Versatile nitrate-respiring heterotrophs are previously concealed contributors to sulfur cycle

Domain: earth_science
Published: 2025-01-31
Editorial outcome: minor_revision

Abstract
Heterotrophic denitrifiers play crucial roles in global carbon and nitrogen cycling. However, their inability to oxidize sulfide renders them vulnerable to this toxic molecule, which inhibits the key enzymatic reaction responsible for reducing nitrous oxide (N2O), thereby raising greenhouse gas emissions. Here, we applied microcosm incubations, community-isotope-corrected DNA stable-isotope probing, and metagenomics to characterize a cohort of heterotrophic denitrifiers in estuarine sediments that thrive by coupling sulfur oxidation with denitrification through chemolithoheterotrophic metabolism. Remarkably, ecophysiology experiments from enrichments demonstrate that such heterotrophs expedite denitrification with sulfur acting as alternative electron sources and substantially curtail N2O emissions in both organic-rich and organic-limited environments. Their flexible, non-sulfur-dependent physiology may confer competitive advantages over conventional heterotrophic denitrifiers in detoxifying sulfide, adapting to organic matter fluctuations, and mitigating greenhouse gas emissions. Our study provides insights into the ecological role of heterotrophic denitrifiers in microbial communities with implications for sulfur cycling and climate change.

Introduction
Denitrification converts nitrate into dinitrogen gas via nitrite, nitric oxide (NO), and nitrous oxide (N2O). This process has been widely recognized as a primitive energy-conserving metabolism that profoundly influenced the early evolution of life prior to the oxygenation of the atmosphere1,2. Denitrification has significant environmental implications, as it helps regulate the nitrogen content in ecosystems and contributes to the nitrogen balance in the biosphere. Within marine and terrestrial ecosystems, denitrification contributes substantially to the loss of bioavailable nitrogen, therefore constraining global primary productivity, soil fertility, and crop yields3–5. Incomplete denitrification is one of the two major sources of N2O6, a potent greenhouse gas that contributes to climate warming. Additionally, N2O emissions contribute to ozone layer depletion and acid rain formation7.

In many estuaries, coastal regions, and marine oxygen-deficient zones, the sulfur cycle plays a crucial role in regulating biogeochemical processes. This is primarily due to the dominance of sulfate as a terminal electron acceptor and the subsequent release of hydrogen sulfide (H2S) from benthic sediments8,9. Studies have reported that sulfide, as a toxic chelator, strongly inhibits the copper-dependent metalloenzymes nitrite reductase (NirK) and N2O reductase (NosZ)10, adversely impacting denitrifying activity and potentially leading to increased N2O emissions11–13. At present, ubiquitous chemolithoautotrophic denitrifiers are recognized as the dominant biological contributors responsible for the oxidation of sulfur compounds and detoxification of sulfide14–19. Some of these chemolithoautotrophs are also involved in dissimilatory nitrate reduction to ammonium (DNRA), thereby supplying ammonium to support microbial growth of indigenous bacteria20,21. Most known heterotrophic denitrifiers are unable to detoxify sulfide due to their absence of enzymes necessary for sulfide oxidation13,22. However, recent evidence of bacterial isolates suggests that certain chemolithoheterotrophic denitrifiers, designated here as facultative sulfide-oxidizing heterotrophic denitrifiers (F-SOHDs), can oxidize sulfide, allowing them to thrive in sulfidic environments23,24. A notable example is the recently identified chemolithoheterotroph Arcobacter peruensis23, which has been shown to play a significant role during sulfidic events off the coast of Peru. Despite the energetic advantage and detoxification potential of F-SOHDs, their presence and ecological roles in other natural ecosystems remain poorly understood. Thus, questions remain regarding how heterotrophic denitrifier consortia adapt to sulfide stress within complex microbial communities and their ecological impact on regulation of N2O emissions.

In this study, we investigate the response of heterotrophic denitrifiers to sulfide in the highly productive Songhua River Estuary, a major tributary within the largest water system in East Asia. By employing a combination of microcosm and lab culture experiments, metagenomics, and DNA stable-isotope probing (DNA-SIP) approach, we describe a previously hidden cohort of heterotrophic denitrifiers that use sulfur compounds as supplemental electron donors. This capability allows them to detoxify sulfide and achieve complete denitrification, thereby mitigating the N2O emission. This work enhances our understanding of heterotrophic denitrifiers capable of sulfide detoxification in aquatic environments, with broader implications for regulating carbon, nitrogen, and sulfur cycles, including the mitigation of N2O emissions in both natural and engineered system.

Results
Both sulfide and organic matter fuel nitrate reduction in estuarine sediments
Estuarine sediments were sampled at the junction of the Songhua River and effluent from municipal treatment in 2019 (Supplementary Fig. 1a). Despite the low concentration of sulfide in situ (<0.02 mM), sulfur cycling was active with 46.2 μM of sulfide produced in 20 mL (~24.2 g) of surface sediments over 52 h (Fig. 1a). These sediments contained 2.17 ± 0.64 × 104 copies of dsrA, encoding dissimilatory sulfite reductase subunit A, per gram of sediments (Fig. 1b). Sulfide addition (53.8 μM) resulted in a 30.2% increase in nitrate consumption (Fig. 1c). We therefore conjectured that microbes inhabiting the community may be capable of reducing nitrate with sulfide as an electron donor. As the incubation reached a point of relatively slow nitrate reduction, potentially due to the depletion of electron donors, organic matter (OM) was introduced, triggering an instant decrease in nitrate (Fig. 1d). The concomitant increase of dinitrogen gas (94.2 μM), N2O (167.3 μM), and ammonium (14.6 μM) suggests that OM promoted both complete and incomplete denitrification, as well as DNRA. The high abundance of narG, nirS, norB, nosZ, and nrfA genes reinforces this inference (Fig. 1b). Thus, provision of the electron donors sulfide and OM simultaneously boost nitrate respiration in the estuary.

Unexpected sulfur oxidation by heterotrophic denitrifiers
To better understand interactions between the carbon, nitrogen, and sulfur cycles, we conducted DNA-SIP in microcosms with estuarine sediments that were amended with combinations of carbon (C, with both inorganic and organic carbon added), nitrate (N, with NO3− added), and sulfide (S, with S2− added): +C+N+S, +C+N, +Ci+N+S, +Ci (only inorganic carbon provided), and +Co (only organic carbon provided) (Supplementary Fig. 2). To determine whether chemolithoautotrophs or heterotrophs drive nitrate reduction, we conducted DNA-SIP in microcosms with either inorganic (13Ci, 13CO2 and bicarbonate-13C) or organic composite (13Co, glucose-13C, propionate-13C, and acetate-13C) carbon labeled, along with unlabeled carbon substrates (12Ci and 12Co). These incubations led to increases in bulk DNA buoyant density along with 16S rRNA gene copies as determined by qPCR, showing that both inorganic 13Ci and organic 13Co were incorporated into cells in both +C+N+S and +C+N microcosms (Fig. 2a, b). In comparison to the heavy fraction without 13C labeling in the +C+N+S microcosm, 13Co labeling led to increases of 1.3%, 21.6%, and 3.8% in the relative abundances of Thauera, Azoarcus, and Pseudomonas in the heavy DNA fraction, but these taxa decreased by 0.1%, 6.1%, and 1.5% in relative abundance when provided with 13Ci, suggesting a possible heterotrophic lifestyle (Fig. 2c). Similar increases (0.7–29.8%) in these taxa were observed in +C+N microcosm accompanied by 13Co assimilation (Fig. 2d). When starved of nitrate, very little increase in these three genera was observed in the heavy DNA fraction regardless of the presence of OM (Fig. 2e). We therefore conjectured that in addition to heterotrophy, they might be also capable of reducing nitrate. Unlike most other studies suggesting that sulfide oxidation coupled to denitrification is primarily driven by chemolithoautotrophs in freshwater, estuarine, and coastal environments14,25–27, the identified heterotrophic denitrifiers including Azoarcus and Pseudomonas increased 13Co assimilation in response to sulfide addition, with their relative abundances increasing by 64.1% and 8.0% in (12Ci+13Co)+N+S treatment compared to (12Ci+13Co)+N treatment (Fig. 2c, d). This appears to suggest that these heterotrophs contribute to both sulfur oxidation and denitrification in our system. Despite this, interestingly, the metabolic trait of these two genera in 13Co assimilation with reliance on nitrate in non-sulfide-involved (12Ci+13Co)+N treatment (Fig. 2d, e), indicates that these heterotrophs are capable of denitrification irrespective of sulfide addition. In contrast, the provision of 13Ci in +Ci+N+S and +Ci+N+S microcosms consistently led to 4.5- and 4.6-fold increases in Thiobacillus in the heavy fraction (Fig. 2c, f), whereas Thiobacillus did not increase in that of the +Ci microcosm (Fig. 2g), suggesting a primarily chemolithoautotrophic lifestyle with reliance on nitrate. Collectively, these findings underscore the metabolic versatility of Azoarcus and Pseudomonas as heterotrophic denitrifiers capable of sulfur oxidation and nitrate reduction. Meanwhile, Thiobacillus has a more specialized chemolithoautotrophic role, highlighting the diverse metabolic strategies present within the microbial community.

In addition to 16S rRNA genes, we tracked the gene abundance of biomarkers for nitrate reduction (napA, narG), nitrite reduction (nirS, nirK), nitrous oxide reduction (nosZ), sulfide oxidation (sqr), and thiosulfate oxidation (soxB) along DNA density gradients (Fig. 2h–m and Supplementary Fig. 3). The high proportion of these genes in the heavy fractions in (13Ci+12Co)+N+S and (12Ci+13Co)+N+S treatments illustrated that both chemolithoautotrophs and heterotrophs contributed to nitrate reduction and sulfur oxidation (Supplementary Fig. 3a–g). Compared to the treatment amended only with 13Co, the significantly higher abundance of sqr and nirS gene copies in heavy fractions of (12Ci+13Co)+N+S treatment suggests the possible contribution of heterotrophic denitrifiers to sulfur oxidation (Fig. 2i, j, l, m). Intriguingly, in the (12Ci+13Co)+N treatment, both sqr and soxB were highly enriched in heavy DNA fractions, with 30.9% and 17.8% abundance (Fig. 2i and Supplementary Fig. 3m, n). This indicates that the sqr-/soxB-encoding heterotrophic denitrifying microorganisms are stimulated when organic carbon and nitrate are provided even without sulfide (Supplementary Fig. 3j–n). These findings suggest that heterotrophic denitrifiers could potentially engage in sulfur oxidation without relying on it, as potential F-SOHDs. By linking element cycling functions inferred from isotope incorporation in the microcosms to relative abundance of individual genera, we found that 60.8% of the denitrifying sulfur-oxidizer community was composed of heterotrophs, while only 12.1% were chemolithoautotrophs (Supplementary Fig. 4). Compared to the +C+N microcosm, the relative abundance of Thiobacillus, the most abundant chemolithoautotroph in the community, decreased by an average of 3.8% in the +C+N+S microcosm (Fig. 2c, d). This suggests that F-SOHDs outcompete chemolithoautotrophs when sulfide is present. Taken together, these microcosm data imply that heterotrophs, rather than obligate chemolithoautotrophs, may dominate sulfur oxidation and denitrification in this ecosystem.

Metabolic potential of F-SOHDs revealed by metagenomics
To better understand the metabolic potentials of key microbes identified by DNA-SIP, metagenomic sequencing was conducted on heavy DNA fractions from microcosms and natural sediments. A total of 63 high- and medium-quality representative metagenome-assembled genomes (MAGs) were reconstructed (Fig. 3a–c and Supplementary Table 1). Functional annotation revealed that 15 MAGs, primarily from Desulfobacterota, Betaproteobacteria, Gammaproteobacteria, Bacillota, and Bacteroidota, represent heterotrophic sulfur oxidizers harboring near-complete denitrification or DNRA pathways (Fig. 3a, c). Virtually all of these MAGs encode either periplasmic nitrate reductases (napAB) or membrane-bound nitrate reductases (narGHI), indicating the widespread potential for nitrate reduction (Supplementary Fig. 5). Approximately 60% of those MAGs harbor one or two nitrite reductases encoded by nirS and nirK, accompanied by norBC, nosZ, and accessory/regulatory genes (e.g., nosR), pointing to denitrification capacity. In contrast to most known heterotrophic denitrifying bacteria that are unable to use sulfur compounds as electron donors28–30, these MAGs also encode various enzymes for sulfur oxidation (Fig. 3c and Supplementary Fig. 5), the most prevalent being sulfide:quinone oxidoreductase (SQR), a key enzyme that can catalyze sulfide detoxification in nitrate-reducing chemolithoautotrophs19,31. Additionally, genes encoding thiosulfate:cyanide sulfurtransferase (TST) and periplasmic sox gene clusters encoding for thiosulfate oxidation and sulfane dehydrogenation were also detected15,32. Flavocytochrome c sulfide dehydrogenase (FccAB), which has a role in oxidative sulfur metabolism in photolithoautotrophic and chemolithoautotrophic sulfur bacteria33,34, was also abundant. The presence of sulfur dioxygenase (ETHE1) in partial MAGs suggests that zero-valent sulfur or polysulfide can also be oxidized to sulfite by glutathione35.

Unlike well-studied chemolithoautotrophs including Sulfurimonas19, Thiobacillus36, and SUP05 groups37 that are widely distributed and capable of both denitrification and sulfur oxidation, most denitrifiers in the present study lack known CO2 fixation pathways (Fig. 3c and Supplementary Fig. 5), herein identified as F-SOHDs. Consistent with the absence of known autotrophic pathways, M4.bin9 (Pseudomonas), M2.bin44 (Ralstonia), M5.bin12 (Dechloromonas), M5.bin61 (Thauera), M5.bin41 (UBA2357), M4.bin17 (Azoarcus), and M5.bin1 (Azoarcus) MAGs were more abundant in metagenomes of heavy DNA fractions amended with 13Co than those with 13Co and unlabeled-12C treatments (Fig. 3b), implying their enrichment on OM and incorporation of 13Co to support growth. While these results point to a predominantly heterotrophic lifestyle, some members of these genera might possess the capacity for CO2 fixation under specific conditions, though this possibility cannot be confirmed due to the incomplete genomes reconstructed from the metagenomic sequencing. The detection of genes diagnostic of heterotrophy including fermentations and carbohydrate mineralization in many of the MAGs is also consistent with the heterotrophic lifestyle of these microbes (Supplementary Figs. 5, 6). Most of them also encode enzymes to synthesize acetyl-CoA from acetate via acetyl-CoA synthetase (Acs) and/or acetate kinase-phosphate acetyltransferase (Ack-Pta), both of which could be coupled with the formation of ATP38,39. The presence of genes encoding glycoside hydrolases, glycolysis, β-oxidation, and propionyl-CoA production in these MAGs suggests that organic molecules such as carbohydrates, long-chain fatty acids, and propionate can be utilized as carbon sources to conserve energy and support heterotrophic growth40–42. Our findings suggest a strong genomic potential for F-SOHDs within these sediments at the study site, supporting their responses in the microcosm experiments. This highlights the importance of previously unrecognized heterotrophic denitrifiers in the cycling of both sulfur and nitrogen. Apart from denitrifiers, several heterotrophs encoding DNRA and sulfur oxidation were also identified (Supplementary Fig. 7). Some ammonium was produced in nitrate respiration assays (Fig. 1d), and nrfA genes were uncovered by qPCR (Fig. 1b) and in MAGs (Fig. 3c), confirming DNRA in certain microbes.

Long-term enrichment cultures confirm the non-sulfur-dependent physiology of F-SOHDs
A custom bioreactor system was used to enrich F-SOHDs from sediments based on alternating medium with and without sulfide (Fig. 4a). Two enrichments named ENR_C4 and ENR_U2 were successfully maintained for 612 days. Kinetic tests showed that approximately 140–300 μM of sulfide, soluble zero-valent sulfur, thiosulfate, and sulfite could be oxidized to sulfate in ENR_C4, accompanied by the consumption of OM and an increase in cell numbers (Supplementary Figs. 8a–I, 9). Though both enrichments reduced nitrate along with OM consumption, the poor net cell growth suggested that microbes inhabiting the ENR_U2 community were only able to oxidize sulfide, but not other sulfur compounds (Supplementary Fig. 10). Two representative MAGs identified as Pseudomonas (ENRC.bin3) and Thauera (ENRC.bin26), with a cumulative relative abundance of 88.6% in the ENR_C4 enrichment culture (Fig. 3b; Supplementary Fig. 8k), were interpreted to be F-SOHDs. Interestingly, Thauera (ENRC.bin26) was enriched in response to sulfide addition and possessed a complete sulfide oxidation pathway despite decreasing in relative abundance in the 13Co-incorporated DNA fraction (Figs. 2c, d, 3c). We speculate that the pattern of enrichment of Thauera (ENRC.bin26) may be attributed to the competition with other microorganisms, such as Pseudomonas (ENRC.bin3), under certain environmental conditions. It is possible that high sulfide concentrations lead Thauera to be outcompeted by Pseudomonas, thereby lowering its relative abundance and contribution to the overall community. This pattern has been observed in benthic marine waters, where the sulfide oxidizers Arcobacter, SUP05, and Sulfurovum co-exist at low sulfide levels, but Arcobacter and Sulfurovum dominate over SUP05 when sulfide concentrations exceed 10 μM10.

To further verify the coupling of denitrification with sulfur oxidation, we performed three consecutive incubations, each with different substrates provided (Fig. 4b). In the first stage, the addition of OM and nitrate into ENR_C4 led to a complete removal of nitrate and rapid decrease in OM. Then in the second stage, sulfide and nitrate were added. About half of the sulfide was completely oxidized to sulfate, which was concomitant with further consumption of both OM and nitrate, consistent with sulfide oxidation coupled with heterotrophic denitrification. When harvested cells from this enrichment were transferred to sulfide-free medium in the third stage, heterotrophic denitrification capacity in the absence of sulfide was restored. This experiment suggests key denitrifiers in enrichments modify their metabolism to adjust to the presence or absence of sulfide. OM was necessary for sulfur oxidation since nitrate was stable in the absence of OM, whereas it decreased upon replenishment of OM (Fig. 4c). This was further supported by the higher net increases in cell numbers when cultures were amended with OM (Student’s t test, P = 0.004 and 0.002; Fig. 4d, e).

Overlooked significance of F-SOHDs in sulfide detoxification and reducing N2O emission
To verify the ecological significance of F-SOHDs, we examined biochemical cycling within different laboratory cultures (Supplementary Fig. 11a–x). As OM flux is known to control nitrogen loss43, experiments were conducted under either OM-rich or OM-limited conditions. Although minimal sulfide was consumed in enrichment cultures of conventional heterotrophic denitrifiers (Fig. 5a and Supplementary Fig. 11f, t), sulfide was completely consumed within 32 h in enrichment cultures dominated by F-SOHDs (enrichments ENR_C4 and ENR_U2), regardless of OM concentrations. The contributions of OM and sulfide as electron donors for denitrification were evaluated (Fig. 5b, c). Analysis of electron flow balance indicated that although OM dominates (Supplementary Fig. 11a, d, p, r), approximately 24.6–27.1% of electrons supporting denitrification could be generated by the oxidation of sulfide to zero-valent sulfur (Fig. 5b). With the provision of sulfide, less OM was consumed by F-SOHDs than by conventional denitrifiers (Fig. 5d). To a certain extent, this implies that despite the necessity of OM for growth, sulfide might be an important supplemental electron donor. When fed with limited OM, two F-SOHDs can theoretically incorporate an additional 34.7–102.4% of electrons from sulfide (Fig. 5c), promoting complete denitrification (Supplementary Fig. 11w, x). With this, we conjecture that, rather than simply suppressing heterotrophic denitrifiers, sulfide might serve as an alternative electron source for certain denitrifiers that can oxidize it to less toxic sulfate/sulfur, thereby mitigating or preventing sulfide toxicity and potentially accelerating the denitrification process. These extra electrons could overcome a bottleneck imposed by limited reducing power under low OM conditions that can lead to incomplete denitrification by conventional heterotrophic denitrifiers. For example, OM-limited conditions led to only 16.3% of nitrate being reduced to dinitrogen gas in conventional heterotrophic denitrifier enrichment (Supplementary Fig. 11s), whereas complete denitrification was observed in an OM-limited culture containing F-SOHDs when they were supplemented with sulfide (Supplementary Fig. 11w).

