open in new tab Figure 4. Depth distribution and temporal variability of the main taxa of N and S cycling microorganisms. Selected nitrifiers (a-c) , anammox-performing bacteria (d) , organotrophic- (e, f) , sulfidotrophic- (g, h, i) and methanotrophic N reducers (j) , and S-reducers (k-l) are presented. Data are based on relative abundances of 16S rRNA gene amplicons. Nitrogen transformation incubation experiments In the unlabeled incubations with water from the RTZ ( Fig. 5 ) with NO 3 - added only, NO 3 - consumption occurred, but was, in most cases, incomplete (reduction by ∼3-11 µM NO 3 - ). Most of the NO 3 - was only converted to NO 2 - ( Fig. 5d ). Complete removal NO 3 - (and subsequently also of NO 2 - ) occurred only in one replicate experiment in February 2017 and one replicate experiment in April 2018 ( Fig. 5A and D ). Download figure Open in new tab Figure 5. Concentrations of NO 3 - (a-c) and NO 2 - (d-f ) in unlabeled incubation experiments performed with water samples from the redox transition zone (RTZ) in the North Basin of Lake Lugano, with added NO 3 - (a, d ), with NO 3 - and acetate (b, e) , or with NO 3 - and H 2 S (c, f) . Inserts show nitrate concentration changes within the first 3 days of the experiment. The addition of acetate led to the relatively rapid and complete reduction of NO 3 - within 4-5 days, after an initial lag phase ( Fig. 5b ). The addition of H 2 S caused complete NO 3 - consumption within ∼10-30 days ( Fig. 5c ). The NO 3- drawdown was fastest, and almost linear, in water samples from the lower RTZ (105 m) in October 2017 and April 2018. NO 2 - accumulation was transient in the incubations with acetate and H 2 S ( Fig. 5e-f ). The initial (i.e., first 2-3 days of incubation) NO 3 - reduction rates (Fig. S12a) were highest for the NO 3 - + H 2 S treatment with 1.4 ± 0.3 (standard error, SE) µmol NO 3 - L -1 d -1 . For the NO 3 - -only (0.6 ± 0.2 SE µmol NO 3 - L -1 d -1 ) and the NO 3 - + acetate treatment (0.9 ± 0.4 SE µmol NO 3 - L -1 d -1 ), initial NO 3 - reduction rates were lower. Whereas after the initial phase (Fig. S9b), NO 3 - consumption rates in the NO 3 - -only treatment (0.7 ± 0.4 SE µmol NO 3 - L -1 d -1 ) and the NO 3 - + H 2 S treatment (1.7 ± 0.3 SE µmol NO 3 - L -1 d -1 ) remained similar, NO 3 - consumption rates in the NO 3 - + acetate treatment increased to 3.2 ± 0.6 SE µmol NO 3 - L -1 d -1 (ANOVA; F 1,22 = 9.8, R 2 = 0.31, p = 0.005). During the course of the incubations, H 2 S was consumed and S 0 and S 2 O 3 2- accumulated, but no substantial amounts of SO 4 2- were produced (Table S3, Fig. S13). 15 N incubation rate measurements In the 15 N-incubation experiments conducted in October 2017 and April 2018, denitrification rates (i.e., N 2 production by denitrification) ranged between 28.8 and 113.0 nmol N-N 2 L -1 d -1 in the treatments with 15 NO 3 - and no added electron donors ( Table 2 ). Hence, N 2 production rates were almost an order of magnitude lower than NO 3 - consumption rates determined in parallel unlabeled experiments (see above), which can be explained by the accumulation of NO 2 - , which had also occurred in the unlabeled incubations ( Fig. 5d ). A representative example of a 15 N-label incubation experiment from April 2018 is presented in Fig. S14. Amended acetate and H 2 S did not significantly change the measured denitrification rates (ANOVA; F 2,6 = 2.1, R 2 = 0.41, p = 0.21), which ranged between 19.4 and 31.7 nmol N-N 2 L -1 d -1 for the 15 NO 3 - + acetate treatment and between 39.1 and 71.0 nmol N-N 2 L -1 d -1 for the 15 NO 3 - + H 2 S treatment. Qualitatively consistent with the observed NO 3 - reduction dynamics in the unlabeled incubations, acetate-supported 30 N 2 production started only after a lag phase of more than three days (Fig. S14e). In the 15 NO 3 - and 15 NO 3 - + acetate incubations, DNRA rates were mostly undetectable. Only in April 2018 (95 m, NO 3 - treatment: 142.2 nmol 15 N-NH 4 + L -1 d -1 and 105 m, 15 NO 3 - + acetate treatment: 161.3 nmol 15 N-NH 4 + L -1 d -1 ), they were significant and contributed more than denitrification to the total NO 3 - turnover. In contrast, the addition of H 2 S resulted in high DNRA rates between 471.1 and 1706.6 nmol 15 N-NH 4 + L -1 d -1 , representing 87-97% of the total 15 N-N 2 + 15 N-NH 4 + produced. Anammox rates were generally low in all treatments, ranging from 0.1 up to 12.5 nmol 15 N-N 2 L -1 d -1 . They accounted for 0 to 30% (but mostly <10%; Table 2 ) of the produced 15 N-N 2 + 15 N-NH 4 + , without any significant differences between treatments (ANOVA; F 2,6 = 1.5, R 2 = 0.33, p = 0.30), and were not considered further in the discussion. Discussion The results from unlabeled and 15 N-label incubation experiments and 16S rRNA gene sequencing data provide conclusive evidence for active N-cycling around the RTZ, closely coupled to the S- and C-cycles. The overlapping concentration profiles of NH 4 + and NO 3 - with complete consumption of both substrates within the RTZ suggest that ammonia oxidation (aerobic and anaerobic) and nitrate reduction take place simultaneously, and in close vicinity. In the following sections, we discuss the relevance of H 2 S and organic compounds (including CH 4 ) for N reduction, the N reduction pathway they fuel, as well as potential microbial players involved. We thereby elucidate the importance of closely coupled (and possibly cryptic) N- and S-cycling within the RTZ. Role of organic electron donors for N reduction Limited availability of reactive organic material (OM) for canonical denitrification was indicated in the unlabeled incubation experiments, where, in most cases, only a relatively small fraction of the added NO 3 - was reduced. In two incubations, NO 3 - was consumed completely within about one week, but NO 3 - reduction rates were low until two to three days into the experiment. In these bottles, we may have trapped sinking organic aggregates. In situ, they would have passed the RTZ in less than 2 days, based on sinking rates reported by Grossart and Simon (1998) , but in the incubations, the organic substrate within the aggregates could be exploited more efficiently by the organotrophic denitrifying microorganisms present. The 15 N-label incubation experiments confirm an overall relatively low denitrification activity. Even during the algal bloom in April 2018 (Fig. S2), when large amounts of OM were produced (46.5 g C m -2 ; www.cipais.org ), 15 NO 3 - added to water samples did not result in higher denitrification (or DNRA) rates. Also, the addition of acetate (an important intermediate of central metabolisms and an important compound during OM degradation) as electron donor only enhanced NO 3 - reduction after a lag phase, which was similarly observed by Wenk et al. (2013) . This suggests that organotrophic denitrifiers are present within the Lake Lugano RTZ, but are not very active. They respond, however, to organic-substrate addition (e.g., acetate) with growth and, hence, increased N reduction. We hypothesized that in spring/summer, during periods of higher OM production and export, organotrophic denitrification rates are higher compared to other seasons. However, this was not evident in our data set. Readily degradable OM, produced in the primary production zone at ∼5-25 m depth, as indicated by maximum O 2 and Chl a concentrations and low-nitrate conditions during algae blooms in April (Fig. S1, Fig. S2), is likely scarce in the RTZ for several reasons. A large portion of the freshly produced OM is already consumed in subsurface waters through aerobic respiration before even reaching the RTZ at 80-110 m depth. This is indicated by a strong decline of dissolved O 2 in and below the primary production zone in summer and autumn (Fig. S1a). As a consequence, only a small fraction of the exported OM, with a less accessible (i.e., less susceptible to hydrolysis and microbial degradation), organic compound composition, makes it to the RTZ in the relatively deep northern basin of Lake Lugano, also during periods of high primary productivity. Moreover, the residence time of sinking detrital OM in the RTZ might be too short for microorganisms to exploit the OM ( Grossart and Simon 1998 ). Indeed, an unrestrained transit of particles through the RTZ is likely because the RTZ in the Lake Lugano North Basin is not associated with a notable density gradient and has a weak water column stability (Fig. S1e), thus preventing any significant retention of sinking OM aggregates ( Alldredge