Contribution of ammonia-oxidizing
bacteria and archaea to nitrification in
estuarine sediments along a eutrophic
gradient
Emily Stone
Clark University
Advisors: Julie Huber and Anne Giblin
Abstract
Eutrophication, the enrichment of ammonium and phosphate, is a major concern for
coastal aquatic systems such as estuaries. Eutrophication may hinder the coupling of
nitrification-denitrification in sediment, which is responsible for removing large amounts of
nitrogen in aquatic ecosystems. The first and rate-limiting step in nitrification is catalyzed by the
enzyme ammonia monooxygenase (amo) and is performed by two distinct groups of
microorganisms: ammonia-oxidizing bacteria and ammonia-oxidizing archaea. Presence of
ammonia-oxidizing bacteria and archaea was determined by amplifying the amoA gene.
Sediment slurries were used to measure potential nitrification rates from estuarine sediments
taken along a eutrophic gradient. Results showed lower potential nitrification rates at more
eutrophic sites and higher nitrification rates at less eutrophic sites. Similarly, the relative
abundance of ammonia oxidizing archaea was lowest at the most eutrophic site and highest at the
least eutrophic site. The bacterial inhibitor, 1-octyne, which has been shown to inhibit ammonia
oxidizing bacterial activity in soils, did not inhibit these bacteria in sediment. Although
ammonia-oxidizing bacteria were detected in sediments, the relative abundance was not
determined. However, results suggest that ammonia-oxidizing archaea are more abundant in
estuarine sediments than ammonia-oxidizing bacteria. Trends indicate that the relative
abundance of ammonia-oxidizing archaea increases as eutrophication decreases but no trends
were detected for ammonia-oxidizing bacteria.
Key words: eutrophication, nitrification rates, ammonia oxidizing bacteria, ammonia
oxidizing archaea
Introduction
Anthropogenic inputs on ecosystems influence the nitrogen cycle and have resulted in
many environmental problems including loss of biodiversity, global water acidification,
increased levels of the potent greenhouse gas, nitrous oxide, and eutrophication (Gruber and
Galloway 2008). Coastal regions receive much of the nitrogen-rich runoff from municipal and
industrial wastewaters as well as urban and agricultural fertilizers. These regions are also
susceptible to nutrient enrichment. Nutrient enrichment is mainly in the form of ammonium and
leads to eutrophication in the water. Eutrophication, in particular, is a serious concern for
estuaries. More than half of the estuaries in the US experience some type of effect from it
(Meyer-Reil and Köster 2000). However, nitrogen levels in estuaries can typically be reduced
through tightly coupled microbial processes in the nitrogen cycle. Nitrification converts
ammonium to nitrite and then nitrate, which then can be converted to dinitrogen gas by
denitrification or anammox. These microbially-mediated processes thus determine recycling and
removal of dissolved inorganic nitrogen from aquatic systems. Denitrification in marine
sediments is dependent on the supply of nitrate and is driven almost exclusively from sediment
nitrification (Jensen et al. 1993). Coupling of nitrification-denitrification removes up to 50% of
external dissolved inorganic nitrogen from ammonium inputs to estuaries (Seitzinger et al. 2006).
Nitrate production requires oxygen, thus nitrification is restricted to aerobic environments (Ward
et al. 2011). Because both ammonia-oxidizing bacteria and archaea require oxygen in order to
nitrify, these microorganisms may be affected by the environmental changes brought about from
nutrient loading by eutrophication such as reduced oxygen concentration in sediments. Thus,
eutrophication may limit aerobic regions for nitrification, and thereby, may decrease nitrate
production and removal of inorganic nitrogen in aquatic environments.
The first and rate-limiting step in nitrification is ammonia oxidation catalyzed by the
ammonia monooxygenase (amo) gene (Beman et al. 2007). Nitrification was long thought to be
solely carried out by ammonia-oxidizing bacteria in soils and sediments (Bernhard et al. 2010).
However, recent studies have shown that archaea from the phylum Thaumarchaeota can also
oxidize ammonium and are present in various soils and sediments (Taylor, et al. 2013).
