Protocol Handbook for
NICE
Nitrogen Cycling In Estuaries
A project under the EU research programme:
Marine Science and Technology (MAST III)
Protocol handbook for NICE - Nitrogen Cycling in Estuaries
A project under the EU research programme: Marine Science
and Technology (MAST III)
Tage Dalsgaard (ed.), Lars Peter Nielsen, Vanda Brotas, Pierluigi Viaroli, Graham
Underwood, Dave Nedwell, Kristina Sundbäck, Søren Rysgaard, Alison Miles, Marco
Bartoli, Liangfeng Dong, Daniel C. O. Thornton, Lars D. M. Ottosen, Giuseppe Castaldelli,
Nils Risgaard-Petersen
Protocol handbook for NICE - Nitrogen Cycling in Estuaries
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Authors: Tage Dalsgaard (ed.), Lars Peter Nielsen , Vanda Brotas , Pierluigi Viaroli , Graham
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J. C. Underwood , David B. Nedwell , Kristina Sundbäck , Søren Rysgaard , Alison Miles ,
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Marco Bartoli , Liangfeng Dong , Daniel C. O. Thornton , Lars D. M. Ottosen , Giuseppe
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Castaldelli , Nils Risgaard-Petersen .
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Department of Lake and Estuarine Ecology
National Environmental Research Institute
Vejlsøvej 25, PO Box 314
DK-8600 Silkeborg
Denmark
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Department of Microbial Ecology
University of Aarhus
Ny Munkegade, Building 540
DK-8000 Aarhus C
Denmark
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Instituto de Oceanografia
Faculdade de Ciencias,
Campo Grande
1749-016 Lisboa
Portugal
Dipartimento di Scienze Ambientali,
University of Parma,
Parco area delle Scienze,
43100, Parma.
Italy.
Department of Biological Sciences,
John Tabor Laboratories,
University of Essex,
Colchester, Essex. CO4 3 SQ
England
Department of Marine Botany
University of Göteborg
Box 461
SE-405 30 Göteborg
Sweden
Publisher:
Ministry of Environment and Energy
National Environmental Research Institute, Denmark ©
Department of Lake and Estuarine Ecology
Date of Publication:
March 2000
Layout:
Pia Nygaard Christensen
Please cite as:
Dalsgaard, T. (ed.), Nielsen, L.P., Brotas, V., Viaroli, P., Underwood,
G., Nedwell, D. B., Sundbäck, K., Rysgaard, S., Miles, A., Bartoli, M.,
Dong, L., Thornton, D.C.O., Ottosen, L.D.M., Castaldelli, G. &
Risgaard-Petersen, N. (2000): Protocol handbook for NICE - Nitrogen
Cycling in Estuaries: a project under the EU research programme:
Marine Science and Technology (MAST III). National Environmental
Research Institute, Silkeborg, Denmark. 62 pp.
Reproduction is permitted, provided the source is explicitly
acknowledged.
ISBN:
87-7772-535-2
Price:
100 DKr
Paper quality:
Printed by:
Multi Art Silk
Silkeborg Bogtryk, Silkeborg, Denmark
EMAS registration number DK-S-0084
Number of pages:
62 pages
Circulation:
500
Funding:
This handbook has been prepared in the framework of the project:
Nitrogen Cycling in Estuaries - NICE. We acknowledge the support
from the EU research programme "Preserving the Ecosystem" under
contract MAS3-CT96-0048. This is ELOISE contribution number 132.
Can be obtained at:
National Environmental Research Institute
Department of Lake and Estuarine Ecology
Vejlsøvej 25, PO Box 314
DK-8600 Silkeborg
Denmark
Fax: +45 89 20 14 14
Att: Tage Dalsgaard
E-mail: [email protected]
European Commission
Research Directorate-General
Directorate D.I - Preserving the ecosystem I
Marine ecosystems, infrastructure
Rue de la Loi 200
B-1049 Bruxelles
Belgique
Fax.: +32 2 296 3024
Att: Elisabeth Lipiatou
E-mail: [email protected]
Internet:
The handbook can be downloaded in PDF format at:
http://www.dmu.dk/LakeandEstuarineEcology/nice/
Introduction
This protocol handbook has been prepared in order to standardise sampling, analysis
and calculations within the project "Nitrogen Cycling in Estuaries" (NICE). The
protocol handbook is based on a workshop held from 30 September to 9 October,
1996 at the Aarhus University, Marine Biology Field Station in Rønbjerg, Denmark.
The project aimed at investigating benthic nitrogen cycling in shallow coastal
European waters with special emphasis on nitrogen removal via denitrification. In
order to elucidate the regulating mechanisms for denitrification on a European scale
a number of other variables and processes were also measured. Two of the major
variables on a European scale affecting nitrogen cycling were expected to be tidal
amplitude and climate and sites were selected to represent these differences.
The sediments of shallow waters are very often inhabited by benthic primary
producers and previous studies indicated that they have the capacity to control the
sediment nitrogen cycling. One of the major aims of the project was therefore to
investigate the role of benthic primary producers on the sediment nitrogen cycle.
The benthic primary producers were divided into three functional groups: the
benthic microalgae, the floating macroalgae and the rooted macrophytes. The
experimental procedures needed to measure nitrogen cycling and associated
variables in these three groups of primary producers are very different. Therefore
there is a chapter for each of the groups detailing the experimental procedures. The
chemical analysis and calculations are the same for all the groups and they are dealt
with in the next chapters.
We hope that this handbook can help to standardise methodologies among scientists
already working in this field and help to introduce new scientists to the field.
More detailed information about the project can be obtained at:
www.dmu.dk/LakeandEstuarineEcology/NICE
Contents
1
General incubation conditions .................................................. 9
1.1
1.2
1.3
1.4
2
Handling of sediment cores and flux chambers.............................9
Replication ........................................................................................9
Preincubation of sediment samples with primary producers ......9
Light ................................................................................................10
Microalgae.................................................................................. 11
2.1
2.2
2.3
Summary of activities ..................................................................11
General experimental set-up .......................................................12
Primary production ......................................................................12
2.3.1
2.3.2
2.4
2.5
Nitrogen flux measurement ..........................................................13
Denitrification measurement........................................................14
2.5.1
2.6
Extraction of samples for N2 ............................................................ 14
Additional variables ......................................................................15
2.6.1
2.6.2
3
Method 1: Oxygen flux .................................................................... 12
Method 2: Microelectrode profiles ................................................... 12
Species composition ......................................................................... 15
Microalgal biomass .......................................................................... 16
Macro algae................................................................................ 19
3.1
3.2
3.3
3.4
3.5
3.6
Summary of experiments..............................................................19
General experimental set-up ........................................................20
Obtaining sediment samples.........................................................21
Primary production ......................................................................22
Flux measurements .......................................................................23
Denitrification................................................................................24
3.6.1
3.6.2
3.7
Addition of 15NO3- ............................................................................ 24
Extraction of samples for N2 ............................................................ 24
Additional variables ......................................................................25
3.7.1
3.7.2
3.7.3
Species composition ......................................................................... 25
Biomass and C/N content of macroalgae.......................................... 25
Biomass and C/N content of microalgae .......................................... 25
4
Rooted macrophytes ................................................................... 27
4.1
4.2
4.3
4.4
4.5
4.6
Summary of experiments..............................................................27
General experimental set-up ........................................................28
Obtaining sediment samples.........................................................30
Primary production ......................................................................30
Flux measurements .......................................................................32
Denitrification................................................................................32
4.6.1
4.6.2
4.7
Additional variables ......................................................................33
4.7.1
4.7.2
4.7.3
5
Biomass ............................................................................................ 33
Macrophyte content of nitrogen ....................................................... 34
Sediment content of nitrogen............................................................ 34
Chemical analysis ...................................................................... 35
5.1
Nutrients ........................................................................................35
5.1.1
5.1.2
5.1.3
5.1.4
5.1.5
5.1.6
5.2
5.3
Nitrate............................................................................................... 35
Nitrite ............................................................................................... 36
Ammonia.......................................................................................... 36
Urea .................................................................................................. 36
Phosphate ......................................................................................... 36
Relative abundance of 15NO3- ........................................................... 36
Chlorophyll a .................................................................................37
Gases...............................................................................................37
5.3.1
5.3.2
5.3.3
5.3.4
6
Denitrification associated with the sediment surface ....................... 32
Denitrification associated with the rhizosphere................................ 32
Oxygen concentration by the Winkler technique ............................. 37
Oxygen by the microelectrode method............................................. 38
Isotopic composition of N2 ............................................................... 39
Nitrous oxide .................................................................................... 40
Calculations................................................................................ 43
6.1
6.2
6.3
6.4
Sediment-water fluxes...................................................................43
Chlorophyll a concentrations .......................................................43
Oxygen penetration depth ............................................................44
Fluxes of O2 from the photosynthetic zone .................................44
6.5
Denitrification................................................................................45
6.5.1
6.5.2
6.5.3
6.6
Incubation time and -types ............................................................... 45
Calculating denitrification ................................................................ 47
Rhizosphere associated denitrification ............................................. 49
Leaf marking technique................................................................50
7
Infauna densities........................................................................ 51
8
Sediment characteristics ............................................................ 53
8.1
8.2
8.3
9
Density............................................................................................53
Porosity ..........................................................................................53
Grain size distribution ..................................................................53
Frequency of measurements...................................................... 55
9.1
9.2
9.3
10
Microphytes ...................................................................................55
Rooted macrophytes......................................................................56
Floating macroalgae......................................................................57
References ............................................................................... 59
Index................................................................................................... 61
Chapter 1
1 General incubation conditions
1.1
Handling of sediment cores and flux chambers
All handling of sediment cores, flux chambers, site water and water samples in the
laboratory must be done wearing clean gloves. Never put your bare hands into
incubation water, or ammonia and urea concentrations will be overestimated.
Handling of cores and site water must be done at a temperature close to that in situ.
This is especially critical in the winter time as cores handled at room temperature
very rapidly warm up. After such a warming the metabolic processes never return to
the in situ level.
1.2
Replication
Measurements of fluxes and denitrification are carried out on a minimum of 3
parallel cores or flux chambers in light and in darkness.
1.3
Preincubation of sediment samples with primary producers
After sampling the cores are brought back to the laboratory where they are left over
night and the incubations are started the following day. All sediment cores, whether
containing benthic microphytes, rooted macrophytes or floating macroalgae must
upon return to the laboratory be submersed in site water at in situ O2 concentration
and temperature. The top of the cores must be open and the stirring system must be
on. All cores (both those intended for light and dark experiments) must be exposed
to light until sunset on the day of sampling. The light level and source is identical to
that used during the light incubation of cores the following day (section 1.4).