and Cracker 1995 ). In summary, also during the productive season, availability of C org in the RTZ is likely low, limiting heterotrophic denitrification. We note that our 15 NO 3 - addition experiments without any additional electron donors, while consistent with previous reports by Wenk et al. (2014) , might slightly underestimate the in situ rates of denitrification, because volatile electron donors such as CH 4 and H 2 S were likely removed during the purging. Nevertheless, denitrification rates in the Lake Lugano North Basin (up to 113 nmol N-N 2 L -1 d -1 ) are at least one order of magnitude lower compared to several other lakes ( Table 1 ). For instance, maximum denitrification rates reached >8000 nmol N-N 2 L -1 d -1 in tropical Lake Kivu ( Roland et al. 2018 ), up to ∼4800 nmol N-N 2 L -1 d -1 in Lake Tanganyika ( Schubert et al. 2006 ), and up to 1000 nmol N-N 2 L -1 d -1 in Wintergreen Lake ( Burgin et al. 2012 ). The higher rates observed in other lakes could be due to increased temperature (e.g., Lake Kivu) and/or a shorter distance between the primary production zone and the RTZ, where less OM is degraded before reaching the active denitrification zone. For example, in Lake Kivu the RTZ lies between 40 and 50 m ( Roland et al. 2018 ) and in Wintergreen Lake at 4-5 m depth ( Burgin et al. 2012 ). Moreover, in relatively shallow RTZs, light penetration can enable OM production in situ, and sustain organotrophic denitrification, as observed in Lake La Cruz. (Oswald et al. 2016; Tischer et al. 2022). Lake Tanganyika has a similar O 2 -depletion depth as Lake Lugano of 80-90 m ( Schubert et al. 2006 ) and similar mean annual daily rates of primary production of 1 g C m -2 d -1 ( Hecky and Fee 1981 ). Yet, also in Lake Tanganyika, there is evidence for a deep secondary, anoxic Chl a maximum suggesting phototrophic growth near the RTZ, at least in the northern part of the basin, which may partly support denitrification (and anammox) (Ehrenfels et al. 2023). Moreover, besides the higher Chl a content at greater depths in Lake Tanganyika, and higher water column temperatures, other factors may be particularly conducive to higher rates of heterotrophic denitrification. More specifically, a relatively deep thermocline at 70 m depth ( Mziray et al. 2018 ) likely subserves an increased OM availability at the RTZ by promoting the retention of OM aggregates along the density gradient close to the RTZ. Sulfur-dependent N reduction We demonstrate a high potential of S-dependent DNRA in the 15 N-label experiments with up to 1707 nmol NH 4 + L -1 d -1 . Active S-dependent N reduction is also indicated by the presence of intermediates of S-oxidation, especially of S 0 and S 2 O 3 2- , which accumulated in the unlabeled incubation experiments and were present in situ with up to ∼0.25 µM S 0 and ∼0.10 µM S 2 O 3 2- . Contrary to S-dependent DNRA, S-dependent denitrification showed low potential to contribute to nitrate reduction, as in situ denitrification rates were generally low (up to 71 nmol N-N 2 L -1 d -1 ), and were not stimulated in the incubations with added 15 NO 3 - plus H 2 S, compared to the 15 NO 3 - -only treatments. Measured rates in the water column of other lakes revealed significantly different relative and absolute importance of DNRA and denitrification coupled to S-oxidation and OM-oxidation, respectively. For example, in Lake Kivu ( Roland et al. 2018 ) and in Wintergreen Lake ( Burgin et al. 2012 ), both organotrophic and S-dependent denitrification and DNRA (partly coupled to S-oxidation) have been detected. Yet, in both cases, DNRA accounted for only up to ∼15% of the total 15 N-turnover, while in Lake Lugano DNRA potentially contributed up to 97%. The OM limitation in the RTZ in Lake Lugano may competitively favor the activity of S-dependent N reduction. However, ambient sulfide concentrations were below the detection limit within the RTZ, and high DNRA rates were only observed upon H 2 S addition. Hence, the dominance of S-dependent DNRA in the water column under in situ conditions remains to be confirmed, for instance, by quantifying the in situ expression of the nrfA gene, a molecular marker for DNRA ( Pandey et al. 2020 ). The 16S rRNA gene sequencing analysis revealed a