Molecular studies have identified genes putatively encoding archaeal ammonia monooxygenase
subunits (amoA, amoB, and amoC). The amoA subunit is homologous to the amoA gene found in
ammonia-oxidizing bacteria (Mosier and Francis 2008). Thaumarchaeota carrying the archaeal
amoA gene appear to be ubiquitous and have been detected in coastal and marine waters, cold
sediments, subsurface of radioactive thermosprings, microbial mats, coral reefs, as well as,
terrestrial forest soils and grasslands (Erguder, Boon et al. 2009). Several studies have found that
ammonia-oxidizing archaeal amoA gene copy numbers are more abundant than the bacterial
amoA gene in soil, estuaries, and open waters (Mosier and Francis 2008). Beman et al. (2008)
determined that ammonia-oxidizing archaea dominated ammonia-oxidizing bacteria by a factor
of 37-217 in surface waters from the Gulf of California while Caffrey et al. (2007) found an
abundance ratio of archaeal to bacterial amoA gene around 80 in estuarine sediments. Although
the archaeal amoA gene has been found in higher abundance than the bacterial amoA gene, the
relative contributions of ammonia-oxidizing bacteria and archaea to nitrification are still not fully
understood.
Recent studies suggest that selectively blocking ammonium oxidation by ammonia
oxidizing bacteria may help estimate ammonia-oxidizing archaea nitrification rates. This, in
return, may aid in determining the relative contribution of both ammonia-oxidizing bacteria and
archaea to nitrification in soils and sediments. For example, Taylor et al. (2013) found that 1octyne inhibited a range of ammonia-oxidizing bacterial activity in soils by inactivating the amo
enzyme. It had no effect on ammonia-oxidizing archaeal activity in the soil. They also found that
1-octyne inhibited the bacterial activity in less than 48 hours. With its relative efficiency, this
inhibitor may also help differentiate potential nitrification rates attributable to either ammoniaoxidizing bacteria or archaea in estuarine sediments.
Using a combination of potential nitrification rate measurements and PCR-based
methods, this paper investigates i) the effects of eutrophication in estuarine sediments around
Falmouth, Massachusetts on nitrification rates and ii) the relative contribution of ammoniaoxidizing bacteria and archaea to nitrification in eutrophic sediments.
Methods
Sampling Sites
Four inch diameter sediment cores were taken in duplicates from four estuaries located
around Falmouth, Massachusetts in early November 2015. Sediment taken from Little Pond and
Eel pond did not show a clear separation of aerobic and anaerobic regions. Sediment cores
collected from these locations were near black and viscous indicating more anaerobic conditions
and higher eutrophication. However, sediment collected from Quissett Harbor and Waquoit Bay
was less viscous and had more distinction between the aerobic and anaerobic regions indicating
less eutrophic sediments. Additionally, amphipod tubes were present in sediment taken from
Waquoit Bay, which indicates well-oxygenated sediment. To determine level of eutrophication
in sediment, subsamples of sediment from cores from each of the four sites were burned at
500ºC in a muffle furnace (Thermo Scientific™ Thermolyne™ Industrial Benchtop Muffle
Furnaces). Percent organic matter within sediments was calculated by subtracting original weight
of sediment from post-burned sediment. The four sites were then organized along a eutrophic
gradient based on percent organic matter in sediment subsamples with higher percent organic
matter reflected more eutrophic sediment and lower percent organic matter reflected less
eutrophic sediment.