For sediments with benthic microphytes or rooted macrophytes the cores are
exposed to light for 1 hour and 4 hours respectively, before the actual measurements
are started.
9
Introduction
Atmospheric air used to maintain the correct oxygen concentration can be bubbled
through pure water to remove ammonia prior to being bubbled through the site
water.
One hour before closing the cores for measurement of rates the water above the
sediment is replaced with additional site water, collected at sampling, at in situ
oxygen concentration and temperature. If the oxygen consumption in the water is
suspected to be high enough to lower the oxygen concentration to below half of air
saturation the water should be bubbled when stored overnight. It is important to
shake the containers with the site water before using it to bring any sedimented
matter back into suspension. The activity of filter feeding infauna is very much
dependant on the level of suspended matter in the water. In order to realistically
include the activity of infauna in the rate measurements the level of suspended
matter must be close to in situ levels.
1.4
Light
For incubations requiring light the level of photosynthetically available radiation
(P.A.R.) used is the mean daily P.A.R. level reaching the benthic primary producers
for that month, determined as the average irradiation for the last 3 years data and the
extinction coefficient on the day of sampling. The mean is calculated from sunrise to
sunset. The light level at the sediment surface is calculated as:
Iz = I0 x e -kz
where:
Iz = light level at the sediment surface
I0 = light level at the water surface
(from meteorological data)
z = water depth (m)
k = extinction coefficient
Halogen or metal halide lamps (e.g. Osram Powerstar HQI-T 400W/D) are used as
light source. Cooling may be provided by fans or water. Cooled water is circulated
through a Plexiglas or glass tray positioned between light source and cores.
10
Chapter 2
2
Microalgae
2.1
Summary of activities
Sampling strategy
Fluxes of oxygen, nitrous oxide and nutrients in light and dark as well as
measurements of microalgal biomass must be carried out on the same set of cores,
in order to give paired data of light and dark fluxes and biomass. Two other sets of
cores are then needed for denitrification 1 for light and 1 for dark measurements.
Microprofiles of oxygen and microalgal species composition are measured on a
different set of cores.
General experimental set up
Cores
Lids
Light source
Light intensity
Stirring
Sediment samples
Sampling technique
Transportation
Treatment of cores in the lab
Sampling to measurement time
Primary production
Flux measurement
Denitrification measurement
Transparent Plexiglas tubes, i.d. = 8 cm
Transparent floating lids or transparent fixed
lids which allow for sampling
Halogen or metal halide lamps
Monthly in situ average (see section 1.4)
Central rotating Teflon coated magnet driven
by external rotating magnet
3 cores from each station for light and 3 for
dark. Same cores for flux and denitrification
measurements
By hand
Cores filled and stoppered. Darkness at in situ
temperature or slightly below.
Cores are left open submersed in site water
kept at in situ temperature and O2
concentration with the stirring turned on. See
also section 1.
Measurements are started on the day after
sampling
Measured as the rate of change in O2
concentration with time.
Measured as the rate of change in
concentration with time at the time this is
linear
Isotope pairing technique
11
Chapter 2
2.2
General experimental set-up
Cores are made of Plexiglas tubing with 8 cm internal diameter and a wall thickness
of 0.5 cm. The working depth above the sediment should be 10-20 cm, to give a
volume of 0.5-1 l. Stirring in the cores is created by a 4 cm long Teflon coated
magnetic stirring bar suspended 6 cm above the sediment surface. An external
rotating magnet drives the stirring bars inside the cores. Lids are made of either
Plexiglas or Lexan. The lids can be made to float by gluing a small glass petri disk
to the lower side.
2.3
Primary production
2.3.1 Method 1: Oxygen flux
The sediment-water oxygen flux is measured by closing the sediment cores and
measuring the change in oxygen concentration in the water overlying the sediment
with time. Oxygen concentration is measured by the Winkler technique as described
in section 5.3.1.
At each incubation a blank core tube with only site water is used as a control for
water column production/consumption. The blank core is sampled exactly like the
sediment containing cores. The production/consumption rate of O2 per volume of
water can be calculated and the fluxes measured in the sediment containing cores
can be corrected.
On intertidal flats where there is pronounced diatom migration measurements in the
light should be made in the two hours surrounding low tide.
Primary production
Benthic respiration = dark O2 flux
Benthic primary production = light O2 flux - Dark O2 flux
Benthic net primary production = light O2 flux
Efflux is positive; uptake is negative.
2.3.2 Method 2: Microelectrode profiles
Cores are illuminated (see section 1.4) for 1 hour prior to the start of measurements.
For subtidal systems, the overlying water is bubbled with air mixed with the amount
of N2 required to maintain the in situ O2 concentration. Intertidal cores should be
water saturated but have no overlying water present. For intertidal sediments, the
measurement period must fall within the natural low water period due to the tidal
migration of the microphytobenthos.
The cores should be placed in a set-up giving the microelectrodes easy access to the
sediment surface and maintaining the cores at in situ oxygen concentration and
temperature.
12
Microalgae
Microprofiles can be measured on other cores than those used for flux and
denitrification measurements. The dimension of these cores can be different from
the 8 cm cores.
Measurement
The oxygen microelectrodes (Revsbech & Jørgensen, 1986) must be calibrated (see
section 5.3.2) and give a stable reading in air saturated water before measurements
can be undertaken. Microelectrodes are then introduced stepwise into the sediment
from above with the aid of a micromanipulator. Steps of 50-100 Pm should be made
until a constant low value indicates that the anoxic part of the sediment has been
reached (this can also be compared to the zero obtained from nitrogen bubbled
water). At each depth the current/voltage should be recorded; either logged to a
computer, read from a picoammeter or recorded to a strip chart recorder. The speed
of sampling at any one depth will depend on the response time of the individual
electrode. A constant current must be obtained before the oxygen concentration can
be recorded for a given depth.
Net primary production
From the oxygen concentration profiles measured, net primary production in the
sediment can be determined. The upwards and the downwards oxygen fluxes are
calculated separately and summed to give the total net primary production rate. The
thickness of the photosynthetic zone can be determined and the mean net primary
production rate per volume of sediment can be calculated (section 6.4).
2.4
Nitrogen flux measurement
The exchange rate of nutrients and N2O between sediment and water is measured the
same way as the oxygen flux. The measurements can be carried out in parallel with
the oxygen flux measurements; that is on the same cores at the same time. Every
time a blank core tube with only site water is used as a control for water column
production/consumption (see 2.3.1).
Sampling
Initial water samples (time zero) are taken from the circulated water surrounding the
cores as the cores are closed. Water samples are collected using a clean plastic
syringe which has not been in contact with 15NO3- enrichment experiments. Water
samples are filtered through a glass fibre (GF/C) or cellulose acetate filter into
plastic vials and immediately frozen (-18 qC) (section 5.1). Samples for N2O are
transferred to vacutainers and preserved and stored as described in section 5.3.4.
Samples are extracted as a time series to monitor concentration changes within the
cores. The incubation time for light and dark incubated cores is set at the time
required for dark incubated cores to lower the O2 concentration by 10 - 20% of air
saturation. For example if air saturation at the given salinity and temperature is 250
13
Chapter 2
PM O2 the decrease in oxygen concentration during incubation must be 38 - 50 PM.
This will under “normal” conditions ensure that the concentration change with time
can be measured with a reasonable precision.
If the nutrient concentration in the water is very high, the cores may need to be
incubated for longer time to obtain a detectable concentration change. It is then
necessary to bubble the water with a gas mixture to keep the O2 concentration within
the above mentioned limits throughout the incubation.
2.5
Denitrification measurement
Denitrification is measured in both light and darkness using the isotope pairing
technique. Measurements are performed on one set of cores in light and one set in
darkness.
The general set-up is the same as described above for flux measurements.
At the start of the experiment 15NO3- is added to a final concentration of at least 20%
of the oxygen concentration and a final enrichment of at least 30 atom % in the NO3pool. The NO3- concentration is measured before addition of 15NO3- and at the time
the cores are closed in order to calculate the 14N/15N ratio in the NO3- pool. The
15
NO3- will diffuse towards the denitrification zone and after a certain time the flux
of 15NO3- into the denitrification zone and the evolution rate of 15N2 will be constant.
The produced 15N2 can either be extracted as time series (6.5.1.2) or as start-end
incubations (6.5.1.3). To check the assumptions underlying the isotope pairing
technique it is necessary to run a concentration series (6.5.1.4).
The labelled N2 is traditionally sampled by mixing the whole core as described
below (2.5.1). Alternatively this can be done using a subcore technique as described
in 3.6.2.
2.5.1 Extraction of samples for N2
One ml ZnCl2 (50% w/v) is added to the water and the sediment and water is stirred
using a 5 - 10 mm thick rod. It is very important that all porewater is mixed well
with the overlying water so that the labelled N2 that has been produced is homogeneously distributed. Stirring must be gently in order to minimise exchange of
gases between water and atmosphere. When the stirring is completed the core is left
for a short period (< 2 min) to allow the coarser sediment particles to settle out. A
sample is then taken from the upper part of the water column where almost no
sediment is left. Sampling and preservation is described in section 5.3.3.
14
Microalgae
2.6
Additional variables
2.6.1 Species composition
The aim is to get a semi-quantitative measure of the composition (relative abundance) of the microphytobenthic community on each sampling occasion, at each
site. This is done by counting living (fluorescing) algal cells under a microscope.
Preferably an epifluorescence microscope should be used, but a standard research
microscope is adequate for counting the lens-tissue fraction (see below).
The microalgae are assigned to the main taxonomic groups (diatoms, cyanobacteria,
dinoflagellates, coccolithophorids, other flagellates etc.).Dominating species, genera
and/or size groups are also identified.
Safety check
Scan a fresh sediment sample under an epifluorescence microscope. Pay particular
attention to the presence of small epipsammic (attached) algae on sediment particles
(mineral grains, as well as flocs and faecal pellets) and motile flagellates. Time
spent per sample 5-10 minutes
2.6.1.1 Sampling the microphytes
One or a combination of two methods is used depending on the sediment
characteristics and the structure of the community. The lens-tissue technique is
used to sample motile algal cells (“the epipelic fraction”). Ultrasonicated sediment
samples are used for counting attached cells (“the epipsammic fraction”). Check
which method is applicable for the sediment to be studied. For muddy tidal areas,
the lens-tissue method may be adequate. Previous studies have shown that up to
90% of the algal biomass can be sampled by this method (Eaton and Moss 1966,
Jonge 1980). In sandy sediments, the combination of both methods must be used
because a large part of the algal biomass is often attached to sediment particles (e.g.
