Introduction
Anthropogenic doubling of the supply of biologically available nitrogen
(Galloway et al., 2008;
Fowler et al., 2013) has
increased the importance of understanding the multiple components of the
nitrogen cycle. In marine ecosystems, microbial activity has been shown to
be a key driver in the nitrogen cycle, and while phototrophs can dominate
uptake in the water column (Flombaum et
al., 2013), chemolithotrophs and chemoheterotrophs have also been shown to
be quantitatively significant to nitrogen cycling
(Capone et al., 2008; Francis et al.,
2007; Zehr and Ward, 2002). In coastal marine areas, the large biomass of
macrofauna and macrophytes presents the opportunity for microbial taxa to
form associations where microbes have habitat as well as a predictable
nitrogen supply (Moulton et al., 2016). Many of these macrobiota
are restricted in movement, making them reliable substrates for microbial
populations.
There are many quantitative estimates of microbial nitrogen fluxes,
including ammonium oxidation (nitrification), in seawater from disparate
open-ocean locales (Beman et al.,
2011; Ward and Bouskill, 2011). Comparatively, there is little knowledge of
the microbially mediated nitrogen fluxes associated with nearshore species,
including whether the presence of animal and plant hosts enhance the
diversity and/or intensity of microbial functions. With the harvest and loss
of many marine species (Maranger
et al., 2008; Worm et al., 2006), the importance of determining the
biogeochemical role of microbes associated with macrobiota becomes more
urgent. Here, we quantify microbial nitrogen processing in coastal and
offshore water and in association with two key coastal species. Because
dissolved organic matter is one of the microbial resources supplied by
macrobiota in aquatic systems (Hansell and Carlson,
2015), we also manipulated dissolved organic carbon (DOC) concentration to
examine the effect of carbon availability on microbial nitrogen processing.
Across diverse aquatic ecosystems, the metabolic activities of animals and
plants can generate the environmental niches necessary for a variety of
microbial metabolisms
(Allgeier
et al., 2014; Croll, 2005; Layman et al., 2011; Schindler et al., 2001;
Subalusky et al., 2015; Vanni, 2002). Animals and plants create strong
gradients in oxygen and inorganic and organic nutrients such that processes
that vary over hundreds of meters or kilometers in the open ocean can change
over scales of millimeters in proximity to an animal (de Goeij et
al., 2013) or over a scale of meters relative to species aggregations
(Clasen and Shurin, 2015). With respect to temporal variation,
the photosynthetic and respiratory processes of coastal biota can drive wide
fluctuations in oxygen, possibly leading to both oxidizing and reducing
microbial metabolisms in the same location over a diel cycle (e.g.,
de Goeij et al., 2013; Pfister et al., 2016).
While strong gradients in biogeochemical cycling and associated microbial
activity have been demonstrated in soft sediment communities
(Eyre et al., 2011; Grossmann and Reichardt,
1991; Murray et al., 2015), there is increasing documentation that microbial
nitrogen processing, including nitrification and nitrate reduction, is
enhanced in proximity to surface-dwelling animals in marine systems
(Heisterkamp et al., 2013;
Pfister et al., 2014a, b, 2016a; Stief, 2013; Welsh and Castadelli,
2004). These enhanced nitrogen metabolisms also contribute to nitrous oxide
production (Heisterkamp et al., 2010, 2013), as well as
to the retention of nitrogen (Pfister et al., 2016).
A further influence of macrobiota on microbial metabolisms, particularly
seaweeds, is the production of dissolved organic matter
(Reed et al., 2015). Macroalgae produce DOC, likely
influencing microbial metabolism. DOC release by macroalgae may stimulate
heterotrophic nitrate reduction, where microbes respire DOC with nitrate
(NO3-) or nitrite (NO2-) as alternative electron
acceptors. DOC can also stimulate the oxidation of NH4+ through
heterotrophic nitrification (e.g., Joo et al., 2005). In
addition to promoting microbial transformations between NH4+ and
NO2/NO3, enhancing the DOC supply can result in competition
between different microbial metabolisms for dissolved inorganic
nitrogen (DIN). Work in streams suggests
heterotrophic bacteria may compete with chemolithotrophs for DIN
(Butturini and Sabater, 2000), a
result that may depend upon the ratio of C:N of available substrates, where
increasing DOC increases C:N and promotes nitrogen competition
(Strauss and Lamberti, 2000). In sum, increasing the supply
of DOC to marine microbes could have opposing effects on nitrification
rates. While an increase in ammonium oxidation would indicate stimulation of
heterotrophic nitrifiers, a decrease in NH4+ oxidation rate
would be consistent with increased competition for NH4+ with
heterotrophic microbes. Though the precise role of DOC in nitrogen
metabolisms is likely varied and still not fully described, DOC contributes
greatly to heterotrophy in microbes and fuels the quantitatively significant
“microbial loop” (Azam, 1998).