Interestingly, further experiments suggests that the F-SOHDs could significantly lower the release of the greenhouse gas N2O. In these experiments, conventional heterotrophic denitrifiers produced 55.8% more N2O and less dinitrogen gas when exposed to only 4.83 μM sulfide (Fig. 5e, f and Supplementary Fig. 11c–f). This is consistent with the reported observations of excess N2O accumulation reported by sulfide suppression in the Shinji Estuary11. Remarkably, negligible N2O emission and comparable levels of nitrogen in the form of dinitrogen gas and initial nitrate were detected in two F-SOHD enrichments, suggesting a complete denitrification process when sufficient OM is provided (Fig. 5g and Supplementary Fig. 11o, q). Although slow, chemolithoautotrophic denitrifiers also achieved the full nitrate reduction to dinitrogen gas (Supplementary Fig. 11g). In contrast, in the presence of sulfide, the conventional heterotrophic denitrifiers released more N2O, accounting for approximately 50% of the reduced nitrogen (Supplementary Fig. 11e). When OM is deficient, the additional electrons released from the complete oxidation of sulfide to sulfate, compared to partial oxidation to sulfur, can give rise to lower 72.6% N2O emission and 42.0% higher production of dinitrogen gas by F-SOHDs (Supplementary Fig. 11u, w). Our results highlight that F-SOHD is a crucial sink for mitigating greenhouse effects by reducing N2O emissions through harnessing various substrates as electron donors to support complete denitrification.

Methods
Estuary sampling and dynamic incubation
Estuarine sediments were collected in the main nutrient-exchanging hotspot zone belonging to Eastern Asia’s largest Songhua River System in August of 2019 (45°49′12.81″N, 126°43′22.51″E). Five replicate sediment cores were sampled at 0–20 cm depth, with a water depth of 3.7 m. The surface sediments contained 48.35% sand, 22.57% total organic matter, 1.34% total nitrogen, 0.07% total phosphorus, and had a pH of 7.06. One overlying water sample was fixed immediately using sulfide antioxidant buffer73 to prevent sulfide oxidation, while in another sample sulfide was precipitated by zinc chloride to filter for other chemical analyses. The anoxic bottom water contained 0.75–0.98 mM sulfate, 0.04–0.06 mM nitrate, 0.002–0.005 mM nitrite, 0.08–0.13 mM ammonium, and 1.03–1.79 mM total organic carbon at pH 7.13. The concentrations of sulfate, nitrate, and nitrite were measured using an ICS-3000 ion chromatograph (Dionex, USA), with detection limits of 0.2 μM for sulfate, 0.3 μM for nitrate, and 0.2 μM for nitrite. Ammonium was determined by the standard Nessler reagent method, with a detection limit of 1.8 μM. Total organic carbon was determined using a multi N/C 3100 (Analytik Jena, Germany), with a detection limit of 0.4 μM.

Sediment incubation experiments were performed in-situ to test the activity of element cycling in the estuary. Sediments for the measurements of sulfate reduction rate were rapidly filtered on-site through a homogenized mesh and then transferred into 340 mL glass-bottles with overlying water. A homemade assembly, featuring a three-way valve connected to an airbag pre-filled with 15 L pure helium and a vacuum pump, was employed for negative vacuum gas replacement. After inoculation, sediment microcosms were then incubated on-site on a thermostatic shaker at 22°C before transportation to the lab. Liquids were periodically sampled and stored on dry ice for later physiochemical analysis. Sulfate reduction activity in estuarine sediments was assessed by sulfate consumption and sulfide production over a per unit time using 20 mL of sediments. To determine relationships between nitrate respiration and sulfur cycle, equivalent volume of sediments was inoculated into two bottles of estuarine water, and then the nitrate concentration was initialized by diluting a stock solution. After deoxygenation, a group of bottles was additionally amended with a mixed sulfur stock solution with a 1:1 molar ratio of sulfate to sulfide to reach a concentration of ~50 μM sulfide compared with the control. While sediments for OM additions were similarly inoculated in anaerobic estuarine water that was initialized with nitrate and then incubated for 44 h at which time nitrate consumption became sluggish based on the limited dissolved organic carbon in the estuarine water. At that point, 800 μM of a composite OM stock solution, comprising equimolar glucose, propionate, acetate, and lactate, was added. The decrease in nitrate concentration and the production of dinitrogen gas and ammonium before and after OM amendments were then compared to determine the rate of nitrate respiration coupled to OM utilization in the estuary. All field sediment incubations were conducted in three repetitions.

Quantitative PCR of biomarkers
DNA was extracted from 0.25 g of sediment using PowerSoil DNA isolation kit (MoBio, USA). Ten genes involved in sulfate reduction (dsrA), sulfide oxidation (sqr, soxB), nitrate reduction (napA, narG), denitrification (nirS, nirK, norB, nosZ), and DNRA (nrfA) were quantified and details regarding the primers are described in Supplementary Table 2. Cell numbers from each density gradient were estimated by counting the absolute 16S rRNA gene copies. All qPCR assays were performed in triplicate in a 20 μl reaction, containing 10 μL of TB-Green Premix Ex Taq (Takara, Japan), 0.4 μL of ROX Reference Dye, 0.4 μL of each forward/reverse primer (10 μM), 2 μL of standardized DNA template (17.5 ng μL−1), and 6.8 μL of DNAase-free water. The reactions were carried out on a StepOnePlus ABI 7500 PCR system (Applied Biosystems, USA). DNA concentrations were determined using the Quant-iT PicoGreen dsDNA kit (Thermo Scientific, USA). Recombinant plasmids with a cloned target gene were ten-fold diluted for an eight-point standard calibration curve ranging from 10 to 108.

Isotope incubation experiments
Prior to conducting microcosm experiments, sediments were rinsed with deoxygenated Milli-Q water within the BACTRONI-2 Anaerobic Workstation (Shel Lab, USA) to eliminate insoluble sandy particles and extracellular residues22. To stimulate microbial activity within the communities, the sediments underwent a dark preincubation in estuarine water, within a headspace of N2 to CO2 gas ratio of 4:1, maintained at 22°C for a period of 5 days. Microcosms were amended with various combinations of inorganic carbon (Ci, CO2 in gas-phase and bicarbonate in liquid-phase), organic carbon (Co, 5 mM mixture of acetate, glucose, and propionate), nitrate (0.30 mM), and sulfide (0.05 mM) along with the basic additions of ammonium (0.10 mM), sulfate (0.75 mM), disodium phosphate (0.10 mM), and HEPES buffer. To delineate the contributions of chemolithoautotrophs and heterotrophs, 13Ci (98–99% labeling percent, Sigma-Aldrich, USA) and 13Co (99% labeling percent, Cambridge Isotope Laboratories, USA) were used in SIP incubation experiments (Supplementary Fig. 2). No copper ligand complex was supplied in trace elements to prevent the selective enrichment of denitrifiers with NirK74. During incubation, 700 μL of liquid was routinely collected for chemical determination, while 400 μL of headspace was taken for N2, N2O and methane analysis. Each microcosm was incubated up to three generation times (~10–12 days). During medium replacement after each cycle, a custom-designed hose with on-off switch was connected between microcosm vessel outlet and a washing bottle containing deoxygenated phosphate buffer solution to maintain the strict anoxic environment22. Following a negative pressure exchange, fresh isotope substrates slowly flowed into the incubation bottle, and the headspace was simultaneously refilled with helium gas.

Function identification and community isotope response correction
Approximately 3 μg of DNA was mixed by vortexing with CsCl solution to achieve a standard density of 1.725 g mL−1. Isopycnic ultracentrifugation was performed with a VTi 65.2 vertical rotor (Beckman, USA) at 20°C, 174,000 × g for 55 h. DNA from each fraction of the gradients was collected through sterile water displacement, resulting in 14 individual fractions. Buoyant density was determined by calculating the calibrated refractive index measured with a AR200 refractometer (Reichert, USA). To recover the trace DNA mixed with CsCl solution, DNA purification was performed through precipitation using glycogen (Roche, USA) and polyethyleneglycol 6000 (Sigma-Aldrich, USA). Both 16S rRNA and functional genes were quantified across varying DNA buoyant densities in each microcosm. The identified heavy fractions were pooled for subsequent DNA extraction and high-throughput sequencing.

We introduced a approach to identify and quantify functions of individual microbes within a community and their roles in the cycling of all three elements. The fundamental principle of this approach involves comparing variations in the relative abundance of a given microorganism in heavy DNA fractions across different treatments, where the disparity in treatments serves as an indicator of the microbe’s metabolism in each of the treatments. For instance, a significant increase in the relative abundance of species A in the heavy fractions of a +C+N+S microcosm compared to a +C+N microcosm would lead to the conclusion that species A is capable of sulfide oxidation. Through simultaneous assessments of community isotope response, the functions of each taxon, encompassing carbon (inorganic carbon fixation or OM assimilation), nitrogen (nitrate reduction), and sulfur (sulfur oxidation) metabolisms, were determined within the complex community. For a given taxon, we performed following comparisons: (1) microbes are considered chemolithoautotrophs if they increase in relative abundances in heavy fractions in both (13Ci+12Co)+N+S compared to (12Ci+12Co)+N+S and 13Ci+N+S compared to 12Ci+N+S treatments; (2) microbes are considered heterotrophs if they increase in relative abundances in heavy fractions of both (12Ci+13Co)+N+S compared to (12Ci+12Co)+N+S and (12Ci+13Co)+N compared to (12Ci+12Co)+N treatments; (3) microbes are capable of nitrate reduction and sulfur oxidation if they increase in relative abundances in +Ci+N+S compared to +Ci microcosms; (4) microbes have the capacity to oxidize sulfide if they increase in relative abundances in +C+N+S compared to +C+N microcosms. Here, an increase in (13Ci+12Co)+N+S compared to (13Ci+12Co)+N and (12Ci+12Co)+N+S treatments suggests a chemolithoautotroph capable of sulfide oxidation, while an increase in (12Ci+13Co)+N+S compared to (12Ci+12Co)+N+S and (12Ci+13Co)+N treatments suggests a heterotroph capable of sulfide oxidation.

16S rRNA gene sequencing
After DNA extraction, the V3–V4 regions of bacterial 16S rRNA gene were amplified by using the primer set 341F (CCTACGGGNGGCWGCAG) and 805R (GACTACHVGGGTATCTAATCC)75. Amplicon sequencing was performed on an Illumina MiSeq sequencer to generate 2 × 300 bp paired-end reads, with raw reads ranging from 54,985 to 104,902 per sample. Sequences were clustered into OTUs at the 99% sequence similarity level using Mothur (v1.34.3) (https://github.com/mothur/mothur/releases/). Representative sequences were taxonomically assigned by searching against SILVA (v132) rRNA reference database (https://www.arb-silva.de/) and NCBI nucleotide collection (nt) database (https://www.ncbi.nlm.nih.gov/) using the Naïve Bayesian algorithm in RDP classifier. For the enrichment sample, near full-length 16S rRNA gene was amplified using primer set 27F (AGRGTTYGATYMTGGCTCAG) and 1492R (RGYTACCTTGTTACGACTT)76. After blunt-end-ligation, the SMRTbell library (PacBio, USA) was constructed and then purified for sequencing via dedicated PacBio Sequel II 8M cells. Circular consensus sequencing (CCS) reads were generated using a PacBio Sequel II sequencer.

Metagenomic assembly and genome binning
DNA samples from heavy fractions following incubations with 13Ci-/13Co-labeled compounds and 12Ci-/12Co-natural abundance compounds, raw sediments, short-term isotope-inoculated sediments, and ENR_C4 enrichments were collected for metagenome sequencing. Trace amounts of heavy DNA in SIP experiments were amplified by using the multiple displacement amplification technique77. Chimeras were removed based on the general approach of Chen and colleagues78 using Phi29 DNA polymerase and S1 nuclease. Metagenomic sequence data for each sample were generated using Illumina HiSeq X Ten instruments (PE150 mode) with an insert size of ~450 bp. Raw reads with adapters, duplicated sequences, and low-quality nucleotides at both ends (average phred value < 20) were preprocessed using homemade scripts79. Metagenomic sequences for each sample were de novo assembled using SPAdes80 (v3.15) with parameters “-k 21, 33, 55, 77, 99, 127 --meta”. The sequence depth was calculated by mapping quality reads to the assembled scaffolds using BBMap (https://sourceforge.net/projects/bbmap/) with the following parameters: minid=0.97, local=t. Scaffolds with length ≥ 2,500 bp were used for binning using MetaBAT281, MaxBin282, and CONCOCT83. Default parameters were applied and both tetranucleotide frequency and sequence depth were considered. The best MAGs were refined using DAS Tool84 (v1.1.1). MAGs assembled from different samples were combined and dereplicated using dRep85 to generate representative bins. To optimize the genome quality, clean reads were recruited using BBmap with the same parameters as above. Genome bins were subsequently reassembled by SPAdes with following options: “--careful -k 21, 33, 55, 77, 99, 127”. All bins were manually curated by removing scaffolds with multiple copies of marker genes and abnormal sequence depth. A total of 63 representative genomes with relatively high completeness (>80%) and low contamination (<5%) evaluated by CheckM86 were retained for further analysis.

Gene calling was conducted for each MAG using Prodigal (v. 2.6.3) with the “-p single” option. Functional annotations were performed by searching predicted genes against the KEGG, NCBI-nr, carbohydrate-active enzymes, and eggNOG databases using DIAMOND87 with 1e−5 as the E-value cutoff. Predicted genes were also used to search against the KOfam88 database using kofamscan v1.3.0 for metabolic pathway reconstruction. Genes associated with heterotrophic lifestyle and environmental adaptation were validated by BLASTing against a recent dataset40. Relative abundance of MAG was determined by dividing the sum of coverage for each scaffold in a single MAG by the sum of coverage for all scaffolds in the belonging sample.

Genome- and SIP-informed enrichment of F-SOHDs
A combination of bioreactors and serum bottle cultures were employed to enrich for F-SOHDs. To sustain the relatively high biomass, the cultivation was first conducted in a custom-designed sequencing batch reactor (SBR). Approximately 120 mL of sediments was inoculated into a 950 mL of the SBR for a five-month (~152 days) continuous enrichment (Fig. 4a). Considering the functional flexibility of F-SOHDs, which are able to grow under both sulfide and sulfide-free conditions, the medium was periodically alternated between HED and CNS conditions. Here, HED refers to culture medium containing OM (composite OM, including glucose, propionate, acetate, and fermentative DL-lactate) and nitrate, while CNS represents the same medium supplemented with additional sulfide. During the enrichment process, sterile deionized water was used instead of filtered river water, along with essential salts and other necessary nutrients (see media composition in the “Isotope incubation experiments” section), to maintain greater control and prevent impurities from interfering with the growth of F-SOHDs. Throughout long-term operation, pure argon gas was continuously bubbled from the bottom of the reactor to expel CO2 released through microbial respiration, thereby preventing the undesired growth of chemolithoautotrophs.

After 152 days of cultivation in a SBR, we switched to a dilution-transfer approach to further purify F-SOHDs. This process was carried out within 120 mL serum bottles sealed with butyl rubber stoppers. A 20 mL of the culture was transferred to flasks provided with HED medium. Following growth in HED medium, 2 mL of culture was inoculated in triplicate into anaerobic Hungate tubes containing 18 mL of CMD medium to test for chemolithoautotrophic metabolism. This medium replicates the optimal growth conditions for the well-known chemolithoautotrophic sulfide-oxidizing denitrifier, Thiobacillus denitrificans ATCC 25259 (450 T2 medium), where sulfide, nitrate, and bicarbonate are provided. Viable cells were counted using Vi-Cell XR coulter (Beckman, USA) with trypan blue dyeing, and the results were validated through FACSCalibur flow cytometry analysis (Becton Dickinson, USA). For treatments with low cell numbers, the one exhibiting a lowest sulfide oxidation rate was regarded as a negative response to the CMD condition. It was then chosen and inoculated into the fresh CNS and then HED medium at a volume dilution ratio of 1:6 for the next enrichment cycle. Other treatments showing a positive response were discarded in the subsequent enrichment. After 67 cultivation cycles (~460 days), two stable enrichments named ENR_C4 and ENR_U2 were ultimately acquired. For these two enrichments, various sulfur sources, including sulfide, zero-sulfur, thiosulfate, and sulfite, were separately provided for kinetic tests under both heterotrophic and chemolithoautotrophic conditions.

Physiological experiments
Experiments were conducted with seven different cultures to determine the effect of F-SOHDs on sulfur and nitrogen cycles under controlled conditions through tracking the biochemical transformation and greenhouse gas emission. The cultures were: HD, an enrichment culture containing conventional heterotrophic denitrifiers with no sulfur-oxidation activity (previously enriched from sediments in laboratory); CD, a pure culture of the chemolithoautotrophic sulfur-oxidizing denitrifier T. denitrificans ATCC 25259 (purchased from The Global Bioresource Center); ENR_U2 and ENR_C4, two enrichment cultures dominated by F-SOHDs with incomplete and complete sulfide oxidation capacity; and CE_H, CE_E, and CE_L contain mixtures of the HD culture with the CD T. denitrificans in ratios of 3:1, 1:1, and 1:3 (synthetic microbial communities). To establish environments with rich and limited OM, we maintained carbon/nitrogen ratios of 3.52 and 0.70. An automatic sampling needle, and two precalibrated ultra-trace N2O-100 and SULF-100 microsensors (Unisense, Denmark) were used for continuous monitoring the concentration of soluble N2O and H2S in liquid phase. Greenhouse gases (N2O, CO2, and CH4) in the headspace were analyzed using an Agilent 7890B gas chromatography (USA) equipped with two series-wound packed columns (J&W G3591-81121 and G3591-81004). The detection limits were 0.8 ppm for CH4, 0.2 ppm for CO2, and 0.05 ppm for N2O. Dinitrogen gas was measured using a separate Agilent 7890A gas chromatography (USA) equipped with a thermal conductivity detector and J&W G3591-81121 packed column, with a detection limit of 2.6 ppm. The completion extent of denitrification process was calculated with the following equation:

N0 = ΔNO3− / (ΔNO3− + ΔNO2− + ΔN2O + ΔNH4+) × 5 + ΔNO2− × 2 + ΔN2O × 4 + ΔNH4+ × 8 / CiNO3− × 5 (1)

where N0 denotes the completion extent of denitrification. ΔNO3− indicates the net nitrate concentration consumed, and other nitrogen compounds after triangle indicate the net concentration produced. CiNO3− denotes the initial nitrate concentration, and its value multiplied by 5 represents the theoretical total electron demand for complete reduction of nitrate to dinitrogen gas.

The proportion of electrons contributed by sulfide to facilitate the complete denitrification was calculated with the following equation:

PS = ΔS0 × 2 + ΔSO32− × 4 + SO32− × 6 + ΔSO42− × 8 / CiNO3− × 5 (2)

where PS denotes contribution of sulfide, as an electron donor to promote the complete reduction of nitrate to nitrogen. All sulfur compounds after triangles indicate the net produced concentration. The concentrations of the anions sulfate, sulfite, and thiosulfate were determined using an ICS-3000 ion chromatograph (Dionex, USA) equipped with an IonPac AS11-HC analytical column. Sulfide was measured by the standard methylene blue method at a wavelength of 665 nm89, while zero-valent sulfur was quantified through ultraviolet-visible spectrophotometric analysis (Shimadzu, Japan) at a custom wavelength of 282 nm90.