number of bacterial genera in the RTZ, which likely perform the observed S-dependent N reduction, like Sulfuritalea , with a relatively high abundance of up to 1%, as well as less abundant taxa such as Sulfuricurvum and Sulfurimonas . We speculate that the N-reducing Dechloromonas , which also possesses genes for the Sox sulfur-oxidation enzyme system ( Luo et al. 2018 ), and proliferated in the unlabeled incubations with added NO 3 - and H 2 S (Tischer et al. 2024/in prep.), may also be an important S-oxidizer in the RTZ, in addition to performing organotrophic denitrification. Based on 16S rRNA gene sequencing data, the observed sulfur speciation in the water column, and the H 2 S-addition experiments, we suggest that the most active zone for S-oxidizing N reducers may be a “cryptic-sulfur-cycling” zone, where S-oxidation and reduction are so tightly coupled that free H 2 S is undetectable. Cryptic sulfur cycling has already been reported for marine oxygen minimum zones, albeit with much lower rates ( Canfield et al. 2010 ; Callbeck et al. 2018). In the North Basin of Lake Lugano, in the 10 to 40 m thick anoxic water layer between the water depths where O 2 disappears and where H 2 S begins to accumulate, respectively, we detected S 0 , a high-rate potential for S-dependent DNRA, and maximum relative abundances of Sulfuritalea . We hence argue that H 2 S diffusing towards the RTZ is quantitatively oxidized by nitrate-reducing microbes performing DNRA. The volumetric sulfide oxidation rate that would be required to explain the observed flux of H 2 S toward the RTZ (85 to 304 µmol d -1 m -2 ) was, on average, 0.013 µmol H 2 S L -1 d -1 (assuming a 15 m thick layer of S-dependent denitrification activity; Table S2). The H 2 S consumption in the unlabeled anoxic incubation experiments, however, shows, with an average of 4.6 µmol H 2 S L -1 d -1 , a more than 350-times higher potential of H 2 S utilization than what would be required to completely oxidize the upward diffusing H 2 S in the water column. Furthermore, the presence of a community of S-reducing taxa, dominated by Desulfobacca ( Fig. 4k ), within, and below the RTZ, underscores the high potential for producing reduced S species in the same water-column region where the high S oxidation potential was observed. Hence, even in the non-sulfidic anoxic water column, microbial sulfate reduction represents a continuous source of sulfide, which is immediately consumed by S-oxidizing microorganisms. This process thus supports cryptic S-dependent DNRA, and to a lesser extent, S-dependent denitrification. Scope for methane-dependent N reduction CH 4 may serve as a viable alternative electron donor for microbial N reduction, indicated by its depletion within the RTZ and the elevated abundance and activity of Ca. Methylomirabilis in link with CH 4 oxidation stimulation by NO 2 - ( Su et al. 2023 ). Methane and nitrate concentration profiles overlap in a ∼10 m-thick water layer ( Fig. 1 ) where peaking relative abundances of Ca. Methylomirabilis were observed, underscoring that the basic hydro-biogeochemical and microbiological requirements for nitrite/nitrate reduction with methane are fulfilled. In addition to anaerobic N reduction by Ca. Methylomirabilis, micro-aerobic CH 4 -oxidation coupled to N reduction might occur, potentially by Crenothrix , which is abundant in the upper RTZ, as discussed in Su et al. (2023) . We did not perform CH 4 addition experiments to determine the potential for CH 4 -dependent nitrate/nitrite reduction in the water column. Nevertheless, some indication for such a microbial pathway is provided by the methane oxidation rates reported by Su et al. (2023) , which were ∼0.05-0.06 µmol L -1 d -1 , translating to a maximum CH 4 -dependent NO 2 - reduction rate of ∼0.08 µmol NO 2 - L -1 d -1 (assuming a stochiometric ratio of 3:5 for CH 4 :NO 2 - consumption; Ettwig et al. 2010). This rate would be in the same range as some of the denitrification rates determined in the presence of H 2 S. However, it must be considered as an upper-limit estimate because it assumes that all CH 4 is oxidized with NO 3 - as electron acceptor, and even this maximum possible rate is clearly lower than the observed DNRA rates determined under the same