Molecular evidence for sediment nitrogen fixation in a temperate New England estuary

Introduction

Nitrogen (N) limitation in marine waters is controlled, in large part, by the balance between denitrification and nitrogen fixation (N₂-fixation). While denitrification removes biologically reactive N from the environment, N₂-fixation converts dinitrogen gas (N₂) into biologically available ammonium. Although N₂ is the most abundant element in the atmosphere, the majority of organisms on earth cannot use N in this form. Thus, N-fixing organisms, known as diazotrophs, are unique in that they can grow without external sources of reactive N by drawing upon the high concentrations of N₂ dissolved in seawater (∼400–574 µmol L⁻¹ N₂: Sharp, 1983; Zehr & Paerl, 2008). Through the fixation of di-nitrogen gas, diazotrophs have the ability to increase reactive N availability and, subsequently primary and secondary production (Montoya, Carpenter & Capone, 2002; Ryther & Dunstan, 1971).

The genetic potential for N₂-fixation is widespread among the Domains of Bacteria and Archaea (Zehr et al., 2003). In fact, geochemists argue that N cannot be the ultimate limiting nutrient, as diazotrophic microorganisms possess the ability to reduce N deficiencies given sufficient time (Redfield, 1958; Tyrrell, 1999). Despite this genetic potential, filamentous cyanobacteria (e.g., Trichodesmium) and diatom symbiotic cyanobacteria (e.g., Richelia) were traditionally thought to be the only marine diazotrophs that produce a significant source of new N to marine environments (Capone et al., 1997; Karl et al., 1997; Zehr et al., 2001). Additionally, N₂-fixation was thought to be of only minor importance compared to N inputs from upwelling and advection of nitrate rich water in the open ocean (Zehr & Paerl, 2008) and river N loading in coastal systems (Howarth et al., 1988). Furthermore, heterotrophic sediment N₂-fixation was generally considered to be of minimal significance to subtidal benthic N cycling, contributing as little as 0.1% of total new N to the environment (Howarth et al., 1988).

This view however, has changed dramatically over the last decade, with observations of significant N₂-fixation by both unicellular autotrophic and heterotrophic diazotrophs in a range of marine environments. For example, unicellular cyanobacteria are now thought to contribute substantially to N₂-fixation in temperate (Rees, Gilbert & Kelly-Gerreyn, 2009; Bentzon-Tilia et al., 2015b), subtropical (Zehr et al., 2001), and tropical (Goebel et al., 2007) water columns. Significant unicellular heterotrophic N₂-fixation has also been reported in water columns from the Mediterranean Sea (Rahav et al., 2013), the Baltic Sea (Bostrom et al., 2007), and the hypoxic Southern California Bight (Hamersley et al., 2011). Additionally, N₂-fixation has been found at hydrothermal vents (Mehta & Baross, 2006), in cold seep sediments (Dekas, Poretsky & Orphan, 2009), and associated with the decomposition of a large marine macrophyte (Hamersley et al., 2015). Recently, substantial fluxes of sediment N₂-fixation has been observed through direct N₂ uptake (Dias et al., 2012; Grosskopf et al., 2012; Rao & Charette, 2012) and nifH gene expression (Fulweiler et al., 2013; Andersson et al., 2014). In some cases, the reported sediment N₂-fixation comprises a significant portion of the system N budget (Fulweiler & Heiss, 2014).

In order to expand our understanding of the potential role of N₂-fixation as a nitrogen source in marine environments, we examined the expression of nifH, a gene encoding for nitrogenase (the enzyme associated with N₂-fixation), in the sediments of a temperate New England estuary (Waquoit Bay, MA). The aim of this research was to investigate the spatial distribution of nifH transcript diversity at three sites with different environmental characteristics and to link these results to concurrently collected biogeochemical data. We had three specific questions: (1) Which organisms appear to be responsible for the observed nifH expression? (2) How did the active community composition and diversity change at these three estuarine sites? (3) Does nifH expression occur simultaneously with net sediment denitrification as has been reported for sediments in a nearby estuary (Fulweiler et al., 2013)? If so, this would indicate that these two opposite processes overlap in space and time highlighting the dynamic nature of the marine nitrogen cycle and how much more we need to learn about what controls its major processes. In turn, these findings will help us better inform estuarine N budgets, as we hypothesize that this process may provide a significant and yet overlooked source of N.