Phylogenetic tree and gene analysis
The phylogenomic tree was inferred based on 31 conserved protein markers91 identified by AMPHORA292. Genomes belonging to the phylum Planctomycetota were downloaded from GenBank (74 available in July 2021) and were chosen as outgroup. For each marker, multiple sequence alignment was performed using MUSCLE93 (v3.8.31) and poorly aligned regions were trimmed using TrimAL with options “-gt 0.95 -cons 50”. Maximum-likelihood-based phylogeny was reconstructed based on concatenated markers using IQ-TREE94 with ultrafast bootstrapping (-bb 1000 -m LG + R6). The best-fitting model was determined by ModelFinder95.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This manuscript found the important contribution of heterotrophic denitrifiers on N2O reduction in natural ecosystem,\nespecially in estuaries environments. Based on authors data, heterotrophic denitrifiers seemed to exert a more significant\nrole compared to chemolithoautotrophic denitrifiers, which was completely beyond folks’s traditional understanding. These\nfindings help folks further comprehend the interaction between C, N and S cycles. Before its publication, the followings\nshould be addressed:\n1. Line 88-89, why authors sampled the Estuarine sediments at the junction of the Songhua River and effluent from\nmunicipal treatment? And in Fig.S1 the detailed sample site was not provided.\n2. line 91, what was the quality of 20 mL of surface sediments?\n3. In Fig.1d, the N amounts were balanced or not?\n4. line 171-177, please provide more details on this deduction of heterotrophs may dominate sulfur oxidation and\ndenitrification in this ecosystem.\n5. line 222-229, author mainly focused on the contribution of denitrification processes, and they also found DNRA process in\nthe sediments. Thus, how about the DNRA process contribution on both NO3- reduction and S2- oxidation?\n6. line 299-303, more details information are required to support authors’s deduction.\n7. line 351-353, please provide more similar sites, I do not believe that only 3 similar sample were enough for supporting\nauthors’s opinion.", "reviewer_2": "The study by Shao et al., explores the presence of facultative sulfur-oxidizing heterotrophic denitrifiers, abbreviated by the\nauthors as F-SOHDs, in estuary sediments of the Songhua River System. Specifically, their study employed DNA-SIP\nexperiments, in combination with metagenomic sequencing and enrichment cultures to identify F-SOHD members in a\nmixed microbial community comprising chemoorganoheterotrophs and chemolithoautotrophs. Using this range of\ntechniques, they identified environmental genera affiliated with Thauera, Pseudomonas, and Azoacus as likely F-SOHDs in\ntheir samples. Lastly, they demonstrate using F-SOHD enriched bioreactors, that such members are capable at better\nsuppressing N2O emissions than their heterotrophic denitrifying counterparts. In turn, this led to the suggestion that F-\nSOHDs could be useful in industrial settings to mitigate N2O emissions.\nAs someone who has previously studied F-SOHDs, which I refer to as chemolithoheterotrophs, I find the work to be of high\nvalue as it expands on the potential for chemolithoheterotrophy in the environment. Moreover, the authors employed an\nimpressive range of tools to identify these microbes in situ. Overall, I find the results fairly compelling, albeit the manuscript is\nnot well-rounded at times, and does tend to overstate claims. Namely, the introduction did not provide the reader with state-\nof-the-art background information concerning chemolithoheterotrophs. The results I generally find compelling, although I\nhave some points that need further clarification, and the authors should discuss some of the discrepancies in their data.\nWhile I don’t disagree with all the points raised in the discussion, it does need some more work/reframing, to present a more\nwell-rounded case for chemolithoheterotrophy in the environment. In addition, some errors in their estimates of Gibbs free\nenergy need to be addressed in the discussion. The method section was at times thorough, but other parts were fairly vague,\nto the point that the reader would not be able to reproduce the experiment without additional clarification. Ending on an\noptimistic note, the figures are nicely polished, and the data/conclusions (while overstated at times) appear generally sound.\nComments\nLines 40-41: I find this statement to be a little vague, namely because heterotrophic denitrifiers operate in environments\nwhere sulfide is present. But what the authors could specify is that sulfide inhibits a key enzymatic reaction that catalyzes the\nreduction of N2O to N2. Hence, heterotrophic denitrifers are sensitive unless sulfide is removed, either by cooperation with\nchemolithoautotrophs or with chemolithoheterotrophs. This type of denitrifying community structure is indeed what we find in\norganic and sulfide-rich ecosystems.\nLine 43: This statement comes across as a little too strong as their role in the environment, including aquatic ecosystems,\nhas been noticed by some and even empirically demonstrated (Sorokin 2003; Boltyanskaya et al. 2007; Callbeck et al.\n2019; Wang and Shao 2021). As I mentioned above, the value of the study is not in that it discovered sulfide oxidation by\nheterotrophs in the environment, it is that it demonstrates that numerous microbes in estuary sediments could thrive via a\nchemolithoheterotrophic metabolism greatly expanding its potential. I would therefore consider re-framing this part.\nLines 48-51: I agree with this statement, but this largely repeats the findings of Callbeck et al., 2019, which I detail a little\nlater in this review. Moreover, their justification of this statement based on their Gibbs free energy calculations appears to be\nflawed (see later discussion).\nLines 51-53: I’m in favor of this work generally, but this is an overstatement. Firstly, environmental examples of sulfide\noxidation by heterotrophs exist, thus the use of “paradigm shift” is over the top. Secondly, I understand that if you work on\nN2O you can make a link to climate change, however, to say that his work has “profound implications” for climate change is a\nstretch too far. For example, it is not the case that the authors have tested/modeled their reactors under various climate\nchange scenarios. In my view, the work is a good piece because it advances the idea that chemolithoheterotrophs form a\npersistent part of the microbial community in an organic matter-rich estuary.\nLines 62-64: For clarity, denitrification is one of two major processes, nitrification being the other. The statement makes it\nseem as though denitrification is the only one.\nLines 65-68: Fully agree, for a comprehensive review of sulfur cycling and its influence over the nitrogen cycle in oxygen\nminimum zones please see (Callbeck et al. 2021).\nLines 71-73: The authors only superficially mention that it impacts N2O but how is this the case? Some have speculated that\nsulfide also inhibits the last step of denitrification, as sulfide acts as a Cu-chelator.\nLines 68-70: Certainly, chemolithoautotrophs are well-recognized, but work by Callbeck et al., 2019 has shown that both\nchemolithoautotrophs, and chemolithoheterotroph (aka. F-SOHD) can co-exist to oxidize sulfide during a sulfidic event off\nthe coast of Peru.\nLine 72: I understand the point the authors are making, however, for clarification sulfide is toxic to all denitrifiers, including\nnitrate-reducing chemolithoautotrophs, chemolithoheterotrophs and organoheterotrophs. The reason for this I explain a little\nlater in the discussion section below.\nLines 73-75: This statement, which is effectively the premise of the manuscript, is simply not up to date with the state-of-the-\nart. As I mentioned above, it’s not as though we know nothing about this topic of F-SOHD/chemolithoheterotrophs. The\nauthors have partly relayed this information in the discussion (on lines 368-370), but choose not to interject this information\nhere or above when background information is needed on this specialized metabolism. Again, the work is valuable not\nbecause it is the first to discover F-SOHD, but because it expands the potential for this metabolism to operate in the aquatic\nrealm. The authors should consider reworking this paragraph to include the state-of-the-art, and perhaps frame the premise\nin a slightly different way.\nLines 80-83: To be specific, the study identifies new F-SOHDs at a sediment site. Somewhat misleading is the use of\n“ubiquitous” as this implies that the authors provided evidence of F-SOHDs widespread distribution across multiple\nsampling sites, although their study site is positioned at one location (45°49′12.81″N, 126°43′22.51″E). Again, the authors\nhave a habit of overstating claims, when in fact, there is no need.\nLines 82-85: I find this overly broad and introduces a bit of speculation, especially on the climate change front; can the\nauthors be a bit more specific?\nLines 98-101: If I have this right, this would amount to roughly two-thirds of the original 300  M of nitrate being recovered as\nN2 and NH4+. But roughly one-third is not recovered, hence, both complete and incomplete denitrification are evident in this\nexperiment.\nLine 108: The with and without sulfide addition experiments are somewhat confusing. I understand that the authors added\nadditional sulfide, but it should be perhaps reiterated that sulfide is going to be produced in the background, unless the\nauthors removed the sulfate for sulfate reducers, in incubations like C+N, or Ci, or Co etc…\nLines 117-121: The increases of 1.3% and 3.8% don’t seem to be substantial for Thauera and Pseudomonas. Thus, can the\nauthors assume that their heterotrophs according to these values? And in the second part, how much did these members\ndecrease when amended with 13Ci? In light of chemolithoautotrophic abundances increasing with the addition of 13C,\ncould this not artificially dilute such heterotrophs like Thauera and Pseudomonas, if it’s based on relative abundances?\nLines 121-122: Can the authors comment on why Thauera comprised a large fraction of the microbial community in the\nexperiment void of sulfide (C+N) when compared to the C+N+S experiments? It would appear that this member is sensitive\nto sulfide inhibition. In addition, an increase of 0.7% is practically negligible. Moreover, it appears that Pseudomonas shows\na different pattern that is opposite to your statement. Also in Fig. 2d, Pseudomonas and Azoarcus both have 13Ci which is\nhigher than the control in grey. Hence, this goes against the idea that no 13C-DIC is incorporated. Part of the problem the\nauthors might encounter in their analysis is that heterotrophs can also assimilate 13C-DIC (e.g. Erb 2011), which might\ncomplicate their analysis, or at least require a note.\nLines 125-129: Thiobacillus increased in Fig. 2d (C+N) when sulfide was not present, which runs counter to their\nchemolithoautotrophic lifestyle. And also what about other chemolithoautotrophic members such as Chromatiales\n(M5.bin31) and Sulfurivermis (M5.bin64) shown in Fig. 3?\nLines 130-154: It seems that this method distinguishes Thiobacillus, but what about other chemolithoautotrophic members\nsuch as Chromatiales (M5.bin31) and Sulfurivermis (M5.bin64) shown in Fig. 3, where do they position in Fig. 2i? In\naddition, organisms like Dechloromonas, which according to Fig. 3 is an F-SOHD, are positioned in the bottom left of Fig. 2i\nalongside Thiobacillus the chemolithoautotroph. I also have difficulties in reconciling the very negligible increases the\nauthors reported on lines 118 and 121 and how this results in very clear patterns for Thauera and Pseudomonas in Fig. 2i.\nI think the authors should add that this method might not be perfect at disentangling all F-SOHDs from other community\nmembers, but it may work for more active microbes.\nLines 138-142: The exception would be Arcobacter peruensis, which reduces nitrate, oxidizes sulfide, and simultaneously\nassimilates/oxidizes organic matter into biomass for energy/growth (Callbeck et al. 2019). Arcobacter peruensis was also\nabundant in sulfidic and organic matter-rich shelf waters alongside chemolithoautotrophs like SUP05.\nLine 142-144: In the methods section, on lines 488-490, the authors state that “microbes have the compacity to oxidize\nsulfide if they increase in relative abundances in +C+N+S compared to +C+N microcosms”, but the issue with Thauera is\nthat its abundance decreases, and hence, by their logic above is not a sulfide oxidizer. The authors could further clarify this.\nLine 171: The authors coin the new acronym facultative sulfur-oxidizing heterotrophic denitrifiers (F-SOHDs) for a metabolic\nprocess that has been described previously as chemolithoheterotrophy (Callbeck et al. 2019). For consistency’s sake with\nprevious work, why not term F-SOHDs as chemolithoheterotrophs which include microbes that oxidize sulfide, and reduce\nnitrate while simultaneously utilizing organic carbon as an energy/carbon source for growth? This would reduce the number\nof acronyms in the literature, and make for a more comparable term to chemolithoautotrophy, which the authors readily apply\nin their manuscript.\nLines 204-207: The authors mention Arcobacter, but it was later been shown to grow as a chemolithoheterotroph (Callbeck et\nal., 2019). Hence, Arcobacter peruensis had the capacity to assimilate organic carbon, reduce nitrate, and oxidize sulfide,\nmuch like what the authors describe in their paper for Thauera and other members.\nLines 208-213: I checked the literature on some of the closest affiliated microbes in Figure 3 (see below), and some of them\nhave capacities for CO2 fixation, despite being labeled as having no capacity in Fig. 3c.\n- Azoarcus (see https://www.kegg.jp/pathway/aza00710+AZKH_p0225)\n- Ralstonia pickettii DTP0602 (https://www.kegg.jp/pathway/rpj00710+M00166)\n- Pseudomonas yangmingensis (Wong and Lee 2014)\nThus, while I believe that some are probably chemolithoheterotrophs, the authors should stipulate some caveats and\nunknowns here.\nLines 222-225: As mentioned above the authors should apply some caution here. Moreover, the authors have not measured\nthe nitrate turnover rates of the specific members, thus to say that this cluster is more biogeochemical relevant is rather\nspeculative. It’s fair to say that strong potential for chemolithoheterotrophy exists at their study site.\nLines 244-249: I find it difficult to take the position that Thauera is sensitive to sulfide. If the authors take this position then\nthey have to reconcile in their discussion that both nitrate-reducing chemoorganoheterotrophs and chemolithoheterotrophs\nare sensitive to sulfide. Presently, they hold only the former to be true. However, I would suggest that the authors explain this\ndifferently. For example, sulfide-oxidizers like Arcobacter, SUP05 and Sulfurorvum can co-exist in the presence of sulfide\nhowever Arcobacter and Sulfurovum tend to dominate over SUP05 when sulfide concentrations exceed 10 uM (Callbeck et\nal., 2021). So could it not be the case that Pseudomonas outcompetes Thauera at a certain sulfide threshold, thereby\nlowering its contribution to the overall community (based on relative abundances). Taking this position means that Thauera\nis not necessarily sensitive but rather outcompeted under certain environmental conditions by others.\nLines 266-267: This is a bit confusing here as it is not clear why being a heterotrophic denitrifier leads to N2O production,\nand also why chemolithoautotrophs do not. In my mind it’s not so clear cut, for example, SUP05 identified in Peru can\nperform complete denitrification to N2 (Callbeck et al. 2018), while other SUP05 species in the Saanich Inlet can only do\nnitrate reduction to N2O (Walsh et al. 2009). Might also be worth mentioning here that Arcobacter peruensis is an exception,\nplus others exist as mentioned earlier.\nLines 285-287: This line contradicts the author’s statement above that heterotrophic members like Thauera could be\nsensitive to sulfide.\nLines 305-321: I agree with the authors that DNA-SIP technology is important to unraveling microbial ecology-related\nquestions. But I find a dedicated paragraph to this in the opening discussion is a little excessive, as the method is in\nprinciple the same with the difference being that the authors did an additional correlative analysis (Fig. 2i, g). In addition,\norganisms like Dechloromonas, which according to Fig. 3 is an F-SOHD, are positioned in the bottom left of Fig. 2i\nalongside Thiobacillus the chemolithoautotroph. Hence, this method is not perfect, and the authors (if they decide to keep\nthis piece) will need to address the caveats. As a recommendation, I would instead suggest the authors shorten or drop to\nmake way for a more detailed discussion of the specific chemolithoheterotroph members they identified in their study, which\nwas lacking in my opinion (see below).\nLine 323: I would add, “Epsilonproteobacteria and the Gammaproteobacteria SUP05 clade”\nLine 331: There are more pertinent studies to reference here that discuss sulfidic events in marine environments, which is\nsummarized in the review by Callbeck et al., 2021.\nLines 332-333: I believe this to be an overstatement by the authors “…uncovered here have evolved multiple sulfur oxidation\npathways to cope with sulfide stress.” The authors have not empirically demonstrated sulfide stress on the community and\nhow it leads to the evolution of multiple sulfur oxidation pathways. Rather this statement could be framed as a hypothesis.\nLines 322-340: I see the importance of a wider description of sulfidic events, but this really appears as a zoomed-out\nintroduction piece especially since the nosZ gene discussion below brings more detail to this idea. Instead, I think the\nopening of the discussion would be better served reiterating what defines a F-SOHDs/chemolithoheterotroph. Namely,\nchemolithoheterotrophs couple sulfide oxidation to nitrate reduction while simultaneously utilizing organic carbon as an\nenergy/carbon source (hence the assimilation of 13C into biomass in your DNA-SIP experiments). Next, I would reiterate the\nkey putative chemolithoheterotrophs mentioned in the results section and Fig. 3. Subsequently, the authors need to spend\nmore time describing what is already known about these species in the literature, so that the reader can assess what new\ninformation is added in this study. For example, my cursory inspection of some of the literature finds that some of these\nmembers (identified in Fig. 3) are known sulfide oxidizers like Pseudomonas stutzeri (Li et al. 2022). Thauera is well-studied\nand has been isolated from sulfide-rich environments and bioreactors before, implicating it already with a role as a putative\nsulfide oxidizer. Others are capable of fixing CO2 autotrophically such as Azoarcus (see\nhttps://www.kegg.jp/pathway/aza00710+AZKH_p0225), Ralstonia pickettii DTP0602\n(https://www.kegg.jp/pathway/rpj00710+M00166) and Pseudomonas yangmingensis (Wong and Lee 2014). I recommend\nthe authors incorporate a much more thorough discussion of the key members they identified.\nLines 345-346: I agree that sulfide has an influence over N2O production, but the statement here that sulfide only affects\nheterotrophic denitrifiers and not denitrifiers more broadly – chemolithoautotrophs included – is not evidenced in the\nliterature. Much of this discussion has taken place in the review by Callbeck et al., 2021, but to reiterate their points sulfide is\na chelator of copper, and the nosZ gene is a copper-rich metalloenzyme. The sequestration of copper by its abiotic reaction\nwith sulfide will induce copper limitations that result in nitrate reduction terminating at N2O in microbial cultures (Granger\nand Ward 2003; Manconi et al. 2006; Moffett et al. 2012; Felgate et al. 2012). The accumulation of N2O has therefore been\nlinked to copper limitations induced by sulfide, which would affect nitrate-reducing bacteria (with a nosZ) irrespective of\nwhether they are chemolithoautotrophs, chemolithoheterotrophs or heterotrophs.\nLines 346-349: I agree with this statement, but as above this is not exclusively a heterotrophic denitrifier issue but will rather\naffect denitrifiers broadly.\nLines 349-353: Indeed, sulfide-driven denitrification is likely responsible for major N2O hotspots (Naqvi et al. 2000, 2006;\nArevalo-Martinez et al. 2015; Kock et al. 2016; Bourbonnais et al. 2017; Arévalo-Martínez et al. 2019), which likely occur as\na result of the mechanism outlined above. However, the difficulty I have with this line is the authors claim that these\nN2O/H2S-rich systems are dominated by heterotrophic denitrifiers, when in fact, the cited studies do not make that claim\n(mainly because they also pre-date large community metagenomic surveys). Secondly, in light of more recent analyses of\nthese same environments, such as Peru, where we have major N2O hotspots along with the presence of sulfide, the\nmicrobial community is comprised of a consortium of nitrate-reducing chemolithoautotrophs, chemoorganoheterotrophs and\nchemolithoheterotrophs. Thirdly, from a logical perspective, if such systems have high organic matter, sulfide, and nitrate,\nwouldn’t they offer the perfect ecological niche for chemolithoheterotrophs to thrive? To that point, Callbeck et al. 2019, and\nthis study enriched for chemolithoheterotrophs from exactly these environments.\nLines 353-356: I generally agree with this line, but perhaps clarify that this occurs because chemolithoheterotrophs can\nremove sulfide and therefore remove its ability to chelate copper, enabling N2O reduction to N2 to continue (via the Cu-rich\nnosZ).