BB 201
AQUATIC MICROBIAL AND
MOLECULAR ECOLOGY
2016
LABORATORY MANUAL
Institute of Biology
University of Southern Denmark
EACH TEAM’S TIME SCHEDULE FOR THE VARIOUS EXERCISES
The codes refer to chapters in the ”Procedures” section of each exercise.
KF indicates Kærby Fed, FS indicates Fællesstrand,
Date
Team 1
KF
Team 2
KF
Team 3
FS
Team 4
FS
Mo 01.08.
intro
1A
8ABC
intro
1A
8ABC
intro
1A
8ABC
intro
1A
8ABC
Tu 02.08.
1B
2ABD
8DE
1B
4AB
8DE
1B
6ABD
8DE
1B
5,1A
5,2AB
8DE
We 03.08.
1C
2E
4AB
8F
1C;D
2ABD
8F
1C;D
5,1A
5,2AB
8F
1C
6ABD
8F
Th 04.08.
1D
2C
4CDE
2CE
5,1A
5,2AB
4AB
1D
2ABD
Fr 05.08.
7AB
4CDE
2ABD
2E
4AB
Mo 08.08
5,1A
5,2AB
6ABD
2C
4CDE
2C
3AB
Tu 09.08.
5BD
3AB
5BD
2F,5C,6 C
5BD
7AB
5BD
2F,5C,6C
We 10.08
6ABD
7AB
2E
3AB
4CDE
Th 11.08.
2F,5C,6C
3AB
2F,5C,6C
7AB
Fr 12.08
8H
8H
8H
8H
Mo 15.08
8GI
8GI
8GI
8GI
Tu 16.08.
------------- Biogeochemistry – data analysis--------------
We 17.08.
--------------------Report preparation------------------------
Th 18.08.
--------------------Report preparation------------------------
Fr 19.08.
--------------------Report presentation-----------------------
2
Aquatic Microbial and Molecular Ecology 2016
LABORATORY MANUAL
CONTENTS
General introduction ................................................................................................................4
EXERCISE 1: Determination of sediment characteristics (density, water content, organic
content and chlorophyll a content) ...................................................................9
EXERCISE 2: Measurement of sediment oxygen, carbon dioxide and dissolved
inorganic nitrogen exchange in light and darkness ........................................13
EXERCISE 3: Determination of O2 microdistribution and diffusive O2 exchange in
surface sediments ...........................................................................................18
EXERCISE 4: Measurement of sulfate reduction by the 35S technique .................................22
EXERCISE 5,1: Measurement of denitrification in the sediment by the 15N isotopepairing technique ............................................................................................27
EXERCISE 5,2: Measurement of potential nitrification (ammonium oxidation) in
sediment slurry. ..............................................................................................31
EXERCISE 6: Determination of vertical TCO2, DIN (NH4+ and NO2-+NO3-) and SO42profiles in the sediment ..................................................................................33
EXERCISE 7: Determination of particulate iron and redox profiles in the sediment ............35
EXERCISE 8: Molecular analysis of microbial community structure and function ..............39
Reference list ........................................................................................................................50
Appendix 1: Sediment collection ............................................................................................52
Appendix 2: Tables for the data obtained during the exercises ..............................................53
Appendix 3: How to do tricky calculations.............................................................................65
Appendix 4: Safety regulations for laboratory work...............................................................67
Appendix 5: Manual for report writing ...................................................................................71
3
Introduction
GENERAL INTRODUCTION
Organic matter enters coastal sediments either by sedimentation of phytoplankton and
terrestrial derived particles (sewage and rivers) or via primary production by benthic
microalgae (diatoms and cyanobacteria) and macrophytes (algae and seagrasses). Organic
particles located at the sediment surface will gradually be buried within the sediment by
continued sedimentation and faunal particle reworking (bioturbation). The rate by which the
burial occurs depends on the sedimentation/erosion regime and the density of benthic
macrofauna at any specific location. Organic matter in sediments is degraded (mineralized) by
an array of aerobic and anaerobic microbial processes with a concurrent release of inorganic
nutrients. The actual rates of decay depend primarily on organic matter quality (i.e., the
content of protein, cellulose, lignin etc.), age (decomposition stage) and temperature (season).