Materials and Methods

Description of study area

Waquoit Bay is a shallow (mean depth 1.8 m) estuary on the southwestern shore of Cape Cod, Massachusetts (Fig. 1). Although the estuary is mostly landlocked, it opens to Vineyard Sound through a narrow (100 m wide) channel along its southern edge. Within Waquoit Bay we focused on three sites with varying in situ nutrient conditions and sediment net N₂ fluxes (Table 1). Two of these sites, Metoxit Point (MP) and South Basin (SB), are located within the main estuary of Waquoit Bay and the third, Sage Lot Pond (SLP), is located just to the east (Fig. 1). MP is located directly downstream of the Quashnet River, which has a mean annual load of 50 × 10³ kg N km⁻² y⁻¹ (Bowen & Valiela, 2001; Foster & Fulweiler, 2014; Valiela et al., 1997). It is characterized as having extensive macroalgae blooms and extreme fluctuations in diel summer oxygen conditions. SB is located close to the mouth of the estuary where tidal flushing with Vineyard Sound water occurs and it is therefore presumably less impacted by anthropogenic N loading. Finally, SLP is our least N impacted site as it is located in an undeveloped, forested watershed and has a low annual N loading rate (2.1 × 10³ kg N km⁻² y⁻¹; Valiela et al., 1997; Valiela et al., 2000).

Table 1:

Physical and biogeochemical characteristics for the Waquoit Bay sediment cores used for the molecular analyses in this study.

Inorganic nutrient concentrations and salinity are from the water overlying each sediment core before the start of the incubations. The net N₂–N fluxes across the sediment water interface are calculated for each core using a 5-point linear regression and rates were prorated for the volume and area of the cores. All data from Foster (2012) and Foster & Fulweiler (2014).

Site	Core-ID	Incubation date	Incubation temperature, °C	Salinity	${\mathrm{NH}}_4^{+}\mathrm{\mu }\mathrm{mol}{\mathrm{L}}^{-1}$	${\mathrm{NO}}_2^{-}+{\mathrm{NO}}_3^{-}\mathrm{\mu }\mathrm{mol}{\mathrm{L}}^{-1}$	${\mathrm{PO}}_4^{3-}\mathrm{\mu }\mathrm{mol}{\mathrm{L}}^{-1}$	Net N2–N flux μmol m−2 h−1
Metoxit Point	16	24-Aug-2011	25	30	11.35	0.11	0.58	−50
	7	11-Oct-2011	20	31	34.10	0.76	1.94	0
	18	11-Oct-2011	20	31	18.49	0.13	0.92	−51
	8	11-Oct-2011	20	31	98.91	0.30	5.36	nd
South Basin	9	7-Jul-2011	26	32	7.37	0.00	0.67	0
	1	24-Aug-2011	25	32	1.00	0.12	0.44	−51
	5	24-Aug-2011	25	32	1.87	0.15	0.41	28
	1	11-Oct-2011	20	31	nd	0.12	0.57	23
	6	11-Oct-2011	20	31	nd	0.12	0.54	75
	4	11-Oct-2011	20	31	nd	0.11	0.60	0
	5	11-Oct-2011	20	31	nd	0.12	0.49	15
Sage Lot Pond	8	29-Jul-2011	25	30	9.66	0.28	0.54	24
	8	24-Aug-2011	25	30	5.19	0.27	0.56	0
	19	24-Aug-2011	25	29	9.85	nd	0.76	191

DOI: 10.7717/peerj.1615/table-1

Notes:

nd

non-detect

Statistical analysis

Operational taxonomic units (OTUs) were defined as nifH sequence groups in which sequences differed by ≤5% and ≤15% over the aligned 360-bp region (Geneious alignment) using the MOTHUR 10.6 program (Schloss et al., 2009; Table S1). We used MOTHUR 10.6 to calculate the non-parametric estimate of the Shannon diversity index, which is used when the number of species and the species abundances are unknown and there are a significant number of undetected species (Chao & Shen, 2003), and the Simpson’s evenness calculation, which is also less impacted by sample size, because it is weighted toward the most abundant OTUs (Hughes & Bohannan, 2004). MOTHUR was also used to determine the Morisita–Horn index, which provides a (dis)similarity measurement based on presence–absence data only (Magurran, 1988) and is not influenced by sample size or richness (Wolda, 1981). Rarefaction analysis was also performed in MOTHUR. All tests were performed for OTU groups at both the 5% and 15% cutoff levels. All figures and tables in the main text include both the 5% and the 15% OTU cutoff values. nifH transcript abundance data were log-transformed for normalization and differences among sites were determined by one-way ANOVA and paired Student’s t-test.