\nLines 357-359: I agree, but it would make sense to mention that sulfide provides an additional electron donor (on top of\norganic matter) to reduce available nitrate to N2 rather than to N2O.\nLines 356-363: I agree with this statement, but it should be pointed out that chemolithoautotrophs have the same advantage\n(as your work also demonstrates in Fig. 5g). In other words, this feature is not unique to chemolithoheterotrophs.\nLines 363-365: As I stipulated above, this is complicated and not so clear-cut. The environment where Arcobacter peruensis\nwas identified, was from a sulfidic event off the coast of Peru (Callbeck et al., 2019). Here, the environment was defined as\nbeing sulfide-, N2O-, and organic matter-rich. Hence, the conditions where we find chemolithoheterotrophs are also\nenvironments where relatively high N2O concentrations persist. To better frame this one could say that\nchemolithoheterotrophs/chemolithoautotrophs might contribute to suppressing some N2O in sulfidic environments that are\nassociated with high N2O production rates.\nLines 368-370: I think the introduction, results, and discussion would be improved if the authors made a greater attempt to\ndraw parallels between the microbes identified here, and the recently described Arcobacter peruenesis as well as the other\nones mentioned earlier. I found it somewhat unfortunate that the authors when discussing organic matter and sulfide-rich\nenvironments, failed to mention Arcobacter peruenesis in that context, but rather discussed chemlithoautotrophs like\nSUP05.\nLines 370-375: Could the authors be a little more specific? For example, what pathways did they identify in these microbial\ngenomes, that made them comparable to the chemolithoheterotrophs described here?\nLines 375-378: The authors did not touch on what features make chemolithoheterotrophs more adaptable over\nchemolithoautotrophs, so can they be more specific? Moreover, it appears that the authors have mixed up the Gibbs free\nenergy values from their figure. According to the authors Figure 6, the authors have calculated a Gibbs free energy of -494\nand -579 for F-SOHDs related processes (rxns. C1 and C2), and with chemolithoautotrophy estimated at -566 (rxn. b). But in\nthe text, they list the Gibbs free energy of heterotrophy instead of chemolithoautotrophy. This mix-up makes it seem that F-\nSOHDs activity is more energetically favorable when in reality it’s not according to Figure 6.\nWith that said the authors have overlooked an important feature that gives chemolithoheterotrophs an advantage over\nchemolithoautotrophs. As detailed by Callbeck et al., in 2019, the assimilation of acetate into biomass is energetically more\nfavorable than the assimilation of CO2 via a chemolithoautotrophic metabolism. This is because CO2 fixation pathways\nrequire substantial amounts of ATP (Erb 2011). The favorable energetics of a chemolithoheterotrophic metabolism explain\nhow Arcobacter can rapidly bloom in the sulfide- and organic-rich upper shelf waters off the coast of Peru and Namibia\n(Lavik et al. 2009; Callbeck et al. 2019, 2021). This is a theme the authors could consider exploring in the context of the\nmicrobes identified in their study.\nLines 378-381: Their exceptional performance hinges on their ability to rapidly bloom over chemolithoautotrophs for\navailable sulfide, this is something to keep in mind. Overall, I find that the authors could be a bit more specific regarding their\nindustrial application reference. Do the authors mean to use chemolithoheterotrophs to remove N2O in sulfide, and organic\nmatter-rich engineered systems?\nLines 381-383: Fully agree, and I think this is where you could end the discussion. I find the next line regarding climate\nchange and its evolutionary trajectories highly speculative and rather vague.\nMethods:\nLine 389: What month specifically did sampling take place? What was the water depth of the study site?\nLines 390-392: Do the authors have a reference to support its biogeochemical significance to the nitrogen cycle?\nLine 393: Why did the authors pick the depth between 0-20 cm? What is the nitrate and oxygen penetration depth?\nLine 394: What is sulfide antioxidant buffer, please be specific.\nLines 396-398: I would convert from mg L-1 to uM/mM to be consistent with the rest of the manuscript. Also, how were these\nnutrient concentrations measured? And by which methods? What were the detection limits?\nLines 404-407: What were the incubation temperatures? And do they differ from in situ temperatures?\nLine 420: How was dinitrogen gas measured?\nLines 424: How much sediment was used for the extraction? Keep in mind that the readers need to be able to repeat the\nexperiments.\nLines 424-428: The authors provide no mention of references either in the qPCR section or supplementary Table 2\nregarding the primers. Surely, these primers were designed elsewhere and if so, the authors need to indicate this\ninformation.\nLines 428-429: Is this common to do in qPCR, because cells often have more than one gene copy? So, the cell numbers are\ngoing to be overestimated?\nLines 430-432: What qPCR chemistry was used, and how much DNA template was used? Were DNA concentrations also\nverified, and how? For the standards what concentration range was used in genes per uL? Was melt curve analysis also\ndone?\nLines 434-445: Is this a completely novel technique or is there other studies that have done the same?\nLines 443-445: What was the 13C labeling percent for the incubation experiments?\nLine 488: Change “compacity” to “capacity”\nLines 495-497: Please provide a reference to the study that designed these primers. And also indicate if these are universal\nprimers or primers used to only amplify the Bacterial community.\nLines 497-500: Can the authors indicate the number of raw reads generated?\nLines 504-506: Please provide a reference to the study that designed these primers.\nLines 514-515: Is there a reference for this line?\nLines 551-5552: Perhaps I missed this somewhere, but what is the composition of the medium in terms of salts and other\nnutrients. Or is the medium just filtered river water with amended substrates? Please clarify.\nLines 555-558: If CO2 is removed, how is the system then pH buffered? Based on the principle that chemolithoautotrophs\nare slower growing compared to chemolithoheterotrophs I don’t fully understand the reasoning behind removing CO2 to\nhinder chemolithoautotrops. As we know, chemolithoheterotrophs when supplied sulfide, nitrate, and organic matter will\nnaturally outcompete chemolithoautotrophs for available sulfide (Callbeck et al., 2019; 2021). So, actively removing CO2\ncould have downstream consequences for pH buffering in the system. Do the authors have pH measurements?\nLine 580: This could be shortened to read, “Physiological experiments”.\nLines 583-585: Do the authors know which heterotrophic denitrifiers were enriched? And do they lack a capacity for sulfide\noxidation and have the nosZ gene? I’m wondering how distinct they are from the sulfide-oxidizing heterotrophs.\nLines 595-596: The authors should provide more information regarding the instruments and standards used for gas analysis.\nLines 608-611: This is too vague. Please provide citations and additional information on how these sulfur species were\nquantified.\nFigures:\nFigure 6: How was Gibbs free energy calculated and where did they get the formation energies from? Under what conditions\nwas this calculated, for example, pH, sulfide, nitrate concentrations? Reaction c2 is driven by mostly sulfide (1:5 ratio), with a\nlittle bit of organic matter, which seems more like a chemolithoautotrophy with a little bit of heterotrophy. What does this look like if the\nstoichiometry is a bit more balanced?\nLiterature cited in my review:\nArevalo-Martinez, D. L., A. Kock, C. R. Loscher, R. A. Schmitz, and H. W. Bange. 2015. Massive nitrous oxide emissions\nfrom the tropical South Pacific Ocean. Nat. Geosci 8: 530–533. doi:10.1038/ngeo2469\nArévalo-Martínez, D. L., T. Steinhoff, P. Brandt, A. Körtzinger, T. Lamont, G. Rehder, and H. W. Bange. 2019. N2O\nEmissions From the Northern Benguela Upwelling System. Geophys. Res. Lett. 46: 3317–3326.\ndoi:10.1029/2018GL081648\nBoltyanskaya, Y. V, V. V Kevbrin, A. M. Lysenko, T. V Kolganova, T. P. Tourova, G. A. Osipov, and T. N. Zhilina. 2007.\nHalomonas mongoliensis sp. nov. and Halomonas kenyensis sp. nov., new haloalkaliphilic denitrifiers capable of N2O\nreduction, isolated from soda lakes. Microbiology 76: 739–747. doi:10.1134/S0026261707060148\nBourbonnais, A., R. T. Letscher, H. W. Bange, V. Échevin, J. Larkum, J. Mohn, N. Yoshida, and M. A. Altabet. 2017. N2O\nproduction and consumption from stable isotopic and concentration data in the Peruvian coastal upwelling system. Global\nBiogeochem. Cycles 31: 678–698. doi:10.1002/2016GB005567\nCallbeck, C. M., D. E. Canfield, M. M. M. Kuypers, P. Yilmaz, G. Lavik, B. Thamdrup, C. J. Schubert, and L. A. Bristow. 2021.\nSulfur cycling in oceanic oxygen minimum zones. Limnol. Oceanogr. 66: 2360–2392. doi:https://doi.org/10.1002/lno.11759\nCallbeck, C. M., G. Lavik, T. G. Ferdelman, and others. 2018. Oxygen minimum zone cryptic sulfur cycling sustained by\noffshore transport of key sulfur oxidizing bacteria. Nat. Commun. 9: 1729. doi:10.1038/s41467-018-04041-x\nCallbeck, C. M., C. Pelzer, G. Lavik, and others. 2019. Arcobacter peruensis sp.nov., a chemolithoheterotroph isolated from\nsulfide and organic rich coastal waters off Peru. Appl. Environ. Microbiol. 85: e01344-19. doi:10.1128/AEM.01344-19\nErb, T. J. 2011. Carboxylases in Natural and Synthetic Microbial Pathways. Appl Env. Microbiol 77: 8466–8477.\ndoi:10.1128/aem.05702-11\nFelgate, H., G. Giannopoulos, M. J. Sullivan, A. J. Gates, T. A. Clarke, E. Baggs, G. Rowley, and D. J. Richardson. 2012.\nThe impact of copper, nitrate and carbon status on the emission of nitrous oxide by two species of bacteria with\nbiochemically distinct denitrification pathways. Environ. Microbiol. 14: 1788–1800. doi:10.1111/j.1462-2920.2012.02789.x\nGranger, J., and B. B. Ward. 2003. Accumulation of nitrogen oxides in copper-limited cultures of denitrifying bacteria. Limnol.\nOceanogr. 48: 313–318. doi:10.4319/lo.2003.48.1.0313\nKock, A., D. L. Arévalo-Martínez, C. R. Löscher, and H. W. Bange. 2016. Extreme N 2 O accumulation in the coastal oxygen\nminimum zone off Peru. Biogeosciences 13: 827–840. doi:10.5194/bg-13-827-2016\nLavik, G., T. Stührmann, V. Brüchert, and others. 2009. Detoxification of sulphidic African shelf waters by blooming\nchemolithotrophs. Nature 457: 581–584. doi:10.1038/nature07588\nLi, P., W. Yuan, Y. Huang, C. Zhang, C. Ni, Q. Lin, Z. Zhu, and J. Wang. 2022. Complete genome sequence of\nPseudomonas stutzeri S116 owning bifunctional catalysis provides insights into affecting performance of microbial fuel cells.\nBMC Microbiol. 22: 137. doi:10.1186/s12866-022-02552-8\nManconi, I., P. van der Maas, and P. Lens. 2006. Effect of copper dosing on sulfide inhibited reduction of nitric and nitrous\noxide. Nitric oxide Biol. Chem. 15: 400–407. doi:10.1016/j.niox.2006.04.262\nMoffett, J. W., C. B. Tuit, and B. B. Ward. 2012. Chelator-induced inhibition of copper metalloenzymes in denitrifying\nbacteria. Limnol. Oceanogr. 57: 272–280. doi:10.4319/lo.2012.57.1.0272\nNaqvi, S. A. W., H. Naik, D. A. Jayakumar, M. S. Shailaja, and P. V Narvekar. 2006. Seasonal oxygen deficiency over the\nwestern continental shelf of India , p. 195–224. In N.L. Neretin [ed.], Past and Present Water Column Anoxia. Springer\nNetherlands.\nNaqvi, S. W. A., D. A. Jayakumar, P. V Narvekar, H. Naik, V. V. S. S. Sarma, W. D’Souza, S. Joseph, and M. D. George.\n2000. Increased marine production of N2O due to intensifying anoxia on the Indian continental shelf. Nature 408: 346–349.\ndoi:10.1038/35042551\nSorokin, D. Y. 2003. Oxidation of Inorganic Sulfur Compounds by Obligately Organotrophic Bacteria. Microbiology 72: 641–\n653. doi:10.1023/B:MICI.0000008363.24128.e5\nWalsh, D. A., E. Zaikova, C. G. Howes, Y. C. Song, J. J. Wright, S. G. Tringe, P. D. Tortell, and S. J. Hallam. 2009.\nMetagenome of a versatile chemolithoautotroph from expanding oceanic dead zones. Science (80-. ). 326: 578–582.\ndoi:10.1126/science.1175309\nWang, L., and Z. Shao. 2021. Aerobic Denitrification and Heterotrophic Sulfur Oxidation in the Genus Halomonas Revealed\nby Six Novel Species Characterizations and Genome-Based Analysis. Front. Microbiol. 12: 652766.\ndoi:10.3389/fmicb.2021.652766\nWong, B.-T., and D.-J. Lee. 2014. Pseudomonas yangmingensis sp. nov., an alkaliphilic denitrifying species isolated from a\nhot spring. J. Biosci. Bioeng. 117: 71–74. doi:https://doi.org/10.1016/j.jbiosc.2013.06.006", "reviewer_3": "This study identifies a previously unknown group of heterotrophic denitrifiers capable of coupling sulfur oxidation with\ndenitrification autonomously. Traditionally, heterotrophic denitrifiers have been thought to be hampered by sulfide, which\nnegatively affects their growth and increases nitrous oxide (N₂O) emissions, a potent greenhouse gas. However, in this\nstudy, these bacteria were found to use sulfide as an alternative electron source, enhancing denitrification and reducing\nnitrous oxide (N₂O) emissions. Through microcosm incubations, DNA stable-isotope probing, and metagenomics, the\nresearchers demonstrated that these versatile heterotrophs exhibit a flexible physiology allowing them to thrive in sulfide-\nrich environments and compete effectively with chemolithoautotrophic denitrifiers. This discovery shifts our understanding of\nsulfur cycling and suggests new implications for mitigating climate change.\nSpecific Comments\n1. There are a few grammatical errors and some sentences are lengthy and complex, which might hinder comprehension.\nSimplify and break down long sentences where possible.\n2. Line 62-64: suggested revision: “Incomplete denitrification is a major source of N₂O, a potent greenhouse gas that\ncontributes to climate warming. Additionally, N₂O emissions contribute to ozone layer depletion and acid rain formation.”\n3. Line 65-68: Please revise to “In many estuaries, coastal regions, and marine oxygen-deficient zones, the sulfur cycle\nplays a crucial role in regulating biogeochemical processes. This is due to the dominance of sulfate as a terminal electron\nacceptor and the subsequent release of hydrogen sulfide (H₂S) from benthic sediments.”\n4. Line 68-75: This part should include a more detailed introduction to sulfide-driven dissimilatory nitrate reduction to\nammonium (DNRA) (Canfield et. al, 2010 (Science); Wang et. al, 2021 (Water Research), etc.).\n5. Line 71-73: Please revise to “Sulfide is highly toxic to heterotrophic denitrifiers, adversely affecting microbial activity and\npotentially increasing N₂O emissions.”\n6. Line 76-80: “In this study, we investigate the response of heterotrophic denitrifiers to sulfide in the highly productive\nSonghua River Estuary, a major tributary within the largest water system in East Asia. We employed a combination of\nmicrocosm and lab culture experiments, metagenomics, and a novel community-isotope-corrected DNA stable-isotope\nprobing (DNA-SIP) approach.”\n7. In sediment microcosms, how do community compositions shift over time under different treatment conditions? How is the\nstability and resilience of these microbial communities in response to fluctuating environmental factors?\n8. How do the lab-based findings translate to natural estuarine environments? Are the conditions in the microcosm\nexperiments reflective of those in natural settings, and how might environmental variability influence the activity of F-\nSOHDs? Can modelling approaches be used to predict their contributions to nutrient cycling and greenhouse gas emissions\nunder various environmental scenarios?\n9. Given the ability of F-SOHDs to reduce N2O emissions, how can this be applied in wastewater treatment? What are the\npractical steps to harness these microbes in engineered systems to mitigate greenhouse gas? Can the activity of F-SOHDs\nbe enhanced in situ through biostimulation strategies? What types of amendments (e.g., organic carbon, sulfide) would be\nmost effective, and what are the potential ecological side effects of such interventions?\n10. While metagenomics provides a snapshot of potential metabolic capabilities, it does not confirm actual activity. How can\nthe findings from metagenomics be integrated with other omics approaches to provide a more comprehensive understanding\nof microbial functions in these environments?"}, "author_response": "Response to Reviewers’ Comments\nWe sincerely appreciate the insightful comments and suggestions from the reviewers and editor,\nwhich have greatly helped improve the quality of the manuscript (NCOMMS-24-34488A). We have\nnow carefully revised the manuscript and addressed each comment raised by the reviewers with a\npoint-by-point response. The reviewer comments are provided in italic black font, and our responses\nand the changes made within the revised manuscript are indicated in blue font color.\nResponses to Reviewer #1\nReviewer #1 (Remarks to the Author):\nThis manuscript found the important contribution of heterotrophic denitrifiers on N O reduction in\n2\nnatural ecosystem, especially in estuaries environments. Based on authors data, heterotrophic\ndenitrifiers seemed to exert a more significant role compared to chemolithoautotrophic denitrifiers,\nwhich was completely beyond folks’s traditional understanding. These findings help folks further\ncomprehend the interaction between C, N and S cycles. Before its publication, the followings should\nbe addressed:\nResponse: We sincerely thank the reviewer for the positive assessment and insightful comments on\nour work! We have carefully revised the manuscript, fully addressing all the comments/suggestions\nprovided. Please find our detailed responses below.\n1. Line 88-89, why authors sampled the Estuarine sediments at the junction of the Songhua River\nand effluent from municipal treatment? And in Fig.S1 the detailed sample site was not provided.\nResponse: We conducted sampling at the estuarine junction site, a critical hotspot for element\nexchange, due to its key role as a regulator of nutrient flow between human communities and natural\necosystems. Annually, over 300 billion cubic meters of anthropogenic wastewater, about one-\nseventh of the global river volume, are generated, with around 60% being discharged into natural\nwaters after treatment (Mateo et al. 2015; Wu et al. 2019). This contributes large amounts of nitrogen,\nphosphorus, sulfur, and organic matter to rivers, influencing global biogeochemical cycles (Shao et\nal. 2022). In this study, we focused on the junction between the Taiping Wastewater Treatment Plant,\nthe largest wastewater treatment system in Heilongjiang Province, and the Songhua River, the\nlargest river in East Asia. This site was chosen to examine the impact of anthropogenic activities on\nbiogeochemical cycling. Per the reviewer’s suggestion, we added detailed geographic coordinates\nand sampling procedures in Revised Supplementary Information Lines 42–45 and Fig. S1a.\n2. line 91, what was the quality of 20 mL of surface sediments?\nResponse: We are unsure what the reviewer means by “quality” in this comment. If it refers to\nweight, the 20 mL of surface sediments weighs 24.2 grams. If the intent was to inquire about the\nphysicochemical parameters, they were measured as follows: the collected 20 mL surface sediment\ncontained 48.35% sand, 22.57% total organic matter, 1.34% total nitrogen, 0.07% total phosphorus,\nand had a pH of 7.06. Both weight and physiochemical parameters have been integrated in the main\ntext (see Revised Manuscript Lines 122 and 544–545).\n3. In Fig.1d, the N amounts were balanced or not?\nResponse: The total nitrogen remained stable at around 300 µM. In Fig. 1d, we displayed only\nnitrate, nitrogen gas and ammonium concentrations to focus on how much nitrate has been reduced\nvia denitrification and DNRA, producing nitrogen gas and ammonium respectively. Intermediate\nproducts, including nitrite and nitrous oxide, were also measured (see Figure R1 below). Total\nnitrogen, calculated by summing different nitrogen states, ranged from 293.6 to 304.4 µM,\nindicating a balanced nitrogen amounts.\nFigure R1. Concentration of nitrite, and nitrous oxide and calculated total nitrogen.\n4. line 171-177, please provide more details on this deduction of heterotrophs may dominate sulfur\noxidation and denitrification in this ecosystem.\nResponse: As suggested, we have added further clarification to strengthen our inference. Combined\nwith the feedback from Reviewer #2, we have made substantial revisions to this section. We now\nbelieve there is strong evidence supporting the dominance of F-SOHDs over chemolithoautotrophs\nin sulfur oxidation and denitrification. The details are as follows:\n1) The F-SOHDs constitute 60.8% of the denitrifying sulfur-oxidizing community, whereas\nchemolithoautotrophs account for only 12.1% (see Revised Manuscript Lines 219–221).\n2) In the microbial community of the +C+N+S microcosm, Azoarcus is the most dominant\ngenus, accounting for up to 61.5% of the relative abundance (Fig. 2c). Its relative abundance\nincreased by 21.6% with the addition of 13C , but decreased by 10.1% with the addition of 13C.\no i\nAdditionally, this genus exhibited enhanced 13C assimilation in response to sulfide addition, as its\no\nrelative abundance increased by 64.1% in the (12C+13C )+N+S treatment compared to\ni o\n(12C+13C )+N treatment. These findings support its functional identification as an F-SOHD (see\ni o\nRevised Manuscript Lines 149–153 and 158–164).