Potential Nitrification Rates
Potential rates of nitrification in sediments were measured using a shaken sediment-slurry
method with added nutrient solution (Bernhard et al. 2007). Two treatments were analyzed to
look at total nitrification and archaeal nitrification: (i) environmental control and (ii) with
addition of the bacterial inhibitor, 1-octyne (10 uM). Octyne was used as in inhibitor to block
ammonia-oxidizing bacteria activity while allowing ammonia-oxidizing archaea activity to
continue (Taylor et al. 2013). Cores were sectioned off to isolate the aerobic region of the
sediment by removing the top 2 centimeters of sediment. Approximately one gram of wet weight
of sediment from each site was added to 50 mL falcon tubes for destructive sampling at time
points 12 hours, 24 hours, 48 hours, and 72 hours, and several blanks in duplicates. Archaeal
nitrification from Little Pond and Waquoit Bay had additional time points at 120 hours and 168
hours for a total 7 day incubation period to determine if the archaeal population would rapidly
increase without bacterial ammonia oxidizers. Subsamples from cores of each site were weighed,
dried at 60ºC for two days and re-weighed to obtain wet/dry conversion factors. Seawater was
collected from each site and filtered through GF/F filters and re-filtered again with 0.20
nanometer pore sized membrane filters to remove microorganisms. A nutrient solution (300 µM
NH4, 60 µM PO43) was created with filtered seawater from each location and 30 mL was added
to each tube to create a slurry mix. For the bacterial inhibitor treatment, 700 µL of 1-octyne was
added initially to the slurry mix and 300 µL was added subsequently each day of the three-day
incubation period. All tubes were shaken at low speed at room temperature. Tubes were taken
after 12 hours, 24 hours, 48 hours, and 72 hours and centrifuged for 2 minutes at 10,000 rpm.
Approximately 15 mL of supernatant was collected from each time point from each of the sites
and measured for nitrate concentration using QuickChem 8500 Series 2 FIA system (Lachat
Instruments, Milwaukee, WI). Sediment from initial sectioning of the cores and 72 hour time
points were saved and frozen at -80ºC for DNA extractions.
The change in nitrate concentration was plotted for each site for both the control and the
treatment with the bacterial inhibitor. The nitrate concentration between the two replicates for
both treatments was averaged together at each location to determine potential nitrification rates.
Wet weight of sediment was converted to dry weight of sediment to give potential nitrification
rates in nmol dry weight of sediment-1 hour-1. Standard error was found for each treatment for
each site.
DNA Extraction and Polymerase Chain Reaction
DNA was extracted from approximately 1 gram of sediment that was initially collected
and from sediment collected after 72 hour incubation for nitrification from all four sites using a
MoBio PowerSoilTM DNA isolation kit following the manufacturer’s protocol (MoBio
Laboratories Inc., Carlsbad, CA). Extracted DNA was quantified with a NanoDrop™ Micro
Volume Spectrophotometer and stored at -200C (Thermo Scientifiic, Wilmington, DE).
Polymerase chain reaction (PCR) was used to determine the presence of bacteria and archaea by
targeting the conserved 16s rRNA bacterial and archaeal gene. 16s rRNA was amplified using
the bacterial forward primer 8F (5’-AGA GTT TGA TCM TGG CTC AG) and reverse primer
1492R (5’-GGT TAC CTT GTT ACG ACT T) and the archaeal forward primer 21F ( 5’-TTC
CGG TTG ATC CYG CCG GA) and reverse primer 958R (5’-YCC GGC GTT GAM TCC ATT
T). Due to the conserved phylogeny and its functional significance in nitrification, the amoA
gene was amplified to test for both ammonia oxidizing bacteria and archaea using the bacterial
amoA gene forward primer AmoA1F (5’-GGG GTT TCT ACT GGT GGT; Rotthauwe et al.
1997) and reverse primer AmoA2R (5’-CCC CTC KGS AAA GCC TTC TTC; Rotthauwe et al.
1997) and archaeal amoA gene forward primer ArchAmoAF (5’-STA ATG GTC TGG CTT
AGA CG; Francis et al. 2005) and reverse primer ArchAmoAR (5’GCG GCC ATC CAT CTG
TAT GT; Francis et al. 2005). All PCR reactions were performed in duplicate for all four sites in
a total volume of 25 µL containing 12.5 µL Taq DNA Polymerase (Thermo Scientific PCR
Master Mix, ThermoFisher Scientific), 4 µL of bovine serum albumin (0.4 g/ 100 mL), 1 uL of
forward and reverse primer (10 uM), 1 µL of template DNA and 5.5 µL of molecular water. The
standard thermal profile used for both bacterial and archaeal 16s rRNA gene amplification was
as follows: 3 min at 94°C; then 35 cycles consisting of 40 s at 94°C (denaturation), 1.5 min 55°C
(annealing), and 2 min at 72°C (extension); and a final extension at 72°C for 10 min. The
standard thermal profile for archaeal amoA gene amplification consisted of 10 min at 95°C and
then 35 cycles consisting of 30 s at 95°C (denaturation), 30 s 54°C (annealing), and 45 s at 72°C
(extension); and a final extension at 72°C for 10 min (Bernhard 2007). A similar profile was
used for bacterial amoA gene amplification but with 42 cycles and an annealing temperature at
57°C (Benrhard 2007, Rotthauwe et al. 1997). A separate PCR cycle was run for archaeal amoA
gene for a total of 35 cycles but stopped at cycle 15, 20, 25 and 30 to plot difference in gene
abundance to determine an appropriate cycle number to quantify relative abundance of archaeal
amoA gene abundance among the four sites.
