Introduction
Only 6 % of nitrogen (N) in seawater is bioavailable
(Gruber, 2008). This bioavailable N is mainly present in
the form of nitrate (NO3-), whereas the large pool of atmospheric
dinitrogen gas (N2) is only available for N2 fixing
microorganisms (diazotrophs). N often limits marine productivity
(Ward and Bronk, 2001;
Gruber, 2008) and the largest source of bioavailable N (i.e., ammonium;
NH4+) in the marine environment is N2 fixation
(Falkowski et al.,
1998; Strous et al., 1999; Brandes and Devol, 2002).
To date, the quantitative contribution of diazotrophs in the marine N cycle
remains unclear and numerous estimates of global sources and sinks of global
N have led to an unbalanced budget with deficits of around 200 Tg N yr-1 (Codispoti, 2007). This
suggests that either previous N2 fixation rate determinations have
been underestimated (Großkopf et al.,
2012) or that N loss processes are overestimated (Codispoti, 2007). However,
also balanced budgets such as ∼265 Tg N yr-1 for N sources and
∼275 Tg N yr-1 for N sinks exist (Gruber, 2004).
These budget discrepancies illustrate that the current knowledge on
diazotrophy and the marine N cycle is still limited.
Recent investigations argue that N2 fixation in the water column cannot
be totally attributed to phototrophic cyanobacteria, but that also
heterotrophic prokaryotes contribute substantially
(Riemann
et al., 2010; Farnelid et al., 2011; Dekaezemacker et
al., 2013;
Löscher et al., 2014; Fernandez et al., 2015). This was shown for the
Peruvian oxygen minimum zone (OMZ), where proteobacterial clades dominated
with heterotrophic diazotrophs, indicating that cyanobacterial diazotrophs
are of minor importance in this area
(Löscher et al., 2014).
Cross-section of dissolved O2 concentrations (µM) along
the continental margin of the Peruvian OMZ at 12∘ S. The vertical
lines represent CTD cast for O2 measurement during the cruise M92.
Stations 1 to 10 for multicorer (MUC) sampling are indicated by station
numbers according to Dale et al. (2015).
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Pelagic N2 fixation has been studied mostly in the oligotrophic surface
oceans, but it was not until the past decade that benthic habitats began to
receive more attention (Fulweiler et al., 2007; Bertics et al., 2010, 2013). Most studies on benthic N2 fixation focused on
coastal environments (Capone et al., 2008 and references
therein). For example, subtidal sediments in Narragansett Bay (Rhode Island)
were found to switch from being a net sink in the form of denitrification to
being a net source of bioavailable N by N2 fixation, caused by a
decrease of organic matter deposition to the sediments
(Fulweiler et al., 2007). Shallow brackish-water
sediments off the Swedish coast revealed benthic N2 fixation along with
a diverse diazotrophic community (Andersson et al., 2014).
N2 fixation was positively influenced by a variety of environmental
factors, such as salinity and dissolved inorganic Nitrogen, while wave
exposure had a negative influence. Recent work revealed that benthic N2
fixation is often linked to sulfate-reducing bacteria. For instance,
bioturbated coastal sediments showed enhanced N2 fixation activity
mediated by sulfate-reducing bacteria, adding new dissolved inorganic N to
the system (Bertics et al.,
2010; Bertics and Ziebis, 2010). Further coupling of N2 fixation to SR
was observed in organic-rich sediments of the seasonal hypoxic
Eckernförde Bay (Baltic Sea, Bertics et al.,
2013), as well as in the sub-tidal, heterotrophic sediments of Narragansett
Bay (Rhode Island, USA; Fulweiler et al.,
2013). Several sulfate-reducing bacteria carry the functional gene marker
for N2 fixation, the nifH gene
(Sisler and ZoBell, 1951;
Riederer-Henderson and Wilson,
1970; Zehr and Turner,
2001) and were shown to actively fix N2 in culture experiments
(Riederer-Henderson and Wilson, 1970). However, information
on sulfate-reducing bacteria and their contribution to N2 fixation in
the environment is still sparse and restricted to a small selection of
environments.
So far, the distribution of benthic N2 fixation and its relevance for N
cycling in the Peruvian oxygen minimum zone (OMZ), defined by dissolved
oxygen < 20 µmol kg-1
(Fuenzalida et al.,
2009), are unknown. The shelf and the upper slope in the Peruvian OMZ
represent recycling sites of dissolved inorganic N with dissimilatory
NO3- reduction to NH4+ being the dominant process
(∼ 15 mmol N m-2 d-1) in the benthic N cycle
(Dale et al., 2016). This
process is mediated by the filamentous sulfide-oxidizing Thioploca bacteria
(Schulz, 1999; Schulz and Jørgensen,
2001). Benthic denitrification, which is mediated by foraminifera at water
depth between 80 and 250 m of the Peruvian OMZ, represent a sink for
bioavailable N in sediments, accounting for a potential NO3- flux,
i.e., N loss, of 0.01 to 1.5 mmol N m-2 d-1
(Glock et al., 2013; Dale
et al., 2016).
The high input of labile organic carbon to Peruvian OMZ sediments
(Dale et al., 2015) and subsequent SR should
favor benthic N2 fixation. Sulfate-reducing bacteria could considerably
contribute to N2 fixation in these organic-rich OMZ sediments, given
that several sulfate-reducing bacteria (e.g., Desulfovibrio spp.;
Riederer-Henderson and Wilson, 1970; Muyzer
and Stams, 2008) carry the genetic ability to fix N2, and provide an
important bioavailable N source for non-diazotrophic organisms
(Bertics
et al., 2010; Sohm et al., 2011; Fulweiler et al., 2013). We therefore
hypothesize a possible coupling of N2 fixation and SR in sediments off
Peru. The aim of the present study was to identify and quantify benthic
N2 fixation along a depth transect through the Peruvian OMZ, together
with SR, and compare its distribution with environmental factors, such as
organic matter, to study its control mechanisms. The identification of
bacteria carrying the genetic ability to perform N2 fixation should
further deliver information about benthic diazotrophic community structures
at the different stations. The overall knowledge gained is needed
to better constrain benthic N cycling in OMZs and to improve our knowledge
on sources and sinks of fixed N.