The chemical composition of organic matter in marine environments can be generalized by
the following formula:
(CH2O)x(NH3)y(H3PO4)z
where x, y and z vary depending on the origin and age of the material.
Decomposition in the upper part of the sediment occurs under aerobic conditions with
oxygen as electron acceptor by the following stoichiometry:
(CH2O)x(NH3)y(H3PO4)z + xO2 + yH2O → xHCO3- +yNH4+ + zPO43- + (x+3z)H+ + yOHAs the oxic (oxygen containing) zone in coastal sediments usually is limited to the
upper 1-5 mm, a large fraction of the organic matter will be buried more or less
undecomposed into anoxic layers. In these layers the large and normally complex organic
molecules will be split into smaller and water-soluble units under the production of energy by
hydrolyzing and fermenting bacteria. These smaller units will then be degraded completely by
an array of microorganisms using a variety of oxidized inorganic compounds as electron
acceptors. These processes generally occur in the following sequence with depth in the
sediment: Mn4+ ≈ NO3-, Fe3+, SO42- and CO2 respiration (Fig. 1). The actual sequence is
determined by the ability of each electron acceptor to receive electrons, and thus the energy
output per degraded organic carbon atom. Nitrate respiration (denitrification):
(CH2O)x(NH3)y(H3PO4)z + 4/5xNO3- + 4/5xH+ + yH2O →
xHCO3- + yNH4+ + zPO43- + (x+3z)H+ + yOH- + 2/5xN2 + 2/5xH2O
is therefore favored energetically compared to sulfate reduction:
(CH2O)x(NH3)y(H3PO4)z + 1/2xSO42- + xH+ + yH2O →
xHCO3- + yNH4+ + zPO43- + (x+3z)H+ +yOH- + 1/2xH2S
4
Introduction
Most of the major microbial processes are carried out by specific groups of organisms,
each of which is specialized in one or a few processes only. No single organism can catalyze
all the different processes. Oxygen-respiring bacteria may be facultative aerobes that grow by
denitrification or fermentation when oxygen is absent. Most manganese- and iron-reducing
bacteria, on the other hand, as well as all sulfate-reducing bacteria and methanogenic archaea
are strict anaerobes and are specialized in their use of electron acceptor. Furthermore, the
strict anaerobes cannot oxidize complex organic matter but rely on fermenting bacteria to
produce their substrates, of which acetate, other short-chain fatty acids, and hydrogen are the
most important. The sequence of anaerobic processes observed in the sediment (Fig. 1) is the
result of competition between the different respiratory groups for these common substrates.
The transition from one electron acceptor to the other downwards in the sediment will occur
when the most favorable is exhausted. ”When the best is gone, one has to accept something
less good.” Nevertheless, the usually observed decreasing degradation rate with depth in
sediments is not caused by the less efficient electron acceptors in the deeper layers, but rather
by decreasing quality of the organic matter (lability or degradability) with depth. However,
the variety of organic substrates which can be degraded seems to be narrower when moving
through the sequence of electron acceptors from O2 to CO2.
RESP
Oxygen
Mangan.
Concentration (arbitrary scale)
O2
Mn
4+
-
NO3
Nitrate
Fe
3+
Iron
Sulfate
-
HCO3
Depth in sediment
2-
SO4
Methanogen.
Figure 1. Idealized depth distribution of electron acceptors in marine sediment. The sequence of
respiration processes is indicated along the left vertical axis.
5
Introduction
The quantitative significance of each of the various aerobic and anaerobic processes
for the total decomposition in any sediment is determined by several local conditions, such as
sedimentation rate, bioturbation rate, temperature etc. Oxygen respiration and sulfate
reduction are usually considered to contribute by up to 50% each. However, the latter process
can be responsible for up to almost 100% in organic-rich sediments. A process like
denitrification is usually of limited quantitative significance for mineralization (few percent)
due to low availability of NO3-. Although sulfate reduction is one of the least favorable
respiration processes, the high concentration of SO42- in seawater (300-1000 times higher than
O2) is responsible for its deep vertical distribution and thus quantitative importance. Inorganic
C, N and P compounds (HCO3-, NH4+ and PO43-), which are released by the mineralization
processes, will accumulate in the sediment porewater and gradually be transported upwards
by molecular diffusion (or bioturbation). Eventually they will all end up in the overlying
water column and once again act as substrate for primary producers (plants).