Because the effects that macrobiota have on both nitrogen excretion and DOC
release remain poorly understood, we tested how the presence of the
California mussel (Mytilus californianus), a red alga
(Prionitis sternbergii), and the proximity to shore affected
microbial nitrogen transformations during both daylight and nighttime
periods. We know that both species have a diverse microbial community
(Pfister et al., 2014b) and that mussels are a source
of nitrogen. Further, Prionitis is a common tide pool alga in
association with animals. We thus hypothesized that macrobiota (both mussels
and algae) would enhance microbial nitrogen cycling and nearshore seawater
would have greater microbial activity compared with offshore because it was
in close proximity to the macrobiota. We used gas-tight chambers and added
15N-enriched ammonium or nitrate to estimate the flux of ammonium
and nitrate attributable to microbial activity. Further, gas-tight chambers
allowed us to test whether microbial denitrification resulted in loss of
nitrogen via N2 gas. By quantifying nitrification, nitrate reduction, and
denitrification, we could distinguish nitrogen retention versus loss. We then
manipulated DOC (in the form of glucose) to test its specific effects on
nitrogen transformations. In sum, we asked the following questions. (1) How does the presence of
mussel, red algal tissue, or inert substrates affect microbial nitrogen
cycling? (2) Does the microbial activity in seawater differ between
nearshore and 2–5 km offshore? (3) Are there diel cycles in these
microbial nitrogen transformations? And (4) does the addition of dissolved
organic carbon alter microbial nitrogen metabolism?
Materials and methods
Chambers for assaying microbial metabolisms
In order to quantify how microbial nitrogen transformations contribute to
loss and retention of dissolved nitrogen we enclosed seawater and some
components of the rocky shore environment within two gas-tight Plexiglas
chambers. Each 2.26 L chamber measured approximately 15 cm in diameter,
30 cm in height, and contained two ports at the top: one for an o-ring-sealed
connection to an oxygen probe and the other with a septate lid for gas-tight
sampling of seawater. From 29 June to 22 August 2012, 54 assays (27 pairs) were
done in the chambers either in situ in tide pools 2 km east of Neah Bay, WA, USA; at
Second Beach, WA (n=19) (48.23∘ N, 124.40∘ W); at the
shore at Tatoosh Island, WA (n=26, 48.39∘ N, 124.74∘ W);
or onboard the R/V Clifford A. Barnes using seawater 2–5 km from each of
these shore-based sites (n=4). By using paired statistical contrasts to
compare the treatments described below, we minimized the possibility that
environmental variation drove differences between treatments. The Second
Beach site is described in Pather et al. (2014) and has
tide pools at a height of 1.2 to 1.5 m above Mean Lower Low Water (MLLW),
with a diversity of species (described in
Pfister, 2007; Pfister et al., 2016). The
chambers were anchored into a number of these tide pools for 3–5 h at a
time during periods of low tides when the tide pools were emergent. Thus, the
chambers contained tide pool water and were incubated to approximately half
of their height (15 cm) under natural light and temperature conditions.
Temperature could differ depending on the amount of sun and the air
temperature during a given day and ranged from 10.9 to 21.4 ∘C, though
it was most often in the range of 13.0 to 14.0 ∘C. Experimental trials
included tide pool seawater only (n=5), seawater with the California
mussel Mytilus californianus (n=9, estimated mean dry mass = 4.4 g), or seawater with bioballs
and ceramic rings (n=5 or 6 each). Bioballs are topographically complex
26 mm plastic balls used in commercial aquaria to provide substrate for
microbes with an estimated 15–20 cm2 surface area, while
Filstar™ ceramic rings (1 cm diam) are also used in filtration
and have an estimated 6 cm2 surface area (Aquatic
Eco-systems™). Both inert substrates were anchored in the
tide pools for 1 month to enable a natural microbial community to develop
prior to the experiments.
Because wave action was more significant at Tatoosh Island, chambers were
placed on shore in a 15 cm water bath within a shaded styrofoam cooler
rather than in tide pools. The cooler, with shading, protected the seawater
from reaching high temperatures. The chambers were filled with seawater at
the shore of Tatoosh Island and contained seawater only (n=8), seawater
with the California mussel Mytilus californianus (n=9, mean estimated dry mass = 6.2 g
+ 2.6SD), seawater with mussel shells only (n=3, mean
estimated dry mass = 3.9 g + 1.6SD), seawater with the red
alga Prionitis sternbergii (n=3, estimated wet mass = 40.0 + 12.5SD g), or
seawater with bioballs (n=3). Bioballs had been incubated at the lower
edge of the mussel bed for 1 month prior to use in the chambers. For all
experiments, the wet mass of Prionitis was weighed with a Pesola™ spring
scale, while the mussel dry mass was estimated from individual length
measurements of the mussels (Wootton, 2004). For the treatment
with mussel shells only, animal tissue was removed immediately prior to the
assay, thus testing the role of microbes residing only on the shell. By
doing the experiments at each site, mussels and Prionitis were always collected at
the site of the experiment, inert substrates were incubated at each site,
and seawater was local. We were thus able to make chamber incubations as
realistic as possible. Some of the above paired replicates of seawater
versus mussels were run during night hours to test for diel differences in
microbial activity.