Sabbe 1993, Sundbäck and Snoeijs1991). Note, naked flagellates are destroyed by
ultrasonication.
Lens-tissue method (epipelic fraction)
x Place two layers (e.g. 2 x 2 cm area) on the surface of “drained” sediment.
x For cores from macrotidal areas, the topmost lens tissue layer is sampled at the
time of the normal low tide. For microtidal sites, the lens tissue is sampled on the
following day approximately at noon.
x Place the pieces of lens tissue in a beaker containing a mixture of filtered
seawater and glutaraldehyde (final concentration 2.5% glutaraldehyde).
x Tear the lens tissue into small pieces using for example a needle and filter the
lens tissue/algal mixture through e.g. bandage to remove the lens tissue fibres.
x Shake the sample and take a few drops with a disposable pipette and place on a
microscope slide.
15
Chapter 2
Dilution of the sample might be necessary. Samples can be saved for later
identification and counting by preserving the lens tissue including algae with
glutaraldehyde.
Ultrasonicated sample (epipsammic fraction)
Take a 0.5 cm deep sample with a cut-off 20-ml disposable syringe.
Place the sample in a small vial (e.g. a scintillation vial or test tube), add some
filtered seawater.
Treat the sample in a ultrasonication bath filled with ice to avoid warming up of the
sample (use for instance a 35 Hz Sonorex bath and sonication times between 6 and
12 minutes). Note! The sonication time needed has to be checked. You should be
able to remove most cells from the sediment particles, but without destroying them.
Use the epifluorescence microscope to check this. Sonication destroys naked
flagellates!
Before pipetting out a sample for relative cell counts, shake the slurry, and let sand
grains sink to the bottom of the vial. Dilution of the sample may be necessary.
Samples saved for later identification and counting should be preserved with
glutaraldehyde before sonication.
Counting of the epipsammic fraction is easier if it can be made using living material,
but if this is not possible, preserve the sample with glutaraldehyde (2.5% final
concentration of sample). This method probably underestimates the number of living
cells, but this should not be a problem when aiming at relative counts.
2.6.1.2. Counting
Count 300 fluorescing cells (or filaments, if filamentous cyanobacteria are present)
and try to group these into major taxonomic groups(see above). Within each
taxonomic group, try to identify the most common species or genera and/or assign
cells to size groups. Time spent per lens tissue sample 15-30 minutes (after some
training) and between 30-60 minutes for the epipsammic fraction.
Note: Fluorescence of chloroplasts fades rapidly (within seconds) if glutaraldehydepreserved samples are counted. This is particularly a problem for sediments in which
small epipsammic algae are dominant.
2.6.2 Microalgal biomass
Microalgal biomass is determined as chlorophyll a and is measured on all cores
used for flux measurement. The method is based on extraction of pigments with 90
% acetone and spectrophotometric determination of chlorophyll a and phaeopigment concentration (Lorenzen, 1967; Lorenzen and Jeffrey, 1980).
Sampling
Sediments should be cored using 20 ml plastic syringes with bevelled ends. The
sediment core should then be subsectioned, using a razor blade, to obtain the top 5
16
Microalgae
mm of the sediment. Sections may need to be pooled to obtain sufficient sediment
chlorophyll to be determined spectrophotometrically.
Samples should be frozen immediately (-18/-20 qC) and then freeze dried within a
few days. All actions must be carried out in the dark. Freeze dried samples must be
stored in the dark and analysed within 1 month. Analysis of chlorophyll a is
described in section 5.2.
It is acknowledged that a single pigment extraction will underestimate the
chlorophyll concentration within a sample. This underestimation may become
serious where high pigment concentrations are present in the sample. Under such
circumstances it is recommended that a series of extractions is carried out to
quantify this underestimation for each sediment type.
17
Chapter 3
3
Macro algae
3.1
Summary of experiments
Sampling strategy
At each site 3 flux chambers are sampled. Fluxes of nutrients, nitrous oxide and
oxygen as well as denitrification and biomass are measured on each chamber.
Measurements are performed in the following order: fluxes of oxygen, nitrous
oxide and nutrients, denitrification and biomass. Flux and denitrification are first
measured in light and then in darkness.
General experimental set up
Flux chambers
Lids
Light source
Light intensity
Stirring
Sediment samples
Sampling technique
Transportation
Treatment of cores in the lab
Sampling to measurement
time
Primary production
Flux measurement
Denitrification measurement
Transparent Plexiglas
Transparent floating lids
Halogen or metal halide lamps
Monthly in situ average (see 1.4)
Water circulation system with external pump
3 flux chambers from each station to be used for
light and dark incubation for both flux and
denitrification
Hand held box corer cuts block of sediment.
Placed undisturbed in flux chamber
Flux chambers totally filled and closed.
Darkness at in situ temperature or slightly below
Flux chambers are left open submersed in site
water kept at in situ temperature and O2 concentration with the stirring turned on
Measurements are started on the day after
sampling
Measured as the rate of change in O2 concentration with time at the time this is linear
Measured as the rate of change in concentration
with time this is linear
Isotope pairing technique
19
Chapter 3
3.2
General experimental set-up
Flux and denitrification measurements on sediments dominated by floating
macroalgae are performed in Plexiglas chambers (Figure 1) in which both sediment,
macroalgal mat and water column are represented.
The base of chamber is 20 x 20 cm on the inside and the working height is 40 cm.
The chamber is made of 5 mm thick Plexiglas. Stirring is created by drawing water
out through a diffuser (Figure 2) placed at one side of the chamber and pumping it
back in at the other side through an identical unit.
The slit in the unit is 3 mm x 19.5 cm and placed 2 cm above the macroalgal mat or
0.5 cm above the sediment surface if macroalgae are not present. As water leaves the
dispersal unit it passes almost horizontally over the macroalgal mat. Water passes
through the slit at the same velocity throughout the whole slit. Water is pumped
through tygon tubing from one slit to the other by a centrifugal pump at a rate of
high enough to keep the water column stirred and so low that flushing the algal mat
is avoided. The length of the tubes is kept at a minimum to minimise exchange of
gases between water and atmosphere. During measurement of change in gas
concentration with time, gas exchange between water and atmosphere is prevented
by a 2 mm thick transparent Lexan plate floating on the water surface. It covers
more than 95% of the water surface. Flotation is provided by a glass petri dish glued
to the lower side of the Lexan plate.
Side view
Front view
Centrifugal
pump
Tygon
tubing
Lexan
lid
Glass petri
dish
Diffusor
40 cm
Algal mat
Sediment
Rubber
stopper
20 cm
20 cm
Figure 1. Flux chamber with the sediment, algal mat and water column.
20
Macroalgae
Front view
Side view
9.5 cm
19.5 cm
0.5 cm
3.3
Figure 2. Diffuser used to circulate
the water in the flux chambers.
Obtaining sediment samples
Sampling is done with a hand held box corer made of 1 mm thick steel plate fitting
precisely inside the flux chamber (Figure 3). The corer can be opened or closed at
the top with two 5 cm rubber stoppers. After pushing the corer 10 to 15 cm into the
sediment it is stoppered, dug out with a shovel and placed carefully on the sediment.
A Plexiglass plate (19 x 19 cm, 2 mm thick) is then used to cut the sediment across
the end of the corer. This plate is left in the opening of the corer and held by hand
when the corer is lifted to prevent the sediment from falling out. The flux chamber is
placed on the sediment next to the corer and a wooden stick is pushed into the
sediment through a hole in the bottom of the flux chamber and extending 10 cm
above the chamber. The corer is placed above the flux chamber resting on the
wooden stick which then holds the Plexiglass plate in place and the flux chamber is
lifted up around the corer. The flux chamber is stoppered in the bottom and the corer
is removed.
Side view
Handle
Stoppers
Bottom view
Plywood
24 x 39 x 1 cm
Handle
Stoppers
Steel
frame
50 cm
1 mm stainless steel plate
Outer dimension: 19.5 x 19.5 cm
Figure 3. Stainless steel box corer
used for sampling intact sediment
blocks and transferring them to the
flux chambers.
21
Chapter 3
When removing the corer a gap of 2 to 3 mm is left between the sediment and flux
chamber walls, however, the sediment block expands horizontally and sinks correspondingly to fill this gap within a few seconds.
Transportation
Chambers with sediment and algal mats are kept at or slightly below in situ
temperature in an insulated box. During transport the chambers are filled with water
and a transportation lid is in place in order to prevent strong water motion which
could disturb the sample. The lid is a Plexiglas plate fitting exactly in between the
two long sides and rests on top of the two short sides of the chamber.
3.4
Primary production
The primary production is measured as the difference between steady state algae water fluxes of oxygen in light and darkness. Oxygen fluxes are measured as change
in oxygen concentration in the water with time with a lid floating on the water
surface which prevents exchange of oxygen between water and air. The
measurements should continue until the change in concentration is linear with time
and the slope of the linear part can be safely determined. Measurements after bubble
formation starts will underestimate the primary production. Oxygen concentration is
measured by the Winkler technique (section 5.3.1) or by O2 electrode.
Concentrations of O2 within a macroalgal mat are different in light and in darkness.
When shifting from light to darkness or vice versa the pool within the mat will
hence change. The O2 flux between algal mat and water does, therefore, not reflect
the actual production/consumption within the mat, until this pool has reached steady
state. This is normally the case when O2 concentration change over time in the water
is linear.
Bubbles are often formed within the mat when illuminated due to the very high
oxygen production rates. Measurement of oxygen concentration only includes
dissolved oxygen and oxygen present in bubbles is therefore not measured and the
oxygen production is underestimated. Furthermore, the presence of bubbles within
the mat when shifting from light to darkness represents a pool of oxygen which is
not measured and oxygen uptake in darkness is underestimated. Bubble formation
can be delayed by lowering the total gas pressure within the water. The procedure
described below only lowers the partial pressure of nitrogen leaving the partial
pressures and concentrations of oxygen and 6CO2 unchanged.
Avoiding bubble formation
A container is filled half with water leaving the volumes of water and headspace
equal. The headspace is flushed with pure O2 until the partial pressure of O2 in the
headspace is 1 atm. The container is closed and water and headspace is allowed to
equilibrate. Equilibration needs to be speeded up by shaking, stirring, by pumping
the water into the headspace or by pumping gas from the headspace through the
water. After equilibrium is obtained the total pressure in the container is lowered to
22
Macroalgae
0.2 atm and the container is again closed and the gas- and water phase are again left
to equilibrate. Equilibration must again be speeded up as mentioned above.