A large fraction of sediment oxygen uptake is not caused by aerobic respiration
(decomposition), but rather by reoxidation of reduced inorganic metabolites (e.g., H2S and
NH4+). About 50% or more of the sediment oxygen uptake is usually consumed by
reoxidation, which occurs when reduced compounds diffuse from deeper layers and up to the
oxic sediment surface. Reoxidation can be a pure chemical process, but usually it is mediated
by microorganisms. The oxidation of reduced compounds releases energy, which drives the
metabolism of chemoautotrophic Bacteria and Archaea. The most important of these
processes in sediments are nitrification and sulfide oxidation. Although the latter process is
quantitatively most important in terms of oxygen uptake, only the former process will be dealt
with here, since it is very important for nitrogen cycling in sediments. Nitrification is a twostep process driven by aerobic Bacteria and Archaea:
Ammonium oxidation:
Nitrite oxidation:
NH4+ + 3/2O2 → NO2- + H2O + H+
NO2- + 1/2O2 → NO3-
The produced NO3- will diffuse either downwards into the sediment or upwards to the
overlying water column. If downwards, NO3- will be denitrified to N2 in anoxic layers and
thus become unavailable for most living organisms. If upwards, NO3- will be an essential
nutrient for primary producers. About 30-50% of all NH4+ being produced in sediments will
normally be lost by coupled nitrification-denitrification. Despite its limited influence on the
carbon cycling, denitrification is very important for the nitrogen cycling in sediments.
Until very recently it was difficult to obtain reliable information about the nature of
the microorganisms responsible for the chemical processes in sediments. Unlike animals and
plants, most bacteria cannot be identified simply by their size and shape. Only a minority of
the bacteria present in a particular environment can be cultivated using currently available
techniques. This makes a comprehensive description of bacterial diversity in the environment
using classic microbiologic methods impossible. The solution to this problem is the use of
molecular sequence information to investigate microbial diversity and identify prokaryotic
6
Introduction
species, an approach known as “molecular microbial ecology approach”. The sequence of
DNA bases in a particular region of the genome is characteristic of a particular type of
microbe and can be used for identification. The most commonly used genes are those
encoding the ribosomal RNA molecules, which are structural components of the ribosomes,
particularly the 16S rRNA of the small subunit. This is certain to be present in all bacterial
cells whatever their type of energy metabolism or ecological niche.
The purpose of these laboratory exercises is to quantify processes driven by the most
important microorganisms and their diversity in 2 different coastal sediments. Sediment will
be obtained from a relatively organic-rich site, Kærby Fed in Odense Fjord, and from an
organic-poor site, Fællesstrand at Fyns Hoved. You will be split into 4 teams - team 1 and 2
will work with Kærby Fed sediment and team 3 and 4 with Fællesstrand sediment. During the
exercises, oxygen, carbon dioxide and dissolved inorganic nitrogen (DIN) exchange caused
by microalgal primary production and heterotrophic respiration in the sediment will be
measured (exercise 2). Subsequently, the small-scale dynamics of oxygen near the sedimentwater interface in light and darkness will be illustrated and related to microalgal biomass
(exercise 1 and 3). The rates of oxygen uptake and carbon dioxide release in the dark will be
related to rates of sulfate reduction (exercise 4) and denitrification (exercise 5,1) in order to
determine the quantitative role of aerobic respiration, denitrification and sulfate reduction (see
Appendix 3). The distribution of oxidized and reduced particulate iron pools (Fe3+ and Fe2+)
will be examined (exercise 7) and compared with measurements of vertical redox profiles in
the sediment in order show the influence of processes related to iron on the oxidation level of
sediments. The oxidized iron pool will also be used to quantify the role of iron reduction for
the total sediment metabolism (see Appendix 3). Exercise 6 (porewater profiles of TCO2,
dissolved inorganic nitrogen and SO42-) and exercise 1 (porosity and organic content) will
contribute with essential data for the final calculation and interpretation of results obtained in
the other exercises. From the porewater profiles of TCO2 and NH4+ combined with fluxes
from exercise 2, rough estimates can be done of nitrification and denitrification (see Appendix
3). These can be compared with the measured rates in exercise 5. Potential nitrification will be
determined in exercise 5,2 and related to the results from exercise 8, where marker genes for
ammonia oxidation will be quantified.