Sampling deployments, including station number according to Dale et
al. (2015), core ID, sampling date and coordinates. Water depth (m) recorded
by the ship's winch and bottom water temperature (∘C) and bottom water O2 concentration (µM; bdl = below detection
limit: 5 µM) measured on the CTD.
Station
Core ID
Date (2013)
Latitude (S)
Longitude (W)
Depth (m)
Temp. (∘C)
O2 (µM)
1
MUC 13
January 11
12∘13.492′
77∘10.511′
70
14
bdl
4
MUC 11
January 9
12∘18.704′
77∘17.790′
144
13.4
bdl
6
MUC 6
January 7
12∘23.322′
77∘24.181′
253
12
bdl
8
MUC 23
January 15
12∘27.198′
77∘29.497′
407
10.6
bdl
9
MUC 17
January 13
12∘31.374′
77∘35.183′
770
5.5
19
10
MUC 28
January 19
12∘35.377′
77∘40.975′
1025
4.4
53
Materials and methods
Study area
The most extensive OMZ worldwide is found in the eastern tropical south
Pacific Ocean at the central Peruvian coast
(Kamykowski and Zentara,
1990). The Peruvian OMZ ranges between 50 and 700 m water depth with
oxygen (O2) concentrations below the detection limit in the mid-waters
(Stramma et al., 2008). The mean water
depth of the upper OMZ boundary deepens during intense El Niño Southern
Oscillation years and can reach a depth of 200 m
(Levin et al.,
2002) with oxygenation episodes reaching concentrations of up to 100 µM O2 (Gutiérrez et
al., 2008). O2 concentrations (Fig. 1, Table 1) off Peru are modulated
by coastal trapped waves
(Gutiérrez et al., 2008),
trade winds (Deutsch et al., 2014) and
subtropical–tropical cells (Duteil et al., 2014),
and can vary on monthly to interannual timescales
(Gutiérrez et al., 2008).
Biogeochemical porewater profiles in MUC cores from sampling
stations along the 12∘ S depth transect. Graphs show
NH4+ (µM), SO42- (mM), sulfide (µM),
organic carbon content (Corg, wt %) and the C / N ratio (molar). Water
depths and bottom water O2 concentrations (BW O2, µM) are
detailed on the right.
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At 12∘ S, the OMZ extends from water depths between 50 and 550 m
(Dale et al., 2015; Fig. 1). During our field work, bottom water O2
concentrations varied greatly with water depth and were below the detection
limit (5 µM) at stations from 70 to 407 m water depth. Bottom water
O2 increased to 19 µM at 770 m water depth and 53 µM at
1025 m water depth, indicating the increase of dissolved O2 below the
lower boundary of the OMZ (Dale et al., 2015). Between 70 and 300 m water
depth, the sediment surface was colonized by dense filamentous mats of
sulfur-oxidizing bacteria, presumably of the genera Marithioploca spp. These bacteria are able
to glide up to 1 cm h-1 through the sediment in order to access
hydrogen sulfide
(Fossing et al., 1995;
Jørgensen and Gallardo, 1999; Schulz, 1999). Sediments at the lower
boundary (770 and 1025 m) of the OMZ host a variety of macrofaunal
organisms,
e.g., ophiuroids, gastropods, and crustaceans (Mosch et al.,
2012).
The 12∘ S region is in the center of an extensive upwelling zone
and features high primary productivity
(Pennington et al., 2006). Sediments at
12∘ S have higher rates of particulate organic carbon accumulation
(2–5 times) compared to other continental margins and a high carbon burial
efficiency, indicating preferential preservation of organic matter in the
Peruvian OMZ (Dale et al., 2015). The shelf (74 m) of the Peruvian OMZ is
characterized by high sedimentation rates of 0.45 cm yr-1, while
mid-waters and below the OMZ show rates between 0.07 and
0.011 cm yr-1.
Sampling
Sediment samples were taken in January 2013 at six stations (70, 144, 253,
407, 770, and 1025 m) along a depth transect at 12∘ S in the OMZ
off Peru (Fig. 1) during an expedition on RV Meteor (M92). January
represents austral summer, i.e., the low upwelling, high productivity season
in this area (Kessler, 2006). Samples were
retrieved using a TV-guided multiple corer (MUC) equipped with seven core
liners. The core liners had a length of 60 cm and an inner diameter of 10 cm. Location, water depth, temperature, and O2 concentration (from Dale
et al., 2015) at the six sampling stations are listed in Table 1. Retrieved
cores for microbial rate measurements were immediately transferred to cold
rooms (4–9 ∘C) for further processing.
Geochemical analyses
Porewater analysis and the determination of sediment properties and
geochemical data have been previously described in detail by Dale et al. (2015). In short, the first core was subsampled under anoxic conditions
using an argon-filled glove bag, to preserve redox sensitive constituents.
NH4+ and sulfide concentrations were analyzed on a Hitachi U2800
UV/VIS spectrophotometer using standard photometric procedures
(Grasshoff et al., 1999), while sulfate (SO42-)
concentrations were determined by ion chromatography (Methrom 761).
The second replicate core was sampled to determine porosity by the weight
difference of the fresh sediment subsamples before and after freeze-drying.
Particulate organic carbon and particulate organic nitrogen contents were
analyzed using a Carlo-Erba element analyzer (NA 1500).