Microbial nitrogen metabolisms were compared in shore-based seawater
collections versus seawater collected offshore in 2012. The offshore samples
were collected with a CTD Rosette system with 10 L Niskin bottles on the R/V
Clifford A. Barnes 2–5 km offshore from Tatoosh Island (48.432∘ N,
124.73∘ W) or Second Beach (48.37∘ N, 124.57∘ W) at a depth of 1 m and
adding 15N-enriched ammonium. The offshore
assays were done with the chambers in a cooler with a water bath onboard the
ship deck, again with a depth of 15 cm. Water temperature ranged from 10.7 to
15.5 ∘C, with most assays in the range of 11.0 to 12.0 ∘C. We
compared ammonium oxidation in four replicates of each shore and offshore
chambers during June and July of 2012 by pairing our offshore assays done
aboard the R/V Clifford A. Barnes with assays run at the shore at Tatoosh Island or
Second Beach within a week of two of the offshore assays.
We initiated each run by filling the chamber with seawater and any macrobiota
or bioballs. Oxygen and temperature were immediately recorded by a probe that
remained in the chamber through the duration of the experiment. We added
sufficient volume of 0.05 M of 15N-labeled ammonium chloride
solution (15NH4Cl) or sodium nitrate solution
(Na15NO3) to achieve an approximate enrichment of δ15N of
10 000 ‰ (Cambridge Isotopes).
We thus increased 15N-NH4+ or 15N-NO3- by a
factor of 10 with the intention of maximizing our ability to detect the
enriched signal in dissolved N2 gas. Both ammonium and nitrate
concentrations in seawater in this region are typically high (>2 and >10 µmol L-1, respectively), minimizing any
concentration-related effects from a tracer addition that ranged from only 8
to 40 µL. The chamber was agitated by hand to mix the tracer and then
agitated three to four more times during the 3 to 5 h incubation period
(approximately once per hour). No samples were taken during the incubation so
that we did not compromise the gas-tight nature of the chambers. At the end
of the incubation, we inserted a needle attached to a gas-tight syringe
through a rubber septa, drew out seawater, and injected this into a 30 mL
serum vial with a rubber stopper that had been evacuated to 160 mTorr with a
Welch 8905 vacuum pump. Samples had no head space and were stored at room
temperature.
Testing the effects of adding DOC
We hypothesized that heterotrophic microbes in association with phototrophs
would have the capacity to increase nitrification with added DOC. We thus
compared bioballs and the red alga Prionitis to seawater in an
additional experiment in 2014. We tested whether DOC additions enhanced
microbial nitrogen processing by increasing the concentration of DOC
approximately 6 times above the ambient nearshore concentration to 1000 µM DOC. We added 1.0 mL of a 1.96 M glucose solution to one chamber at the
beginning of the experiment while the other served as a control across all
paired experiments. We used glucose as a source for DOC because it is general
carbon and energy source for many organisms, facilitating comparison with
other published studies. All paired experimental runs with added glucose were
performed at Tatoosh Island and resulted in eight paired runs with seawater, and
four paired experiments each with either bioballs or Prionitis (mean
wet mass = 14.13 g + 4.61SD). We used an enrichment target
of 2000 ‰ of δ15NH4 (as 0.001 M
ammonium chloride, 15NH4Cl), a decreased enrichment
compared to those described above because we were not trying to detect an
enriched signal in N2 gas. This tripling of 15N-NH4+
allowed us to test whether ammonium oxidation changed with added DOC; an
increase in ammonium oxidation would indicate stimulation of heterotrophic
nitrifiers, while a decrease would be consistent with increased competition
for nitrogen by heterotrophs.