Exercise 8 illustrates how molecular techniques based on the analysis of
environmental DNA can be used to obtain information about microbial communities,
avoiding the need to culture the microbes. We will be using two approaches: Terminal
Restriction Fragment Length Polymorphism, (T-RFLP; usually pronounced "trif-lip") to
profile the bacterial community and quantitative PCR (qPCR) to determine the copy numbers
of the functional gene ammonia monooxygenase (amoA). For both analyses we will extract
metagenomic DNA (also called community DNA) from sediment samples. We will use a
special method that avoids the co-extraction of contaminating substances like humic acids.
For the T-RFLP analysis we will subsequently amplify a portion of the 16S ribosomal RNA
gene with polymerase chain reaction (PCR). The resulting PCR products will have similar
fragment lengths but originate from different organisms in the community DNA and therefore
have different sequences in the variable parts of the gene. Cutting the DNA with restriction
7
Introduction
enzymes, which recognize characteristic short sequences, gives a mixture of DNA molecules
of different lengths. If one of the PCR primers is labeled with a fluorescent molecule then it is
possible to separate the PCR products by capillary electrophoresis and obtain an
electropherogram containing fragments starting with the labeled primer. This can be used as a
fingerprint of a microbial community and gives in addition semi-quantitative abundance
measurements for the different groups of organisms. It is also possible to compare
characteristic fragment lengths obtained with particular restriction enzymes with the results
expected from 16S rRNA gene sequences in databases. For the qPCR analysis of the bacterial
and archaeal amoA genes we will use the same metagenomic DNA as previously. Both types
of amoA genes will be amplified by PCR and the amount of PCR product will be determined
during each amplification cycle using a double-stranded DNA binding fluorophore and a
detection system in the qPCR machine. Using this information will allow us to calculate back
to the copy numbers of amoA genes in the original sample and compare them with the
potential nitrification measurements from exercise 5,2.
You will find a precise time schedule for the work of each team during the various
exercises in the front of this manual (Appendix 1). The laboratory setup for the experimental
work has been done in advance, and each exercise has a specific location in the laboratory and
is indicated by its number. Please follow the instructions and avoid moving equipment around
in the laboratory without informing the instructors.
The sediment cores to be used during the course will be sampled by you students on
selected dates under guidance of the teachers. Make your own choice of when to participate in
this essential part of the course, but everyone must go at least one time. The schedule for
sediment sampling is listed in the back of this manual (Appendix 1). Please read the safety
instruction sheets (Appendix 4) – particularly the radiosafety instructions before performing
exercise 4.
All data obtained in exercise 1, 2, 4, 6 and 7 should be filled into the appropriate tables
found in Appendix 2. They must be sent by e-mail to the teachers when completed. At the
end, the results obtained by all teams will be available at the course web-site for your use
when writing the final report. Reports should be submitted as team-reports. All members of
the teams must participate on equal terms during laboratory work and result treatment as well
as during report planning, writing (guidelines for writing a report is given in Appendix 5) and
oral presentation during the final synthesis.
8
Exercise 1
EXERCISE 1: Determination of sediment characteristics (density, water content,
organic content and chlorophyll a content)
Water content, density, organic content and chlorophyll a content of sediments are important
supporting parameters to fully understand many aspects of microbial ecology in sedimentary
environments. When related to sediment density, the water content describes the volume of
pore spaces in the sediment (porosity, φ = volume water/total volume). This parameter is
usually used when microbial rates have to be transformed from per weight unit to per unit
volume of sediment. Raw data from laboratory experiments are, of practical reasons, usually
determined per unit weight, but they have a better ecological meaning when presented per unit
volume.