Benthic nitrogen fixation
At each of the six stations, one MUC core was sliced in a refrigerated
container (9 ∘C) in 1 cm intervals from 0 to 6 cm, in 2 cm
intervals from 6 to 10 cm, and in 5 cm intervals from 10 to 20 cm. The
acetylene reduction assay (Capone, 1993; Bertics et
al., 2013) was applied to quantify nitrogenase activity. This application is
based on the reduction of acetylene (C2H2) to ethylene
(C2H4) by the nitrogenase enzyme
(Dilworth, 1966;
Stewart et al., 1967; Capone, 1993). To convert from nitrogenase activity to
N2 fixation, a conversion factor of 3 C2H4 : 1 N2 was
applied
(Patriquin
and Knowles, 1972; Donohue et al., 1991; Orcutt et al., 2001; Capone et
al., 2005), which was previously used to measure N2 fixation in
sediments (Welsh et al.,
1996;
Bertics et al., 2013).
Serum vials (60 mL) were flushed with N2 and filled with 10 cm3
sediment from each sampling depth (triplicates). The samples were flushed
again with N2, crimp sealed with butyl stoppers and injected with 5 mL
of C2H2 to saturate the nitrogenase enzyme. Serum vials were
stored in the dark at 9 ∘C, which reflected the average in situ
temperature along the transect (compare with Table 1). Two sets of
triplicate controls (10 cm3) were processed for every station. Sediment
was collected from each core liner from 0 to 5, 5 to 10, and from 10 to 20 cm and placed in 60 mL serum vials. One set of controls was used to
identify natural C2H4 production without the injection of
acetylene, and the second control set was fixed with 1 mL 37.5 %
formaldehyde solution.
The increase of C2H4 in each sediment slice was measured onboard
over 1 week (in total five time points, including time zero) using gas
chromatography (Hewlett Packard 6890 Series II). From each serum vial, a 100 µL headspace sample was injected into the gas chromatograph and the
results were analyzed with the HP ChemStation gas chromatograph software.
The gas chromatograph was equipped with a packed column (Haye SepT, 6 ft,
3.1 mm ID, Resteck) and a flame ionization detector. The carrier gas was
helium and the combustion gases were synthetic air (20 % O2 in
N2) and hydrogen. The column had a temperature of 75 ∘C and
the detector temperature was 160 ∘C.
Standard deviation of individual N2 fixation rates was calculated from
three replicates determined per sediment depth in one multicorer. Standard
deviation of depth-integrated N2 fixation was calculated from the three
replicate integrated rates.
It should be mentioned that the incubation with C2H2 can
potentially lead to a lack of fixed N caused by the saturation of the
nitrogenase enzyme, which leads to a reduction of cell viability and
consequently N2 fixation (Seitzinger and Garber, 1987).
These effects are expected to cause an underestimation of N2 fixation
rates. However, the acetylene reduction method is to the best of our
knowledge still the standard method for the determination of benthic N2
fixation (Bertics et al., 2013). The δ15N rate determinations are not feasible in sediments, as they would
require incubation times of several weeks to months to achieve signals that
are statistically above the natural δ15N abundance of
sediments.
We are further aware that our samples might have experienced a potential
microbial community shift during the N2 fixation determination, which
was shown to be driven by the addition of C2H2
(Fulweiler et al., 2015). Again, a community
shift would be expected to cause rather an underestimation of absolute
N2 fixation rates.
Sulfate reduction rates
One MUC core per station was used for determination of SR activity (same MUC
cast as for N2 fixation, but different core). First, two replicate push
cores (length 30 cm, inner diameter 2.6 cm) were subsampled from one MUC
core. The actual push core length varied from 21 to 25 cm total length. Then,
6 µL of the carrier-free 35SO42- radio tracer
(dissolved in water, 150 kBq, specific activity 37 TBq mmol-1) was
injected into the replicate push cores in 1 cm depth intervals according to
the whole-core injection method (Jørgensen, 1978). The
push cores were incubated for ∼ 12 h at 9 ∘C. After
incubation, bacterial activity was stopped by slicing the push core into
1 cm intervals and transferring each sediment layer into 50 mL plastic
centrifuge tubes filled with 20 mL zinc acetate (20 % w/w). Controls were
done in triplicates from different depths and first fixed with zinc acetate
before adding the tracer. Rates for SR were determined using the cold
chromium distillation procedure according to
Kallmeyer et al. (2004).
It should be mentioned that the yielded SR rates have to be treated with
caution due to long (up to 3 half-life times of 35S) and unfrozen
storage. Storage of SR samples without freezing has recently been shown to
result in the re-oxidation of 35S-sulfides
(Røy et al., 2014). In this reaction, FeS is
converted to ZnS. The released Fe2+ reacts with O2 and forms
reactive Fe(III). The Fe(III) oxidizes ZnS and FeS, which are the major
components of the total reduced inorganic sulfur species, resulting in the
generation of SO42- and hence an underestimation of SR rates.
However, because all SR samples in the present study were treated the same
way, we trust the relative distribution of activity along sediment depth
profiles and recognize potential underestimation of absolute rates.
<italic>nifH</italic> gene analysis
Core samples for DNA analysis were retrieved from the six stations and were
sliced in the same sampling scheme as described for benthic N2
fixation. Approximately 5 mL sediment from each depth horizon was
transferred to plastic whirl-paks® (Nasco, Fort Atkinson, USA),
frozen at -20 ∘C and transported back to the home laboratory. To
check for the presence of the nifH gene, DNA was extracted using the
FastDNA® SPIN Kit for Soil (MP Biomedicals, CA, USA) following
the manufacturer's instructions with a small modification. Sample
homogenization was done in a Mini-Beadbeater™ (Biospec Products,
Bartlesville, USA) for 15 s. PCR amplification, including primers and
PCR conditions, was done as described by Zehr
et al. (1998), using the GoTaq kit (Promega, Fitchburg, USA) and
additionally 1 µL bovine serum albumin (20 mg mL-1; Fermentas).