Quantifying enrichment results
In all experiments, a water sample was collected prior to tracer addition
(To) to quantify concentrations of ammonium, nitrate, nitrite,
phosphorus, and silica, as well as natural abundance isotope levels of
δ15NH4, δ15NO2, and δ15NO3. For all experiments, we used these initial measures to
calculate the subsequent change in enrichment in unlabeled N pools for rate
determination. For labeled forms of N, these data were used to calculate
enrichment levels after the addition of tracer. We collected the To
sample by filtering ∼180 mL of source water through a
syringe filter (Whatman GF/F) into HDPE (high-density polyethylene) bottles, which we kept frozen until
analysis. For the final sample (Tf) after 3–5 h of incubation, we
filtered directly from the individual chamber. All nutrient concentrations
were analyzed at the University of Washington Marine Chemistry Laboratory (methods
from UNESCO, 1994), while isotope determinations were done at
the University of Massachusetts Dartmouth using methodology for isotopic
composition reported previously (Pather et
al., 2014; Pfister et al., 2014b, 2016). Briefly, nitrogen stable isotopes
of ammonium were measured according to a modified version of the
NH4+ oxidation method detailed in Zhang et al. (2007). Ammonium is oxidized to nitrite using a hypobromite solution and
then reduced to N2O using a sodium-azide–acetic-acid reagent before
analysis on an IRMS (isotope ratio mass spectrometer). The stable isotope
ratios of nitrate were measured by cadmium reduction to nitrite, followed by
reaction with azide to N2O (McIlvin and Altabet, 2005). For
the DOC analysis, an additional 25 mL were filtered into a 40 mL VOA vial
(Shimadzu Corp.) and analyzed at the University of Washington Marine Chemistry
Laboratory. We tested for the presence of enriched N2 gas in the chambers
deployed in 2012 using sample collection and analytical procedures described
in Charoenpong et al. (2014). Chamber
oxygen and temperature were recorded with a Hach™ HQ4D and a
LDO probe. All comparisons of the effect of each substrate type on nitrogen
processing was assessed with ANOVA or paired t tests using R (https://www.r-project.org/, last access: November 2018).
Due to heteroscedastic variance in rate estimates, log
transformations were used.
Quantifying microbial transformations
Stable isotope enrichment experiments quantify nitrogen processing in marine
environments by tracking the transfer of the tracer between its source and
product pools (Glibert et al.,
1982; Lipschultz, 2008). The traditional isotope tracer transfer model
generally involves estimating a single rate parameter from time 0 to time t
(Lipschultz, 2008) and has the general form
Rate=(Rkt-Rko)/[(Rso-Rko)⋅Δt]⋅[k‾],
where k is the sink or product pool at time t (or the average k‾),
s is the source pool, and R designates the atom %
(15N/(15N+14N)×100) of either the source or sink.
The source-product model (Eq. 1) is thus used to estimate individual
nitrogen transformation rates. We estimated ammonium oxidation by quantifying
15N enrichment in nitrite following 15NH4+
addition. Similarly, nitrate reduction to nitrite was estimated from
15N enrichment in nitrite following 15NO3 addition.
A previous study of enrichment in tide pools showed substantial oxidation and
reduction in inorganic nitrogen that was best described with differential
equation models fit to multiple time points and underestimated with
source-product models (Pfister et al., 2016). Source-product
models likely underestimated the oxidation of ammonium here too because
remineralization by species within the chamber diluted the
15NH4+ tracer. We nevertheless used the simpler
source-product models because we had only a two-sample protocol, at the
beginning and the end of the experiment, to prevent gas escape. Isotope
dilution is important and indicates ammonium remineralization by species
within the chamber. We quantified ammonium remineralization in chambers with
15NH4+ tracer using the methods of
Pather et al. (2014). Briefly by fitting an exponential
model to the decline in δ15NH4 from the beginning to the end
of the experiment y=ae-bx. The parameters a and b were fitted
where b was the exponential decay constant in the δ15NNH4
enrichment. Remineralization rates were thus calculated as
NH4+remineralization=-b⋅[NH4‾]
in nmol L-1 h-1, where [NH4‾] was the mean concentration of
ammonium in nM at the beginning and end of the experiment.
The change in the concentrations of DIN (a ammonium, b nitrite,
c nitrate) in µM h-1 over the course of daytime experiments when only
seawater was present versus the addition of bioballs, mussels, or
Prionitis displayed as boxplots. Nitrogen transformation rates (in nmol L-1 h-1)
for (d) ammonium oxidation, (e) nitrate reduction, and (f) the
ammonium remineralization rate. Letters indicate statistical differences
with ANOVA and Tukey HSD. The box shows 50 % of the data, the horizontal
line is the median, and the vertical lines represent the first and fourth
quartiles. Where vertical lines are absent, they are contained within the
boxes. Outliers are shown as individual points outside the vertical lines
and are 1.5 times the value beyond the top or bottom of the box.
Results
Dynamics of nutrients and isotopes in chambers
The presence of either the California mussel or the red alga Prionitis amplified net
changes to ammonium and nitrate concentration in the experimental chambers
compared with chambers that contained bioballs or only coastal seawater
during daylight hours (Fig. 1). Chambers during daylight hours with
Prionitis and mussels had increased ammonium over the course of the experiment
compared with the relatively unchanged coastal seawater and bioball
treatments (Fig. 1a, F5,51=6.150, p<0.001), while nitrate
decreased with Prionitis and increased with mussels (Fig. 1c, F5,51=3.512,
p=0.008). Changes in nitrite did not differ among treatments (Fig. 1b,
F5,51=0.66, p=0.659).
The dynamics of δ15NNH4, δ15NNO2, and
δ15NNO3 within the chambers revealed transfer of 15N
isotope and thus microbial transformations. When 15N-NH4+
was added, enrichment in δ15NNO2 and δ15NNO3 and any dilution
in the δ15NNH4signal was measured (Fig. A1a, b). Similarly, enrichment in
δ15NNH4 and δ15NNO2 followed the addition of
15N-NO3- (Fig. A1c, d). Deviations in our target of
initial enrichment (10 000 ‰ and 2000 ‰) occurred due to
natural variation in nutrient concentrations at the time of tracer addition, which was factored into rate calculations.