experimental conditions ( Table 2 ). Moreover, parallel investigations by our group assessing the role of CH 4 in N 2 O formation during denitrification in the same lake basin (T. Einzmann, unpubl.), showed that reductive N 2 O production is not stimulated by CH 4 , suggesting, indeed, a subordinate role of CH 4 as electron donor for denitrification. Cryptic N cycling through nitrification and DNRA coupling We suggest that N recycling via S-dependent DNRA and nitrification is a key mechanism in the RTZ of the Lake Lugano North Basin, which helps to retain large amounts of bioavailable N, while N 2 production by denitrification and anammox removes only a minor portion of the fixed N. High activity of ammonia and nitrite oxidation close to the oxic-anoxic interface is evidenced by the presence of a diverse and abundant community of ammonia and nitrite-oxidizers, including Ca. Nitrosopumilus. This archaeal ammonia oxidizer is common in marine environments ( Könneke et al. 2005 ), but has also been observed more recently in deep freshwater bodies such as Lake Constance and Lake Maggiore ( Coci et al. 2015 ; Klotz et al. 2022 ). Although nitrification rates remain to be quantified, maximum relative abundances close to the upper boundary of the RTZ indicate that Nitrosomonas or Nitrosospira encounter excellent conditions for nitrification, with replete O 2 and a constant supply of NH 4 + . Nitrospira might perform complete ammonia oxidation (comammox) from NH 4 + to NO 3 - ( Sakuola et al. 2021 ), and both Nitrosospira and Nitrosomonas can perform nitrifier denitrification, where NO 2 - produced from NH 3 is subsequently reduced to NO and N 2 O under low O 2 conditions ( Wrage et al. 2004 ; Shaw et al. 2006 ). Studies on other stratified lakes have shown evidence of active nitrification occurring in the respective RTZs ( Christofi et al. 1981 ; Pajares et al. 2017 ). More specifically, in the RTZ of a monomictic tropical lake, Pajares et al. (2017) demonstrated the co-occurrence of nitrification, DNRA, and denitrification genes, highlighting the importance of internal N recycling in the lake. Nitrification in the RTZ of the Lake Lugano North Basin is likely more important than previously reported by Wenk et al. (2013) , who interpreted the non-overlapping profiles of O 2 and NH 4 + (i.e., the quantitative consumption of NH 4 + below the redoxcline) as evidence that ammonium oxidation primarily occurs anaerobically, driven by anammox bacteria. However, overlapping O 2 and NH 4 + concentration profiles for 2015 to 2018 (this study), recent results from dual nitrate N and O isotope measurements (Tischer et al. 2024/in prep.), and an observed community shift of certain nitrifying taxa like Nitrosospira and Nitrosomonadaceae (Table S1), point towards an increased role of nitrification in the RTZ of Lake Lugano’s North Basin over the past decade. We also argue that, independent of the relatively low turbulent-diffusive fluxes of N compounds towards to RTZ (Table S2), the overall N turnover may be quite high, yet cryptic, due to efficient recycling between the ammonium and nitrate pools via DNRA and nitrification. Indeed, an average net consumption rate of not more than 0.055 µmol NO 3 - L -1 d -1 would have been necessary to explain the low observed nitrate flux towards the RTZ. In the NO 3 - -only treatment of the unlabeled incubations, however, potential NO 3 - reduction rates were much higher (average 0.7 µmol NO 3 - L -1 d -1 ), suggesting that the flux-based rate estimates underestimate the potential for NO 3 - reduction and that nitrate reduction is closely coupled to microaerobic nitrate regeneration. The latter, in turn, may be supported by DNRA, especially by S-dependent DNRA, which showed a high potential in the 15 N-label incubations. Cryptic N cycling, i.e., without measurable intermediates as nitrite or ammonium, has been demonstrated in oxic riverbeds ( Ooyang et al. 2021 ). In addition, Lam et al. (2009) demonstrated that a substantial fraction of anammox in the Peruvian oxygen minimum zone is supported by DNRA, without significant accumulation of NH 4 + . In the Lake Lugano North Basin of today, anammox arguably plays a subordinate role. Here, it is a close coupling between DNRA, nitrification, and