\n3) Compared to the +C+N microcosm, the relative abundance of Thiobacillus, the most\nabundant chemolithoautotroph in the community, decreased by an average of 3.8% in the +C+N+S\nmicrocosm (Fig. 2c,d). This suggests that F-SOHDs outcompete chemolithoautotrophs when sulfide\nis present (see Revised Manuscript Lines 221–224).\n4) Furthermore, the subsequent metagenomic analyses further confirm the functional\ninferences for the microbes mentioned above, specifically identifying Azoarcus and Pseudomonas\nas F-SOHDs and Thiobacillus as a chemolithoautotroph.\n5. line 222-229, author mainly focused on the contribution of denitrification processes, and they\nalso found DNRA process in the sediments. Thus, how about the DNRA process contribution on both\nNO3- reduction and S2- oxidation?\nResponse: The DNRA process plays a relatively minor role in nitrate reduction, as evidenced by\nthree factors: 1) qPCR experiments showed that nrfA gene copies accounted for only 3% of nirS\nand nirK gene copies (Fig. 1b); 2) ammonium release was just 16.63% of dinitrogen gas release\nunder the same conditions (Fig. 1d); 3) metagenomic data revealed that microbes capable of DNRA\nhave a much lower relative abundance (3.76%) compared to F-SOHDs (86.55%) (Fig. 3). Thus, F-\nHOSDs rather than DNRA microbes, are the primary contributors to nitrate reduction. Given the\nimportance of F-SOHDs and to avoid distraction, we focused directly on their role in detoxification\nin the presence of sulfide, without assessing the contribution of DNRA microbes to sulfide oxidation.\n6. line 299-303, more details information are required to support authors’s deduction.\nResponse: Following the reviewer’s suggestion, we have provided additional details to strengthen\nthe deduction (see Revised Manuscript Lines 370–383).\n7. line 351-353, please provide more similar sites, I do not believe that only 3 similar sample were\nenough for supporting authors’s opinion.\nResponse: Thank you for your insightful comment. We have provided additional sample sites,\nincluding bay and freshwater sediments (Aelion et al. 2009; Miller et al. 1986), agricultural soils\n(Tam et al. 1979), marine and soil pure cultures (Sørensen et al. 1987; Sørensen et al., 1980), and\nwastewater treatment-related activated sludges (Pan et al. 2013; Schönharting et al. 1998), to further\nsupport the discussion. Relevant references have also been included (see Revised Manuscript Lines\n478–482). This sentence has been rephrased in response to the comments provided by Reviewers\n#1 and #2.\nResponses to Reviewer #2\nReviewer #2 (Remarks to the Author):\nThe study by Shao et al., explores the presence of facultative sulfur-oxidizing heterotrophic\ndenitrifiers, abbreviated by the authors as F-SOHDs, in estuary sediments of the Songhua River\nSystem. Specifically, their study employed DNA-SIP experiments, in combination with metagenomic\nsequencing and enrichment cultures to identify F-SOHD members in a mixed microbial community\ncomprising chemoorganoheterotrophs and chemolithoautotrophs. Using this range of techniques,\nthey identified environmental genera affiliated with Thauera, Pseudomonas, and Azoacus as likely\nF-SOHDs in their samples. Lastly, they demonstrate using F-SOHD enriched bioreactors, that such\nmembers are capable at better suppressing N2O emissions than their heterotrophic denitrifying\ncounterparts. In turn, this led to the suggestion that F-SOHDs could be useful in industrial settings\nto mitigate N2O emissions.\nAs someone who has previously studied F-SOHDs, which I refer to as chemolithoheterotrophs,\nI find the work to be of high value as it expands on the potential for chemolithoheterotrophy in the\nenvironment. Moreover, the authors employed an impressive range of tools to identify these\nmicrobes in situ. Overall, I find the results fairly compelling, albeit the manuscript is not well-\nrounded at times, and does tend to overstate claims. Namely, the introduction did not provide the\nreader with state-of-the-art background information concerning chemolithoheterotrophs. The\nresults I generally find compelling, although I have some points that need further clarification, and\nthe authors should discuss some of the discrepancies in their data. While I don’t disagree with all\nthe points raised in the discussion, it does need some more work/reframing, to present a more well-\nrounded case for chemolithoheterotrophy in the environment. In addition, some errors in their\nestimates of Gibbs free energy need to be addressed in the discussion. The method section was at\ntimes thorough, but other parts were fairly vague, to the point that the reader would not be able to\nreproduce the experiment without additional clarification. Ending on an optimistic note, the figures\nare nicely polished, and the data/conclusions (while overstated at times) appear generally sound.\nResponse: We sincerely appreciate the reviewer’s positive and encouraging feedback on our work.\nAll the comments are highly constructive and thorough, which helped us substantially improve the\nquality of the manuscript. Following suggestions, we have revised the sentence description\nthroughout the manuscript to avoid overstatements and have rewritten the Introduction to include\nmore research background on chemolithoheterotrophs. Regarding the Gibbs free energy estimates,\nwe have recalculated these values and corrected the relevant sentences/figures in the Revised\nManuscript. To clarify, we have added more details to the Methods section as recommended, to\nprovide a clearer explanation for readers. We have carefully revised the manuscript, fully addressing\nall the comments/suggestions provided. Please find our detailed responses below.\nComments\n1. Lines 40-41: I find this statement to be a little vague, namely because heterotrophic denitrifiers\noperate in environments where sulfide is present. But what the authors could specify is that sulfide\ninhibits a key enzymatic reaction that catalyzes the reduction of N O to N . Hence, heterotrophic\n2 2\ndenitrifers are sensitive unless sulfide is removed, either by cooperation with chemolithoautotrophs\nor with chemolithoheterotrophs. This type of denitrifying community structure is indeed what we\nfind in organic and sulfide-rich ecosystems.\nResponse: We fully agree with the reviewer’s comment that heterotrophic denitrifiers are active but\ncarry out incomplete denitrification in the presence of sulfide, owing to the inhibition of NosZ. To\nclarify this point, we have revised the sentence as follows: “However, their inability to oxidize\nsulfide renders them vulnerable to this toxic molecule, which inhibits the key enzymatic reaction\nresponsible for reducing nitrous oxide (N O) to dinitrogen gas, thereby raising greenhouse gas N O\n2 2\nemissions.” (also see Revised Manuscript Lines 40–43).\n2. Line 43: This statement comes across as a little too strong as their role in the environment,\nincluding aquatic ecosystems, has been noticed by some and even empirically demonstrated\n(Sorokin 2003; Boltyanskaya et al. 2007; Callbeck et al. 2019; Wang and Shao 2021). As I\nmentioned above, the value of the study is not in that it discovered sulfide oxidation by heterotrophs\nin the environment, it is that it demonstrates that numerous microbes in estuary sediments could\nthrive via a chemolithoheterotrophic metabolism greatly expanding its potential. I would therefore\nconsider re-framing this part.\nResponse: As suggested, we have revised the sentence to emphasize the detection of\nchemolithoheterotrophs and their ability to thrive, without referring to them as “previously\nunnoticed” (see Revised Manuscript Lines 43–48).\n3. Lines 48-51: I agree with this statement, but this largely repeats the findings of Callbeck et al.,\n2019, which I detail a little later in this review. Moreover, their justification of this statement based\non their Gibbs free energy calculations appears to be flawed (see later discussion).\nResponse: We agree that a similar phenomenon has been observed in other studies. As the reviewer\npointed out, while this statement aligns with previous findings and may appear repetitive, we have\nkept it in the abstract and revised it to accurately represent the results of our study (see Revised\nManuscript Lines 52–55). Additionally, we cited the paper the reviewer mentioned in the discussion\nto support our findings. Regarding the concerns about Gibbs free energy calculations, we have\nrecalculated these values and revised the statement accordingly in Revised Manuscript Lines 514–\n519 and Revised Figure 6 (please see Comments 42 and 69).\n4. Lines 51-53: I’m in favor of this work generally, but this is an overstatement. Firstly,\nenvironmental examples of sulfide oxidation by heterotrophs exist, thus the use of “paradigm shift”\nis over the top. Secondly, I understand that if you work on N2O you can make a link to climate\nchange, however, to say that his work has “profound implications” for climate change is a stretch\ntoo far. For example, it is not the case that the authors have tested/modeled their reactors under\nvarious climate change scenarios. In my view, the work is a good piece because it advances the idea\nthat chemolithoheterotrophs form a persistent part of the microbial community in an organic matter-\nrich estuary.\nResponse: We appreciate the reviewer’s thoughtful comments and are pleased to hear that the\nreviewer considers our work as advancing the current understanding of chemolithoheterotrophs. As\nsuggested, we have rephrased the sentence to avoid overstatement (see Revised Manuscript Lines\n57–60).\n5. Lines 62-64: For clarity, denitrification is one of two major processes, nitrification being the\nother. The statement makes it seem as though denitrification is the only one.\nResponse: Revised as suggested (see Revised Manuscript Lines 69–71). The sentence has been\nrephrased in response to the comments provided by Reviewers #2 and #3.\n6. Lines 65-68: Fully agree, for a comprehensive review of sulfur cycling and its influence over the\nnitrogen cycle in oxygen minimum zones please see (Callbeck et al. 2021).\nResponse: This is a great citation. We have carefully studied this paper and have incorporated\nrelevant key insights into the Introduction and Discussion sections of the Revised Manuscript.\nPlease see the detailed changes in Lines 76–79, 86–91, 447–451, and 466–473.\n7. Lines 71-73: The authors only superficially mention that it impacts N2O but how is this the case?\nSome have speculated that sulfide also inhibits the last step of denitrification, as sulfide acts as a\nCu-chelator.\nResponse: Per the reviewer’s suggestion, we have added details regarding the specific inhibitory\nmechanism of sulfide in the Revised Manuscript Lines 76–79 and 466–471.\n8. Lines 68-70: Certainly, chemolithoautotrophs are well-recognized, but work by Callbeck et al.,\n2019 has shown that both chemolithoautotrophs, and chemolithoheterotroph (aka. F-SOHD) can\nco-exist to oxidize sulfide during a sulfidic event off the coast of Peru.\nResponse: We have provided additional sentences to review previous findings regarding\nchemolithoheterotrophic denitrifier Arcobacter peruensis in coastal waters off Peru (see Revised\nManuscript Lines 86–91).\n9. Line 72: I understand the point the authors are making, however, for clarification sulfide is toxic\nto all denitrifiers, including nitrate-reducing chemolithoautotrophs, chemolithoheterotrophs and\norganoheterotrophs. The reason for this I explain a little later in the discussion section below.\nResponse: Similar to comment 6, we have revised the relevant sections to specify sulfide toxicity\ntoward Cu-dependent enzymes, rather than limiting the description to conventional heterotrophic\ndenitrifiers (see Revised Manuscript Lines 76–79 and 466–469).\n10. Lines 73-75: This statement, which is effectively the premise of the manuscript, is simply not up\nto date with the state-of-the-art. As I mentioned above, it’s not as though we know nothing about\nthis topic of F-SOHD/chemolithoheterotrophs. The authors have partly relayed this information in\nthe discussion (on lines 368-370), but choose not to interject this information here or above when\nbackground information is needed on this specialized metabolism. Again, the work is valuable not\nbecause it is the first to discover F-SOHD, but because it expands the potential for this metabolism\nto operate in the aquatic realm. The authors should consider reworking this paragraph to include\nthe state-of-the-art, and perhaps frame the premise in a slightly different way.\nResponse: We greatly appreciate the reviewer’s constructive comment. Following your suggestions,\nwe have thoroughly rewritten this paragraph to incorporate more relevant research background on\nchemolithoheterotrophs and sulfide toxicity toward Cu-dependent enzymes. Additionally, we have\nclarified the knowledge gap and refined the research objective (see the second paragraph of\nIntroduction section in the Revised Manuscript).\n11. Lines 80-83: To be specific, the study identifies new F-SOHDs at a sediment site. Somewhat\nmisleading is the use of “ubiquitous” as this implies that the authors provided evidence of F-SOHDs\nwidespread distribution across multiple sampling sites, although their study site is positioned at one\nlocation (45°49′12.81″N, 126°43′22.51″E). Again, the authors have a habit of overstating claims,\nwhen in fact, there is no need.\nResponse: As suggested, we have deleted the word “ubiquitous” from the manuscript to avoid\noverstating (see Revised Manuscript Line 107).\n12. Lines 82-85: I find this overly broad and introduces a bit of speculation, especially on the climate\nchange front; can the authors be a bit more specific?\nResponse: We have removed the speculated description of climate change and rewritten these two\nsentences to make it more specific. The revised sentences are pasted below for convenience (also\nsee Revised Manuscript Lines 108–114).\n“This capability allows them to detoxify sulfide and achieve complete denitrification, thereby\nmitigating the N O emission. This work enhances our understanding of heterotrophic denitrifiers\n2\ncapable of sulfide detoxification in aquatic environments, with broader implications for regulating\ncarbon, nitrogen, and sulfur cycles, including the mitigation of N O emissions in both natural and\n2\nengineered system.” (Lines 108–114)\n13. Lines 98-101: If I have this right, this would amount to roughly two-thirds of the original 300\nM of nitrate being recovered as N2 and NH4+. But roughly one-third is not recovered, hence, both\ncomplete and incomplete denitrification are evident in this experiment.\nResponse: We agree with the reviewer’s observation that both complete and incomplete\ndenitrification are occurring simultaneously. As shown in Figure R1 below, 56.1% of the consumed\nnitrate was converted to N O, rather than being fully reduced to N . We have revised this sentence\n2 2\nto reflect this clarification (see Revised Manuscript Lines 129–132).\nFigure R1. Supplementary data of total nitrogen amounts, nitrite, and nitrous oxide for Fig.1d.\n14. Line 108: The with and without sulfide addition experiments are somewhat confusing. I\nunderstand that the authors added additional sulfide, but it should be perhaps reiterated that sulfide\nis going to be produced in the background, unless the authors removed the sulfate for sulfate\nreducers, in incubations like C+N, or Ci, or Co etc…\nResponse: We appreciate the reviewer’s concerns. In these experiments, we did not specifically\nexclude sulfate or sulfate reducers. Our experiment was designed to study the effect of excess sulfide\non denitrification without excess manipulation of the community. While we acknowledge that\nsulfide produced by background sulfate reduction could potentially contribute to nitrate reduction\nthrough the re-oxidation of sulfide, the addition of excess sulfide led to a significant increase in the\nrate of nitrate reduction compared to the group without added sulfide (Figure 1c), suggesting that\nthis supplemental sulfide may serve as an electron donor to enhance nitrate reduction. Thus, our\nresults provide evidence of a sulfide-supported nitrate reduction process within the sediment\ncommunity. To clarify the experimental design, we have revised the label in Revised Figure 1c from\n“sulfide free” to “no sulfide added”.\n15. Lines 117-121: The increases of 1.3% and 3.8% don’t seem to be substantial for Thauera and\nPseudomonas. Thus, can the authors assume that their heterotrophs according to these values? And\nin the second part, how much did these members decrease when amended with 13Ci? In light of\nchemolithoautotrophic abundances increasing with the addition of 13C, could this not artificially\ndilute such heterotrophs like Thauera and Pseudomonas, if it’s based on relative abundances?\nResponse: We understand that the reviewer has three concerns: 1) Whether a heterotrophic lifestyle\ncan be inferred from relatively small changes in relative abundance in the (13C +12C)+N+S and\no i\n(12C +12C)+N+S treatments; 2) The detailed changes in the relative abundances of these three\no i\ngenera when comparing the (12C +13C)+N+S and (12C +12C)+N+S treatments; 3) The reliability of\no i o i\ninferring lifestyle solely from relative abundance data.\nResponse to point 1: The modest increases observed in fast-growing heterotrophs like Thauera\n(1.3%) and Pseudomonas (3.8%) are indeed a common outcome in DNA-SIP analysis, particularly\nwithin complex microbial communities, especially given relatively slow growth under anaerobic\nconditions and short-term isotope labeling. Similar small increases in abundance have been widely\nreported in other studies (e.g., Sathyamoorthy et al. 2018; Sun et al. 2021). In our study, the\nrelatively small increases in Thauera and Pseudomonas can partially be attributed to their low\nbackground abundances (2.0% and 5.1%). These two genera exhibited 63% and 74% additional\nincreases in relative abundance in the heavy DNA fraction, compared to the 35% increase observed\nin Azoarcus. This suggests that, despite their low initial abundance, their growth may be higher than\nAzoarcus. The dominance of Thauera and Pseudomonas in the 612-day long-term enrichment\nexperiment further supports their active growth.\nResponse to point 2: Compared to the heavy fraction without 13C labelling in the +C+N+S\nmicrocosm, the relative abundances of Thauera, Azoarcus, and Pseudomonas decreased by 0.1%,\n10.1%, and 1.5% when amended with 13C. These values have been added for clarity (see Revised\ni\nManuscript Lines 151–152).\nResponse to point 3: We acknowledge the limitations of inferring microbial lifestyles solely\nfrom relative abundance data, as changes in chemolithoautotrophic populations could dilute the\napparent relative abundance of heterotrophs like Thauera and Pseudomonas. This limitation is\ninherent to DNA-SIP when paired with 16S rRNA amplicon sequencing. DNA-SIP alone provides\npreliminary insights into active microorganisms but lacks the resolution for definitive lifestyle\ncharacterization. To enhance accuracy, integrating DNA-SIP with qPCR, metagenomics, or\nmetatranscriptomics could provide advantages. We have moderated the description regarding\nlifestyle inference and emphasized the methodological limitations in the Revised Manuscript Lines\n153 and 404–407.\n16. Lines 121-122: Can the authors comment on why Thauera comprised a large fraction of the\nmicrobial community in the experiment void of sulfide (C+N) when compared to the C+N+S\nexperiments? It would appear that this member is sensitive to sulfide inhibition. In addition, an\nincrease of 0.7% is practically negligible. Moreover, it appears that Pseudomonas shows a different\npattern that is opposite to your statement. Also in Fig. 2d, Pseudomonas and Azoarcus both have\n13Ci which is higher than the control in grey. Hence, this goes against the idea that no 13C-DIC is\nincorporated. Part of the problem the authors might encounter in their analysis is that heterotrophs\ncan also assimilate 13C-DIC (e.g. Erb 2011), which might complicate their analysis, or at least\nrequire a note.\nResponse: Thank you for your suggestion and relevant literature. The concern regarding the\nincreased abundance of Thauera in C+N compared to C+N+S has been fully addressed in Comment\n25 (please refer to that section for details). Regarding the concern of 13C-DIC assimilation,\nwe\ni\nacknowledge that some heterotrophic microorganisms might incorporate some DIC, as noted for\nAzoarcus and Pseudomonas, which partially incorporated 13C isotope in the (13C+12C )+N\ni i o\ntreatment compared to (12C+12C )+N treatment (Fig. 2d). This observation aligns with the\ni o\nreviewer’s comment that some heterotrophs are capable of assimilating 13C-DIC, as described in the\nreference provided (Erb, 2011). To address this potential complication, we have noted this issue\nexplicitly in the main text and cited the relevant reference to provide readers with a balanced and\ncomprehensive understanding of the data interpretation (see Revised Manuscript Lines 407–409).\n17. Lines 125-129: Thiobacillus increased in Fig. 2d (C+N) when sulfide was not present, which\nruns counter to their chemolithoautotrophic lifestyle. And also what about other\nchemolithoautotrophic members such as Chromatiales (M5.bin31) and Sulfurivermis (M5.bin64)\nshown in Fig. 3?\nResponse: It is true that Thiobacillus was slightly less abundant and apparently slightly less active\nwhen sulfide was added in the presence of organic matter (comparing Fig. 2c and Fig. 2d). We\nreason that in the presence of OM, Thiobacillus may be outcompeted by chemolithoheterotrophs,\nbecause heterotrophy is energetically more favorable than autotrophy. But our results in Fig. 2f and\nFig. 2g in comparison with each other and with Fig. 2c and Fig. 2d show a strong stimulation of\nThiobacillus when sulfide is added in the absence of added OM. This supports our inference that\nThiobacillus is primarily chemolithoautotrophic in our experiments, a conclusion further\ncorroborated by metagenomic data.