The TopoTA Cloning® Kit (Invitrogen, Carlsbad, USA) was used
for cloning of PCR amplicons, according to the manufacturer's protocol.
Sanger sequencing (122 nifH sequences) was performed by the Institute of
Clinical Molecular Biology, Kiel, Germany. For the sampling sites 70, 144, 253, 407, 770, and 1025 m water depth the number of obtained
sequences was 22, 24, 24, 13, 18, and 21, respectively. Negative controls
were performed using the PCR mixture as described without template DNA; no
amplification was detected. Sequences were ClustalW aligned in MEGA 6.0
(Tamura et al., 2007), and a maximum
likelihood tree was constructed on a 321 base pair fragment and visualized
in iTOL (Letunic and Bork, 2007, 2011).
Reference sequences were obtained using BlastX on the NCBI database.
Sequences were submitted to Genbank (Accession numbers: KU302519 –
KU302594).
Statistical analysis
A principle component analysis (PCA) was applied to microbial rates and
environmental parameters to determine most likely explanatory variables for
active N2 fixation at the sampling St. 1 to 9. The deepest St. 10 was
excluded from the analysis because at this site SR rates were below the
detection limit and the PCA only allows complete datasets, which otherwise
would have resulted in the exclusion of all SR rates. Prior to PCA, the
dataset was Hellinger transformed in order to make it compatible with PCA.
The PCA was performed in R v3.0.2 by using the R package “Vegan”
(Oksanen et al., 2013) according to the approach described in
Löscher et al. (2014).
For the depth profiles of N2 fixation rates (mmol m-2 d-1)
the variables water depth (m), sediment depth (cm), sulfate reduction (mmol m-2 d-1), organic carbon content (wt %), C / N ratio (molar),
ammonium (µM), and sulfide (µM) were tested. A PCA of
integrated (0–20 cm) N2 fixation rates (mmol m-2 d-1) and
environmental parameters could not be done due to the lack of sufficient
data points.
Finally, two biplots for the depth profiles were produced, which allowed
having two different views from two different angles, i.e., one biplot for
principle component 1 and 2, and one biplot for principle component 2 and 3.
These biplots graphically reveal a potential negative, positive or zero
correlation between N2 fixation and the tested variables.
Results
Sediment properties
Although sediments were sampled down to the bottom of the core, the focus
here is on the 0–20 cm depth interval where benthic N2 fixation was
investigated.
Sediments at the shelf station (St.) 1 (70 m) were black between 0 and 1 cm
and then olive green until 20 cm. Only a few metazoans (polychaetes) were
observed in the surface sediment. The sediment surface was colonized by
dense filamentous mats of sulfur-oxidizing Marithioploca spp. These bacteria extended down to
a sediment depth of 36 cm. The sediment on the outer shelf St. 4 (144 m) was
dark olive green from 0 to 13 cm and dark grey until 20 cm. At St. 6 (253 m), which was located within the core of the OMZ, the sediment appeared dark
olive green between 0 and 17 cm and olive green with white patches between 17 and 20 cm. At this station, Marithioploca spp. was abundant.
Uniquely, surface sediments (0–3 cm) at St. 8 (407 m), consisted of a fluffy, dark olive-green layer mixed
with white foraminiferal ooze. This layer also contained centimeter-sized
phosphorite nodules with several perforations (ca. 1–3 mm in diameter).
Below 2 cm, the sediment consisted of a dark olive green, sticky clay layer.
No Marithioploca mats were found here. St. 9 (770 m) was below the OMZ, and sediments
were brown to dark olive green with white particles between 0 and 12 cm, and
brown to olive green without white particles below this depth. Organisms
such as anemones, copepods, shrimps and various mussels were visible with
the TV-guided MUC and in the sediment cores. The deepest St. (10; 1025 m)
had dark olive green sediment from 0 to 20 cm and black patches from 17 to 20 cm. The sediment was slightly sandy and was colonized with polychaete
tubes at the surface and organisms that were also present at St. 9. For
further sediment core descriptions see also Dale et al. (2015).
Sediment profiles of N2 fixation (nmol N2 cm-3 d-1, average of three replicates) and sulfate reduction rates (SR,
nmol SO42- cm-3 d-1, two replicates; R1 and R2) from 0 to 20 cm at the six stations. The upper x axis represents the N2 fixation,
while the lower x axis represents the SR. Error bars indicate standard
deviation of N2 fixation.
![]()
Integrated nitrogen fixation (mmol N2 m-2 d-1, grey bars,
average of three replicates) and integrated sulfate reduction (mmol
SO42- m-2 d-1, green bars, two replicates) from 0 to 20 cm, including dissolved inorganic carbon flux (DIC, mmol m-2 d-1,
red curve from Dale et al., 2015) and bottom water O2 (µM,
blue curve) along the depth transect (m). Error bars indicate standard
deviation of N2 fixation.
![]()
Geochemical porewater profiles of NH4+, SO42-, sulfide,
organic carbon content, and organic C / N ratio between 0 and 20 cm at the six
stations are shown in Fig. 2. In all cores, NH4+ concentrations
increased with sediment depth. The highest NH4+ concentration was
reached at St. 1 (70 m), increasing from 316 µM in the upper cm to
2022 µM at 20 cm. St. 4 and 6 showed intermediate NH4+
concentrations between 300 and 800 µM at 20 cm,
respectively. At St. 8 (407 m) the NH4+ concentration increased
from 0.7 µM at the surface to 107 µM at 20 cm. The two deep
stations (St. 9 and 10) had the lowest NH4+ concentrations with 33 and 22 µM at 20 m sediment depth, respectively.
The SO42- concentrations remained relatively constant in the
surface sediments along the transect. A decrease was only observed at St. 1;
from 28.7 µM in the surface layer to 19.4 µM at 20 cm. In
parallel with the decrease in SO42-, only St. 1 revealed
considerable porewater sulfide accumulation, whereby sulfide increased from
280 µM at the surface sediment to 1229 µM at 20 cm.