Organic content (loss on ignition) provides an estimate of the potential substrate for
decomposing bacteria. There is, however, rarely a simple relationship between organic
content and sediment metabolism. Sediment metabolism is more related to the quality and
degradability of the organic matter. Most of the organic matter determined as loss on ignition
is actually composed of refractory organic compounds of low degradability.
Chlorophyll a content is a measure of microalgal biomass (diatoms and cyanobacteria)
and should be related to benthic primary production as measured in Exercise 2. The most
important input of labile organic matter at the two study locations is from benthic microalgae.
The biomass of benthic microalgae will here be quantified as chlorophyll a content in the
uppermost sediment layers. Chlorophyll a present in deeper layers is not expected to
participate in overall sediment metabolism.
I. Materials:
2 sediment cores from the study sites (5 cm i.d.)
Slicing plates + stands + rulers + pistons
Alu-trays + alu-foil
4 cut-off syringes (5 ml, new every year)
8 centrifuge tubes (15 ml plast)
90% ethanol
2 M HCl
Plast spoons
Plast cuvettes
Pasteur pipets
Balance
Centrifuge
Spectrophotometer
Oven, trays from oven
16 crucibles
Muffle furnace.
Dessicator + separator plates
9
Exercise 1
II. Procedures.
A. Slicing cores and determination of density
Slice two sediment cores, which has been taken with 5 cm i.d. core liners at the selected
locality, one at the time in the following intervals: 0-1, 1-2, 2-3, 3-4, 4-5, 5-6, 6-8, 8-10 cm.
Place the piston in the core liner (tight, to avoid loss of porewater) instead of the lower rubber
stopper (the upper stopper has to remain tightly in place). The core is then fixed on the stand
and the upper stopper is removed (Fig. 2). The first 6 slices are cut in 1 cm intervals and the
remaining 2 slices in 2 cm intervals by the use of the ruler and cutting plate. The slicing is
done by carefully pushing the sediment core inside the liner up to the chosen thickness as
indicated by the ruler. Then the cutting plate is pushed along the upper edge of the core liner
leaving the intact sediment slice on the cutting plate (use a finger to prevent the slice to slide
of during cutting). The sediment slice is then transferred into a pre-weighed and -labeled
(with a pencil) alu-tray, which is kept ready. Before the next slice is pushed up, the cutting
plate should be cleaned with a paper towel. Cut 4 slices at the time, and handle these as
described below - before cutting the next 4 slices. This will minimize evaporation, and thus
reduce errors in the water content results
The density of the sediment is determined as the weight of a known volume. Fill a preweighed and -labeled 5-ml cut-off (i.e., the tip is cut off) plastic syringe with sediment to the
4-ml mark (avoid any air in the syringe) and weigh it again. Density is calculated as follows:
d = m/4 (g cm-3), where m is the sediment weight (i.e., final weight minus syringe weight).
Weigh the alu-tray + the remaining sediment and place it in an oven at 105ºC.
Ruler
Cutting plate
Sediment
Core tube
Piston
Figure 2. Schematic drawing of set-up for core slicing
10
Exercise 1
B. Water content
When the alu-trays have been in the oven for about 12 hours (the teachers will take them out),
they are transferred to a desiccator. Weigh the alu-trays with the dried sediment.
Water content (β) is calculated from wet weight (ww) and dry weight (dw) as follows
(remember to subtract the weight of the alu-tray):
β = [(ww - dw)/ww] ⋅ 100
Porosity (φ) is calculated according to:
(%)
φ = β ⋅ d/100.
Transfer subsamples of the dried sediment to pre-weighed and -labeled crucibles (do
not fill them more than 2/3). Weigh the crucibles + sediment and placed it in a muffle furnace
at 520ºC.
Plot water content as a function of sediment depth.
C. Organic content
When the crucibles have been in the muffle furnace for 12 h (the teachers will take them out),
they are transferred to a desiccator. Weigh all crucibles + sediment and dump the sediment in
the waste bucket.
Organic content or loss on ignition (LOI) is calculated from dry weight (dw) and ash
weight (aw) as follows (remember to subtract the weight of the crucibles):
LOI = [(dw - aw)/dw] ⋅ 100
(%)
Plot organic content as a function of sediment depth.