Table 2:

Diversity and evenness indices for each site.

	Metoxit Point	South Basin	Sage Lot Pond
5% cutoff
Non-parametric Shannon diversity	2.75	3.65	2.94
Simpson evenness	0.76	0.63	0.62
Morisita–Horn index	1	1	0.99
15% cutoff
Non-parametric Shannon diversity	2.6	3.32	2.76
Simpson evenness	0.7	0.66	0.74
Morisita–Horn index	0.75	0.8	0.7

DOI: 10.7717/peerj.1615/table-2

Results

Of the 18 cores analyzed, nifH expression was detected and ultimately sequenced from 14 cores (3, 8, 3 from MP, SB, and SLP, respectively), revealing diverse nifH expression in Waquoit Bay sediments. In total, we successfully sequenced 307 cDNA clones. Of these, 289 were unique sequences, which phylogenetically grouped into three of the major clusters (Cluster I, II, and III) previously used to characterize nifH phylogeny (Chien & Zinder, 1994; Fig. 2). The 5% sequence variation was selected for the operational taxonomic unit (OTU) cut-off to analyze phylotype richness and diversity of nifH bacteria found at each site. 85 OTUs were identified at the 5% cutoff, with most expressed sequences grouping together by date and site location (Table S1). South Basin showed the greatest diversity (non-parametric Shannon diversity was 3.65) of the three sites, and it also had the greatest number of unique sequences (133 vs. 55 at MP and 101 at SLP) (Table 2). The active nifH community at MP was the least diverse (2.75). Rarefaction analysis showed the greatest richness to be at SB, with MP and SLP less rich (Fig. 3). A significant difference was detected between SB (the most diverse and rich) and MP (the least diverse and rich site with the fewest unique sequences) at both the 5% and 15% cutoffs (ANOVA, p = 0.01 and p = 0.02 respectively). The Morisita–Horn index, indicating dissimilarity between communities, shows very different communities at the 5% cutoff between all three sites, as there was only a single shared OTU (Table 2). However, at the 15% cutoff, greatest dissimilarity is between MP and SLP (Morisita–Horn value of 0.80), while the greatest similarity (lowest Morisita–Horn value) is between SB and SLP (0.70) (Table 2).

Expressed nifH sequences at each site were distributed throughout our phylogenetic tree (Fig. 2). MP nifH sequences were primarily contained in Cluster I. The largest OTU in MP (7) contained sequences closely related (87–94% similar) to other uncultured nifH sequences from forest soil (Rosch & Bothe, 2009). The second and third largest OTUs (15 and 17) contained sequences most closely related (87–93% similar) to Acinetobacter sp. Z21 (EU693341), Agrobacterium tumefaciens (FJ822995) nifH, and Bradyrhizobium japonicum (GQ289582) (Fig. 4). We also observed the presence of a close relative (88–93% similar) of Thiocystis violascens, a purple sulfur bacteria that can grow with sulfide or sulfur as an electron donor under anoxic conditions (Lucas et al., 2012). MP nifH sequences were also >87% similar to sequences from uncultured bacteria found in a wide range of environments, including the Mediterranean Sea (Yogev et al., 2011), mangrove roots (Flores-Mireles, Winans & Holguin, 2007), and soil (Berthrong et al., 2014).

SB nifH sequences were most closely related to uncultured bacteria and Cluster II or Cluster III sequences. The largest SB OTU (3) encompassed sequences most closely related (82–95% similar) to sequences from uncultured bacteria from soil (Lopez-Lozano et al., 2012) and a Dutch intertidal microbial mat (Severin & Stal, 2010). The second largest OTU (4) contained sequences related (89–93% similar) to Desulfovibrio sp. and Desulfobulbus sp. nifH sequences (Fig. 4). One major OTU (9) did contain Cluster 1 sequences, most closely related (98–99% similar) to the nifH sequences from Bradyrhizobium denitrificans. Several of the small OTUs also contained Desulfo-related sequences.