\nDue to limited space, we decided to not show low-abundance taxa in Figure 2, such as\nChromatiales (M5.bin31) and Sulfurivermis (M5.bin64), which are both below 0.08%. This decision\nalso allowed us to focus the related text on the most abundant microbes. We included them in Figure\n3 because that’s a more comprehensive figure. We think this decision strikes the right balance of\nkeeping the focus on the most abundant taxa while still showing data for these other\nchemolithotrophs.\n18. Lines 130-154: It seems that this method distinguishes Thiobacillus, but what about other\nchemolithoautotrophic members such as Chromatiales (M5.bin31) and Sulfurivermis (M5.bin64)\nshown in Fig. 3, where do they position in Fig. 2i? In addition, organisms like Dechloromonas,\nwhich according to Fig. 3 is an F-SOHD, are positioned in the bottom left of Fig. 2i alongside\nThiobacillus the chemolithoautotroph. I also have difficulties in reconciling the very negligible\nincreases the authors reported on lines 118 and 121 and how this results in very clear patterns for\nThauera and Pseudomonas in Fig. 2i. I think the authors should add that this method might not be\nperfect at disentangling all F-SOHDs from other community members, but it may work for more\nactive microbes.\nResponse: We agree with the reviewer’s comment and acknowledge the limitations of this approach.\nWhile it effectively highlights the more abundant and active microbes within the community, it may\nnot fully capture the functional roles of all microbial members, particularly those of low abundance.\nIn response to these considerations, we have removed the Fig. 2h and 2i, along with the associated\ndescriptions, and opted for a more straightforward analysis of abundance changes for functional\nidentification in the main text (see Revised Manuscript Lines 158–200). Moreover, we have\nsignificantly shortened the overstated description of DNA-SIP (see Revised Manuscript Lines 388–\n404) and explicitly addressed its limitations to provide readers with a clearer understanding (see\nRevised Manuscript Lines 404–407). For the concerns raised in Lines 150 and 154, these have been\naddressed in our response to Comment 15 above.\n19. Lines 138-142: The exception would be Arcobacter peruensis, which reduces nitrate, oxidizes\nsulfide, and simultaneously assimilates/oxidizes organic matter into biomass for energy/growth\n(Callbeck et al. 2019). Arcobacter peruensis was also abundant in sulfidic and organic matter-rich\nshelf waters alongside chemolithoautotrophs like SUP05.\nResponse: As stated, Arcobacter peruensis is indeed an very important bacterium with metabolic\npotential similar to F-SOHDs identified in this study. We acknowledge that in certain environments,\nsuch as organic carbon-rich waters, Arcobacter peruensis can play a significant role alongside\nchemolithoautotrophs. However, in our study, we aimed to emphasize the most general case rather\nthan focus on specific exceptions. As the reviewer noted, we have added more detailed description\nof the role of Arcobacter peruensis in both the Introduction and Discussion sections (see Revised\nManuscript Lines 89–91 and 423–426).\n20. Line 142-144: In the methods section, on lines 488-490, the authors state that “microbes have\nthe compacity to oxidize sulfide if they increase in relative abundances in +C+N+S compared to\n+C+N microcosms”, but the issue with Thauera is that its abundance decreases, and hence, by their\nlogic above is not a sulfide oxidizer. The authors could further clarify this.\nResponse: We apologize for the confusion caused by the improper sentence structure and the\ninaccurate inference regarding the sulfide sensitivity of Thauera in the manuscript. The original\nsentence “This appears to show that heterotrophs might also contribute to sulfur oxidation and\ndenitrification in our system” was intended to emphasize the genera Azoarcus and Pseudomonas,\nrather than Thauera. Based on the abundance changes observed, here we could not infer that\nThauera act as a sulfide oxidizer in this context; this will be discussed in detail in Comment 25\nbelow. Following the reviewer’s comment, we have rephrased the relevant sentences to improve\nclarity (see Revised manuscript Lines 158–165).\n21. Line 171: The authors coin the new acronym facultative sulfur-oxidizing heterotrophic\ndenitrifiers (F-SOHDs) for a metabolic process that has been described previously as\nchemolithoheterotrophy (Callbeck et al. 2019). For consistency’s sake with previous work, why not\nterm F-SOHDs as chemolithoheterotrophs which include microbes that oxidize sulfide, and reduce\nnitrate while simultaneously utilizing organic carbon as an energy/carbon source for growth? This\nwould reduce the number of acronyms in the literature, and make for a more comparable term to\nchemolithoautotrophy, which the authors readily apply in their manuscript.\nResponse: We appreciate the reviewer’s suggestion. However, we believe that the term\n“chemolithoheterotrophy” is somewhat broad and may not accurately convey the specific\ncharacteristics of the microorganisms we are studying. While chemolithoheterotrophs are defined\nas microbes that derive energy from oxidizing inorganic compounds while using organic matter as\na carbon source, this category includes a wide variety of microbes that utilize diverse inorganic\nelectron donors, such as hydrogen, manganese, or iron (Miroshnichenko et al. 2003; Kompantseva\net al. 2017; Price et al. 2018), and is not limited to sulfur. Moreover, the term\nchemolithoheterotrophy does not necessarily imply a connection to denitrification or the fact that\nthese microorganisms can also grow without added reduced sulfur (this “facultative”). Our findings\nindicate that microbes detected in our study represent a subset of heterotrophic denitrifiers with a\nunique ability to oxidize sulfur, distinct from microbes that utilize other organic matter. More\nimportantly, they catalyze complete denitrification and mitigate N₂O emissions in the presence of\nsulfide. To capture these specific features, we propose the term “facultative sulfur-oxidizing\nheterotrophic denitrifiers” (F-SOHDs), providing a clearer and more precise description for readers.\n22. Lines 204-207: The authors mention Arcobacter, but it was later been shown to grow as a\nchemolithoheterotroph (Callbeck et al., 2019). Hence, Arcobacter peruensis had the capacity to\nassimilate organic carbon, reduce nitrate, and oxidize sulfide, much like what the authors describe\nin their paper for Thauera and other members.\nResponse: Following the suggestion, we have removed “Arcobacter” here to ensure the accuracy\nof the results (see Revised manuscript Lines 256).\n23. Lines 208-213: I checked the literature on some of the closest affiliated microbes in Figure 3\n(see below), and some of them have capacities for CO2 fixation, despite being labeled as having no\ncapacity in Fig. 3c.\n- Azoarcus (see https://www.kegg.jp/pathway/aza00710+AZKH_p0225)\n- Ralstonia pickettii DTP0602 (https://www.kegg.jp/pathway/rpj00710+M00166)\n- Pseudomonas yangmingensis (Wong and Lee 2014)\nThus, while I believe that some are probably chemolithoheterotrophs, the authors should stipulate\nsome caveats and unknowns here.\nResponse: We acknowledge that some of our inferences might be biased due to the incomplete\nnature of the reconstructed genomes, although the majority are of high quality. Regarding the\npossibility that some of these organisms may be chemolithoautotrophs, it is important to note that\nsignificant metabolic differences can exist even among closely related microbes. For instance,\nAzoarcus sp. CIB and Azoarcus sp. BH72 have been reported as a heterotroph (Durante et al. 2019;\nHurek et al. 1996). To address this uncertainty and potential biases, we have added a sentence\nclarifying these limitations in our findings (see Revised manuscript Lines 266–269).\n24. Lines 222-225: As mentioned above the authors should apply some caution here. Moreover, the\nauthors have not measured the nitrate turnover rates of the specific members, thus to say that this\ncluster is more biogeochemical relevant is rather speculative. It’s fair to say that strong potential\nfor chemolithoheterotrophy exists at their study site.\nResponse: As suggested, we revised the sentence to reflect the presence of F-SOHDs in present\nstudy (see Revised Manuscript Lines 279–282).\n25. Lines 244-249: I find it difficult to take the position that Thauera is sensitive to sulfide. If the\nauthors take this position then they have to reconcile in their discussion that both nitrate-reducing\nchemoorganoheterotrophs and chemolithoheterotrophs are sensitive to sulfide. Presently, they hold\nonly the former to be true. However, I would suggest that the authors explain this differently. For\nexample, sulfide-oxidizers like Arcobacter, SUP05 and Sulfurorvum can co-exist in the presence of\nsulfide however Arcobacter and Sulfurovum tend to dominate over SUP05 when sulfide\nconcentrations exceed 10 uM (Callbeck et al., 2021). So could it not be the case that Pseudomonas\noutcompetes Thauera at a certain sulfide threshold, thereby lowering its contribution to the overall\ncommunity (based on relative abundances). Taking this position means that Thauera is not\nnecessarily sensitive but rather outcompeted under certain environmental conditions by others.\nResponse: We sincerely appreciate the reviewer’s insightful comments. From a microbial\necophysiology perspective, the reviewer’s theory on competition among different microorganisms\nunder high-sulfide conditions provides a more reasonable explanation for reconciling the observed\ndecrease in abundance with the detected sulfur oxidation capacity of Thauera. Given the insufficient\nevidence, we cannot definitively conclude that Thauera is a sulfide-sensitive microorganism. After\nthorough discussion, we have fully incorporated the reviewer’s suggestion and revised the sentences\nto present the results in a more reasonable manner (see Revised Manuscript Lines 302–313).\n26. Lines 266-267: This is a bit confusing here as it is not clear why being a heterotrophic denitrifier\nleads to N2O production, and also why chemolithoautotrophs do not. In my mind it’s not so clear\ncut, for example, SUP05 identified in Peru can perform complete denitrification to N2 (Callbeck et\nal. 2018), while other SUP05 species in the Saanich Inlet can only do nitrate reduction to N2O\n(Walsh et al. 2009). Might also be worth mentioning here that Arcobacter peruensis is an exception,\nplus others exist as mentioned earlier.\nResponse: According to the Comments 33 and 34 raised by the reviewer, we agree that sulfide not\nonly poisons heterotrophs but also impacts chemolithoautotrophs by interacting with Cu-dependent\nenzymes, such as NosZ. However, we interpret that sulfide-oxidizing chemolithotrophs may be less\nsensitive to sulfide toxicity due to their ability to oxidize sulfide into less toxic compounds. To avoid\nconfusion, we have deleted this sentence (see Revised Manuscript Lines 333–334) and added more\ndetails (see Comments 33 and 34 below) regarding the sulfide sensitivity of complete denitrification\nin conventional heterotrophic denitrifiers (see Revised Manuscript Lines 466–476). Also, we have\nexpanded the revised Discussion section to elaborate on the relationships between F-SOHDs,\nheterotrophic denitrifiers and broader chemolithoautotrophic denitrifiers, including Arcobacter\nperuensis.\n27. Lines 285-287: This line contradicts the author’s statement above that heterotrophic members\nlike Thauera could be sensitive to sulfide.\nResponse: As mentioned above, several Cu-dependent enzymes such as NosZ, encoded by\nheterotrophic denitrifiers, are sensitive to sulfide. Therefore, the previous description was inaccurate,\nand we have revised it accordingly. The revised sentence reflects that the sulfide oxidation capacity\nof F-SOHDs leads to the consumption of sulfide, thereby mitigating or preventing its toxic effects\n(see Revised Manuscript Lines 352–356).\n28. Lines 305-321: I agree with the authors that DNA-SIP technology is important to unraveling\nmicrobial ecology-related questions. But I find a dedicated paragraph to this in the opening\ndiscussion is a little excessive, as the method is in principle the same with the difference being that\nthe authors did an additional correlative analysis (Fig. 2i, g). In addition, organisms like\nDechloromonas, which according to Fig. 3 is an F-SOHD, are positioned in the bottom left of Fig.\n2i alongside Thiobacillus the chemolithoautotroph. Hence, this method is not perfect, and the\nauthors (if they decide to keep this piece) will need to address the caveats. As a recommendation,\nI would instead suggest the authors shorten or drop to make way for a more detailed discussion of\nthe specific chemolithoheterotroph members they identified in their study, which was lacking in my\nopinion (see below).\nResponse: Following the reviewer’s suggestion, we have substantially shortened this paragraph and\nhighlighted the method’s limitations (see Revised Manuscript Lines 387–404). Additionally, to\nsharpen the study’s focus, we have concentrated the Discussion section on F-SOHDs identified in\nour study rather than on the DNA-SIP methodology. We have also expanded on the ecophysiological\nsignificance and distribution of F-SOHDs in the main text (see Revised Manuscript Lines 417–441).\n29. Line 323: I would add, “Epsilonproteobacteria and the Gammaproteobacteria SUP05 clade”\nResponse: Done as suggested (see Revised Manuscript Lines 411).\n30. Line 331: There are more pertinent studies to reference here that discuss sulfidic events in marine\nenvironments, which is summarized in the review by Callbeck et al., 2021.\nResponse: Following the reviewer’s valuable suggestion and relevant reference, we have revised\nthe citations with the work published by Callbeck et al. 2021, and Caffrey et al. 2018. Additionally,\nthe more pertinent studies of marine sulfidic events and their impacts have also been incorporated\nin Revised Manuscript Lines 443–451.\n31. Lines 332-333: I believe this to be an overstatement by the authors “…uncovered here have\nevolved multiple sulfur oxidation pathways to cope with sulfide stress.” The authors have not\nempirically demonstrated sulfide stress on the community and how it leads to the evolution of\nmultiple sulfur oxidation pathways. Rather this statement could be framed as a hypothesis.\nResponse: Done as suggested. The revised sentence is pasted below for convenience (also see\nRevised Manuscript Lines 451–453).\n“Remarkably, the F-SOHDs may have developed multiple sulfur oxidation pathways to cope with\nsulfide stress, a hypothesis that merits further exploration.” (Lines 451–453)\n32. Lines 322-340: I see the importance of a wider description of sulfidic events, but this really\nappears as a zoomed-out introduction piece especially since the nosZ gene discussion below brings\nmore detail to this idea. Instead, I think the opening of the discussion would be better served\nreiterating what defines a F-SOHDs/chemolithoheterotroph. Namely, chemolithoheterotrophs\ncouple sulfide oxidation to nitrate reduction while simultaneously utilizing organic carbon as an\nenergy/carbon source (hence the assimilation of 13C into biomass in your DNA-SIP experiments).\nNext, I would reiterate the key putative chemolithoheterotrophs mentioned in the results section and\nFig. 3. Subsequently, the authors need to spend more time describing what is already known about\nthese species in the literature, so that the reader can assess what new information is added in this\nstudy. For example, my cursory inspection of some of the literature finds that some of these members\n(identified in Fig. 3) are known sulfide oxidizers like Pseudomonas stutzeri (Li et al. 2022). Thauera\nis well-studied and has been isolated from sulfide-rich environments and bioreactors before,\nimplicating it already with a role as a putative sulfide oxidizer. Others are capable of fixing CO2\nautotrophically such as Azoarcus (see https://www.kegg.jp/pathway/aza00710+AZKH_p0225),\nRalstonia pickettii DTP0602 (https://www.kegg.jp/pathway/rpj00710+M00166) and Pseudomonas\nyangmingensis (Wong and Lee 2014). I recommend the authors incorporate a much more thorough\ndiscussion of the key members they identified.\nResponse: We sincerely appreciate the reviewer for the insightful comments, which are greatly\nimportant in improving the logical flow and clarity of the discussion in our manuscript. Following\nyour suggestion, we have revised the beginning of the discussion section to better define the concept\nof F-SOHDs. Additionally, we have incorporated more comprehensive comparisons between the F-\nSOHDs identified in our study and previously characterized species reported in the literature. With\nthese updates, we aim to emphasize the significance of our findings as suggested, particularly the\ndiscovery that F-SOHDs represent a persistent part of the microbial community in organic matter-\nrich estuaries, with important implications for the sulfur cycle and N O reduction. Please refer to\n2\nthe detailed revisions in the Revised Manuscript Lines 415–441.\n33. Lines 345-346: I agree that sulfide has an influence over N2O production, but the statement\nhere that sulfide only affects heterotrophic denitrifiers and not denitrifiers more broadly –\nchemolithoautotrophs included – is not evidenced in the literature. Much of this discussion has taken\nplace in the review by Callbeck et al., 2021, but to reiterate their points sulfide is a chelator of\ncopper, and the nosZ gene is a copper-rich metalloenzyme. The sequestration of copper by its abiotic\nreaction with sulfide will induce copper limitations that result in nitrate reduction terminating at\nN2O in microbial cultures (Granger and Ward 2003; Manconi et al. 2006; Moffett et al. 2012;\nFelgate et al. 2012). The accumulation of N2O has therefore been linked to copper limitations\ninduced by sulfide, which would affect nitrate-reducing bacteria (with a nosZ) irrespective of\nwhether they are chemolithoautotrophs, chemolithoheterotrophs or heterotrophs.\nResponse: After carefully reviewing the relevant literature, we fully agree with the reviewer’s\nsuggestion. We have revised our discussion to more accurately address the broader impacts of\nsulfide on denitrifiers and its abiotic copper chelation mechanism (see Revised Manuscript Lines\n466–473).\n34. Lines 346-349: I agree with this statement, but as above this is not exclusively a heterotrophic\ndenitrifier issue but will rather affect denitrifiers broadly.\nResponse: Revised as suggested. Please see the changes in Revised Manuscript Lines 466–468.\n35. Lines 349-353: Indeed, sulfide-driven denitrification is likely responsible for major N2O\nhotspots (Naqvi et al. 2000, 2006; Arevalo-Martinez et al. 2015; Kock et al. 2016; Bourbonnais et\nal. 2017; Arévalo-Martínez et al. 2019), which likely occur as a result of the mechanism outlined\nabove. However, the difficulty I have with this line is the authors claim that these N2O/H2S-rich\nsystems are dominated by heterotrophic denitrifiers, when in fact, the cited studies do not make that\nclaim (mainly because they also pre-date large community metagenomic surveys). Secondly, in light\nof more recent analyses of these same environments, such as Peru, where we have major N2O\nhotspots along with the presence of sulfide, the microbial community is comprised of a consortium\nof nitrate-reducing chemolithoautotrophs, chemoorganoheterotrophs and chemolithoheterotrophs.\nThirdly, from a logical perspective, if such systems have high organic matter, sulfide, and nitrate,\nwouldn’t they offer the perfect ecological niche for chemolithoheterotrophs to thrive? To that point,\nCallbeck et al. 2019, and this study enriched for chemolithoheterotrophs from exactly these\nenvironments.\nResponse: As the reviewer pointed out, the cited studies do not explicitly indicate that these\nmicrobial communities are dominated by heterotrophic denitrifiers. Instead, such environments\nlikely host a diverse consortium of microorganisms, including chemolithoautotrophs,\nchemoorganoheterotrophs, chemolithoheterotrophs, and others. To prevent potential\nmisinterpretation, we have removed the inaccurate speculative statement and clarified the text\naccordingly (see Revised Manuscript Lines 476–482).\nWe fully agree with the reviewer’s insight regarding ecological niche favoring F-SOHDs (or\nchemolithoheterotrophs) in environments rich in OM, sulfide, and nitrate. These conditions indeed\ncreate an ideal habitat for F-SOHDs to thrive, as exemplified by the Arcobacter peruensis identified\nin Peruvian coastal waters and the microorganisms observed in our estuarine sediments. This\nhighlights an intriguing research avenue deserving of further exploration. Future studies employing\nlarge-scale metagenomic sequencing could help determine the persistence and abundance of F-\nSOHDs within microbial communities in these typical ecosystems.\n36. Lines 353-356: I generally agree with this line, but perhaps clarify that this occurs because\nchemolithoheterotrophs can remove sulfide and therefore remove its ability to chelate copper,\nenabling N2O reduction to N2 to continue (via the Cu-rich nosZ).\nResponse: Done as suggested. The revised sentence is pasted below for convenience (also see\nRevised Manuscript Lines 485–486).