Lowering the pressure in the container to 0.2 atm means that 80% of the gas is taken
out of the headspace and the rest is left to equilibrate. Since most of the CO2 is in the
water its concentration is almost unchanged, whereas most of the N2 is in the
headspace and close to 80% of what was left from equilibration with pure O2 is
removed in this step.
It should be noted that the only gas removed from the water during this treatment, is
that transported into the headspace during equilibration and subsequently removed
when the total pressure is lowered to 0.2 atm.
Example:
Temperature = 5qC, Salinity = 20‰
Concentration in gas phase = Cg
Concentration in water phase = Cw
Equilibrium
constants:
Pure O2 headspace
before equilibrium:
300 PM
42000 PM
After equilibrium
with headspace at 1
atm.:
1510 PM
40790 PM
After equilibrium
with head-space at
0.2 atm.:
302 PM
8158 PM
O2
Cw
Cg
1
28
N2
Cw
Cg
1
54
Cw
Cg
617 PM
0 PM
Cw
Cg
11 PM
606 PM
Cw
Cg
2 PM
121 PM
CO2
Cw
Cg
160
1
Cw
Cg
2000 PM
0 PM
Cw
Cg
1987 PM
13 PM
Cw
Cg
1977 PM
12 PM
3.5
Cw
Cg
Cw
Cg
Cw
Cg
Flux measurements
Concentrations of a given species within a macroalgal mat are different in light and
in darkness. When shifting from light to darkness or vice versa the pool within the
mat will hence change. The flux between algal mat and water does, therefore, not
reflect the actual production/consumption within the mat, until this pool has reached
steady state. This is normally the case when concentration change over time in the
water is linear.
Nitrogen flux
The exchange of NO3- + NO2-, NH4+, urea and N2O between the sediment/algae
system and the water is quantified by monitoring the concentration of these species
over time in the water above the algal mat. The measurements should continue until
the change in concentration is linear with time and the slope of the linear part can be
23
Chapter 3
safely determined. Sampling, preservation and analysis of NO3-, NH4+, urea and N2O
is described is section 5.1 and 5.3.4.
The volume of water in each chamber must be determined either by measuring the
height of the water column or by transferring it to a measuring cylinder. The
dimensions of the algal mat must also be determined.
3.6
Denitrification
Denitrification is measured using a modified version of the isotope pairing
technique. When measuring denitrification in bare sediment and sediment colonised
by microalgae 15NO3- is added only to the water and the added 15NO3- is transported
mainly by diffusion into the sediment and becomes mixed with the 14NO3- already
present and that produced by nitrification within the sediment.
When measuring denitrification in sediments with several cm of macroalgae lying
on the top of the sediment surface it might take longer for the added 15NO3- to reach
the denitrification zone in the anoxic part of the sediment. This is dependant on the
on the water movement within the mat. The denitrification rate can first be
calculated when the production rate of 15N-N2 has reached a constant value. Since
the time this takes is unknown it is necessary to extract samples for N2 several times
during the incubation.
15
-
3.6.1 Addition of NO3
The concentration of 15NO3-must be at least 20% of the oxygen concentration and
the 15NO3- must make up at least 30% of the total NO3- pool. The labelled nitrate is
added to the water column of the flux chamber.
3.6.2 Extraction of samples for N2
Samples for 15N2 are taken at least 4 times during the incubation from each chamber.
For calculating the appropriate incubation time see section 6.5.1. At each sampling
time 3 subsamples are taken with a thin walled steel tube (id = 5-10 mm). The steel
tube is gently pushed through the macroalgal mat and 3 cm into the sediment. The
top is then closed and the tube is gently pulled out and now contains a sub-core with
both water and sediment. All 4 subsamples are transferred to a cylindrical glass
beaker (id < 2 cm) containing 15 Pl of a 50% (w/v) ZnCl2 per millilitre of water and
sediment transferred. The ZnCl2 and samples are immediately but gently stirred to
ensure homogenous distribution of ZnCl2 and 15N2. After the stirring is stopped the
slurry is left for 1 - 2 minutes to allow the more coarse sediment particles to
sediment. The water (still containing some sediment) is then transferred to a
vacutainer as described in section 5.3.3.
24
Macroalgae
There are special problems associated to interpretation of data from experiments
with macroalgae on the sediment surface. Depending on the circulation in the flux
chamber the labelled nitrate may or may not reach the sediment surface. In case the
15
NO3- reaches the sediment surface and the production of labelled N2 proceeds with
a constant rate, there is no problem. However, in the case where no labelled N2 is
produced there might 2 interpretations:
A: Labelled N2 does not accumulate because denitrification does not proceed at a
measurable rate even though 15NO3- reaches the sediment.
B: Labelled N2 fails to accumulate because 15NO3- does not reach the sediment. In
that case it is most probable that there is no denitrification of nitrate from the
water column. However, denitrification of nitrate produced in the sediment
might be taking place and is not registered because 15NO3- fails to reach the
denitrifying zone and 29N2 is not formed.
3.7
Additional variables
3.7.1 Species composition
At each measurement the macroalgal species composition must be determined. If
differences between chambers from the same site exist the species composition for
each chamber must be determined.
3.7.2 Biomass and C/N content of macroalgae
In each chamber the total biomass must be determined as dry weight. If there are
more species in a chamber the species must be separated before drying and the dry
weight of each species must be determined. The algae are dried at 90qC to constant
weight. Before taking out the algae for dry weight determination the dimensions of
the algal mat must be measured.
After drying the macroalgae are homogenised in a mortar and a subsample is
analysed for C and N content on a CHN analyser.
3.7.3 Biomass and C/N content of microalgae
In situations where no macroalgae are present the biomass of benthic microalgae
must be determined as described in section 2.6.2.
25
Chapter 4
4
Rooted macrophytes
4.1
Summary of experiments
Sampling strategy
At each site 6 cores (20 cm diameter) are sampled. On 3 of these fluxes of nutrients,
nitrous oxide and oxygen as well as denitrification are measured in the light and on
the other 3 cores the same variables are measured in the dark. Biomass is measured
on all cores.
General experimental set-up
Cores
Transparent Plexiglas; i.d. 20 cm; height 40 cm
Lids
Opaque or transparent floating lids
Light source
Halogen or metal halide lamps
Intensity
Monthly in situ average (see 1.4)
Stirring
Water circulation system with external pump
Sediment samples
6 cores: 3 for light incubation, 3 for dark
Sampling technique
By hand; Stainless steel sampler
Transportation
In aerated dark box at in situ or lower temperature
Cores maintenance in lab
Open in aerated water bath at in situ temperature
and oxygen concentration and experimental light
level until sunset
Sampling-measurement time
Measurement on the day after sampling
a) 4 hours light or dark pre-incubation; Measured
Primary production
Technique
as change in O2 concentration with time in the
water column in light and darkness.
Change in O2 concentration < ± 20%
Bare sediment O2 production/respiration by 5 cm
i.d. core light and dark incubation
b) Leaf marking in situ
NO3-, NO2-, NH4+, N2O, Urea
Flux measurement
Technique
4 hours light or dark pre-incubation; Measured as
change in concentration with time in the water
column in light and darkness.
Change in O2 concentration < ± 20%
Denitrification measurement a) Isotope pairing for surface denitrification.
Technique
b)15NH4+ perfusion for rhizosphere denitrification
27
Chapter 4
The annual variation in nitrogen cycling in sediments with rooted macrophytes is
investigated. Processes to be measured are the exchange of dissolved nitrogen
compounds between the macrophyte-sediment compartment and the overlying
water, denitrification, N-incorporation and N-retention in the biomass of the rooted
macrophytes and the sediment.
4.2
General experimental set-up
Cores for incubation experiments
Measurements of fluxes, between the sediment-plant system and the water, and
denitrification (associated with the sediment surface) is carried out in Plexiglass
tubes (i.d. 20 cm; height 40 cm). The core tubes are equipped with transparent or
opaque floating lids made of glass or PVC; an adjustable stopper serves as bottom
(Figure 4).
A3
A1
A2
B1
B2
B3
B4
A
B
Figure 4. Core sampling unit (A), with Plexiglas tube inside a stainless steel frame (A2), equipped with a
lid (A1). A valve in the lid (A3) allows water to leave the tube when pushed into the sediment. The
incubation system is shown in B. A pump (B1), connected to perforated acrylic tubes (B3) maintains a
continuous circulation of the water in the incubation tube. The tube is sealed at the top with a
transparent floating lid (B2), made from Plexiglas, and sealed at the bottom with a stopper (B4) made of
an elastic rubber ring squeezed between two layers of PVC with bolts.
28
Rooted macrophytes
Channels
O-ring
Figure 5. The bottom lid for rhizosphere associated nitrification-denitrification measurement is a 3 cm
thick perfusion stopper inserted into the core tube and fixed to the tube with silicone greased o-rings.
The perfusion stopper has concentric channels (0.25 cm wide, 0.5 cm deep), all connected to a radial
channel. The concentric channels are covered by a nylon mesh (mesh size 0.1 mm) preventing
sediment from entering the channels.
For measuring rhizosphere associated denitrification smaller cores (i.d. = 9.5 cm)
with a special bottom stopper (Figure 5) are used. This construction allows replacement of the sediment porewater.
For measurement of oxygen, nitrous oxide and nutrient fluxes across the sedimentwater interface without the influence of macrophytes smaller Plexiglas tubes are
used, i.e. cores similar to those used for incubation of microphytobenthic covered
sediments (section 2).
Stirring
Stirring of the water column in the cores is obtained by a water circulation system,
driven by a centrifugal pump. The flow rate applied must be sufficient to ensure
mixing of the water column and so low that resuspension of the sediment does not
occur. To determine the flow rate to be used, a comparison between oxygen flux
rates in the cores used for microphytobenthic covered sediments (section 2) and the
20 cm diameter cores must be made. The correct flow rate will be the one giving
identical results.
29
Chapter 4
4.3
Obtaining sediment samples
At each station 6 cores (20 cm diameter) are collected for oxygen and N-compounds
flux and denitrification measurements. Three cores are used for light incubation and
3 for dark incubation. The cores are collected by hand, by pushing the Plexiglas
tubes, mounted in a stainless steel coring device, 10-15 cm into the sediment (Figure
4). Before pushing the tube into the sediment the roots below the core walls should
be cut to avoid dragging material deeper into the sediment and to minimise the
disturbance of sediment horizons; for the same reason the leafs of the sampled plants
should be kept inside the core walls by hand.