Sequences from SLP were largely related to Cluster III sequences or uncultured bacteria. The two largest OTUs of all were comprised of SLP sequences most closely related (87–93% similar) to nifH sequences from uncultured bacteria from similar environments: Narragansett Bay (Fulweiler et al., 2013), Chesapeake Bay (GenBank Accession Number DQ098254), and a Dutch intertidal microbial mat (Severin & Stal, 2010). The third largest SLP OTU exclusively contained sequences related to nifHsequences (85–96% similar) from Desulfovibrio vulgaris (Fig. 4). Both SB and SLP also contained nifH transcripts similar to those of the sulfur-oxidizing Sulfiricurvum kujiense and sulfur-reducing Sulfurospirillum multivorans from Cluster II (92% similar).

Finally, we examined the relative percentage of uncultured as well as Cluster I, II, and III sequences for all sampling dates and stations as a function of depth (Fig. 5). We found the highest relative percentage of uncultured bacteria in the surface (0–1 cm) sediments and the highest percentage of Cluster III sequences deeper in the sediments (2–3 cm). For all sampling dates there is a trend of increasing relative abundance of Cluster III sequences accounting for more of the sequences with depth, from 1% at the surface to 25% at depth (Fig. 5).

Discussion

N-fixing microorganisms from Clusters I, II, and III were found in Waquoit Bay sediments, illustrating the diversity of taxa containing nifH sequences, although the gene itself has been well-conserved. The majority of the sequences were most closely related to uncultured environmental sequences. It is worth noting that Cluster II nitrogen fixers are rarely reported in environmental samples (Zehr et al., 2003), yet we found they accounted for ∼3% of the surface sediment sequences (Fig. 5). Organisms falling into this cluster have alternative nitrogenases and maybe associated with methanogenic archaea (Zehr et al., 2003; Dang et al., 2009). Their presence is interesting and suggests these types of organisms can play a key role in environmental nitrogen cycling.

At all stations, we detected the presence of bacteria involved in sulfur cycling. For example, sequences similar to Desulfovibrio sp., a Cluster III sulfate reducing bacteria (SRB), were observed at SB and SLP. And, at MP, we observed Thiocystis violascens, which can grow photolithoautrophically using sulfide, sulfur, and occasionally thiosulfate (Lucas et al., 2012). Over six decades ago, Sisler & ZoBell (1951) measured N₂-fixation via direct N₂ uptake in Desulfovibrio. Since then, SRB have been found fixing N in a variety of environments, including in the above- and below-ground biomass of salt marshes (Lovell & Davis, 2012), seagrass beds (Welsh et al., 1996), microbial mats (Steppe & Paerl, 2002), sediments associated with ghost shrimp Neotrypaea califoriensis (Bertics et al., 2010), and temperate estuarine sediments (Fulweiler et al., 2013). Desulfovibrio-related phylotypes were also found associated macroalgae covered water columns (Zhang et al., 2015). Heterotrophic N₂-fixation has been observed in the anoxic, ammonium rich waters of the Baltic (Farnelid et al., 2013). Our findings in this study are similar to this recent work. Specifically, at MP, the site with the highest in situ water column ammonium concentrations (Table 1) and significant macroalgae blooms, nifH expression was detected at all depths. Additionally, this site had an overall mean net N₂ uptake −34 ± 16 µmol N₂-Nm⁻²y⁻¹ (n = 4), indicating net N₂-fixation (Foster & Fulweiler, 2014). This was the only site to exhibit overall mean net N₂-fixation. This site is closest to a recent study in Waquoit Bay that also observed N₂-fixation in sandy, organic rich sediments in along the north shore of the main basin (Rao & Charette, 2012). The majority of uncultured matches were from soil clones (Rosch & Bothe, 2009), while many of the Cluster I cultured matches were from organisms with the genetic capability to both fix N and denitrify (e.g., Pseudomonas stutzeri and Bradyrhizobium sp.). The active nifH community at this site was the least diverse (non-parametric Shannon diversity = 2.75) and rich. We propose that the organic-rich environment is promoting a small number of dominant, actively N-fixing organisms. Although some studies have reported higher nifH gene diversity in highly N-impacted, organic rich sites (e.g., Chesapeake Bay: Moisander et al., 2007), a study of Danish estuaries with high nutrient loading showed greater nifH diversity at the DNA-level, but generally lower diversity in expressed nifH transcripts (Bentzon-Tilia et al., 2015b). Thus, nitrogen fixers have a diverse approach for dealing with environmental conditions (Bentzon-Tilia et al., 2015a).