\n“By oxidizing sulfide, F-SOHDs prevent it from chelating the copper within Cu-rich NosZ, thereby\nenabling N O reduction to continue.” (Lines 485–486)\n2\n37. Lines 357-359: I agree, but it would make sense to mention that sulfide provides an additional\nelectron donor (on top of organic matter) to reduce available nitrate to N2 rather than to N2O.\nResponse: Thank you for your kind suggestion. We agree and we have revised the sentence\naccordingly. We pasted as below for convenience (also see Revised Manuscript Lines 486–489).\n“Sulfide oxidation could not only rescue the sulfide-sensitive NosZ [Cu:S] active sites but may\nsimultaneously provide an additional electron donor beyond organic matter to reduce available\nnitrate to dinitrogen gas rather than to N O.” (Line 486–489)\n2\n38. Lines 356-363: I agree with this statement, but it should be pointed out that\nchemolithoautotrophs have the same advantage (as your work also demonstrates in Fig. 5g). In\nother words, this feature is not unique to chemolithoheterotrophs.\nResponse: Done as suggested (see Revised Manuscript Lines 491–493).\n39. Lines 363-365: As I stipulated above, this is complicated and not so clear-cut. The environment\nwhere Arcobacter peruensis was identified, was from a sulfidic event off the coast of Peru (Callbeck\net al., 2019). Here, the environment was defined as being sulfide-, N2O-, and organic matter-rich.\nHence, the conditions where we find chemolithoheterotrophs are also environments where relatively\nhigh N2O concentrations persist. To better frame this one could say that\nchemolithoheterotrophs/chemolithoautotrophs might contribute to suppressing some N2O in\nsulfidic environments that are associated with high N2O production rates.\nResponse: We agree that the relationship between F-SOHD activity and N O emissions is complex\n2\nand likely depends on specific environmental conditions, such as sulfidic events and N₂O-rich\nenvironments. In cases like the sulfidic event off the coast of Peru (Callbeck et al., 2019), where\nArcobacter peruensis was identified, high levels of N O coexist with sulfide and organic matter,\n2\nproviding conditions conducive to both chemolithoheterotrophic and chemolithoautotrophic activity.\nGiven these dynamics, we have rephrased our conclusion to emphasize that F-SOHDs could play a\nrole in mitigating N O emissions, particularly in certain sulfidic environments, rather than as a\n2\ngeneral mechanism across all estuarine systems. This aligns with the idea that\nchemolithoheterotrophs and chemolithoautotrophs may help suppress N O in environments where\n2\nboth high N O production rates and sulfide stress coexist. Since chemolithoautotrophs are not the\n2\nprimary focus of our study, we have not specifically mentioned their contribution to N O reduction.\n“Thus, our results suggest that F-SOHDs may contribute to the reduction of N O emissions under\n2\ncertain sulfidic conditions that are associated with high N O production rates, although their role\n2\nmight be context-dependent and potentially overlooked in estuarine systems.” (Lines 496–499)\n40. Lines 368-370: I think the introduction, results, and discussion would be improved if the authors\nmade a greater attempt to draw parallels between the microbes identified here, and the recently\ndescribed Arcobacter peruenesis as well as the other ones mentioned earlier. I found it somewhat\nunfortunate that the authors when discussing organic matter and sulfide-rich environments, failed\nto mention Arcobacter peruenesis in that context, but rather discussed chemlithoautotrophs like\nSUP05.\nResponse: Thank you for your comment. Per the reviewer’s insightful suggestion, we have\nexpanded the Introduction and Discussion sections to include more detailed descriptions of\nArcobacter peruensis and other relevant species discussed earlier (see Revised Manuscript Lines\n86–96 and 423–441).\n41. Lines 370-375: Could the authors be a little more specific? For example, what pathways did\nthey identify in these microbial genomes, that made them comparable to the chemolithoheterotrophs\ndescribed here?\nResponse: Done as suggested. These genomes were identified as heterotrophs, possessing genes for\ncomplete denitrification, OM mineralization and sulfide/sulfur/thiosulfate oxidation (see Revised\nManuscript Lines 509–512).\n42. Lines 375-378: The authors did not touch on what features make chemolithoheterotrophs more\nadaptable over chemolithoautotrophs, so can they be more specific? Moreover, it appears that the\nauthors have mixed up the Gibbs free energy values from their figure. According to the authors\nFigure 6, the authors have calculated a Gibbs free energy of -494 and -579 for F-SOHDs related\nprocesses (rxns. C1 and C2), and with chemolithoautotrophy estimated at -566 (rxn. b). But in\nthe text, they list the Gibbs free energy of heterotrophy instead of chemolithoautotrophy. This mix-up\nmakes it seem that F-SOHDs activity is more energetically favorable when in reality it’s not\naccording to Figure 6.\nWith that said the authors have overlooked an important feature that gives\nchemolithoheterotrophs an advantage over chemolithoautotrophs. As detailed by Callbeck et al., in\n2019, the assimilation of acetate into biomass is energetically more favorable than the assimilation\nof CO2 via a chemolithoautotrophic metabolism. This is because CO2 fixation pathways require\nsubstantial amounts of ATP (Erb 2011). The favorable energetics of a chemolithoheterotrophic\nmetabolism explain how Arcobacter can rapidly bloom in the sulfide- and organic-rich upper shelf\nwaters off the coast of Peru and Namibia (Lavik et al. 2009; Callbeck et al. 2019, 2021). This is a\ntheme the authors could consider exploring in the context of the microbes identified in their study.\nResponse: We sincerely thank the reviewer for your insightful comments and suggestions, which\nhave greatly helped to refine and clarify our manuscript. We understand the reviewer has three main\nconcerns: 1) Adaptability of F-SOHDs; 2) Correction of Gibbs Free Energy Values; 3) ATP\nEnergetics and Carbon Assimilation Pathways.\n1) Adaptability of F-SOHDs\nRegarding this point, we believe that F-SOHDs may be more adaptable than\nchemolithoautotrophs due to their dual capacity to thrive in environments with or without sulfide\nand their ability to metabolize under both OM-rich and OM-limited conditions. This adaptability\nenables F-SOHDs to respond rapidly respond to fluctuating environmental conditions. Per the reviewer’s\nsuggestion, we have added this more specific details in the Revised Manuscript Lines 514–516.\n2) Correction of Gibbs free energy values\nWe sincerely apologize for the error in reporting Gibbs free energy values in the previous\nversion of the manuscript. Upon reviewing the calculations and incorporating the reviewer’s\nfeedback (see Comment 69 below), we found that the Gibbs free energy values for the four reactions\nwere incorrectly listed. We have now recalculated the standard Gibbs free energy changes for each\nreaction, and the corrected values are as follows (Revised Figure 6):\n• Reaction a: ∆G° = -565.8 kJ mol⁻¹\n• Reaction b: ∆G° = -480.1 kJ mol⁻¹\n• Reaction c1: ∆G° = -579.3 kJ mol⁻¹\n• Reaction c2: ∆G° = -522.9 kJ mol⁻¹\nThese results show that F-SOHD-associated reactions (c1 and c2) exhibit a lower Gibbs free\nenergy change compared to chemolithoautotrophic denitrification (b), indicating a thermodynamic\nadvantage. We have revised Figure 6 accordingly and corrected the text regarding the Gibbs free\nenergy values (see Revised Manuscript Lines 517–519).\n3) ATP energetics and carbon assimilation pathways\nWe greatly appreciate the reviewer for highlighting the importance of ATP energetics between\ndifferent carbon assimilation pathways. We fully agree that the assimilation of acetate into biomass\nby chemolithoheterotrophs is energetically more favorable than the ATP-intensive CO fixation\n2\npathways used by chemolithoautotrophs. This provides a compelling explanation for the metabolic\nadvantages of F-SOHDs, such as their ability to proliferate rapidly in sulfide- and organic-rich\nenvironments, similar to those described by Callbeck et al. (2019, 2021) off the coasts of Peru and\nNamibia. Per the reviewer’s suggestion, we have incorporated this discussion into the Revised\nManuscript and cited the relevant references to strengthen the context (see Revised Manuscript\nLines 519–522).\nRevised Figure 6. Proposed ecophysiological advantages and environmental adaptations of F-\nSOHDs.\n43. Lines 378-381: Their exceptional performance hinges on their ability to rapidly bloom over\nchemolithoautotrophs for available sulfide, this is something to keep in mind. Overall, I find that\nthe authors could be a bit more specific regarding their industrial application reference. Do the\nauthors mean to use chemolithoheterotrophs to remove N2O in sulfide, and organic matter-rich\nengineered systems?\nResponse: We believe that F-SOHD microorganisms hold significant potential for industrial\napplication in municipal wastewater treatment plants (WWTPs), particularly within anoxic, OM-\nrich denitrification tanks in engineered systems. Toxic sulfide, commonly around 10 mg L−1, is\nfrequently generated during sewage transport and inevitably introduced into the core anoxic\nbiological treatment tanks (Sharma et al. 2008; Liang et al. 2019; Schönharting et al. 1998). These\ntanks are generally dominated by heterotrophic denitrifiers that lack sulfide oxidation capacity,\nleading to increased N O emissions and reduced nitrogen-removal efficiency (Pan et al. 2013;\nof reducing N₂O, could be effectively utilized through the engineered bioaugmentation. By\nintroducing F-SOHDs into anoxic denitrification tanks of WWTPs, we believe it is possible to\nenhance the microbial resilience to sulfide toxicity, maintain high efficiency in nitrogen and organic\nmatter removal, and significantly mitigate N₂O emissions.\nFollowing the suggestion, we have added additional details on the specific industrial\napplications to clarify this point, along with relevant citations (see Revised Manuscript Lines 525–\n530).\n44. Lines 381-383: Fully agree, and I think this is where you could end the discussion. I find the\nnext line regarding climate change and its evolutionary trajectories highly speculative and rather\nvague.\nResponse: Thank you for your positive feedback. As suggested, we have removed the following\nsentences on climate change and its evolutionary trajectories (see Revised Manuscript Line 532–\n535).\nMethods:\n45. Line 389: What month specifically did sampling take place? What was the water depth of the\nstudy site?\nResponse: Sediment samples from the study site were collected in August 2019, at a depth of 3.7\nm. We have incoporated the information into the sentence accordingly (see Revised Manuscript\nLines 539 and 543–544).\n46. Lines 390-392: Do the authors have a reference to support its biogeochemical significance to\nthe nitrogen cycle?\nResponse: We acknowledge that the sentence overstated its global effect, and we therefore deleted\nit in the Revised Manuscript Lines 540–543.\n47. Line 393: Why did the authors pick the depth between 0-20 cm? What is the nitrate and oxygen\npenetration depth?\nResponse: In this study, we selected a sampling depth of 0–20 cm as it is a commonly used depth\nin similar research (Wang et al. 2024; Laing et al. 2009) which can effectively capture the main\nmicrobial processes occurring in surface sediments. During samplings, we measured the oxygen\npenetration depth, which was approximately 2.04 cm. However, we were unable to obtain the nitrate\npenetration depth due to the lack of an available microelectrode in our lab.\n48. Line 394: What is sulfide antioxidant buffer, please be specific.\nResponse: The sulfide antioxidant buffer is a well-established chemical reagent developed by\nKellerlehmann and his colleagues (Kellerlehmann et al. 2006), used to prevent sulfide oxidation\nduring short-term storage. As suggested, this reference has been cited in the Revised Manuscript\nLine 546.\n49. Lines 396-398: I would convert from mg L-1 to uM/mM to be consistent with the rest of the\nmanuscript. Also, how were these nutrient concentrations measured? And by which methods? What\nwere the detection limits?\nResponse: Per the reviewer’s suggestion, we have adjusted the units accordingly and included\nadditional details regarding the measurement of physicochemical parameters and detection limits\n(see Revised Manuscript Lines 548–556).\n50. Lines 404-407: What were the incubation temperatures? And do they differ from in situ\ntemperatures?\nResponse: The sediment microcosms were incubated at 22 C, which closely matches the in situ\nmeasured temperature, ranging from 21.2–23.6 C. This information has been added in Revised\nManuscript Line 563.\n51. Line 420: How was dinitrogen gas measured?\nResponse: Dinitrogen gas in the headspace was measured using an Agilent 7890A gas\nchromatography (USA) equipped with thermal conductivity detector and J&W G3591-81121\npacked column, with a detection limit of 2.6 ppm. As suggested, we have added this information in\nRevised Manuscript Lines 769–772.\n52. Lines 424: How much sediment was used for the extraction? Keep in mind that the readers need\nto be able to repeat the experiments.\nResponse: Approximately 0.25 g sediment was used for the DNA extraction in our study (see\nRevised Manuscript Line 582).\n53. Lines 424-428: The authors provide no mention of references either in the qPCR section or\nsupplementary Table 2 regarding the primers. Surely, these primers were designed elsewhere and if\nso, the authors need to indicate this information.\nResponse: Per the reviewer’s suggestion, relevant references for the primers has been provided in\nthe Revised Supplementary Table 2 below (also see Revised Supplementary Information).\nRevised supplementary Table 2. Primers and references of the eleven genes selected for qPCR\nquantification.\nTarget\nPrimer Primer sequence (5'-3') References\ngene\nDSR-1F+ ACSCACTGGAAGCACGCCGG (Kondo et al.\ndsrA\nDSR-R GTGGMRCCGTGCAKRTTGG 2004)\nForward GCTCGGCAGCCTCAATAC\nsqr (Yin et al. 2014)\nReverse GGTCGGACGGTGGTTACTG\n710F ATCGGYCAGGCYTTYCCSTA (Tourna et al.\nsoxB\n1184R MAVGTGCCGTTGAARTTGC 2014)\nnrfAF2aw CARTGYCAYGTBGARTA (Welsh et al.\nnrfA\nnrfAR1 TWNGGCATRTGRCARTC 2014)\nV17m TGGACVATGGGYTTYAAYC\nnapA (Bru et al. 2007)\nnapA-4r ACYTCRCGHGCVGTRCCRCA\nnarG-f TCGCCSATYCCGGCSATGTC\nnarG (Bru et al. 2007)\nnarG-r GAGTTGTACCAGTCRGCSGAYTCSG\nnirS1F TACCACCCSGARCCGCGCGT (Braker et al.\nnirS\nnirS3R GCCGCCGTCRTGVAGGAA 1998)\n876F ATYGGCGGVCAYGGCGA (Henry et al.\nnirK\n1040R GCCTCGATCAGRTTRTGGTT 2004)\npF CATGGCGCTGATAACGGG (Dandie et al.\nnorB\npR CTTIACCATGCTGAAGGCG 2007)\nZ2F CGCRACGGCAASAAGGTSMSSGT (Henry et al.\nnosZ\nZ2R CAKRTGCAKSGCRTGGCAGAA 2006)\n16S 341F CCTACGGGNGGCWGCAG (Herlemann et\nrRNA 805F GACTACHVGGGTATCTAATCC al. 2011)\n54. Lines 428-429: Is this common to do in qPCR, because cells often have more than one gene\ncopy? So, the cell numbers are going to be overestimated?\nResponse: Yes, it is common to use this approach in qPCR and is widely applied in many DNA-SIP\nstudies (Wasmund et al., 2021; He et al., 2022; Thomas et al., 2019; Yin et al., 2019). The analysis\nof the 16S rRNA gene across DNA buoyant density fractions helps to determine microbial\ndistribution and the extent of 13C incorporation. While microbial cells can have multiple 16S rRNA\ngene copies, potentially leading to overestimation, this error impacts all density fractions uniformly.\nTherefore, the method remains reliable for comparing shifts in microbial communities along isotope\ngradients under different experimental conditions.\n55. Lines 430-432: What qPCR chemistry was used, and how much DNA template was used? Were\nDNA concentrations also verified, and how? For the standards what concentration range was used\nin genes per L? Was melt curve analysis also done?\nResponse: All qPCR assays were performed in a 20 µl reaction, containing 10 L of TB-Green\nPremix Ex Taq (Takara, Japan), 0.4 L of ROX Reference Dye, 0.4 L of each forward/reverse\nprimer (10 M), 2 L of standardized DNA template, and 6.8 L of DNAase-free water. DNA\nconcentrations were measured using the Quant-iT PicoGreen dsDNA kit (Thermo Scientific, USA),\nand standardized to 17.5 ng L-1 for each DNA template. A melt curve analysis was conducted for\neach qPCR experiment. As suggested, these details have been added in Revised Manuscript Lines\n588–596.\n56. Lines 434-445: Is this a completely novel technique or is there other studies that have done the\nsame?\nResponse: The washing process conducted inside the anaerobic workstation is a customized\nsediment pre-treatment method in our lab. This approach effectively remove undesirable impurities\nand sand from sediments, thereby preserving the biomass-rich portions for subsequent microcosm\nexperiments. To support this procedure, we have cited a recent published work (see Revised\nManuscript Line 600). Regarding the construction of DNA-SIP-based microcosms, this technique\nis relatively common and aligns with approaches used in other studies (Dunford and Neufeld, 2010;\nSathyamoorthy et al. 2018; Thomas et al., 2019).\n57. Lines 443-445: What was the 13C labeling percent for the incubation experiments?\nResponse: The percents of 13C and 13C labeling in the incubation experiments were 98–99% and\ni o\n99%, respectively. This information has been added in the Revised Manuscript Lines 608–609.\n58. Line 488: Change “compacity” to “capacity”\nResponse: Revised as suggested (see Revised Manuscript Line 653).\n59. Lines 495-497: Please provide a reference to the study that designed these primers. And also\nindicate if these are universal primers or primers used to only amplify the Bacterial community.\nResponse: As suggested, we have provided the reference (Herlemann et al. 2011) for the primers\nspecifically designed for amplifying bacterial communities (see Revised Manuscript Lines 660–\n662).\n60. Lines 497-500: Can the authors indicate the number of raw reads generated?\nResponse: Done as suggested (see Revised Manuscript Lines 663–664).\n61. Lines 504-506: Please provide a reference to the study that designed these primers.\nResponse: Done as suggested (Singer al. 2016; see Revised Manuscript Line 672).\n62. Lines 514-515: Is there a reference for this line?\nResponse: Done as suggested (Grob et al. 2015; see Revised Manuscript Line 681).\n63. Lines 551-552: Perhaps I missed this somewhere, but what is the composition of the medium in\nterms of salts and other nutrients. Or is the medium just filtered river water with amended substrates?\nPlease clarify.\nResponse: In addition to the target substrates, such as inorganic carbon (CO and bicarbonate),\n2\ncomposite OM (glucose, propionate, acetate, pyruvate, and fermentative DL-lactate), nitrate, and\nsulfide, the medium also contains essential salts and nutrients like ammonium (NH Cl), sulfate\n4\n(Na SO ), disodium phosphate, HEPES buffer, and trace elements. The medium composition has\n2 4\nalready been described in the “Isotope incubation experiments” section in Methods (see Revised\nManuscript Lines 603–607). While filtered river water was used for in-situ sediment incubation and\nDNA-SIP microcosm experiments, sterile deionized water was employed in the long-term\nenrichment experiments to maintain greater control and prevent impurities from interfering with the\ngrowth of F-SOHDs. Additional details on the medium composition have been added for\nclarification (see Revised Manuscript Lines 721–725).\n64. Lines 555-558: If CO is removed, how is the system then pH buffered? Based on the principle\n2\nthat chemolithoautotrophs are slower growing compared to chemolithoheterotrophs I don’t fully\nunderstand the reasoning behind removing CO to hinder chemolithoautotrops. As we know,\n2\nchemolithoheterotrophs when supplied sulfide, nitrate, and organic matter will naturally\noutcompete chemolithoautotrophs for available sulfide (Callbeck et al., 2019; 2021). So, actively\nremoving CO could have downstream consequences for pH buffering in the system. Do the authors\n2\nhave pH measurements?\nResponse: Thank you for your question. In our experiments, we aimed to enrich the highly pure F-\nSOHDs, which have been confirmed to exhibit an obligate heterotrophic lifestyle through our\npreliminary experiments. While it is understood that chemolithoheterotrophs like F-SOHDs\nnaturally outcompete chemolithoautotrophs in the presence of organic matter, chemolithoautotrophs\nmay still persist by utilizing CO produced by chemolithoheterotrophs, albeit at a slower rate. To\n2\nprevent this, we actively removed CO to suppress the growth of chemolithoautotrophs and enhance\n2\nthe purity of F-SOHDs.\nTo stabilize the pH after CO removal, we employed HEPES buffer, which is effective in the\n2\nneutral pH range (6.8 to 8.2) and commonly applied in microbial cultivation (Kraft et al. 2014),\ninstead of carbonate/bicarbonate buffering. Throughout the enrichment process, pH was\nconsistently monitored and remained stable at 7.10 ± 0.17.\n65. Line 580: This could be shortened to read, “Physiological experiments”.\nResponse: Done as suggested (see Revised Manuscript Line 750).\n66. Lines 583-585: Do the authors know which heterotrophic denitrifiers were enriched? And do\nthey lack a capacity for sulfide oxidation and have the nosZ gene? I’m wondering how distinct they\nare from the sulfide-oxidizing heterotrophs.