D. Chlorophyll a content
We use a rather crude spectrophotometric method, which is easy to perform and reliable. It
may not give the exact concentration of chlorophyll, but it is excellent for comparative
pursposes
Slice two cores into the following depth intervals: 0-0.5, 0.5-1, 1-2 and 2-3 cm. Mix
and homogenize the slices. Transfer a subsample (ca. 0.5 g each) from each depth interval
into a test tube containing 5 ml of 90% ethanol, which is placed on a balance (remember to
zero the balance before adding the sediment) and weigh the sediment sample. Shake or
whirly-mix the test tubes for about 30 seconds. Extract the sediment for 3 h in darkness (5°C).
Shake the test tubes regularly. When the extraction has finished, the test tubes are centrifuged
for 10 min at 3000 rpm. Measure absorbance of the supernatant at 665 and 750 nm before and
after acidification by one drop of 2 M HCl and mixing. Chlorophyll a content is then
calculated as follows (Castle et al., 2011):
Chlorophyll a (μg/g) = 29.6(665o – 665a) v/m
11
Exercise 1
Where 665o is the absorbance at 665 nm subtracted by the absorbance at 750 nm before
acidification; 665a is the absorbance at 665 nm subtracted by the absorbance at 750 nm after
acidification; v is the volume of ethanol (5 ml); and m is the weight (g) of extracted sediment.
Plot the chlorophyll a concentration as a function of depth in the sediment. Compare
the concentrations between the two locations (borrow data from the other teams).
III. Questions, which should be considered in writing the report
1.
2.
3.
4.
5.
Why is it important that there is no air in the syringe during measurement of density?
Why should samples always stay in a desiccator after drying or combustion?
Why are samples for organic content combusted at precisely 520ºC?
Why do we measure absorbance at both 665 and 750 nm for chlorophyll analysis?
Give possible sources of errors.
12
Exercise 2
EXERCISE 2: Measurement of sediment oxygen, carbon dioxide and dissolved
inorganic nitrogen exchange in light and darkness.
Sediment primary production and total heterotrophic metabolism is determined by measuring
oxygen and carbon dioxide exchange across the sediment-water interface. At the same time,
the exchange of dissolved inorganic nitrogen (DIN = NH4+, NO2- and NO3-) between the
sediment and the overlying water is quantified. DIN flux across the sediment-water interface
is a net measure of all nitrogen transformations occurring within the sediment.
I. Materials:
6 sediment cores from the study site (5 cm i.d.)
50 l seawater from the study site
1 tank with core holder and central magnet
Air pumps with tubing and air stones
1 greenhouse lamp
Alu-foil
6 magnet collars and 6 transparent lids
60 ml syringe + tubing
0.45 µm disposable syringe filters
O2 sensor patches
Computerized O2 measuring set up
”Flow injection/diffusion cell“ analyzer.
HCO3- standards (1, 2, 3 mM - made from 100 mM stock solution).
Thermometer.
3 ml Exetainers (6)
Salinity-meter (conductivity).
20 ml scintillation vials (for nutrient samples) + vial holder.
Plast test tubes
Flow Injection Analyzer (Lachat).
Plast bags.
II. Procedures.
A. Sediment cores
Sediment cores have been taken with 5 cm i.d. core liners at the location and pre-incubated in
the storage tank at the selected temperature (15ºC) in 12 h light : 12 h dark cycles (light will
be kept off today until start of the incubation). Select 6 cores for flux incubation among the
stock and assure that the sediment depth is 15-20 cm. Wash hands before or wear gloves when
handling cores in the incubation tanks. Describe the sediment of each core with respect to
colour zonation and any traces of animal activity (i.e. burrows of Nereis diversicolor). This
can be done before or after the incubation. Wrap 3 of the cores in alu-foil (dark cores), but
leave the top open until initial sampling has been completed. Submerge these cores in the core
holder around the magnetic stirring unit of the incubation tank together with the 3 unwrapped
cores (light cores) such, that the water surface is at least 2 cm above the upper edge of the
core liners. The light cores must be placed directly under the greenhouse lamp. Be sure that
13
Exercise 2
there is continuous aeration in the tank. Measure and note water temperature and salinity
(with conductivity detector).