These findings fit well with a recent turning of the tide in our understanding of estuarine nitrogen fixation. Historically, low rates of estuarine N₂-fixation have been attributed to a variety of biogeochemical and physical mechanisms (Howarth, Marino & Cole, 1988a), as well as to the presence of inorganic nitrogen. Specifically, estuarine sediments are typically replete with biologically useable nitrogen such as ammonium, which is known to shut-off N₂-fixation in cyanobacterial diazotrophs (Howarth, Marino & Cole, 1988a). However, a recent review highlighted that diazotroph sensitivity to inorganic nitrogen appears to vary with both the diazotroph type and the geographic location of the N₂-fixation (Knapp, 2012). In fact, Knapp (2012) reports significant N₂-fixation at ammonium concentrations as high as 200 µM, and it appears that benthic N-fixers may be much less sensitive to inorganic nitrogen exposure. Although Andersson et al. (2014) report an overall negative impact of dissolved inorganic nitrogen on nitrogenase activity, they did observe substantial rates of nitrogenase activity at ammonium concentrations up to 350 µmol L⁻¹. In recent culture experiments of heterotrophic water column nitrogen fixers, Bentzon-Tilia et al., 2015a; Bentzon-Tilia et al., 2015b found that bacteria isolates had different responses to environmental characteristics. In particular, they report increased ammonium concentrations stimulate N₂-fixation of one isolate (Rhodopseudomonas palustris, strain BAL398). They propose that this organism is using nitrogenase as an electron sink. Thus, other heterotrophic sediment bacteria may be employing N₂-fixation for a similar reason.

Overall, our findings at the heavily N impacted site, MP, support the idea that at least some benthic, heterotrophic N fixers are less sensitive to dissolved inorganic nitrogen exposure. Specifically, we observed net N₂-fixation at this site in addition to the highest in situ water column ammonium concentrations (98.91 µmol L⁻¹) (Table 1). Other work has also reported similar findings. For example, research in the Chesapeake Bay found the highest nifH richness at the site most heavily impacted by N (Moisander et al., 2007). And high rates of directly measured N₂-fixation have been reported for the nearby Providence River Estuary, another highly N impacted estuarine site (Fulweiler et al., 2007). We hypothesize that in the case of MP, sediment N₂-fixation is occurring at a fast enough and high enough rate to be adding new nitrogen to the system. This is supported by recent experimental work in Waquoit Bay at this same site where ³⁰N₂ was fixed and later released as ¹⁵N-labeled ammonium (SE Newell et al., 2016, unpublished data). If these findings were confirmed in other systems, it would suggest that subtidal heterotrophic N₂-fixation should be considered more often in coastal sediment N budgets. Additionally, it highlights the complex dynamics controlling sediment N₂-fixation and, along with the Knapp (2012) review calls into question the idea that ammonium alone controls rates of sediment N₂-fixation.