The cores must be checked immediately after sampling to ensure that both the leaf
and the root systems are not seriously damaged.
Transportation
Sediment cores are placed with open tops in boxes filled with aerated site water and
then immediately transported to the laboratory. Temperature should be kept at the in
situ level. Plants must be kept submersed during the transport to avoid drying of the
leaves.
Pretreatment in the lab
The lower part of the core where the sediment is located is covered on the outside by
black plastic in order to keep the sediment sides in darkness. Larger animals and
fragments of macroalgae on the sediment surface are removed when possible to
minimise variations not directly due to the rooted macrophyte-sediment system.
Preincubation is described in section 1.3.
4.4
Primary production
Primary production is measured in the lab by the O2 flux method and in the field by
the leaf marking technique (Sand-Jensen 1975). Nitrogen incorporation into the
plants is calculated from primary production rates and C/N ratio.
Oxygen flux method
Using the O2 flux method net primary production rates of the rooted macrophytes
are estimated from the difference between O2 flux rates measured in the plantsediment system and rates measured on bare sediment. Oxygen fluxes are measured
in the light and darkness on the plant sediment-system in the 20 cm cores and on
small cores sampled in between the plants. The small cores are sampled within the
eelgrass bed right next to the spot where the 20 cm cores are sampled. Oxygen
fluxes are determined from concentration changes with time in the water column of
the cores with the air-water gas exchange blocked by floating lids. Five water
samples are collected with glass syringe at regular intervals, until the concentration
of O2 in the water column has changed r20% of the initial value. Oxygen
concentration is determined by the Winkler method (section 5.3.1).
30
Rooted macrophytes
Leaf marking technique
Eelgrass (Zostera marina) growth is also quantified by an in situ leaf marking
technique based on the principles of Sand-Jensen (1975). This technique enables
estimation of the average leaf growth rate, and root-rhizome growth. The latter can
be estimated from the plastochrone interval (the period needed for the appearance of
a new leaf) since in Z. marina a root internode corresponds to a leaf produced.
Within the eelgrass bed 24 plants are chosen and marked with a tag so that the same
plants can be found again. All leaves of each plant are then pierced by a hypodermic
needle a 1 cm above the leaf sheet (Figure 6). The plants are then left to grow for the
period needed for one new leaf to be produced and are then harvested. Plastochrone
intervals are estimated to be about 50 days in the winter and 8 days in the summer
(Pedersen and Borum, 1993). The displacement of the marks on leaf 1-3 relative to
that on the non growing leafs 4-5 and the total length of newly formed leaves is
measured. Individual leaves, rhizomes groups and associated roots are dried at 90°C
to constant weight. The different parts of the plants are homogenised and analysed
for C and N content on a CHN-analyser. See section 6.6 for calculations.
3
Leaves
4
5
2
1
Rhizome
Roots
Figure 6. Leaf marking technique. The leaves and rhizome of the plant are numbered according to
increasing age (see text for details).
31
Chapter 4
4.5
Flux measurements
Fluxes between water and sediment/plant of NO3-, NO2-, NH4+, urea and N2O are
measured in the 20 cm diameter cores. The fluxes are measured as concentration
changes in the water with time. Nutrient and N2O flux measurements are carried out
together with the O2 flux measurements simultaneously on the same cores. Water
samples for nutrients and N2O are taken at time intervals during the incubation
period. If a given species is depleted during the incubation only concentration
measurements obtained before depletion can be used for calculating the flux. If the
concentration is very high relative to the flux rate for a given species it might be
necessary to prolong the incubation. In that case the water should be bubbled with a
gas to keep the O2 concentration close to the in situ level throughout the incubation.
The concentrations of the different nitrogen species are determined as described in
section 5.1 and 5.3.4.
The cores are pre-incubated for 4 hours at the light level applied in the experiment.
4.6
Denitrification
4.6.1 Denitrification associated with the sediment surface
The rate of denitrification associated with the sediment surface (and possibly the
plant surface) is measured separately by the 15N-isotope pairing technique (Nielsen,
1992) in light and dark incubated 20 cm diameter cores as described below (for
calculation see section 6.5.2).
The cores are pre-incubated for 4 hours at the light level applied in the experiment
(section 1.4). To estimate the time needed for the 15NO3- profile to reach steady state
penetration depth of O2 into the sediment is measured by O2 microelectrodes
(section 2.3.2). For calculations se sect. 6.5.1. 15NO3- (99 15N atom%) is added to the
water column of each of the cores or to the open reservoir to a final concentration of
at least 20% of the oxygen concentration and a final enrichment of at least 30
atom% in the final NO3- pool. The cores are left open and aerated at in situ O2
concentration until the 15NO3- porewater profile has reached steady state. Hereafter
the core tubes are closed with floating lids and subsamples of water and sediment for
15
N2 analysis is collected at regular time intervals as subcores using 1 cm i.d. steel or
acrylic tubing. Microbial metabolic activity in the subcores is inhibited by injecting
100 µl ZnCl2 (50% w/v) into the sediments in the steel cores. The O2 concentration
should not be allowed to change more than ± 20% from the initial value during the
incubation.
4.6.2 Denitrification associated with the rhizosphere
Coupled nitrification-denitrification activity associated with the rhizosphere of
rooted macrophytes is estimated by a modified version of the isotope pairing
32
Rooted macrophytes
technique. It is assumed that NO3- for denitrification several centimetres below the
sediment surface is produced by nitrification within the sediment. The oxygen
required for nitrification can at this depth not diffuse in from the surface but must be
released by the plant roots. The assay is hence based on addition of 15NH4+ to the
sediment and quantifying the formation of 15N2 from coupled nitrification and
denitrification. Denitrification is recorded with depth and specific denitrification
activity is related to the vertical distribution of roots.
The assay is performed in the 9.5 cm diameter cores with the perfusion bottom
stopper (Figure 5). Initially the porewater of the core is drawn out through the
perfusion bottom stopper and into a 1 l glass bottle by vacuum. The bottle is then
opened and the water is bubbled with N2 to keep it anoxic and 15NH4+ (99 15N atom
%) is added to the water to a final concentration of 500 PM. The 15NH4+ enriched
porewater is pumped back into the rhizosphere through the perfusion bottom stopper
using a peristaltic pump. Before drawing out the porewater the water overlying the
sediment is bubbled with N2, in order to remove O2, from the water that is being
drawn into the sediment. This procedure ensures that 15NH4+ is homogeneously
distributed throughout the rhizosphere.
For this assay samples are only taken at the start and at the end of the incubation.
Initial samples for 15N2 are then collected with syringe and hypodermic needle
trough silicone rubber sealed holes placed at 1 cm vertical distance in the wall of the
core tubes. Samples of 1 ml water are collected in the water column and in the
sediment with depth intervals of 1-2 cm, from the sediment-water interface down to
10 cm below the sediment surface. The samples for 15N2 MS analysis are transferred
to 6 ml He-purged, pre evacuated, glass vials (Exetainer ®, Labco, High Wycombe,
UK) containing 100 µl 50% (w/v) ZnCl2. At the end of the incubation this procedure
is repeated. The incubation period should be the same as for the assay for surface
associated denitrification.
4.7
Additional variables
4.7.1 Biomass
The biomass and shoot density of the macrophytes is measured by harvesting all
plant material in 3 randomly chosen circular plots (diameter = 30 cm). Plants are
cleaned and separated into leaves and root-rhizomes, and subsequently dried to
constant weight at 90°C. These results are compared with results obtained by
measuring biomass and shoot density in the 20 cm diameter cores. Given that
biomass and shoot density obtained with these 2 sampling techniques are not
significantly different, these variables are routinely measured in the 20 cm diameter
cores.
33
Chapter 4
4.7.2 Macrophyte content of nitrogen
The nitrogen and carbon content of the macrophytes are determined using a CHNanalyser. Before analysis the plant material must be separated into the fractions
needed (old/new leaves, roots, rhizomes etc.) and dried to constant weight at 90qC.
4.7.3 Sediment content of nitrogen
Sediment profiles of total N are determined inside the macrophyte stand. Sediment
cores are taken to 10 cm depth in triplicate using Plexiglas tubes (i.d. 5.5 cm). The
cores are frozen to keep root-rhizomes and dead particulate plant material in a fixed
position and subsequently sliced at 1 cm intervals with a thin saw blade. Living
plant material is removed and quantified. The remaining sediment material is dried
at 60 qC and sieved to separate out particular organic material (defined as particles >
1mm). The samples are stored on plastic vials for later analysis of N and C content
on a CHN analyser.
34
Chapter 5
5
Chemical analysis
5.1
Nutrients
All water samples collected in the field are filtered through GF/C or cellulose
acetate filters, immediately frozen in dry ice and transported back to laboratory. If
transport back to the laboratory takes only a couple of hours samples can just be
kept cold and frozen at -18°C immediately after returning to the laboratory. Samples
are stored at -18qC until analysis.
All water samples taken in the laboratory are filtered through GF/C or cellulose
acetate filters and immediately frozen at -18qC and stored until analysis. All
sampling and handling of water column and sediment cores are performed using
clean gloves. Analysis of water samples should be completed before 3 months of
storage.
Water samples are stored in poly styrol, polyethylene or polypropylene test tubes
with stoppers.
For all nutrient analysis, intercalibration standards are run in all laboratories. Sea
water, collected in Denmark, with a salinity of approximately 25 ‰, is filtered,
dispensed into bottles, autoclaved and sent to all laboratories for analysis. The
results from all laboratories will be compared and serve as a control of methodology
and precision.
All standards are prepared with artificial seawater:
100‰ Stock solution:
NaCl
MgSO4,H2O
NaHCO3
83.34 g/l
26.67 g/l
0.12 g/l
This stock-solution is stored in a closed bottle at 4°C until use. For analysis,
standards are prepared on the same salinity as in situ by dilution of the stocksolution with high purity water.
5.1.1 Nitrate
Determined on autoanalyser/flow injection analyser using standard colorimetric
methods (Grasshoff et al. 1983).
35
Chapter 5
5.1.2 Nitrite
Determined on autoanalyser/flow injection analyser using standard colorimetric
methods (Grasshoff et al. 1983).
5.1.3 Ammonia
Determined manually or automatically using standard colorimetric methods (Bower
and Holm-Hansen, 1980). Ammonia samples must be handled very carefully in
order to avoid contamination and gloves must at all times be used when handling
samples or test tubes. To avoid contamination of the test tubes before use, they must
be stoppered immediately after opening a new package. The stoppered test tubes can
then be stored in the lab for a long time without being contaminated.