Twenty µL of each 16s rRNA gene PCR product was visualized on precast 0.8% agarose
e-gels using E-Gel® Precast Agarose Electrophoresis System. Five µL of bacterial and
archaeal amoA gene PCR product was visualized on 1.5% agarose gels containing ethidium
bromide solution. Relative abundance of bacterial and archaeal amoA was quantified using the
imaging processing and analysis software ImageJ (Rasband, W.S., ImageJ, U. S. National
Institutes of Health, Bethesda, Maryland, USA).
Results
Percent Organic Matter and Potential Nitrification Rates
The four sites, Little Pond, Eel Pond, Quissett Harbor and Waquoit Bay, differed in
bottom water oxygen concentrations according to the Massachusetts Estuary Project suggesting a
eutrophic gradient (Table 1). No data was present for Eel Pond, but given its past history, it was
assumed that the bottom water oxygen concentration was lower than both Quissett Harbor and
Waquoit Bay. Percent organic matter ranged from 16% in Waquoit Bay to 24% in Little Pond
(Fig 2). Higher organic matter present reflects oxygen depleted sediment and more anaerobic
conditions. Using bottom water oxygen data, percent organic matter, and descriptions of
sediment the four locations were placed along a eutrophication gradient from most eutrophic to
least eutrophic: Little Pond, Eel Pond, Quissett Harbor, and Waquoit Bay.
Nifrication rates in the sediments for the control ranged from 3 nmol nitrate dry weight of
sediment-1 hour-1 to 9 nmol nitrate dry weight of sediment-1 hour-1 and 3 nmol nitrate dry weight
of sediment-1 hour-1 to 10 nmol nitrate dry weight of sediment-1 hour-1 for the treatment with the
inhibitor. In general, nitrification rates for the control treatment were higher in the less eutrophic
sites, Quissett Harbor and Waquoit Bay, than the eutrophic sites, Little Pond and Eel Pond (Fig
1). In the control, Little Pond sediment had the lowest potential nitrification rates where oxygen
concentration was the lowest and organic matter was the highest. Similarly, sediment with the
inhibitor had the highest potential nitrification rates from Quissett Harbor but had higher
potential nitrification rates from Little Pond than Waquoit Bay, despite being more eutrophic.
However, no trends were detected for the treatment with inhibitor when comparing potential
nitrification rates along the eutrophication gradient. There was no major difference in potential
nitrification rates between the control and the treatment with the bacterial inhibitor at Eel Pond
and Quissett Harbor. However, there was a slight difference in potential nitrification rates
between the control and inhibitor treatment from Little Pond and Waquoit Bay.
Gene Amplification
Before the bacterial and archaeal amoA gene was amplified, all sediment samples were
tested for presence of bacteria and archaea with the universally conserved 16s rRNA gene. Both
bacterial and archaeal 16s rRNA genes were present and successfully amplified in sediment
samples for all four sites (Fig 2 and 3). The bacterial 16s rRNA gene produced a PCR product of
1450 base pairs and the achaeal 16s rRNA gene produced a PCR product with 950 base pairs.
Archaeal amoA gene was successfully amplified with the expected 600 base pair target (Fig 4).
Band intensity varied over the four sites with Little Pond having the lowest intesity for the
archaeal amoA gene and Waquoit Bay having the highest intensity for the archaeal amoA gene.