Integrated nitrogen fixation (mmol N2 m-2 d-1, grey
bars, average of three replicates), average organic carbon content
(Corg, wt %, orange curve) and the average C / N molar ratio (yellow
curve) from 0 to 20 cm along the depth transect (m). Error bars indicate
standard deviation.
![]()
Organic carbon content decreased with increasing sediment depth at St. 1 (70 m), 9 (770 m), and 10 (1025 m). The highest surface organic carbon content
(∼ 15 wt %) was found at St. 6, whereas the lowest
(∼ 2.6 wt %) was detected at the deep St. 10. The average (0–20 cm) organic carbon content (Fig. 5) increased from St. 1 to St. 6
(15 ± 1.7 wt %) and decreased from St. 6 to the lowest value at St. 10
(2.4 ± 0.4 wt %).
C / N ratios, as a proxy for the freshness of the organic matter, increased
with increasing sediment depth (Fig. 5). The lowest surface C / N ratio (6.2)
was measured at the shallow St. 1, while the highest surface C / N ratio (11)
was found at St. 10.
Benthic nitrogen fixation and sulfate reduction
For a straightforward comparison of SR rates with benthic N2 fixation
only the sediment depths between 0 and 20 cm are considered. Sediment depth
profiles are expressed as N2 fixation, that is, with the conversion
factor of 3 C2H4 : 1 N2.
Highest N2 fixation and SR rates were detected in the surface sediments
(0–5 cm) and both rates tended to decrease with increasing sediment depth
(Fig. 3). N2 fixation and SR rates were high at St. 1, 4, and 6 (70,
144, 253 m) and lowest at the deeper St. 8–10 (407, 770, 1025m).
At St. 1, N2 fixation and SR rates showed different trends in the top
layer of the cores, but depth profiles were more aligned below. Although St.
1 had the highest SR rates of all sites, reaching 248 nmol SO42- cm-3 d-1 at 0–1 cm, N2 fixation was not highest at this
station. At St. 4 (144 m), both N2 fixation and SR revealed peaks close
to the surface. N2 fixation decreased between 0 and 8 cm and increased
below 8 cm. This increase was not observed in SR rates, which were highest
at the surface (181 nmol SO42- cm-3 d-1) and decreased
towards the bottom of the core. St. 6 (253 m) had the highest N2
fixation of all stations, with rates of 4.0 ± 0.5 nmol N2 cm-3 d-1 in the surface centimeter. Yet, although N2 fixation
and SR had overlapping activity profiles, the highest SR rate of all
stations was not detected at St. 6. Very low N2 fixation rates were
measured at St. 8 (407 m; 0.5 ± 0.25 nmol N2 cm-3 d-1
in the surface), as well as very low SR rates (0–4.3 nmol SO42- cm-3 d-1). As mentioned, this station was unique due to the
presence of foraminiferal ooze, phosphorite nodules and a sticky clay layer
below 2 cm. N2 fixation and SR rates showed a peak at 5 and at 7 cm,
respectively. At St. 9 (770 m) N2 fixation was low in the surface and
at 20 cm sediment depth, with a peak in activity at 4–5 cm (0.8 ± 0.08 nmol N2 cm-3 d-1). At St. 10 (1025 m), N2 fixation
rates were low throughout the sediment core, not exceeding 0.16 ± 0.02 nmol N2 cm-3 d-1. This site had the lowest organic carbon
content throughout the core (between 2.6 wt % at the surface and 1.9 wt % at 20 cm), as well as low NH4+ concentrations. At St. 9
(below 9 cm depth) and St. 10 (entire core) SR rates were below detection,
which could point either to the absence of SR or to the complete loss of
total reduced inorganic sulfur due to the long, unfrozen storage (see
methods).
Integrated N2 fixation (0–20 cm) increased from St. 1 to St. 6, with
the highest rate (0.4 ± 0.06 N2 m-2 d-1) at St. 6 (253 m), and decreased from St. 6 (407 m) to St. 10 (1025 m; Fig. 4). Integrated
SR rates (0 to 20 cm) ranged from ∼ 4.6 mmol SO42- m-2 d-1 at St. 1 to below detection at St. 9 (Fig. 4).
Overall, integrated SR rates decreased with increasing water depth.
Integrated N2 fixation rates and SR were in general inversely
correlated between St. 1 and St. 6, and followed the organic carbon content
from St. 1 to St. 6 (70–253 m; Fig. 5). Both parameters had the highest
value at St. 6. This pattern did not hold for the relatively low integrated
SR rate at St. 6. The C / N ratio, averaged over 20 cm, increased with
increasing water depth (Fig. 5). Regarding the three deep stations, the
lowest integrated N2 fixation rate (0.008 ± 0.002 N2 m-2 d-1) was detected at St. 8 (407 m). Also the integrated SR
rate was low at this site (∼ 0.46 mmol SO42- m-2 d-1). At St. 9 and 10 (770 and 1025 m), integrated
N2 fixation was low at 0.05 ± 0.005 and 0.01 ± 0.001 N2 m-2 d-1, respectively, and integrated SR
rates were also lowest at St. 9 (770 m). From St. 8 to 10 a decrease of
integrated N2 fixation and SR together with the average organic carbon
content was detected.
No activity was detected in controls for N2 fixation and SR.
Statistical analysis
The PCA of N2 fixation depth profiles (Fig. 6a and b) showed a weak
positive correlation with sulfate reduction rates (Fig. 6a) and a strong
positive correlation between N2 fixation and the organic matter content
in sediments (Fig. 6b). A negative correlation between N2 fixation and
sediment depth (Fig. 6a), as well as between N2 fixation and sulfide
concentration for St. 1 (Fig. 6b) was found. Furthermore, a weak negative
correlation was detected between N2 fixation and the C / N ratio (Fig. 6a). No correlation was found between N2 fixation and ammonium
concentration and water depth (Fig. 6a and b).