denitrification that efficiently turns over fixed N and ultimately eliminates parts of it as N 2 . As with the “cryptic S cycle” mentioned above, we propose a closely linked “cryptic N turnover” involving S-dependent denitrification and DNRA in the RTZ water layer, where NO 3 - and H 2 S are not detected and NO 2 - usually does not accumulate. Electron donors from sediment fuel water column N cycle We argue that N reduction (denitrification and DNRA) in the North Basin of Lake Lugano is mainly driven by the constant supply of H 2 S and CH 4 , because readily accessible OM for organotrophic denitrification is scarce in the RTZ, as conceptualized in Figure 6 . As mentioned before, most of the OM from primary production is either already degraded in the upper oxic water column, or is not readily available as electron donor in the RTZ because sinking aggregates are not retained, and/or organic particles reaching the deeper hypolimnion are more refractory/stable (i.e., less prone to microbial attack). Instead, OM will accumulate in the permanently anoxic lake sediments, where it is further degraded through fermentation, SO 4 2- reduction, and methanogenesis, effectively transferring electrons to the benthic H 2 S and CH 4 pools (Blees et al. 2014). Microorganisms in the sediment are more abundant and the microbial community is more diverse compared to the water column ( Bartosiewicz et al. 2024 ). The microorganisms have essentially unlimited time to adapt their metabolism to, and exploit, specific compounds. In this context, the sediments and their natural microbial community may be seen as “electron donor refinery”, which turn a non-steady flow of less accessible electron donor compounds from the water column into a sustained production of H 2 S and CH 4 . These metabolites then diffuse out of the sediments towards the RTZ, where they serve as continuous and mostly season-independent electron donors for chemolithotrophic and CH 4 -dependent N reduction, respectively. The sustained benthic flux of electron-rich substrates thus allows for the establishment of a stable assemblage of N- and S-cycling microorganisms structured along a spatial sequence from oxic, to low O 2 , to anoxic conditions, including key organisms such as nitrifying Ca. Nitrosopumilus, Nitrosomonas , and Nitrospira and N reducing Denitratisoma , Sulfuritalea , and Ca. Methylomirabilis. There was little variation within these guilds of microorganisms for the duration of our study, probably due to the stable environmental conditions and reduced or absent grazing pressure in the lower part of the RTZ and the anoxic water layer. The concept of low, yet very constant, supply of H 2 S and CH 4 is supported by the low weighted UniFrac distances within the anoxic zone ( Fig. 3c ), indicating that the microbial community is more stable compared to the oxic water column, where grazing by e.g., protists and fluctuations in OM supply lead to a more variable community composition. We propose that this ultimately explains the dominance of S-, and possibly CH 4 , driven NO 3 - /NO 2 - reduction over canonical organotrophic denitrification within the RTZ of the Lake Lugano North Basin. Download figure Open in new tab Figure 6. Schematic illustration of N formation processes in the Lake Lugano North Basin coupled to the S cycle and the C-cycle. Solid lines indicate transformation processes, dashed lines indicate transport processes or symbolize the flow of electrons. Easily accessible fractions of fresh organic carbon (labile C org ) produced in the primary production zone are mostly degraded in the oxic water column before reaching the redox transition zone (RTZ), while the less accessible or more stable fractions (stable C org ) are incompletely oxidized and sink to the ground where they are degraded under anoxic conditions through fermentation, S-reduction and methanogenesis. Sulfide and methane, along with NH 4 + from organic N mineralization, diffuse to the RTZ, where they are used steadily as electron donors for denitrification and dissimilatory nitrate reduction to ammonium (DNRA). This way, the sediments serve as “electron donor refinery” that generates a steady flux of substrates for denitrification. Within the RTZ, N-cycling via