The presence of the bacterial amoA gene was not clearly detected at 35 cycles. However,
increasing the PCR cycle number to 42 resulted in positive results for presence of the bacterial
amoA gene in both sediments collected initially and sediments collected after nitrification
incubation among the four sites. Unlike the archaeal amoA gene, the bacterial amoA gene
yielded two bands from the PCR product (Fig 5). The two DNA fragments were appoximately
500 base pairs and 250 base pairs. The intensity of the two bands differed indivially and across
the four sites.
Relative amoA gene Abundance
The relative abundance of the amoA gene was determined in relation to the relative band
intensty of the PCR product in the sediment samples. Due to the double banded pattern found for
the bacterial amoA gene, the first band was quantified for relative gene abundance as the 500
base pair product was closer to the expected 491 base pair product noted from Rotthauwe et al.
(1997). There was no apparent pattern of bacterial amoA relative gene abundance for sediment
samples that were initially collected, but it was lowest within sediment collected at Little Pond.
In general, the relative gene abundance for the bacterial amoA gene was highest in sediment that
was collected after 72 hours of nitrification incubation with the added bacterial inhibitor and
increased as sites decreased in eutrophication (Fig 6.) However, since there were major
differences in relative abundance at each site between sediment samples, and no clear pattern
among the four sites, the contribution of ammonia-oxidizing bacteria to nitrification remained
unclear.
The archaeal amoA gene was easily detected but did not show clear difference in band
intensities among each site or between sediment samples initially collected and after nitrification
incubation with 35 PCR cycles (Fig 5). By varying the number of PCR cycles on sediment
samples from Little Pond and Quissett Harbor, it was found that a noteable difference in
abundance of the archaeal amoA gene was detected with 30 PCR cycles (Fig 7). It was
determined that 32 PCR cycles was an approriate number to quantify relative abundance of the
archaeal amoA gene. All sediment samples were reamplifed for archaeal amoA gene with 32
PCR cycles, which showed a greater contrast in band intenstity between sediment samples and
locations (Fig 5). All four sites had the lowest relative abundance of the archaeal amoA gene in
sediment initially collected and highest relative abundance in sediment after the 72 hour
nitrification incubation (Fig 8). Relative abundance was similar between sediment samples taken
at 72 hour nitrification incubation with and without inhibitor. In general, relative abundance for
the archaeal amoA gene increased along the eutrophic gradient as oxygen concentrations
increased. The relative abundance was nearly four times greater within sediment samples
collected from Waquoit Bay, the least eutrophic site, than Little Pond, the most eutrophic site.
Discussion
Nitrification inhibition with the ammonia oxidizing bacterial inhibitor
There was no strong evidence that the bacterial inhibitor, 1-octyne, blocked bacterial
ammonia-oxidizing nitrification activity in estuarine sediments. This directly contradicts
previous studies by Taylor et al. (2013) who found that bacterial amoA was quickly inactivated
with 1 uM of octyne after 2 hours of exposure. Similarly, Lu et al. (2015) were successfully able
to discriminate soil nitrification activity between ammonia oxidizing bacteria and archaea using
4 uM of octyne. However, studies using octyne to inhibit ammonia-oxidizing bacterial activity
have only been performed with ammonia-oxidizing bacteria found in soils. There have been no
studies performed using 1-octyne to differentiate nitrification activity between ammoniaoxidizing bacteria and archaea in estuarine sediments.
The lack of inhibition by 1-octyne on ammonia-oxidizing bacterial activity in the
sediments may have several possible explanations. It may be that the abundance of the bacterial
amoA gene in the sediments was relatively low. Although the relative abundance of ammonia
oxidizing bacteria was not clearly determined, it required 10 more PCR cycles to detect bacterial
amoA gene presence than the archaeal amoA gene. This suggests that the bacterial amoA genes
were in lower abundance than the archaeal amoA gene. In a previous study, it was found that the
ammonia-oxidizing archaeal amoA gene is often more abundant in estuarine sediments, as much
as 80 times greater than the ammonia-oxidizing bacterial amoA gene (Caffrey et al. 2007). Thus,
adding 1-ocytne to sediments with relatively low bacterial amoA gene abundance would not
have as much of a significant impact as it would if the bacterial amoA gene was as close to the
abundance level of the archaeal amoA gene. The lack of inhibition may also suggest that
ammonia-oxidizing bacteria are not contributing as much to nitrification in the sediment as
ammonia-oxidizing archara. Interestingly, Caffrey et al. (2007) found that there was no
relationship between the bacterial amoA gene abundance and the measured nitrification rates.