The mean (and standard error) rates of nitrification and nitrate
deduction in chambers with seawater, bioballs, mussels, and the red alga
Prionitis, expressed as nanomoles (nmol) per liter per hour, nanomoles per gram mass per hour, and as a
daily rate in nanomoles for a square meter of rocky shore, respectively.
Mean nitrification rates (SE)
Mean nitrate reduction rates (SE)
nmol L-1 h-1
nmol g mass-1 h-1
mmol m2 day-1
nmol L-1 h-1
nmol g mass-1 h-1
mmol m2 day-1
Seawater
0.17 (0.09)
–
–
0.71 (0.26)
–
–
Bioballs
0.82 (0.66)
–
–
0.82 (0.25)
–
–
Mussels
14.1 (3.80)
3.21 (0.64)a
2.50 (0.50)
6.70 (1.56)
1.60 (0.41)a
1.20 (0.30)
Prionitis
0.82 (10.31)
1.50 (0.27)b
0.13 (0.08)
63.70 (33.70)
1.56 (0.72)b
0.13 (0.08)b
a designates dry mass, b designates wet mass.
The ammonium oxidation rate (a) and the nitrate reduction rate
(b) contrasted between day and night hours for mussels or seawater at the
shore. Data are log-transformed (from nmol L-1 h-1) to facilitate
comparison. Rates with mussels were always greater (for a
F1,11=52.59, p<0.001; and b F1,14=68.14, p<0.001). Ammonium oxidation rates in association with mussels or in seawater
alone did not differ between day and night (F1,11=0.58, p=0.461),
while nitrate reduction in seawater was greater during the day
(F1,14=5.83, p=0.030). Boxplots as in Fig. 1. Changes to the
isotopes of inorganic nitrogen are shown in Fig. A1.
Nitrogen transformation rates
Microbial nitrogen processing rates increased when either the California
mussel or the red alga Prionitis was present. Ammonium oxidation
rates with the mussel (mean of 14.1 nmol L-1 h-1) or red alga
(32.8 nmol L-1 h-1) were 2 orders of magnitude greater than ammonium
oxidation in seawater only or with bioball surfaces which
were less than 1 nmol L-1 h-1 (Fig. 1d, F5,25=15.19, p<0.001, logged
values; untransformed values in Table 1). Our estimates of nitrate reduction
from the addition of Na15NO3 were also 2 orders of magnitude
greater with Prionitis (63.7 nmol L-1 h-1) and mussels
(6.7 nmol L-1 h-1) compared with bioballs and seawater (Fig. 1e,
F5,19=17.64, p<0.001, logged values; untransformed values in
Table 1). For all these estimates of microbial nitrogen processing, we found
high overlap in the rates estimated with living, intact mussels compared with
mussel shells only, indicating that the responsible microbes reside on the
shell surface, rather than the mussel tissues (Fig. 1d, e). The presence of
mussels was further associated with increased ammonium remineralization and
was double the rate with bioballs and the red alga Prionitis, as well as an
order of magnitude more than that for seawater alone (Fig. 1f).
Our estimates of mussel or algal mass within each chamber resulted in
per-gram estimates of the effect of these macrobiota on nitrogen transformation
rates. For every gram of mussel dry mass, 3.21 nmol (se=0.64) of ammonium
was oxidized per liter per hour, while 1.60 nmol of nitrate was reduced
(se=0.41) (Table 1). A comparable contribution is made per gram of
Prionitis wet mass with 1.50 nmol ammonium oxidized per hour (se=0.27) and 1.56
nmol nitrate reduced per hour (se=0.72).
Day versus night nitrogen transformations
There was no difference between daytime and nighttime nitrification rates in
association with mussels or in seawater alone (Fig. 2, F1,11=0.583,
p=0.461), recalling that mussel-associated ammonium oxidation rates were
2 orders of magnitude higher than for seawater only. Seawater nitrate
reduction rates during the day (1.15 nmol L-1 h-1) were 4 times
greater than those at night (0.26 nmol L-1 h-1, Fig. 2b;
day > night, F1,14=5.83, p=0.030), also keeping in mind
that overall rates were 10 times higher when mussels were present as
compared to only seawater (mussels > seawater, F1,14=68.1,
p<0.001).
DIN uptake in chambers could be due to both microbial transformations or
seaweed uptake. Comparing tracer-based rate estimates with changes in
concentration, we find that nitrate reduction accounted for as little as
4.2 % of the decrease in nitrate concentration during the day, but as much
as 87.2 % at night. Estimates of ammonium oxidation revealed that ammonium
oxidation made up 5.2 %–7.4 % of total ammonium uptake during the day.