Molecular analysis of the <italic>nifH</italic> gene
Sequences for the nifH gene analysis were pooled for each of the six stations,
making about 20 sequences per sample and 120 in total. NifH gene sequences were
detected at all six sampling sites and clustered with Cluster I
proteobacterial sequences and Cluster III sequences as defined by
Zehr and Turner (2001)
(Fig. 7). In Cluster I and Cluster III, three and seven novel clades were
detected, respectively. In general, most of the previously unidentified
clades belonged to uncultured bacteria. One distinct novel clade was found
for St. 1–6. No Cluster I cyanobacterial nifH sequences were detected and no
potential PCR contaminants were present
(Turk et al., 2011). Sequences
clustered with only one identified sulfate-reducing bacterium, Desulfonema limicola (Fukui et
al., 1999, OMZ 253). Other sequences from several stations (OMZ 70, 144,
253, 770) were distantly related to Desulfovibrio vulgaris
(Riederer-Henderson and Wilson, 1970; Muyzer
and Stams, 2008). One cluster (OMZ 144 m) was closely related to the
anaerobic marine bacterium Vibrio diazotrophicus (Guerinot
et al., 1982). Other organisms with which OMZ sequences clustered belonged
to the genera of fermenting bacteria, namely Clostridium beijerincki (Chen, 2005), and
to the genera of iron-reducing bacteria, namely Geobacter bemidjiensis (Nevin et
al., 2005). In addition, several sequences were phylogenetically related to
a gamma proteobacterium
(Zehr and Turner, 2001)
from the Pacific Ocean.
Principle component analysis (PCA) from two different angles of
Hellinger transformed data of N2 fixation and environmental parameters
along vertical profiles. Correlation biplots (a) of principle
components 1 and 2 and of (b) principle components 2 and 3 in a
multidimensional space are shown. Samples are displayed as dots while
variables are displayed as lines. Parameters pointing into the same
direction are positively related; parameters pointing in the opposite
direction are negatively related.
![]()
Phylogenetic tree of nifH genes based on the analysis of 122 sequences
(∼ 20 sequences per sample) from the six sampling stations
between 70 and 1025 m water depth. Novel detected clusters consisting of
several sequences from the same sampling depth are indicated by grey
triangles. Reference sequences consist of the alternative nitrogenase anfD,
anfG, anfK. Cluster III sequences as defined by Zehr and Turner (2001) are
highlighted in blue; Cluster I cyanobacterial sequences are highlighted in
green and Cluster I proteobacterial sequences are highlighted in orange. The
scale bar indicates the 10 % sequences divergence. Sequences marked with
an asterisk represent potential PCR contaminated products, with novel
clusters distant from those clusters. Sequences determined in this study are
termed OMZ plus the corresponding water depth.
![]()
Discussion
Coupling of benthic nitrogen fixation and sulfate reduction
Based on the high organic matter input to Peruvian sediments underneath the
OMZ we hypothesized a presence of N2 fixation and its coupling to
sulfate reduction (SR). We confirmed the presence of N2 fixation in
sediments at all sampled stations along the depth transect. N2 fixation
activity was often enhanced where SR peaked and sometimes both activity
depth profiles revealed similar trends. However, while peaks in SR were very
pronounced, maximum N2 fixation showed a much broader distribution over
depth. These findings are in line with the PCA of depth profiles, which
revealed a weak positive correlation between activities of N2 fixation
and sulfate reduction. But it should be kept in mind that the N2
fixation and SR were determined in replicate MUC cores, which were taken up
to 50 cm apart, depending on where the core liners were situated in the
multicorer. Nonetheless, it appears that the observed N2 fixation is
not exclusively fueled by SR activity.
The coupling between N2 fixation and SR has been previously suggested
for coastal sediments off California, where N2 fixation significantly
decreased when SR was inhibited
(Bertics and Ziebis, 2010). Different
studies confirmed that sulfate-reducing bacteria, such as Desulfovibrio vulgaris can supply
organic-rich marine sediments with bioavailable N through N2 fixation
(Welsh et al., 1996;
Nielsen et al., 2001; Steppe and
Paerl, 2002; Fulweiler et al., 2007;
Bertics et al., 2013;
Fulweiler et al., 2013). Fulweiler et al. (2013) conducted a study in sediments of the Narrangaset Bay and found
several nifH genes related to sulfate-reducing bacteria, such as Desulfovibrio spp.,
Desulfobacter spp. and Desulfonema spp., suggesting that sulfate-reducing bacteria were the dominant
diazotrophs.
The more surprising finding in this study is that integrated rates of
N2 fixation and SR showed opposite trends at the three shallowest
stations, pointing to potential environmental control mechanisms (see Sect. 5.2).
Overall, these findings indicate that N2 fixation might be partly
coupled to processes other than SR or that the two processes are controlled
by different parameters. The nifH gene sequence analyses indicated only a weak
potential of sulfate reducers to conduct N2 fixation in the Peruvian
sediments. Sequences clustered only with the sulfate-reducing bacteria
Desulfonema limicola (Fukui et al., 1999) exclusively at the 253 m Station. D. limicola is known from other
benthic environments through nifH gene analyses
(Mussmann
et al., 2005; Bertics et al., 2010, 2013). A distant relation to the
confirmed diazotrophic sulfate reducer Desulfovibrio vulgaris (Sisler and ZoBell, 1951;
Riederer-Henderson and Wilson, 1970) was detected at several
stations. D. limicola and D. vulgaris clustered with sequences taken from the seasonally hypoxic
Eckernförde Bay in the Baltic Sea (Bertics et al., 2013), suggesting a
major involvement of these sulfate-reducing bacteria in N2 fixation in
organic-rich sediments. Further, sequences related to
Vibrio diazotrophicus were detected, which has the unique ability for a known
Vibrio species to perform N2 fixation and which was found previously in the
water column of the OMZ off Peru
(Fernandez et al., 2011;
Löscher et al., 2014). Interestingly, we detected several new nifH gene
clusters in the Peruvian OMZ that have not been identified yet and which
have, consequently, yet unknown metabolic processes (Fig. 7). Thus, a
coupling of N2 fixation to processes other than SR is also possible,
which might also explain some of the discrepancies between N2 fixation
and SR activity (see above). However, the coupling to heterotrophic
metabolic processes such as denitrification or methanogenesis was not
supported by our molecular data.