Similar studies have observed that ammonia-oxidizing archaea rather than ammonia-oxidizing
bacteria are largly responsible for much of the nitrification activity in estuarine sediments (Lam
et al. 2007, Beman et al. 2006). This may help explain why there was not a significant difference
in nitrification rates between the control and with inhibitor treatment despite the fact that the
concentraion of 1-octyne was nearly twice that of what has been added to soils to successfully
inhibit ammonia oxidizing bacterial activity.
amoA gene abundance and nitrification
It was observed that the relative abundance of the archaeal amoA gene in Waquoit Bay
was 2 to 4 times more abundant than the other sites. This suggests that ammonia-oxidizing
archaea may dominate sediment nitrification and be more important than ammonia-oxidizing
bacteria at this site. Interestingly, Quisset Harbor had the highest potential nitrification rate but
had the second lowest archaeal amoA gene abundance for all four sites. This suggests that
ammonia-oxidizing archaea are not the main contributer to sediment nitrifcation at Quissett
Harbor. However, this inconsistency may also indicate a lack of correlation between potential
nitrification rates and ammonia-oxidizng archaea abundance. Similarly, there was no pattern
detected with bacterial amoA gene abundance between treatments or in relation to nitrification
rates. Thus, the current understanding of the relationship between nitrification activity and
abundance of ammonia oxidizers may be incomplete.
It was long thought that potential nitrifcation rates should correlate with the abundance of
ammonia oxidizers ( Henriksen 1980). However, the findings in this study and past studies
directly contradict that belief (Caffrey et al., 2007 and Moin et al., 2009). There is great
physiological diversity among both ammonia-oxidizing bacteria and archaea despite having a
similar function in the environment. Both ammonia-oxidizing bacteria and archaea require
ammonium, oxygen, and carbon dioxide, which may lead to potential competition among these
microbes. For example, it has been shown that among different species of ammonia-oxidizing
bacteria, there are differences in substrate affinity, ammonium oxidation rates, and distribution in
relation to oxygen concentrations (Bernhard and Bollmann 2010). Thus, diversity and
distribution of ammonia-oxidizing bacteria and archaea reflect differences in the environment,
which may explain the difference in the relative abundance of the amoA gene and nitrification
rates observed. Additionally, potential nitrification rates may not reflect all nitrifies that are
present. Some ammonia-oxidizing bacteria or archaea may have been present but not active
when sediment samples were collected which would also affect nitrification rates and the
relationship to relative gene abundance. Thus, the extent of how much ammonia-oxidizing
bacteria or archaea contribute to nitrification remains unclear.
Effect of eutrophication on nitrification
Seasonal trends of nitrification rates have been observed with lowest nitrification rates
between the summer months of June-September (Hansen et al. 1981). During these months,
nitrification may be reduced from oxygen limitation and organically rich sediment, which
coincide with the months that eutrophication occurs (Henriksen and Kemp 1988). This suggests
that eutrophication may be an important factor affecting sediment nitrification. Although
sediment was collected in November, nitrification rates were lowest in more eutrophic sites and
higher in less eutrophic sites. It is likely that sediments may remain eutrophic past summer
months as phytoplankton and other organisms sink into the sediment and are decomposed.
Additionally, sediments may remain oxygen depleted as oxygen poorly diffuses through the
water column. Thus, eutrophication may have a lasting effect on sediment nitrification by
keeping nitrification rates low.
Organic loading into the sediments has been found as an important factor regulating
distribution and diversity of ammonia oxidizers and thus, nitrification rates (Ward 2011). A
result of eutrophication is increased organic matter, which limits oxygen and may lower the
distribution of ammonia-oxidizing bacteria and archaea. For example, Urakawa et al. (2006)
found a marked decrease in ammonia-oxidizing bacteria communites in relation to changes in
nutrients and organic inputs from river run-off in eutrophic sediments in Tokyo Bay. Similarly, it
was found that the nitrification rates and relative abundance of ammonia-oxiding archaea were
higher at the least eutrophic site, Waquoit Bay, than the most eutrophic site, Little Pond. Little
Pond has approximately 8% more organic matter in the sediment than Waquoit Bay suggesting
eutrophication potentially decreases distrubtion of ammonia oxidizers which directly impacts
nitrification.