(a) The ammonium oxidation rate (nmol L-1 h-1) in surface
seawater collected at the shore versus 2–5 km offshore, based on four trials in
each locale in June and July of 2012. The rates did not differ (t=t3.8=1.65, p=0.177). The change in (b) DIN and (c) silica
(nmol h-1) also did not differ whether seawater was from the shore or
offshore (b DIN t3.8=1.31, p=.260; c silica t3.8=0.68,
p=0.525). Boxplots as in Fig. 1.
Onshore versus offshore microbial nitrogen transformations
The seawater-only chambers showed no difference in ammonium oxidation
(nitrification) rates whether collected at the shore (mean = 0.23 nmol L-1 h-1) or offshore (mean = 1.12, Fig. 3a, t3.8=1.65,
p=0.177), although the sample size was low (n=4). Overall, there was
little change in nutrient concentration when seawater from either offshore
or nearshore was isolated; the overall mean change in DIN was less than 1 µM for both nearshore (-0.70) and offshore (0.63),
with no significant
difference between them (t=1.31, p=0.260). There was also no difference
in silicate uptake between the two regions (t=-0.679, p=0.525),
indicating that diatom activity did not differ in the two regions.
The effect of supplemental DOC on (a) the rate of ammonium
oxidation (in nmol L-1 h-1), (b–g) the change in nutrient
concentrations (µM h-1), and (h) the oxygen concentration (in
mg L-1 h-1). An * indicates a significant difference
(p<0.05) between the control and the DOC addition for
seawater alone, seawater with bioballs, or the red alga Prionitis.
Double dagger indicates 0.10>p<0.05.
Boxplots as in Fig. 1.
Nitrogen transformation rates with added DOC
On average, the coastal seawater that was used in the chambers had a DOC
concentration of 145 µM; replicates with the addition of glucose
increased DOC approximately 6 times that amount to 1000 µM. In the
presence of Prionitis, DOC also increased with a mean of 9.31 mmol L-1 h-1 over
the course of the experiment (n=4). Nitrification rates did not change
significantly when glucose was added (Fig. 4a), although we acknowledge that
our sample size was small and nitrification was not detected in some
instances across both treatments, perhaps impeding a strong test of glucose
effects as nitrification was not detected in some instances across both
treatments. However, DOC addition did change nutrient uptake rates. The
addition of DOC to experimental chambers with Prionitis, bioballs, or in seawater
alone generally resulted in greater uptake of nitrite and nitrate with
Prionitis, bioballs, or in seawater alone while ammonium showed a trend toward greater
uptake only with Prionitis; otherwise there was little overall change in ammonium
concentration (Fig. 4b, c, d). DOC addition was also associated with an
increased uptake of DIN and phosphorus, regardless of the composition of the
chamber (Fig. 4e, f). Silica was unchanged with bioballs or seawater alone,
while there was greater uptake of silicate with Prionitis, suggesting Prionitis hosts diatoms
(Fig. 4g).
The greater uptake of DIN in chambers with supplemental DOC could be due to
increased microbial assimilation and respiration or both with DOC. The
effect of glucose on the uptake of DIN or phosphorus did not differ based on
whether seawater, bioballs, or Prionitis were in the chamber (F2.31=0.645,
p=0.531; and F2.31=0.264, p=0.770, respectively), suggesting that
the background metabolism of heterotrophic bacteria was the same regardless
of the macrobiota or substrate available. If microbial respiration increased
with added DOC, we were unable to detect it by measuring oxygen
concentrations. Whether we pooled treatments for seawater, bioballs, and
Prionitis or examined them separately, dissolved oxygen measurements did not differ
(t=1.125, p=0.277, df=16).
Discussion
Seascape scale importance of macrobiota for microbial N
metabolism
The per-mass estimates of microbial nitrogen transformations that we
measured reveal significant macrobiota-associated microbial processing rates
along coastal shorelines. Studies from Wootton (2004)
estimate that a square meter of mussel bed can contain 32 425 g dry mass of
mussel. Extrapolating from our measurements for both day and night,
microbial nitrification in a square meter of mussel bed would amount to 2.5 mmol day-1 (+0.5SE), with an additional
1.2 mmol of nitrate reduction (+0.3SE) (Table 1). As
a comparison, at this site it would take a volume of seawater of 1 million L to host the same microbial nitrogen metabolism, the equivalent of a
10 m by 10 m area of the ocean to 10 m depth (1000 m3).