nitrification and DNRA and S-cycling via S-oxidation and S-reduction likely takes place. Conclusions and implications We investigated cycling and elimination of fixed N, coupled to the S- and C-cycles, in the RTZ of the Lake Lugano North Basin. Incubation experiments, rate measurements, and 16S rRNA gene sequencing data uncovered a complex interaction of multiple oxidative and reductive N transformation processes, supported by a stable community of relatively few but abundant key microorganisms. We provide evidence that the meromictic North Basin of Lake Lugano, with its deep oxycline, creates conditions that favor sulfur- and methane-driven N reduction over canonical organotrophic denitrification. In particular, H 2 S appears to play an important role in DNRA within the RTZ. The factors that determine the relative importance of denitrification and DNRA under environmentally relevant substrate concentrations can vary and are not fully understood ( Pandey et al. 2020 ). While in other environments (e.g., lacustrine sediments), dissolved H 2 S has been shown to stimulate denitrification but not DNRA ( Cojean et al. 2020 ), we demonstrate here that H 2 S can strongly stimulate DNRA in a water column with limited OM availability. Nitrification, denitrification, DNRA, and reduction of S species take place in basically the same water mass around the oxic-anoxic interface, supporting an interactive, and to large parts “cryptic”, S- and N-cycling. This is the result of a diverse microbial community of S- and N transforming taxa, which can benefit from mutualistic interactions, e.g., through the continuous exchange of reduced and oxidized compounds. The close coupling and partitioning of the different NO 3 - -transforming metabolisms will determine whether fixed-N is recycled and remains in the system or is eliminated from it. Because DNRA (and nitrification), in contrast to denitrification, results in the retention of reactive N in the environment, our findings have important implications for the N budget of the Lake Lugano North Basin. More than 50 years ago, high inputs of P and N have shifted the lake into a meromictic and meso-eutrophic state. Despite a substantial reduction of external P inputs over the last 40 years ( Barbieri and Mosello 1992 ), the trophic state of the lake has only weakly improved ( Lepori et al. 2018 ). The relatively low fixed-N removal potential in the North Basin, coupled with efficient internal N cycling via DNRA, might contribute to maintaining elevated nitrate levels in the water column, thereby slowing down the restoration process of the lake. Data availability Raw sequence data are made available at NCBI under the BioProjectID PRJNA672280 with the accession numbers SRR12936362 through SRR12936382; MH111698 through MH113143 . Water column chemistry and experimental data are available on the Open Science Framework at https://osf.io/gfx7w/ . Author contribution statement JZ and MFL conceived the research project. JT, JZ, and MFL conceptualized research and experimental design. JT, JZ, MFL, GS and FL conducted sampling campaigns and contributed to the data acquisition of physicochemical parameters in the water column of Lake Lugano. The lake water incubation experiments were conducted by JT and analyzed by JT and JZ. JT, JZ, and GS carried out the data analysis of 16S rRNA sequencing data. JT wrote the paper, with substantial input from JZ, MFL, GS, and FL. JT, JZ, and MFL are accountable for the integrity of the data, analysis, and presentation of the findings. Acknowledgements We thank Marco Simona, Stefano Beatrizotti, Adeline Cojean-Egger, Maciej Bartosiewicz, and Lea Steinle for their help during the sampling campaigns on Lake Lugano. Long-term oxygen, nitrate, and primary productivity data were generated within a research program funded by the Dipartimento del Territorio (Department of Environment) of the Canton of Ticino, Switzerland, and the International Commission for the Protection of Italian-Swiss Waters (CIPAIS). We are also grateful to Thomas Kuhn and Judith Kobler for providing technical support in the laboratory. We also thank Teresa Einzmann for providing unpublished experimental data from Lake Lugano. 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