The other two, less N impacted sites (SB and SLP) also exhibited nifHexpression, including Cluster III SRB. In contrast to MP, the mean sediment net N₂ fluxes at SB and SLP in 2011 were positive, indicating net denitrification. SLP exhibited the highest net denitrification rate of 191 µmol N₂–Nm⁻²h⁻¹, followed by 75 µmol N₂-Nm⁻²h⁻¹ at SB. On one occasion (August 2011), a core at SB exhibited net N₂-fixation (−51 µmol N₂-Nm⁻²h⁻¹). Thus, even though nifH was active and, as shown in August for SB N₂-fixation was likely occurring, denitrification dominated. This signal could be driven by the spatial distribution of microbes in the sediment. For example, if denitrification occurred in the sediment surface, closer to the sediment-water interface (Froelich et al., 1979) and the SRB N₂-fixation below, our gas fluxes might be capturing more of the surface net denitrification signal. In fact, co-occurring spatial and temporal nifH and nirS (a gene encoding for denitrification) expression along with directly measured rates of denitrification was recently reported in sediments of a nearby estuary (Narragansett Bay, RI, USA; Fulweiler et al., 2013). And Brown & Jenkins (2014) observed concurrent nifH and nirS expression along a high to low N loading gradient in Narragansett Bay. Together these findings suggest that these two opposing processes, N₂-fixation and denitrification, may routinely co-occur in estuarine sediments. Fulweiler et al. (2013) proposed that while these processes may co-occur they may not be coupled but rather competitive, with N-fixers being able to outcompete denitrifiers for poor quality carbon sources. We also detected sequences similar to those from bacteria that have the capacity to both fix N and denitrify, such as Pseudomonas stutzeri and Bradyrhizobium denitrificans. These results are similar to a recent study of estuarine water column N₂-fixation where Bradyrhizobium, Pseudomonas, and Desulfovibrio were present (Bentzon-Tilia et al., 2015b). Together, these studies suggest that we need to reframe the way we think about estuarine N₂-fixation as the presence and activity of these organisms are found throughout these systems and likely play a key role in estuarine N budgets. Learning how diazotrophs and denitrifiers interact is a key next step toward understanding N limitation in coastal systems.

Of course a key question still remains: since N₂-fixation is considered to be a metabolically costly process, requiring 16 ATP per N₂ reduced, why fix nitrogen in an N-replete environment like an estuary? First, it is possible that nitrogenase may act as an electron sink for certain heterotrophic bacteria. In such a case they are not fixing N to obtain nitrogen but are using it as mechanism to cope with environmental conditions (McKinlay & Harwood, 2010; Bentzon-Tilia et al., 2015a). Additionally, we propose that in many ways sediments may provide an ideal environment for heterotrophic N₂-fixation. One of the primary reasons why N₂-fixation is so energetically expensive is the cost of keeping the highly sensitive nitrogenase enzyme away from oxygen. But most estuarine sediments are anoxic within the top few millimeters thus relieving this constraint and decreasing the overall energy requirements (Paerl & Prufert, 1987; Beman, Popp & Alford, 2012; Bonaglia et al., 2014). Additionally, the variety of marine environments that have shown linkages between N₂-fixation and sulfur and sulfate reducers suggest that conditions that promote sulfur cycling might also promote N₂-fixation (Sisler & ZoBell, 1951; Welsh et al., 1996; Fulweiler et al., 2013). Estuarine sediments are key environments for sulfur and sulfate reduction as they are anoxic, rich in organic matter, and have an abundant supply of sulfate ions from the surrounding seawater. Finally, trace metals critical to N₂-fixation, such as iron and molybdenum, are abundant in estuarine sediments and thus likely do not limit N₂-fixation as they may in estuarine water columns (Howarth, Marino & Cole, 1988a).

Conclusions

We measured nifH expression in the sediments at every site sampled, even when net N₂ fluxes indicated that denitrification was the dominant N₂ pathway. These findings add to a growing body of evidence that suggest these opposing processes, N₂-fixation and denitrification, may routinely co-occur in estuarine sediments. We observed the presence of sulfate reducing bacteria (SRB), as well as bacteria that have the capacity to both fix N and denitrify, such as Pseudomonas stutzeri and Bradyrhizobium denitrificans. The observations of nifH expression occurring with net denitrification, combined with presence of organisms that can both denitrify and fix nitrogen, suggest that the balance between those two processes may occur on the individual cellular level. The research presented here highlights a new perspective emerging from estuarine sediment N₂-fixation studies in which the nitrogen, sulfur, and carbon cycles in coastal marine ecosystems are intimately connected. Additionally, it highlights that heterotrophic N₂-fixation is an important and perhaps too often overlooked source of reactive N in coastal ecosystems. Future studies should also include measuring water column heterotrophic N₂-fixation as recent studies have shown its importance and thus it too may play a critical role in estuarine N budgets.

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