A similar calculation can be done for macroalgae using Paine's (2002) estimates of macroalgal mass in control plots in the intertidal at
Tatoosh Island (8.6 kg m-2). If Prionitis has any functional similarity
to other seaweeds sampled by Paine (2002), then ammonium
oxidation could reach 0.13 mmol day-1 (+0.08SE) for a square meter of seaweed with a mass of 8.6 kg, while nitrate
reduction would occur at approximately the same rate (Table 1). While these
rates are an order of magnitude lower than mussels, the macroalga
contribution is still substantial and comparable to water column
nitrification or nitrate reduction only when we consider a volume greater
than 129 000 L or a sea surface area in excess of a meter on a side
and 10 m depth. Even if Prionitis is exceptional with respect to microbial function
when compared with other seaweeds, the potential contribution of macroalgae
to microbial function could be substantial. Thus, independent of macroalgal
effects on DIN uptake (Fig. 1a, b, c) or ammonium remineralization by mussels
(Fig. 1f), the microbiome of each of these species makes distinct
contributions to nitrogen cycling. Our measurements emphasize the
quantitative importance of common nearshore species to the nitrogen cycle
and highlight how nearshore areas may differ from those offshore.
We demonstrated that seawater isolated from the immediate vicinity of
benthic substrates had similar rates of nitrogen metabolism to offshore
water (Fig. 3). As this measurement indicates no difference in the activity
of suspended microbes, we conclude that microbial metabolism was elevated
due to microbes directly associated with the mussel and the red alga.
Previous analyses of 16s rRNA sequencing of mussel tissue, mussel shell,
Prionitis, seawater, and inert surfaces show that microbial communities can be distinct
on these substrates (Pfister et al., 2014b). Further,
metagenomic analysis of mussel shell microbes indicates DNA sequences
associated with a diversity of nitrogen metabolisms (Pfister
et al., 2010). The similarity of nearshore to offshore microbial function
would appear at odds with our previous work showing that natural isotopes of
ammonium and nitrate (δ15NNH4 and δ15NNO3) are enriched near the shore, indicating increased
microbial processing. However, the nearshore results likely reflect
benthic-associated activity influencing adjacent seawater.
The rates of ammonium oxidation reported here in association with mussels
and red algae greatly exceeded not only our estimates for seawater alone,
but also those reported previously for water column rates. Our
macrobiota-associated rates of ammonium oxidation (Fig. 1) were at least 2 orders of magnitude greater than
a compilation of open-ocean areas
(Beman et al., 2011). Similarly, other seawater assays in
coastal areas of the eastern Pacific Ocean showed nitrification rates on the
order of 1–10 nmol L-1 day-1 (Santoro et al.,
2010; Fernandez and Farías, 2012), comparable to the seawater
rates we report in the absence of benthic species. When mussels were present
in our chambers, the estimated rates of ammonium oxidation were 3.21 nmol g-1, a value comparable to those reported by
Heisterkamp et al. (2013) for snails and mussels and
by Welsh and Castadelli (2004) for bivalves. Thus, through
genetic and biogeochemical analyses, there is increasing evidence for
diverse and quantitatively significant nitrogen metabolisms in association
with macrofauna.
Microbial metabolism and dissolved organic carbon
When considering the effect of DOC in microbial assemblages, there are three
groups of microbes that might be affected. There are nitrifiers that are
either heterotrophic or chemolithotrophic (Ward, 2008), as well
as heterotrophic bacteria that might consume DOC and assimilate ammonium,
but not nitrify (e.g., Kirchman, 1994). Thus, added DOC might be
expected to increase heterotrophic nitrification if DOC was limiting
nitrifier growth. Alternatively, added DOC could decrease nitrification if
generalist heterotrophic bacteria assimilating ammonium were stimulated and
were to then outcompete chemolithotrophs oxidizing ammonium
(Butturini and Sabater, 2000),
although we do not know if ammonium was ever limiting. A third possibility
is that heterotrophic nitrifiers are such a small percentage of
nitrification activity that there is no detectable effect of elevated DOC.
We found mixed evidence for the effects of DOC on nitrification. Ammonium
oxidation was never stimulated by DOC (Fig. 4); if anything, there was a
nonsignificant trend of decreased ammonium oxidation with glucose,
suggesting that general heterotrophic bacteria were consuming the elevated
DOC. Our DOC additions were accompanied by decreased dissolved inorganic
nitrogen and phosphorus in the surrounding seawater, suggesting that
heterotrophic microbial metabolism increased, a result consistent with other
glucose-addition studies with microbes (Zhang et al., 2013).
Bacterial production in seawater has been shown to increase with glucose
addition (Caron et al., 2000; Jacquet et al., 2002), with
heterotrophic bacteria released from carbon limitation when DOC is added
(Jacquet et al., 2002; Joint et al., 2002). In streams,
glucose additions have resulted in decreased nitrification
(Strauss and Lamberti, 2000), a result attributed to
heterotrophic bacteria in direct competition with nitrifiers. While
Strauss and Lamberti (2000) documented decreased oxygen
concentration and increased respiration with added DOC, we detected no
effect of DOC on the change in oxygen within chambers (Fig. 4h). The unknown
contribution of photosynthesis to oxygen concentrations, as well as the
relatively high oxygen content of the seawater in these locales, could have
masked oxygen differences. Nonetheless, DOC stimulated nutrient uptake,
presumably by heterotrophic microbes, and the effect of DOC was the same
whether seawater, bioballs, or Prionitis were in the chamber (Fig. 4b–i). Thus, the
background metabolism of heterotrophic bacteria was unchanged even when
Prionitis was present and reduced chamber DIN concentrations 6.5 µM over the course
of the experimental runs.