Environmental factors controlling benthic N<inline-formula><mml:math display="inline"><mml:msub><mml:mi/><mml:mn mathvariant="normal">2</mml:mn></mml:msub></mml:math></inline-formula> fixation
The observed differences between integrated N2 fixation and SR along
the depth transect indicate potential environmental factors that control the
extent of benthic N2 fixation, which will be discussed in the following
section.
Organic matter
A major driver for microbial processes such as SR and N2 fixation by
potentially heterotrophic organisms is the availability of the organic
material
(Jørgensen,
1983; Howarth et al., 1988; Fulweiler et al., 2007).
Integrated N2 fixation and average organic carbon content showed
similar trends along the Peruvian OMZ depth transect (Fig. 5), and a strong
positive correlation was detected by PCA in the sediment depth profiles
(Fig. 6). Thus, organic matter availability appears to be a major factor
controlling N2 fixation at this study site. Low organic matter content
was previously shown to result in low N2 fixation rates in slope
sediments in the Atlantic Ocean (Hartwig and
Stanley, 1978). Correlation to organic matter was further confirmed by the
study of Bertics et al. (2010), which showed that burrow
systems of the bioturbating ghost shrimp Neotrypaea californiensis can lead to enhanced organic
matter availability in deeper sediment layers, resulting in high rates of
N2 fixation. However, high organic matter availability does not always
result in enhanced N2 fixation rates. Subtidal sediments in the
Narragansett Bay were found to switch from being a net sink via
denitrification to being a net source of bioavailable N via N2 fixation
(Fulweiler et al., 2007). This switch was caused by
a decrease of organic matter deposition to the sediments, which was in turn
triggered by low primary productivity in the surface waters.
Besides quantity also the quality of organic matter in sediments is a major
factor influencing microbial degradation processes
(Westrich and Berner, 1984). In the Peruvian OMZ
sediments, the average C / N ratio increased with water depth indicating that
the shallow stations received a higher input of fresh, labile organic
material compared to the deeper stations. Similar trends were reported for a
different depth transect off Peru
(Levin et al.,
2002). The C / N ratios did not follow the pattern of integrated N2
fixation (Fig. 5), which is in line with the PCA of depth profiles, which
showed a weak negative correlation between N2 fixation and the C / N
ratio. These results indicate that the C / N ratio is not a major factor
controlling N2 fixation in Peruvian OMZ sediments.
DIC fluxes, which were determined in benthic chamber lander incubations at
the same stations and during the same expedition as our study (Dale et al.,
2015), can be used as an indicator for organic matter degradation rates,
e.g., by SR. The DIC flux did not follow the pattern of the integrated
N2 fixation rates (Fig. 4) and thus does not indicate that N2
fixation and SR are coupled. Instead, the benthic DIC flux roughly followed
the pattern of SR rates along the depth transect. The highest integrated SR
rate and DIC flux were found at St. 1 (70 m), whereas the lowest occurred at
St. 10 (1025 m). Assuming that SR is largely responsible for organic matter
remineralization in the sediments below the OMZ (Bohlen et al., 2011; Dale
et al., 2015), the difference between integrated SR and DIC flux is expected
to be mainly caused by the loss of 35S-sulfides during the long
duration of unfrozen storage of the SR samples (see methods).
Ammonium
Interestingly, the highest N2 fixation was measured in sediments
colonized by the sulfur-oxidizing and nitrate-reducing filamentous bacteria
Marithioploca spp. (Schulz, 1999; Schulz and
Jørgensen, 2001;
Gutiérrez et al., 2008; Salman et al., 2011; Mosch et al., 2012).
Marithioploca facilitates dissimilatory NO3- reduction to NH4+, which
preserves fixed N in the form of NH4+ in the environment
(Kartal
et al., 2007). OMZ sediments off Peru are generally rich in NH4+
(Bohlen
et al., 2011; Dale et al., 2016). This co-occurrence of Marithioploca and N2
fixation was puzzling since high concentrations of NH4+ were
expected to inhibit N2 fixation
(Postgate,
1982; Capone, 1988; Knapp, 2012). It remains questionable why microorganisms
should fix N2 in marine sediments, when reduced N species are abundant.
Some doubt remains as to the critical NH4+ concentration that
inhibits N2 fixation and whether the inhibitory effect is the same for
all environments
(Knapp,
2012). For example, NH4+ concentrations up to 1000 µM did
not fully suppress benthic N2 fixation in a hypoxic basin in the Baltic
Sea (Bertics et al., 2013), indicating that
additional environmental factors must control the distribution and
performance of benthic diazotrophs
(Knapp,
2012). We observed high porewater NH4+ concentrations at the
shallow St. 1 with 316 µM at the sediment surface (0–1 cm)
increasing to 2022 µM at 20 cm (Fig. 2), while no inhibition of
N2 fixation was found. This observation is verified by the PCA, which
showed no correlation with ammonium for the N2 fixation depth profiles.
Hence, ammonium did not seem to have a significant influence on benthic
N2 fixation rates in the Peruvian OMZ.