Future Studies
The contribution of ammonia-oxidizing bacteria and archaea to nitrification in estuarine
sediments remains unclear. Future studies should be conducted to better determine the
contribution of ammonia-oxidizing bacteria and archaea to nitrification in estuarine sediments to
understand their role in the environment. Additionally, potential nitrification rates should be
measured during the summer when primary production and eutrophication are highest to
compare to autumn nitrification rates. A comparison study may reveal a better understanding of
how environmental changes from eutrophication affect nitrification. In this experiment, PCR was
used to test for the presence of the bacterial and archaeal amoA gene, but real time quantitative
PCR should be used to better quantify amoA gene abundance in the sediment. This may improve
the understanding of amoA gene abundance in relation to nitrification rates. Additional studies
should also look at bacterial and archaeal amoA gene abundance in relation to actual amoA gene
activity.
Conclusion
As areas around estuaries become more urbanized the amount of nutrient runoff from
anthropogenic sources increases. The risk of eutrophication and environmental changes brought
about from eutrophication may also be on the rise. Results and previous studies indicate that
eutrophication may affect nitrfication by lowering rates. This could have significant impacts on
important metabolic processes within the sediments, most noteably the coupled nitrificationdenitrifcation process. As ammonium levels increase in the water from eutrophication, nitrifiers
may be limited by oxygen concentrations and can not convert ammonium to nitrate at a high rate.
This in return may decouple the nitrification-denitrification process and ultimately reduce the
amount of nitrogen removed from the estuary. Thus, high amounts of ammonium may remain in
the waters of the estuaries which may become harmful to aquatic organisms. However, it is clear
that our understanding of ammonia oxidizing bacteria and archaea in relation to nitrification is
still not complete, but it remains important to understand how these nitrifiers respond to
environmental changes.
Acknowledgements
I would like to thank the Marine Biological Station and the Semester in Environmental
Science (SES) program for funding this project. I would also like to thank Julie Huber and Anne
Giblin for mentoring and supporting my project. Thank you to Emily Reddington, the Bay Paul
Center, and Anne Bernhard for laboratory space, equipment, and primers for DNA amplification.
Lastly, thank you to my fellow SES students and teaching assistants for help in the field and data
analysis.
Tables and Figures
Table 1. Bottom water oxygen concentrations versus site. Eel Pond bottom water oxygen
concentration data unknown. Data obtained from the Massachusetts Estuary Project.
Site
Bottom water oxygen concentration (mg/L)
Little Pond
Eel Pond
Quissett Harbor
Waquoit Bay
0--4
-4-6
>5
Table 2. Percent organic matter found in sediments.
Site
Percent organic matter (%)
Little Pond
24.2
Eel Pond
Quissett Harbor
Waquoit Bay
23.0
17.6
15.9
14
Nitrificiation Rate
(nmol nitrate gr dry weight -1hr-1)
12
10
8
Control
With Inhibitor
6
4
2
0
Little Pond
Eel Pond
Quissett Harbor
Waquoit Bay
Figure 1. Potential nitrification rates across eutrophication gradient. (Little Pond, most eutrophic,
Waquoit Bay, least eutrophic). Standard error given with error bars.
4
Quissett Harbor
Initial Collection
5
6
Eel Pond
Initial Collection
7
8
- Control
Little Pond
Initial Collection
3
- Control
2
+ Control
1
8
1
2
Waquoit Bay
Initial Collection
3
4
Waquoit Bay
72 hr control
Collection
5
6
Waquoit Bay
72 hr bacterial
inhibitor Collection
Figure 2. Amplification of bacterial 16s rRNA gene. Samples from initial collection of
sediment from Little Pond, Eel Pond, Quissett Harbor, and Waquoit Bay and sediment
from 72 hour post nitrification incubation for Waquoit Bay. Band product 1450 base
pairs.