\nResponse: We previously performed 16S rRNA gene amplicon sequencing to examine the microbial\ncomposition of HD-enriched community, which revealed that it is predominantly composed of the\ngenus Pseudomonas (97.9% in relative abundance) (Figure R2 below). To assess the metabolic\npotential for sulfur oxidation and denitrification, we conducted PCR amplification targeting the key\ngenes sqr and nosZ. Electrophoresis results showed a broad and bright band for the N O-reducing\n2\nnosZ gene, while no amplification was observed for the sulfide-oxidizing sqr gene (Figure R3\nbelow). This suggests that the HD community comprises non-sulfur-oxidizing conventional\nheterotrophic denitrifiers capable of reducing nitrate to dinitrogen gas but lacking the capacity to\ndetoxify sulfide. This finding is further corroborated by the results of reaction kinetic experiments\n(Figure R4a–d below). Moreover, earlier studies have demonstrated that Pseudomonas fluorescens\nand Pseudomonas aeruginosa are similarly inhibited by sulfide, leading to increased N O emissions\n2\n(Sørensen et al. 1980; Tam et al. 1979). Taken together, our findings from microbial composition,\nPCR amplification, and reaction kinetics consistently indicate the presence of non-sulfur-oxidizing\nheterotrophic denitrifiers in the HD community.\nFigure R2. Amplicon sequencing of 16S rRNA gene of the enriched HD microbial community.\nOnly the genera with a relative abundance higher than 0.05% were shown.\nFigure R3. Ultraviolet imaging of PCR products after electrophoresis for nosZ (484 bp) and sqr (136\nbp) genes.\nFigure R4. Reaction kinetics of the HD microbial community under sulfide-free (a, b) and sulfide-\npresent conditions (c, d) , respectively.\n67. Lines 595-596: The authors should provide more information regarding the instruments and\nstandards used for gas analysis.\nResponse: Done as suggested (see Revised Manuscript Lines 766–772).\n68. Lines 608-611: This is too vague. Please provide citations and additional information on how\nthese sulfur species were quantified.\nResponse: We have incorporated additional information regarding sulfur determination as\nsuggested, and relevant references have been cited accordingly (see Revised Manuscript Lines 786–\n791).\nFigures:\n69. Figure 6: How was Gibbs free energy calculated and where did they get the formation energies\nfrom? Under what conditions was this calculated, for example, pH, sulfide, nitrate concentrations?\nReaction c2 is driven by mostly sulfide (1:5 ratio), with a little bit of organic matter, which seems\nmore like a chemolithoautotrophy with a little bit of heterotrophy. What does this look like if the\nstoichiometry is a bit more balanced?\nResponse: The standard Gibbs free energy change (∆G0) for each reaction was calculated using the\nstandard method, which involves subtracting the total formation free energies of the reactants from\nthe total formation free energies of the products. The formation energies for each substance were\nobtained from the standard thermodynamic database provided in “Section 5 Thermochemistry,\nElectrochemistry, and Solution Chemistry” of the CRC Handbook of Chemistry and Physics\n(William M. Haynes ed. 2014). These calculations were performed under standard conditions, which\ntypically assume a temperature of 298 K, pressure of 1 atm, and reactant/product concentrations of\ncalculation, as the formation energies used for the calculation of reaction’s ∆G0 are independent of\nthese variables. To improve clarity for readers, we have added an explanation of the calculation\nmethods in Revised Manuscript Lines 1223–1227.\nPer the reviewer’s suggestion, we have adjusted the stoichiometry of organic matter and sulfide\nin the reaction c2 to achieve a more balanced representation and recalculated the corresponding ∆G0.\nThe updated results show that the revised reaction c2 has a more negative Gibbs energy change\n(∆G0(c2) = -522.9 kJ mol-1) compared to that of chemolithoautotrophic denitrifiers (∆G0(b) = -480.1\nkJ mol-1). This indicates a thermodynamic advantage in reaction free energy for F-SOHDs,\nsupporting their metabolic potential in this system (Fig. 6).\nReaction c2:\n0.312 C H O - + 0.313 HS- + NO - + H+ → 0.624 CO + 0.313 SO 2- + 0.5 N + 1.125 H O\n2 3 2 3 2 4 2 2\n∆G0(c2) = -522.9 kJ mol-1\nLiterature cited in my review:\nArevalo-Martinez, D. L., A. Kock, C. R. Loscher, R. A. Schmitz, and H. W. Bange. 2015. Massive\nnitrous oxide emissions from the tropical South Pacific Ocean. Nat. Geosci 8: 530–533.\ndoi:10.1038/ngeo2469\nArévalo-Martínez, D. L., T. Steinhoff, P. Brandt, A. Körtzinger, T. Lamont, G. Rehder, and H. W.\nBange. 2019. N2O Emissions From the Northern Benguela Upwelling System. Geophys. Res. Lett.\n46: 3317–3326. doi:10.1029/2018GL081648\nBoltyanskaya, Y. V, V. V Kevbrin, A. M. Lysenko, T. V Kolganova, T. P. Tourova, G. A. Osipov, and\nT. N. Zhilina. 2007. Halomonas mongoliensis sp. nov. and Halomonas kenyensis sp. nov., new\nhaloalkaliphilic denitrifiers capable of N2O reduction, isolated from soda lakes. Microbiology 76:\n739–747. doi:10.1134/S0026261707060148\nBourbonnais, A., R. T. Letscher, H. W. Bange, V. Échevin, J. Larkum, J. Mohn, N. Yoshida, and M.\nA. Altabet. 2017. N2O production and consumption from stable isotopic and concentration data in\nthe Peruvian coastal upwelling system. Global Biogeochem. Cycles 31: 678–698.\ndoi:10.1002/2016GB005567\nCallbeck, C. M., D. E. Canfield, M. M. M. Kuypers, P. Yilmaz, G. Lavik, B. Thamdrup, C. J. Schubert,\nand L. A. Bristow. 2021. Sulfur cycling in oceanic oxygen minimum zones. Limnol. Oceanogr. 66:\n2360–2392. doi:https://doi.org/10.1002/lno.11759\nCallbeck, C. M., G. Lavik, T. G. Ferdelman, and others. 2018. Oxygen minimum zone cryptic sulfur\ncycling sustained by offshore transport of key sulfur oxidizing bacteria. Nat. Commun. 9: 1729.\ndoi:10.1038/s41467-018-04041-x\nCallbeck, C. M., C. Pelzer, G. Lavik, and others. 2019. Arcobacter peruensis sp.nov., a\nchemolithoheterotroph isolated from sulfide and organic rich coastal waters off Peru. Appl. Environ.\nMicrobiol. 85: e01344-19. doi:10.1128/AEM.01344-19\nErb, T. J. 2011. Carboxylases in Natural and Synthetic Microbial Pathways. Appl Env. Microbiol\n77: 8466–8477. doi:10.1128/aem.05702-11\nFelgate, H., G. Giannopoulos, M. J. Sullivan, A. J. Gates, T. A. Clarke, E. Baggs, G. Rowley, and\nD. J. Richardson. 2012. The impact of copper, nitrate and carbon status on the emission of nitrous\noxide by two species of bacteria with biochemically distinct denitrification pathways. Environ.\nMicrobiol. 14: 1788–1800. doi:10.1111/j.1462-2920.2012.02789.x\nGranger, J., and B. B. Ward. 2003. Accumulation of nitrogen oxides in copper-limited cultures of\ndenitrifying bacteria. Limnol. Oceanogr. 48: 313–318. doi:10.4319/lo.2003.48.1.0313\nKock, A., D. L. Arévalo-Martínez, C. R. Löscher, and H. W. Bange. 2016. Extreme N 2 O\naccumulation in the coastal oxygen minimum zone off Peru. Biogeosciences 13: 827–840.\ndoi:10.5194/bg-13-827-2016\nLavik, G., T. Stührmann, V. Brüchert, and others. 2009. Detoxification of sulphidic African shelf\nwaters by blooming chemolithotrophs. Nature 457: 581–584. doi:10.1038/nature07588\nLi, P., W. Yuan, Y. Huang, C. Zhang, C. Ni, Q. Lin, Z. Zhu, and J. Wang. 2022. Complete genome\nsequence of Pseudomonas stutzeri S116 owning bifunctional catalysis provides insights into\naffecting performance of microbial fuel cells. BMC Microbiol. 22: 137. doi:10.1186/s12866-022-\n02552-8\nManconi, I., P. van der Maas, and P. Lens. 2006. Effect of copper dosing on sulfide inhibited\nreduction of nitric and nitrous oxide. Nitric oxide Biol. Chem. 15: 400–407.\ndoi:10.1016/j.niox.2006.04.262\nMoffett, J. W., C. B. Tuit, and B. B. Ward. 2012. Chelator-induced inhibition of copper\nmetalloenzymes in denitrifying bacteria. Limnol. Oceanogr. 57: 272–280.\ndoi:10.4319/lo.2012.57.1.0272\nNaqvi, S. A. W., H. Naik, D. A. Jayakumar, M. S. Shailaja, and P. V Narvekar. 2006. Seasonal oxygen\ndeficiency over the western continental shelf of India , p. 195–224. In N.L. Neretin [ed.], Past and\nPresent Water Column Anoxia. Springer Netherlands.\nNaqvi, S. W. A., D. A. Jayakumar, P. V Narvekar, H. Naik, V. V. S. S. Sarma, W. D’Souza, S. Joseph,\nand M. D. George. 2000. Increased marine production of N2O due to intensifying anoxia on the\nIndian continental shelf. Nature 408: 346–349. doi:10.1038/35042551\nSorokin, D. Y. 2003. Oxidation of Inorganic Sulfur Compounds by Obligately Organotrophic\nBacteria. Microbiology 72: 641–653. doi:10.1023/B:MICI.0000008363.24128.e5\nWalsh, D. A., E. Zaikova, C. G. Howes, Y. C. Song, J. J. Wright, S. G. Tringe, P. D. Tortell, and S. J.\nHallam. 2009. Metagenome of a versatile chemolithoautotroph from expanding oceanic dead zones.\nScience (80-. ). 326: 578–582. doi:10.1126/science.1175309\nWang, L., and Z. Shao. 2021. Aerobic Denitrification and Heterotrophic Sulfur Oxidation in the\nGenus Halomonas Revealed by Six Novel Species Characterizations and Genome-Based Analysis.\nFront. Microbiol. 12: 652766. doi:10.3389/fmicb.2021.652766\nWong, B.-T., and D.-J. Lee. 2014. Pseudomonas yangmingensis sp. nov., an alkaliphilic denitrifying\nspecies isolated from a hot spring. J. Biosci. Bioeng. 117: 71–74.\ndoi:https://doi.org/10.1016/j.jbiosc.2013.06.006\nResponses to Reviewer #3\nReviewer #3 (Remarks to the Author):\nThis study identifies a previously unknown group of heterotrophic denitrifiers capable of coupling\nsulfur oxidation with denitrification autonomously. Traditionally, heterotrophic denitrifiers have\nbeen thought to be hampered by sulfide, which negatively affects their growth and increases nitrous\noxide (N₂O) emissions, a potent greenhouse gas. However, in this study, these bacteria were found\nto use sulfide as an alternative electron source, enhancing denitrification and reducing nitrous oxide\n(N₂O) emissions. Through microcosm incubations, DNA stable-isotope probing, and metagenomics,\nthe researchers demonstrated that these versatile heterotrophs exhibit a flexible physiology allowing\nthem to thrive in sulfide-rich environments and compete effectively with chemolithoautotrophic\ndenitrifiers. This discovery shifts our understanding of sulfur cycling and suggests new implications\nfor mitigating climate change.\nResponse: We sincerely appreciate the reviewer’s positive and encouraging feedback on our work!\nWe have revised the manuscript by fully considering and addressing all the comments as suggested.\nPlease see our detailed responses below.\nSpecific Comments\n1. There are a few grammatical errors and some sentences are lengthy and complex, which might\nhinder comprehension. Simplify and break down long sentences where possible.\nResponse: Thank you for your comment. Following the reviewer’s subsequent suggestions, we are\ncommitted to making the manuscript clear and digestible to the audience. Additionally, we have\ndone our best to simplify long and complicated sentences throughout the manuscript. Please see\nRevised Manuscript Lines 69–72, 73–76, 80–82, 102–108, 202, 211, 232, 234–237, 279–283, 302–\n304, 346, 394–400, 442–447, 462, 495, 507, 509, 514–519, 568, 594–596, 653, and 1219–1223.\n2. Line 62-64: suggested revision: “Incomplete denitrification is a major source of N₂O, a potent\ngreenhouse gas that contributes to climate warming. Additionally, N₂O emissions contribute to\nozone layer depletion and acid rain formation.”\nResponse: Done as suggested. This sentence has been rephrased in response to the comments\nprovided by Reviewers #2 and #3 (see Revised Manuscript Lines 69–72).\n3. Line 65-68: Please revise to “In many estuaries, coastal regions, and marine oxygen-deficient\nzones, the sulfur cycle plays a crucial role in regulating biogeochemical processes. This is due to\nthe dominance of sulfate as a terminal electron acceptor and the subsequent release of hydrogen\nsulfide (H₂S) from benthic sediments.”\nResponse: Done as suggested (see Revised Manuscript Lines 73–76).\n4. Line 68-75: This part should include a more detailed introduction to sulfide-driven dissimilatory\nnitrate reduction to ammonium (DNRA) (Canfield et. al, 2010 (Science); Wang et. al, 2021 (Water\nResearch), etc.).\nResponse: Thank you for the valuable suggestion and references. We have revised as suggested (see\nRevised Manuscript Lines 82–85). The sentence is pasted below for convenience.\n“Some of these chemolithoautotrophs are also involved in dissimilatory nitrate reduction to\nammonium (DNRA), thereby supplying ammonium to support microbial growth of indigenous\nbacteria20,21.” (Lines 82–85)\n5. Line 71-73: Please revise to “Sulfide is highly toxic to heterotrophic denitrifiers, adversely\naffecting microbial activity and potentially increasing N₂O emissions.”\nResponse: Revised as suggested (see Revised Manuscript Lines 76–79). This sentence has been\nrephrased in response to the comments provided by Reviewers #2 and #3.\n6. Line 76-80: “In this study, we investigate the response of heterotrophic denitrifiers to sulfide in\nthe highly productive Songhua River Estuary, a major tributary within the largest water system in\nEast Asia. We employed a combination of microcosm and lab culture experiments, metagenomics,\nand a novel community-isotope-corrected DNA stable-isotope probing (DNA-SIP) approach.”\nResponse: Revised as suggested (see Revised Manuscript Lines 102–108).\n7. In sediment microcosms, how do community compositions shift over time under different\ntreatment conditions? How is the stability and resilience of these microbial communities in response\nto fluctuating environmental factors?\nResponse: Thank you for your comment. Unfortunately, we do not have data on the dynamic shifts\nin community composition. Whether a time-series investigation is necessary depends on the\nscientific question. In our study, the primary goal was to identify the key contributors in response\nto sulfide stimulation under organic matter-rich conditions, which we addressed using DNA-SIP\ntechnique. The timeframe for DNA-SIP experiments is typically determined by the time required\nfor effective isotope incorporation. Shorter durations risk incomplete labelling, while extended\nincubation can lead to cross-feeding, complicating accurate detection of isotope-assimilating\nmicrobes. Prior to the DNA-SIP-based incubation, our preliminary experiments showed that\nsubstrates were fully consumed within 65–83 hours in the +C+N+S, +C+N, and +C+N+S\ni\ntreatments. Based on this and literature, we chose a 10–12 day incubation (~ three generations), a\nwidely accepted timeframe in previous DNA-SIP-based studies (Cai et. al, 2016; Zhang et. al, 2021;\nWilhelm et. al, 2019; Dunford and Neufeld, 2010). This effectively labels most microorganisms,\nwhile avoiding cross-feeding. Although we did not track microbial community composition over\ntime, we recognize that such data could provide valuable insights into how communities respond to\nfluctuating environmental factors. However, we feel this question is somewhat broad and may\nbeyond the scope of our current study.\n8. How do the lab-based findings translate to natural estuarine environments? Are the conditions\nin the microcosm experiments reflective of those in natural settings, and how might environmental\nvariability influence the activity of F-SOHDs? Can modelling approaches be used to predict their\ncontributions to nutrient cycling and greenhouse gas emissions under various environmental\nscenarios?\nResponse: We acknowledge the complexities in translating lab-based findings to natural estuarine\nenvironments. In our study, we aimed to simulate in situ conditions by using samples directly\ncollected from estuarine environments for microcosm incubations. While this approach allows us to\nclosely mimic natural settings, we recognize that natural estuarine systems are far more complex\nand variable compared to controlled lab environments. As a result, microcosm experiments may not\nfully capture the unpredictable dynamics of natural ecosystems.\nEnvironmental variability, such as fluctuating oxygen levels, temperature, salinity, and nutrient\navailability, can significantly influence the activity of F-SOHDs in situ, which may differ from the\nstable conditions observed in microcosms. In future research, we plan to deploy an on-site chamber\nand conduct in situ incubations with various substrates provided to assess whether F-SOHDs are the\nmain contributors in natural environments. During the incubation, we will leverage static respiration\nchamber measurements, qPCR, DNA-SIP based microcosm assays and metagenomic sequencing to\nexamine nitrate reduction, organic matter consumption and sulfur oxidation, further exploring the\npotential coupling of carbon, nitrogen and sulfur cycling. To better understand these dynamics,\nfuture studies incorporating time-series field investigations could also provide valuable insights into\nhow F-SOHDs adapt and respond over time in natural environments.\nRegarding modeling approaches, they offer a potential tool to bridge the gap between lab-based\nexperiments and real-world conditions. By integrating data from microcosm studies, time-series\nfield observations, and environmental variables, models can help predict the contributions of F-\nSOHDs to nutrient cycling and greenhouse gas emissions under different environmental scenarios.\nThese predictive models could improve our understanding of how microbial activity fluctuates\nacross various estuarine conditions and inform broader ecosystem management strategies.\n9. Given the ability of F-SOHDs to reduce N2O emissions, how can this be applied in wastewater\ntreatment? What are the practical steps to harness these microbes in engineered systems to mitigate\ngreenhouse gas? Can the activity of F-SOHDs be enhanced in situ through biostimulation strategies?\nWhat types of amendments (e.g., organic carbon, sulfide) would be most effective, and what are the\npotential ecological side effects of such interventions?\nResponse: It is a very insightful comment. F-SOHDs offer significant potential for reducing N O\n2\nemissions in wastewater treatment processes. In practical settings, particularly in long-distance\nsewage transport pipelines, sulfide can be generated biologically or chemically at concentrations of\naround 10 mg L−1 (Sharma et. al, 2008; Liang et. al, 2019; Schönharting et. al, 1998). This sulfide\noften enters the anoxic denitrification tanks in wastewater treatment plants, where it can inhibit\nconventional heterotrophic denitrifiers, leading to increased N O emissions (Pan et al., 2013;\nOur findings demonstrate that F-SOHDs, which are essentially heterotrophic denitrifiers with\nthe additional ability to detoxify sulfide, can mitigate these emissions. Rather than biostimulation,\nwe believe bioaugmentation might be a more effective strategy for harnessing their potential in\nengineered systems. Introducing F-SOHDs into anoxic denitrification tanks could unlock their\nability to reduce N O emissions while maintaining efficient nitrogen and COD removal.\nsignificantly reducing the overall greenhouse gas footprint.\nIn terms of amendments, moderate levels of sulfide could be beneficial, helping to\nregulate competition between F-SOHDs and conventional heterotrophic denitrifiers, thus\nensuring the stable presence and activity of F-SOHDs during long-term operation. However,\none potential ecological side effect is the increase in sulfate concentrations in the effluent due\nto the sulfur-oxidizing capabilities of F-SOHDs. Careful monitoring and adjustment of sulfide\nlevels may be necessary to balance N O reduction with the prevention of excessive sulfate\n2\naccumulation.\nFurther research into optimal amendment strategies and the long-term effects of F-SOHDs\nthrough bioaugmentation in wastewater systems would provide valuable insights into\nmitigating both environmental impact and operational challenges.\nFollowing the suggestion, we have added additional details on the potential industrial\napplications of F-SOHDs in wastewater treatment to clarify this point (see Revised Manuscript\nLines 525–530).\n10. While metagenomics provides a snapshot of potential metabolic capabilities, it does not confirm\nactual activity. How can the findings from metagenomics be integrated with other omics approaches\nto provide a more comprehensive understanding of microbial functions in these environments?\nResponse: Thank you for your comment. We acknowledge that relying solely on metagenomics\nprovides only a snapshot of microbial communities and their potential metabolic capabilities. To\ngain a more comprehensive understanding of microbial activity, incorporating other omics\ntechniques, such as metatranscriptomics, metaproteomics, and metabolomics, would be beneficial.\nHowever, we want to emphasize that our study employed a combined approach of metagenomics\nand DNA-SIP. DNA-SIP directly identifies active microorganisms by tracking isotope incorporation\ninto their DNA, ensuring that only metabolically active microbes are detected. By integrating DNA-\nSIP with metagenomics, we can not only identify potential functions but also confirm actual\nmetabolic activity under specific conditions. This, together with various nutrient treatments and\nisotope labeling, allows us to accurately pinpoint key contributors to element cycling in these\nenvironments."}, {"round": 2, "reviewer_comments": {"reviewer_1": "No more questions.", "reviewer_2": "I thank the authors, Shao et al., for making the effort to address my comments in full. The authors made several important\nclarifications across the manuscript regarding the discussion of chemolithoheterotrophic bacteria more generally, added a\nmore nuanced discussion of method caveats, and a more refined assessment of results from the DNA-SIP experiments.\nMoreover, the authors corrected mistakes regarding the Gibbs-free energy calculations. After reviewing the revised\nmanuscript, I did not find any other issues. Again, I commend the authors on their excellent work and look forward to seeing\nthe manuscript published.", "reviewer_3": "No further comments"}, "author_response": ""}]

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