Integrated rates of benthic N2 fixation (mmol N2 m-2 d1) in the Peruvian OMZ sediments from this study compared to other
marine benthic environments. Only the highest and lowest integrated rates
are shown, as well as the integrated sediment depth (cm) and the method used
(ARA = acetylene reduction assay, MIMS = membrane inlet mass spectrometry).
Benthic environment
N2 fixation
Depth of integration
Method
Reference
(mmol N2 m-2 d-1)
(cm)
Peru OMZ
0.01–0.4
0–20
ARA
This study
Coastal region
Baltic Sea, hypoxic basin
0.08–0.22
0–18
ARA
Bertics et al. (2013)
Bioturbated coastal lagoon
0.8–8.5
0–10
ARA
Bertics et al. (2010)
Brackish-water
0.03–3.4
0–1
ARA
Andersson et al. (2014)
Coral reef
6.09 (±5.62)
–
–
Capone (1983)
Eelgrass meadow
0.15–0.39
0–5
ARA
Cole and McGlathery (2012)
Eutrophic estuary
0–18
0–20
MIMS
Rao and Charette (2012)
Mangrove
0–1.21
0–1
ARA
Lee and Joye (2006)
Salt marsh
0.38 (±0.41)
–
–
Capone (1983)
Subtidal
0.6–15.6
0–30
MIMS
Fulweiler et al. (2007)
Zostera estuary
0.39
–
–
Capone (1983)
Open ocean
Atlantic Ocean (2800 m)
0.00008
–
ARA
Howarth et al. (1988)
< 200 m, various sites
0.02 (±0.01)
–
–
Capone (1983)
Mauritania OMZ
0.05–0.24
0–20
ARA
Bertics and Treude, unpubl.
One debated explanation for why diazotrophs still fix N under high
NH4+ concentrations is that bacteria fix N2 to remove
excess electrons and to preserve their intracellular redox state,
particularly with a deficient Calvin–Benson–Bassham pathway, as shown for
photoheterotrophic nonsulfur purple bacteria
(Tichi and Tabita, 2000). Another
explanation could be that microniches, depleted in NH4+, exist
between sediment grains, which we were unable to track with the applied
porewater extraction techniques (Bertics et al.,
2013).
Sulfide
Sulfide is a known inhibitor for many biological processes
(Reis, et al., 1992; Joye and Hollibaugh,
1995) and could potentially affect N2 fixation (Tam
et al., 1982). The shallow St. 1 was the only station with sulfide in the
porewater, reaching 280 µM in surface sediments and 1229 µM in
20 cm (Fig. 2). The presence of relatively high concentrations of sulfide at
St. 1 might explain why N2 fixation was lower at this site when
compared to St. 6, which had the highest N2 fixation rates.
Statistically, depth profiles of N2 fixation and sulfide showed a
negative correlation (Fig. 6b). Generally, interactions of sulfide with
benthic N2 fixation have so far not been investigated, and the PCA did
not provide a complete pattern, as sulfide was not widespread in the
sediments along the transect and thus does not allow robust interpretation.
Oxygen
Dissolved O2 can have a considerable influence on N2 fixation due
to the O2 sensitivity of the key enzyme nitrogenase
(Postgate, 1998; Dixon and Kahn, 2004).
Bioturbating and bioirrigating organisms can transport O2 much deeper
into sediments than molecular diffusion
(Orsi
et al., 1996; Dale et al., 2011). In coastal waters, the bioturbation and
bioirrigation activity of ghost shrimps was found to reduce N2 fixation
when sediments were highly colonized by these animals
(Bertics et al., 2010). While bottom water O2
concentrations in the Peruvian OMZ were below the detection limit at St. 1
to 8 (70 to 407 m), thereby mainly excluding benthic macrofauna, O2
concentrations increased to above 40 µM at St. 10 (1025 m) where a
diverse bioturbating and bioirrigating benthic macrofauna community was
observed (Mosch et al., 2012). Accordingly, St. 10 revealed some of the
lowest N2 fixation activity. We speculate that the low organic matter
content at this St. was mainly responsible for the low N2 fixation
rates and not the high bottom water O2 concentrations, as the
statistics showed a positive correlation between integrated N2 fixation
and organic carbon content.
Comparison of benthic N<inline-formula><mml:math display="inline"><mml:msub><mml:mi/><mml:mn mathvariant="normal">2</mml:mn></mml:msub></mml:math></inline-formula> fixation in different environments
We compiled a list of N2 fixation rates from different marine
sedimentary environments to gain an overview of the magnitude of N2
fixation rates measured in the Peruvian OMZ sediments (Table 2). We found
that N2 fixation rates from the Peruvian sediments exceed those
reported for open ocean sediments (2800 m; Howarth et al.,
1988), bioturbated coastal lagoon sediment (Bertics et
al., 2010) and sediments > 200 m water depth from various sites
worldwide (Capone, 1988). The highest integrated N2
fixation rate determined in our study (0.4 mmol N2 m-2 d-1, St. 6)
closely resembles highest rates found in salt marshes (0.38 mmol N2 m-2 d-1) and Zostera estuarine sediments (0.39 mmol N2 m-2 d-1)
(Capone, 1988). Further, our rates were characterized by a
similar range of N2 fixation rates that were previously measured in an
organic-rich hypoxic basin in the Baltic Sea (0.08–0.22 mmol N2 m-2 d-1, Bertics et al., 2013). In contrast to
the above examples, our N2 fixation rates were 8.5 times lower compared
to shallow (< 1 m) soft-bottom sediment off the Swedish coast
(Andersson et al., 2014) and 17 times lower than coral reef
sediments (Capone, 1988). However, in these environments,
phototrophic cyanobacterial mats contributed to benthic N2 fixation.
Given the dark incubation, N2 fixation of the present study seems to be
attributed to heterotrophic diazotrophs, which is additionally confirmed by
the nifH gene analysis, where none of the sequences clustered with cyanobacteria
(Fig. 7).