The time that ammonium samples are exposed to the atmosphere is minimised in
order to prevent contamination of the samples with ammonia from the air.
5.1.4 Urea
Determined manually or automatically using standard colorimetric methods
(diacetyl monoxime method, Price and Harrison, 1987). The use of clean gloves is
necessary to avoid contamination because human hands always give off urea.
5.1.5 Phosphate
Determined manually or automatically using standard colorimetric methods
(Grasshoff et al. 1983).
15
-
5.1.6 Relative abundance of NO3
Determined by one of the following two ways:
1: Determined from the difference in the NO3- concentration before and after
addition of 15NO3-. A water sample for determination of the NO3- concentration
before the addition of 15NO3- is collected. After addition of 15NO3- another water
sample is collected to determine the concentration of 14NO3- + 15NO3-. Allow the
added 15NO3- to become fully mixed with water before taking the second sample.
The relative labelling with 15NO3- in percent of the total NO3- pool is calculated
as:
15
NO3 labelling
>NO3- @after >NO3 @before
>NO3 @after
2: Direct determination of the relative abundance of 15NO3- in the final NO3- pool is
done on water samples as described by Risgaard-Petersen et al. 1993 and
Risgaard-Petersen and Rysgaard 1995. In short, an enrichment culture of
denitrifying bacteria is used to convert all NO3- into N2 gas composed of 28N2,
29
N2 and 30N2, which subsequently is analysed by mass spectrometry as described
in section 5.3.3. The 15N atom % of the NO3- pool is then calculated from the
29
N2 : 30N2 ratio in the analysed gas as:
36
Chemical analysis
15
N atom %
1 1 - (R 2)(f29C - Rf30C )
R2
R=
where:
f29S
f29C
f30S
f30C
=
=
=
=
f29 S f29 C
f30 S f30 C
ratio of 29N2 to total N2 in the sample
ratio of 29N2 to total N2 in a control sample without 15NO3 added
ratio of 30N2 to total N2 in the sample
ratio of 30N2 to total N2 in a control sample without 15NO3- added.
At atom percentages above the background content (atmospheric
0.366%) the equation above can be reduced to:
15
5.2
N atom % =
15
N content =
2
100%,
R+2
Chlorophyll a
Add 4 ml of 90 % acetone to the freeze dried sample in a glass centrifuge tube (this
volume may vary depending on the sensitivity of the spectrophotometer and the
amount of chlorophyll present in the sample) and cover with parafilm. Thoroughly
mix the contents of the centrifuge tube to ensure that all the sediment comes in
contact with the acetone. Stand the samples in a fridge (4 qC) for 24 hours. Samples
should be agitated thoroughly for a second time during this period to aid extraction
of the pigments. Immediately prior to measurement, centrifuge the samples to ensure
an absorption at 750 nm less than 0.005. Decant the supernatant into a glass cuvette
and then measure the extinction of the extractant at 665 nm and 750 nm. Add 2
drops of 10% HCl and remeasure the extinction at 665 nm and 750 nm. As 750 nm
is a measure of the clarity of the sample, samples should be respun if the extinction
is greater than 0.005 (in a 1 cm cuvette).
For calculations see section 6.2.
5.3
Gases
5.3.1
Oxygen concentration by the Winkler technique
Sampling
Water samples are collected using a glass syringe with an attached length of tubing.
Ensure that the syringe is flushed with sample water prior to the actual sampling and
that no bubbles are present during sampling. Each sample should be transferred to a
37
Chapter 5
12 ml gastight vial, ensuring that the sample is introduced to the bottom of the vial
and allowing the water to overflow as the sampling tubing is slowly pulled out. 150
Pl of Winkler’s reagents I and II (Strickland & Parsons, 1972), respectively should
be added immediately and the vial lids tightly closed. The contents of each vial
should be thoroughly mixed by inverting the vial.
Storage
Samples must be stored in a dark, cool environment and analysed within 2 days.
Winkler titration
Water samples are analysed for dissolved oxygen by Winkler titration (Strickland &
Parsons, 1972). Add 300 Pl 80% phosphoric acid to each 12 ml vial. Close and
shake the vial until all precipitate is dissolved. Following acidification of the sample,
5.000 ml of the sample is titrated with 0.010 M thiosulfate.
Calculations for oxygen concentration
6
>O2@= >Thio@ x Thiovol x 10
Samplevol - Reagvol x 4
where:
[O2]
Thiovol
[Thio]
Samplevol
Reagvol
106
4
5.3.2
oxygen concentration (µmol l-1)
volume of Na2S2O3 5H2O titrated (ml)
concentration of Na2S2O3 5H2O used for titration (mol l-1)
volume of sample titrated (ml)
volume of Winkler reagents I & II in the fraction of the sample
titrated (ml)
= factor to convert from mol l-1 to µ mol l -1
= factor needed because 1 mol O2 reacts with 4 mol of Na2S2O3
5H2O.
=
=
=
=
=
Oxygen by the microelectrode method
Calibration of microelectrodes
x 100% calibration (air saturation) in air bubbled water of the same temperature
and salinity.
x 0% calibration in nitrogen bubbled water, of the same temperature and salinity,
or in anoxic sediment.
Calculating O2 concentration
>O 2 @
38
An A0
x O 2sol
A sat A 0
Chemical analysis
where:
[O2]
An
Asat
A0
O2sol
5.3.3
=
=
=
=
=
oxygen concentration at a single depth (µmol l-1)
current at a given depth (nA)
current for air saturation (nA)
current for zero oxygen (nA)
oxygen solubility for the actual temperature and salinity (µmol l-1).
Isotopic composition of N2
Sampling and preservation
Water and slurry samples (15 ml) are collected by glass syringes equipped with a 10
cm long gas tight Tygon® tube. Ensure that the syringe is flushed with sample water
prior to the transfer of the actual sample and that no bubbles are present during
sampling. The water or slurry is transferred to a gas tight vial (12 ml Exetainer,
Labco, High Wycombe, UK). The vial is totally filled excluding air bubbles and
preserved with 250 µl ZnCl2 (50% w/v).
Analysis
N2 and 30N2 are extracted from the water in the Exetainers by introducing a helium
headspace. This is done by inserting a hypodermic needle in line with a helium flask
through the rubber septum of the Exetainer. With a high precision glass syringe 4 ml
water are removed from the Exetainer and simultaneously replaced by an equivalent
volume of He (Figure 7).
29
The vial is then shaken vigorously for 5 min after which more than 98% of the N2
will be in the headspace (Weiss, 1970). The isotopic composition of N2 in the
He
Open end with
He overflow
Figure 7. Introduction of a headspace in a vacutainer. The hypodermic needle of the high precision
glass syringe is inserted into the water, through the rubber septum, and a volume is drawn out. The
water removed is replaced by helium through the hypodermic needle connected to the tubing.
39
Chapter 5
headspace is then determined by mass spectrometry. The Exetainers are placed in an
auto sampler in line with a gas chromatograph and a mass spectrometer. The entire
headspace (4 ml) of the Exetainer is then carried through the GC columns and into
the MS by a flow of helium (99.9995% purity). The sample passes through a drying
tube (10 mm x 200 mm) packed with Mg(ClO4)2 to remove water vapour and
Carbosorb (10-20 mesh) to remove CO2. After passing a GC column (3 mm x 45
mm, packed with Carbosieve G held at 50qC), the sample goes through a reduction
column (15 mm x 300 mm) packed with Cu wires at 650qC to remove O2. The latter
is performed to minimise formation of NO in the ion source of the mass
spectrometer, which will interfere at m/z 30. After the removal of O2 in the sample,
the N2 is directed to a triple-collector mass spectrometer to obtain the isotopic
composition of N2. The increased abundance of 29N2 and 30N2 in the sample (f29P
and f30P) is obtained by subtracting the abundance of 29N2 and 30N2 in a reference
(f29R and f30R) from the measured abundance in the sample (f29S and f30S).
5.3.4 Nitrous oxide
Sampling
The incubation must be carried out with a lid preventing exchange of gases between
atmosphere and water. A water sample is transferred to a 12 ml Exetainer (Labco,
High Wycombe, UK) using a glass syringe with an attached length of tubing. The
tubing is inserted to the bottom of the Exetainer and the water is gently pushed out
of the syringe. The tubing is gradually pulled out as the syringe is emptied, always
leaving the tip of the tubing below the water surface. The Exetainer is filled totally,
100 Pl of 38% formaldehyde solution is added and the cap screwed on.
Analysis
Nitrous oxide concentration is measured by electron capture gas chromatography
(Rasmussen et al 1976). A gas chromatograph (GC)(model 14A, Shimadzu Ltd, UK)
equipped with an electron capture detector (ECD) and an integrator (Shimadzu CR68) can be used. The column is stainless steel (4m length, internal diameter 2 mm),
packed with Poropak QS (80-100 mesh, Millipore Corporates, Millford, UK).
Carrier gas is helium (25 ml/min). The temperatures of column, injector and detector
are 35, 180 and 200 qC respectively. This method is shown to be linear for nitrous
oxide over the range of 102 to 107 ppb nitrous oxide, with minimum detection limit
of 55 ppb nitrous oxide (Sage 1995)
Before analysing the water sample a headspace must be introduced into the
Exetainer and equilibrium between water and headspace must be obtained.
A headspace of a precise volume is introduced by inserting a hypodermic needle in
line with a helium flask through the rubber septum of the Exetainer. With a high
precision glass syringe 2.0 ml water are removed from the Exetainer and
simultaneously replaced by an equivalent volume of He (Figure 7). The Exetainer is
the shaken vigorously for 5 minutes or until N2O has reached equilibrium between
40
Chemical analysis
water and headspace. A suitable volume of gas can then be withdrawn from the
Exetainer and injected into the GC. The GC is calibrated injecting known
concentrations of N2O. The total amount of N2O originally present in the sample is
calculated from the solubility coefficient for N2O (Weiss and Price, 1980).
41
Chapter 6
6
Calculations
6.1
Sediment-water fluxes
Time series experiment
Flux =
D xV
A
x 10,000
Only initial- and final concentrations are known
Flux =
(C n -C o ) x V
x 10,000
Axt
where:
Flux
C0
Cn
t
A
V
D
6.2
=
=
=
=
=
=
=
flux (µmol m-2 h-1)
concentration at time zero (µmol l-1)
concentration at time n (µmol l-1)
incubation time (hr)
area of sediment surface in core (cm2)
volume of water in core (l)
slope of the linear regression of concentration (Pmol l-1) versus time (hr).