A final explanation for increased DIN uptake with added DOC is that bacteria
are able to compete with any phototrophs for nitrate when an organic carbon
source is increased (e.g., Diner et al., 2016).
Nitrate reduction rates are high with Prionitis and this alga also provisions DOC,
perhaps promoting the coupling of heterotrophy and nitrate reduction.
Whether any of the decreased nitrate concentration associated with
Prionitis in chambers could be attributed to heterotrophic nitrate reduction is
unknown at this time, because our experiments with added DOC did not assay
nitrate reduction, only ammonium oxidation.
In sum, while DOC concentrations can be elevated in nearshore areas compared
with offshore, there was little evidence that enhanced DOC changed
nitrification rates, even in the chambers with Prionitis, where DIN levels were lower
due to seaweed uptake. Whether heterotrophic nitrifiers are present remains
unknown, though previous analysis of microbes at these sites suggests the
presence of taxa associated with heterotrophic nitrification, e.g.,
Arthrobacter (Hynes and Knowles, 1982), Crenarchaeota (Offre et
al., 2013), and Alcaligenes faecalis (Joo et al., 2005), though they were detected
in only a small fraction of samples (Pfister et al., 2014b).
Analyses of 16s rRNA of seawater, mussels, and Prionitis do show sharp distinctions in
β-diversity, with some taxa unique to each (Pfister
et al., 2014b).
Taken together our data suggest that chemolithotrophic nitrifiers are
dominating nitrification in this area. Other heterotrophic bacteria can
noticeably depress DIN and phosphate concentrations when DOC is
supplemented, suggesting there may be some carbon limitation for
heterotrophic microbial metabolisms. If, as suggested by
Strauss and Lamberti (2000), the C:N ratio available to
microbes, either in the water column or in the substrate they are using,
determines the relative fitness of heterotrophic bacteria versus
chemolithotrophic nitrifiers, then the many regions where DIN concentrations
in seawater are lower than they are at our Washington coastal sites may show
a different result.
Of note is that many seaweeds produce detectable amounts of DOC in coastal
areas (Wada and Hama, 2013), with as much as 14 % of net
primary production being released as DOC in a kelp species
(Reed et al., 2015). Among other seaweeds, 20 % to 30 % of
released DOC can be taken up within 2 h (Brylinsky, 1977),
suggesting an active heterotrophic assemblage in proximity. Seaweeds also
have a diverse assemblage of microbial associates
(Lemay et al., 2018;
Marzinelli et al., 2018; Michelou et al., 2013; Pfister et al., 2014b). Which
of these associated microbes benefit from this DOC and whether others are
inhibited is unknown. While we tested the effect of elevated glucose on
nitrification with 15N-enriched ammonium, a next step is to test if
those microbes involved in the nitrate reduction pathways are affected by
glucose addition.
Macrobiota that serve as hosts for microbes provide a predictable substrate
for attachment in a fluid environment and provide dissolved organic matter
in many forms (Carlson and Hansell, 2015). The mussels studied
here also excrete ammonium and likely DON
(Bayne and Scullard, 1977;
Pather et al., 2014). Their filter-feeding activities release DOC in many
forms and continually process organic matter that can be utilized by
microbes (Jacobs et al., 2015). Through filter feeding and mucus
production, there is increasing evidence that marine invertebrates and
microbes are connected through their production and use of dissolved organic
matter (Rädecker et al., 2015; Rix et al.,
2016).
The multiple factors influencing nitrogen availability
Our experiments provide insight into the fate of nitrogen in coastal systems.
While ammonium oxidation and nitrate reduction rates were 2 orders of
magnitude higher than any water column estimates, we have no evidence that
nitrate reduction continued through to denitrification and the release of
N2 gas as we never detected enriched 15N in N2 gas
(e.g., Jensen et al., 2011), despite our ability to detect a 1.0 ‰ enrichment in N2 gas. Thus, nitrogen was being retained in our
experimental system. If ammonium oxidation and nitrate reduction are
occurring relatively constantly, as suggested by our experiments, then a
diversity of microbially mediated DIN dynamics may take place across
microenvironments that differ in oxygen levels. The net result could be
continued microbial use of ammonium and nitrate and the ability for the
microenvironment surrounding the animal or seaweed to sustain a range of
microbial metabolisms, a result obtained for other marine invertebrates
(de Goeij et al., 2013; Heisterkamp et al.,
2013). Research in tide pools containing these same species has also shown
both nitrogen oxidation and reduction processes (Pfister et al.,
2016). In all instances to date, the metabolism of the host macrobiota
results in a daily range of oxygen levels, thus providing a diversity of
environmental niches that favor different microbial transformations through
time.