1
Little Pond
Initial Collection
1
3
2
2
Waquoit Bay
Initial Collection
4
Quissett Harbor
Initial Collection
3
Waquoit Bay
72 hr control
Collection
4
5
6
Eel Pond
Initial Collection
5
6
Waquoit Bay
72 hr bacterial
inhibitor Collection
Figure 3. Amplification of archaeal 16s rRNA gene. Samples from initial collection of
sediment from Little Pond, Eel Pond, Quissett Harbor, and Waquoit Bay and sediment from 72
hour post nitrification incubation for Waquoit Bay. Band product 950 base pairs.
Eel Pond
+72 hr
72 hr
+168 hr
+72 hr
72 hr
Init. hr
cycles
Quissett Harbor
+72 hr
72 hr
Init. hr
cycles
+72 hr
72 hr
Init. hr
cycles
Init. hr
cycles
Little Pond
Waquoit Bay
Figure 4. Bacterial amoA amplification across the four sites. Init. represents amplification from
sediment initially collected, 72 hr from sediment incubated for nitrification for 72 hours and +72
hr from sediment incubated for nitrification with bacterial inhibitor. Target band product ~500 base
pairs.
72 hr 35c
+72 hr 32c
+72 hr 35c
+ Control
- Control
72 hr 35c
+72 hr 32c
+72 hr 35c
+ Control
- Control
72 hr 32c
+72 hr 35c
+72 hr 32c
72 hr 35c
72 hr 32c
72 hr 32c
+72 hr 35c
+72 hr 32c
72 hr 35c
72 hr 32c
Waquoit Bay
Init. 32c
cycles
Init. 35c
Init. 32c
cycles
Init. 35c
Init. 32c
cycles
Init. 35c
Eel Pond
Init. 32c
cycles
Init. 35c
Quissett Harbor
Little Pond
Figure 5. Archaeal amoA amplification across the four sites. Init. represents amplification from
sediment initially collected, 72 hour from sediment incubated for nitrification for 72 hours and +72
hr from sediment incubated for nitrification with bacterial inhibitor. 32c and 35c reference gene
amplification after either 32 PCR cycles or 35 PCR cycles. Band product ~600 base pairs.
3000
Relative Abundance
2500
2000
Initial
1500
72 hour incubation
1000
72 hour incubation with
inhibitor
500
0
Little Pond
Eel Pond
Quissett Harbor
Waquoit Bay
Figure 6. Relative bacterial amoA abundance across a eutrophic gradient. Relative abundance inreference to relative
band intensity analyzed by the software ImageJ. Abundance determined fromsediment initially collected and from
sediment after 72 hours of incubation for nitrification with and without bacterial inhibitor.
18000
16000
Relative abundance
14000
12000
10000
Little Pond 72A
8000
Quissett Harbor 72A
6000
4000
2000
0
10
15
20
25
30
35
40
Cycle Number
Figure 7. Archaeal amoA relative abundance versus PCR cycle number. AmoA amplified from sediment
samples taken from Little Pond and Quissett Harbor after 72 hours of nitrification incubation.
3000
Relative Abundance
2500
2000
Initial
1500
72 hour incubation
1000
72 hour incubation with
inhibitor
500
0
Little Pond
Eel Pond
Quissett
Harbor
Waquoit Bay
Figure 8. Relative bacterial amoA abundance across a eutrophic gradient. Relative abundance in reference to
relative band intensity at 32 PCR cycles analyzed by the software ImageJ. Abundance determined from
sediment initially collected and from sediment after 72 hours of incubation for nitrification with and without
bacterial inhibitor.
3000
Relative Abundance
2500
2000
Initial
1500
72 hour incubation
1000
72 hour incubation with
inhibitor
500
0
Little Pond
Eel Pond
Quissett
Harbor
Waquoit Bay
Figure 8. Relative archaeal amoA abundance across a eutrophic gradient. Relative abundance in reference to
relative band intensity analyzed by the software ImageJ. Abundance determined from sediment initially
collected and from sediment after 72 hours of incubation for nitrification with and without bacterial inhibitor.
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