Chlorophyll a concentrations
Chl a
where:
Chl a
A
K
665b
665a
750b
750a
v
a
l
A x K x ((665b - 750 b ) - (665 a - 750 a )) x v
axl
= chlorophyll a (Pg chl a m-2)
= absorption coefficient of chlorophyll a, 11.0
= factor to equate the reduction in absorbancy to initial chlorophyll
concentration, 2.43
= the extinction at 665 nm before acidification
= the extinction at 665 nm after acidification
= the extinction at 750 nm before acidification
= the extinction at 750 nm after acidification
= volume of acetone extract (ml)
= area sampled (m2)
= path length of cuvette (cm)
For extraction of chlorophyll a from sediment samples see section 5.2.
43
Chapter 6
6.3
Oxygen penetration depth
The penetration depth of O2 is determined from microprofiles (see section 2.3.2). It
is defined as the distance from the sediment surface to the depth at which the O2
signal remains constant with increasing depth.
6.4
Fluxes of O2 from the photosynthetic zone
The diffusive flux is calculated according to Fick's first law of diffusion:
J = - I u D u (wc/wz)
where:
J
I
D
c
z
wc/wz
=
=
=
=
=
=
diffusive flux (nmol cm-2 s-1)
porosity of the sediment
diffusion coefficient (cm2 s-1)
oxygen concentration (nmol cm-3)
depth (cm)
concentration gradient (nmol cm-3 cm-1).
Calculation of upward oxygen flux
Calculate the upward oxygen gradient (wcu/wz) (nmol cm-3 cm-1) by linear regression
through the relevant data points on your concentration profile.
The diffusion coefficient in water (cm2 s-1) is either calculated (e.g. Garcia &
Gordon, 1992) or read from standard tables (Broecker & Peng, 1974).
As the upward gradient is through water the porosity used to calculate the upwards
gradient equals 1.
Upward oxygen flux:
J = -D u (wc/wz)
Calculation of downward oxygen flux
Calculate the downward oxygen gradient (wcu/wz) (nmol cm-3 cm-1) by linear
regression through the relevant data points on your concentration profile.
The diffusion coefficient in water is calculated from a formula (Garcia & Gordon,
1992) or read from standard tables and the sediment (Broecker & Peng, 1974)
porosity is measured as described in section 8.2. However the diffusion coefficient
in Fick’s Law also includes tortuosity which is a measure of the increase in diffusion
path compared to pure water i.e. it describes pore size and pore distribution. For
these calculations use the same value for tortuosity as calculated for the sediment
porosity.
Ds (the substrate diffusion coefficient) = diffusion coefficient for water u tortuosity.
44
Calculations
Downward oxygen flux:
J = - ) u Ds u (wc/wz)
The net production rate is then calculated as
M
where
M =
Ju =
Jd =
Z =
Ju Jd
Z
mean net production rate (nmol cm-3 s-1)
upwards flux of O2 from the photosynthetic zone (nmol cm-2 s-1)
downwards flux of O2 from the photosynthetic zone (nmol cm-2 s-1)
thickness of the photosynthetic zone (cm)
6.5
Denitrification
6.5.1
Incubation time and -types
6.5.1.1.Calculation of optimal incubation time
After addition of 15NO3- to the water the labelled NO3- will start diffusing into the
denitrification zone and denitrification of 15NO3- starts. The rate of 15NO3denitrification will increase until it reaches a constant (steady state) level. The actual
time it takes to reach a certain percentage of 15NO3- denitrification at steady state can
be calculated:
Time(relative) x Z 2
Time(min)
Ds
where:
Time(relative)
Z
Ds
= time on the x-axis of Figure 8
= oxygen penetration depth (mm)
= diffusivity of NO3- / 105
Example
Calculate the time it takes for denitrification of 15NO3- to reach 90% of its steady
state value with a penetration depth of oxygen of 3 mm and a diffusivity of 1 x 10-5
cm2 s-1.
Time (relative) = 5
5 x 32
Time (min) =
= 45 min
1
45
Chapter 6
Denitrification (% of steady state)
100
80
60
40
Current D15
Calculated D15
Calculated D14
20
0
0
2
4
6
8
10
12
14
16
Time (relative)
Figure 8: Results from a numeric model describing the diffusion of 15NO3- from the water into the
sediment and denitrification following an addition of 15NO3- to the water. The calculated rates of denitrification are given relative to the rates at steady state. They are estimated using the production of 14N
and 15N in N2 calculated from the production of 29N2 and 30N2. The "Current D15" denotes the denitrification
of 15NO3- at any given time. The "Calculated D14" and "Calculated D15" denote the rates calculated from
the formula in 6.5.2. The time of addition of 15NO3- is used as time zero and it is assumed that
denitrification occurs immediately below the oxic zone.
When the flux of 15NO3- into the denitrification zone is constant the evolution 15N2
can be expected to be linear with time and denitrification can be calculated from the
production rates of 29N2 and 30N2. It is important to note that denitrification of 15NO3starts right after it is added to the water and its rate increases until it reaches the
steady state rate. The production rates can be determined in two ways:
1: By following the production of 29N2 and 30N2 with time (time series experiment).
2: By determining the concentration of 29N2 and 30N2 at the beginning and the end
of the incubation (start-end experiments).
6.5.1.2 Time series incubation
At least 4 determinations at different times of the labelling of N2 are required. Either
as 4 cores sacrificed at time intervals (2.5.1) or 4 subsamples from each flux
chamber (3.6.2). The first sampling is performed at the time when the evolution of
labelled N2 can be expected to be constant with time (see 6.5.1.1). The last core is
sacrificed or the last subcore is taken at the time required for dark incubated cores to
lower the water column O2 concentration by 15 - 20% of air saturation. The time
series experiment is carried out both in light and darkness and must be done at least
once during winter and once during summer. The sampling for N2 is described in
section 5.3.3.
46
Tc
x
x
x
x
29
Labelled N2
30
( N2 or N2 in mol)
Calculations
Time
15
NO3- added
Figure 9: Concentration of labelled N2 versus time from a time series experiment. 15NO3- is added at
time = 0.
6.5.1.3 Start-end incubation
The production of labelled N2 is determined as the difference between the amount of
labelled N2 before and after the incubation. Incubation time is defined as for the last
core of the time series experiment. Extraction of samples for determination of
labelled N2 is described in section 2.5.1 or 3.6.2. The correct incubation time to be
used in the formula in section 6.5.2.1 is found using the intercept between the x-axis
and the linear regression of labelled N2 with time (Tc, Figur 9) as starting time.
6.5.1.4 Concentration series experiment
Some of the major assumptions in the isotope pairing technique are verified if the
calculated denitrification rates are the same at different 15NO3- concentrations. It is
therefore necessary in a winter and a summer situation to do the measurements at 3
different 15NO3- concentrations.
6.5.2 Calculating denitrification
The basis of this calculation is the mass spectrometric analysis of the isotopic
composition of N2. The mass spectrometer gives the ratio of 29N2 to total N2 and 30N2
to total N2.
6.5.2.1 Production of labelled N2
The absolute amounts of labelled N2 present in a sediment core or flux chamber at
any given time is calculated as:
AM xx R xx u >6N2 @u Volcore
47
Chapter 6
where:
AMxx
Rxx
[6N2]
Volcore
=
=
=
=
the amount of xxN2 present in the core or flux chamber (µmol)
ratio xxN2/total N2 in the sample
concentration of total N2 in the sample (µmol l-1)
volume of water (litre) that the sample represents i.e. the water
volume of the core or flux chamber. If the sample is from a whole
mixed sediment core the whole volume of porewater must be
included. If the sample is from subcores (3.6.2) the volume of
porewater is only included to the depth of the subcore.
The production rate of 29N2 and 30N2 per unit time and area can now be calculated.
The calculation is different depending on the type of incubation.
Time series incubation
The production of 29N2 and 30N2 per unit time is calculated as the slope of the linear
regression line of amount of labelled N2 versus time (Figure 9). The production rate
per unit time and area is the calculated as:
p( xx N2 ) =
D xx
A
x 10,000
Start-end incubation
p( xx N2 ) =
where:
p(xxN2)
Dxx
A
AMi
AMf
t
10,000
AM f AMi
x 10,000
Axt
= production rate of 29N2 or 30N2 (P mol m-2 h-1)
= slope of the regression line of the amount of xxN2 (µmol N) versus
time (h)
= area of sediment surface in the core (cm2)
= initial amount of xxN2 (µmol N)
= final amount of xxN2 (µmol N)
= incubation time. The correct starting time is estimated from a time
series experiment (see Figure 9)
= factor needed to give the result in the specified unit. The variables
must be entered in the units indicated.
Note: AMf, AMi and Dxx represent total amounts of xxN2 in the core/flux chamber,
not concentrations.
6.5.2.2 Isotope pairing calculations
Denitrification rates are then estimated from the production rates of
(Nielsen 1992):
48
15
N isotopes
Calculations
D15 = p( 29 N2 ) + 2p( 30 N2 )
D14 D15
p( 29 N2 )
2p( 30 N2 )
where:
D15 = rate of denitrification of 15NO3- (µmol N m-2 h-1)
D14 = rate of denitrification of 14NO3- (µmol N m-2 h-1)
The part of D14 that is based on NO3- from the water phase (Dw) is calculated from
D15 and the 14N:15N ratio of the water column NO3-:
Dw
D15
>14 NO3 @ w
>15 NO3 @ w
where:
>14NO3-@w = concentration of 14NO3- in the water column
>15NO3-@w = concentration of 15NO3- in the water column.
Finally, in situ denitrification of NO3- produced by nitrification (Dn) is calculated as:
Dn = D14 - D w
All rates are expressed as µmol N m-2 h-1 in dark as well as in light. Diurnal rates
(day + night) are expressed as µmol N m-2 d-1.
6.5.3 Rhizosphere associated denitrification
Calculation of rhizosphere associated denitrification measured with the perfusion
technique (4.6.2).
Volume specific denitrification rates are calculated for each sediment stratum from
the specific accumulation rates of 29N2 and 30N2. This rate is calculated as:
A nxx
I
C ini C final
T
where:
A nxx =
Cini =
Cfinal =
I
=
T
=
volume specific rate of accumulation of 29N2 or 30N2 in the n’th stratum
initial average concentration of 29N2 or 30N2 in the n’th stratum
initial average concentration of 29N2 or 30N2 in the n’th stratum
porosity of the sediment in the n’th stratum
incubation time
49
Chapter 6
Denitrification of 15NH4+ is calculated like D15 above and nitrification-denitrification
of 14NH4+ is calculated like D14 above. The term p(29N2) is replaced with A29 and the
term p(30N2) is replaced with A30 in the formulas for D14 and D15. The volume
specific total rate of coupled nitrification-denitrification is calculated as the sum of
D14 and D15.
6.6
Leaf marking technique
Eelgrass growth is calculated according to Pedersen and Borum (1993). Leaf
production (g dw m-2 d-1) is calculated as the product of average leaf growth rate (cm
shoot-1 d-1), shoot density (shoot m-2) and the specific weight of leaf no 4 (g cm-1).
Root - rhizome production is estimated from the plastochrone interval (the time for
one new leaf to be produced) because one new internode is produced for every leaf
produced. Root-rhizome growth (g dw m-2 d-1) is calculated as the average internode
production multiplied by the average weight of fully grown internode and
associated side roots.
Loss of above and below ground biomass is calculated as:
Loss = Bn+1 - Bn + production
where Bn+1 and Bn are the biomass (g dw m-2) at month n+1 and at month n,
respectively.
Nitrogen incorporation into new leaves is calculated as leaf production multiplied by
the average N content of the fully grown leaf no 3 (Pmol N (g dw)-1). Nitrogen
incorporation into new roots-rhizomes is calculated as root production multiplied by
the N content of the fully grown root-rhizome group.
Loss of plant bound N is calculated as the loss rate multiplied with the N content of
the oldest leaf (leaf No. 6) or the oldest root-rhizome group. Uptake of N is
calculated as the net increase in nitrogen biomass from month n to month n+1 plus
N losses. Reclamation of nitrogen from older to younger plant parts is estimated as
the difference between N incorporation and N uptake.
50
Chapter 7
7
Infauna densities
In order to understand the role of the bioturbating infauna on the biogeochemical
process rates in the sediment ecosystems to be investigated, it is important to have
knowledge of their densities and relative abundance within the sediment.
Information required:
What is the dominating group/genus/species
Relative abundance/densities
Wet weights of the total individuals to give us biomass
On all cores or flux chambers used for measurement of fluxes of oxygen and
nutrients and denitrification the infauna density must be determined. The whole core
is sieved using a sieve with 500 microns mesh. Sieving is carried out using a
continuous flow of water. Infauna retained in the sieve is removed by tweezers and
the wet weight of each group is determined. The weighed fauna is stored in formalin
for later identification. Identification should be to at least group level (bivalve,
polychaete etc.) and if one or a few species or genus are dominating they should be
identified.
The information on infauna must be gathered and stored separately for each core so
that rates measured on a core can be related directly to the infauna data for that
individual core.
51
Chapter 8
8
Sediment characteristics
8.1
Density
The density of the sediment is determined as:
Density =
Wet weight of sediment sample
Volume of wet sediment sample
The sediment is transferred to a graduated cylindrical glass in which 25% of the
volume is filled with water. The weight and the volume of the sediment is
determined as the difference in volume and weight before and after filling in the
sediment. The presence of water in the cylindrical glass ensures that the surface is
smooth when reading the volume after addition of the sediment. The graduated
cylindrical glass must have a volume of approximately 2 times the volume of
sediment that is poured in to obtain good accuracy. Density is determined for the
following depth intervals: 0-0.5 cm, 0.5-1 cm, 1-2 cm, 2-4 cm and 4-6 cm.
8.2
Porosity
The porosity of the sediment is determined as:
Porosity
where:
ww
dw
d
vol
=
=
=
=
ww - dw
vol
wet weight of sample
dry weight of sample
density of the sediment
volume of sample (= ww/d)
Dry weight is determined after drying of the sediment at 105 qC until constant
weight. Porosity is determined for the following depth intervals: 0-0.5 cm, 0.5-1 cm,
1-2 cm, 2-4 cm and 4-6 cm.
8.3
Grain size distribution
Particle size distribution is measured on sediment samples. Segments (10 cm deep)
of the sediment are extruded from a core tube, weighed and dried (60 qC) to a
53
Chapter 8
constant weight and then ground in a pestle and mortar and placed in a series of preweighed sieves (e.g. Endecott, London, UK) with mesh size of 500, 250, 125, and
63Pm corresponding to coarse sand, fine sand, silt and fine silt fractions respectively
(Nedwell et al, 1993). The sediment is shaken for 30 min and each tray is reweighed
to determine the particle size fraction of the sediment at each site. Result can be
expressed as dry mass of material retained by each mesh size as a percentage of total
dry mass (Sage, 1995).
54
Chapter 9
9
Frequency of measurements
Below all variables that should be measured in task 4 of the NICE project are listed.
For each variable the frequency is indicated. The variables indicated as “every
month” should be measured 12 times during one year at approximately one month
interval starting December 1996 or January 1997. Variables indicated as “every six
months” should be measured in a winter and a summer situation.
9.1
Microphytes
Every month
Incubations
x Flux of NO3- light & dark
x Flux of NO2- light & dark
x Flux of NH4+, light & dark
x Flux of urea, light & dark
x Flux of N2O, light & dark
x Primary production (O2 flux)
- Microprofiles of O2, light & dark
x Denitrification, light & dark
- Concentration check, light & dark
- Time course experiment
Variables measured on cores
x Micro algae species composition
x Infauna composition
x Chlorophyll a at sed. surface
x C/N ratio of sediment (depth profile)
x Porosity of sediment (depth profile)
x Density of sediment (depth profile)
x Grain size
x Infauna density and composition
In situ
x Concentration of NO3- + NO2x Concentration of NH4+
x Concentration of urea
x Concentration of PO43x Concentration of O2
x Temperature
x Salinity
x Light at the water- and sediment surface
Every three
months
Every six
months
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
55
Chapter 9
9.2
Rooted macrophytes
Every month
Incubations
x Flux of NO3- light & dark
x Flux of NO2- light & dark
x Flux of NH4+, light & dark
x Flux of urea, light & dark
x Flux of N2O, light & dark
x Primary production, plants & sed. (O2 flux)
- PP of microphytes between plants
x Denitrification, light & dark, 15NO3- in water
- Concentration check, light & dark
- Time course experiment
- 15NH4+ perfusion for rhizosphere. denitrif.
Variables measured on cores
x Biomass (root & leaf)
x C and N content of biomass
x Shoot density
x Macrophyte species composition
x Micro algae species composition
x Infauna composition
x Chlorophyll a at sed. surface
x C/N ratio of sediment (depth profile)
x Porosity of sediment (depth profile)
x Density of sediment (depth profile)
x Grain size
x Infauna density and composition
In situ
x Concentration of NO3- + NO2x Concentration of NH4+
x Concentration of urea
x Concentration of PO43x Concentration of O2
x Temperature
x Salinity
x Light at the water- and sediment surface
56
Every three
months
Every six
months
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
Frequency of measurements
9.3
Floating macroalgae
Every month
Incubations
x Flux of NO3- light & dark
x Flux of NO2- light & dark
x Flux of NH4+, light & dark
x Flux of urea, light & dark
x Flux of N2O, light & dark
x Primary production, plants & sed. (O2 flux)
x Denitrification, light & dark
- Concentration check, light & dark
Variables measured on chambers
x Biomass
x C and N content of biomass
x Macro algae species composition
x Infauna composition
x C/N ratio of sediment (depth profile)
x Porosity of sediment (depth profile)
x Density of sediment (depth profile)
x Grain size
x Infauna density
In situ
x Concentration of NO3- + NO2x Concentration of NH4+
x Concentration of urea
x Concentration of PO43x Concentration of O2
x Temperature
x Salinity
x Light at the water- and macroalgal surface
Every three
months
Every six
months
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
X
57
Index
Chapter 10
10
References
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determining ammonia in seawater. Can. J. Fish. Aquat. Sci 37: 794-798.
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equations. Limnology and Oceanography, 37(6): 1307-1312.
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N2O analysis in the
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59
Chapter 10
Revsbech N. P., Jørgensen B. B. 1986. Microelectrodes: Their use in microbial
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Marine Chemistry 8: 347-359.
60
Index
15
N-nitrate abundance.....................36
N-nitrate addition
macroalgae .................................24
microalgae ..................................14
15
A
ammonia analysis ...........................36
artificial seawater ...........................35
B
biomass
macroalgae .................................25
rooted macrophytes ....................33
bubble formation ............................22
C
C/N ratio
macroalgae .................................25
rooted macrophytes ....................30
chlorophyll a
analysis.......................................37
calculation ..................................43
sampling .....................................16
concentration series ........................47
counting microalgae .................15, 16
D
denitrification
microalgae ..................................14
density ............................................53
dry weight
macroalgae .................................25
rooted macrophytes ....................33
E
epipelic ...........................................15
epipsammic ....................................15
extinction coefficient......................10
F
G
gas chromatograph......................... 40
glutaraldehyde ......................... 15, 16
grain size distribution .................... 53
I
incubation time
denitrification............................. 45
macroalgae........................... 22, 23
microalgae ................................. 13
rooted macrophytes.................... 32
infauna density............................... 51
internode .................................. 31, 50
isotope pairing calculations ........... 48
isotopic composition
N2 ............................................... 39
nitrate......................................... 36
L
leaf marking technique ............ 30, 31
lens-tissue technique...................... 15
light
at preincubation ........................... 9
intensity and source ................... 10
M
macroalgae
denitrification............................. 24
flux............................................. 23
setup........................................... 20
mass spectrometry ......................... 40
microalgae
denitrification............................. 14
flux....................................... 12, 13
setup........................................... 12
microalgal biomass ........................ 16
microelectrodes........................ 12, 38
N
nitrate analysis ............................... 35
nitrous oxide .................................. 40
flux chamber...................................20
frequency of measurements............55
61
O
oxygen concentration
incubation.............................13, 32
preincubation................................9
oxygen flux
calculation ..................................44
primary production.........12, 22, 30
oxygen penetration .........................44
rooted macrophyte
denitrification............................. 32
flux............................................. 32
primary production .................... 30
setup........................................... 28
S
phosphate analysis..........................36
porosity...........................................53
preincubation....................................9
primary production
macroalgae .................................22
microalgae ..................................12
sediment-water fluxes
calculation.................................. 43
sieving ........................................... 51
species composition
infauna ....................................... 51
macroalgae................................. 25
microalgae ................................. 15
sub-core technique......................... 24
R
U
replication.........................................9
rhizosphere denitrification
calculation ..................................49
measurement ..............................32
root - rhizome growth...............31, 50
ultrasonication ............................... 15
urea analysis .................................. 36
P
62
W
Winkler
calculation.................................. 38
sampling .................................... 37
titration ...................................... 38