B. Dark and light flux incubation
After turning the light on, initial (start) samples (20 ml) are taken from the water phase inside
each of the 6 cores while still submerged using a 60-ml syringe for analysis of CO2 and DIN
(Dissolved Inorganic Nitrogen). Transfer 5 ml to a 3-ml Exetainer (carbon dioxide analysis,
see later). Fix a syringe filter to the syringe and filter 15 ml sample to a 20-ml scintillation
vial (DIN analysis, see later). Store the CO2 samples at 5ºC until analysis and freeze the DIN
samples immediately. After finishing the initial sampling, place stirring collars in the middle
of the water of each core and seal the cores air tight with a transparent lid holding a small
patch of oxygen sensitive phorphyrin-derivate fixed to the inside (see later for explanation) on
top of the core liner (do it under water) and wrap the dark cores completely in aluminum foil.
Note the start time. IMPORTANT: The entire procedure is conducted for one core at a time,
i.e., when the start sample has been taken from one core, it is sealed before sampling is started
on another core. Terminate the incubation for one core at a time after 2-3 hours and take the
final samples for CO2 and DIN as described for the initial samples - BUT with the exception
that the cores are lifted above the water level in the tank before opening the lid and during
sampling. Note the precise time at the end of incubation for each core. Measure and note
temperature and the height of the water phase in the cores.
C. Quantifying abundance and biomass of Nereis diversicolor
The respiration contribution to O2 uptake and CO2 production by the most common burrowing
benthic fauna species, Nereis diversicolor, will be estimated from the abundance and biomass
in flux cores. All cores are therefore sieved through a 1 mm mesh to recover the worms.
Recovered worms are transferred to petri dishes (one per core). Worms are counted and
weighed. The weighing is done by blotting worms briefly on a paper tissue to remove excess
water before transferring them one by one to a petri dish containing seawater, which is placed
on a balance.
Respiration (V, µmol h-1) as a function of individual biomass (M, g wet wt.) for N.
diversicolor (15°C) is:
VO2 = 2.15 M0.68
VCO2 = 3.63 M0.91
Use the average individual biomass (M) and multiply by the density (m-2) of N. diversicolor
to get the contribution on an area basis.
D. Photochemical oxygen determination
Oxygen will be measured by a relative new optode approach. Central for the technique is the
ability of oxygen to quench the fluorescence of certain chemical compounds. In other words,
O2 can absorb the energy that otherwise would have been sent out as fluorescence from a
given fluorophore. There exist numerous configurations, chemistries and procedures to
quantify O2 availability by this approach (e.g. Klimant et al. 1997, Kuhl & Polerecky 2008)
14
Exercise 2
and many are custom build and optimized for specific experimental approaches and
procedures – some of these will be demonstrated during the course. The system applied for
flux measurements is a commercial available system that is sold by the company Pyroscience.
The system is called Firesting and has 4 channels that can be applied for simultaneous
measurements.
Small patches of an immobilized phorphyrin-derivate are fixed to the inside of the lids
to the sediment cores. This compound is excited by red light and the O2 sensitive emitted light
has an optimum in the near infrared region. The excitation light is supplied to the sensor patch
via a fibre placed in the small hole of the lids. The same fibre also guides the O2 sensitive
light back to the measuring instrument. The sensors are pre-calibrated and the O2 signal (in %
air saturation) over time during the incubation is directly read out on the computer screen and
can be noted on paper or saved as a text file. Measure temperature and salinity for conversion
from % air saturation to concentration in µM (a small conversion program for PC computers
will be provided).
E. CO2 analysis (Hall & Aller 1992)
TCO2 will be analyzed on a ”flow injection/diffusion cell“ analyzer. Samples (0.1 ml)
containing CO2 are injected into a carrier stream of 45 mM HCl which is pumped into one
side of a diffusion cell equipped with a gas permeable membrane (plumbers tape). A receiver
stream of 10 mM NaOH is pumped simultaneously to the other side of the membrane. All
CO2 from the acid carrier side diffuses into the base receiver side. The amount of CO2 is then
detected as a reduction in conductivity (by a conductivity detector) as a result of base
neutralization in the receiver stream. The signal is recorded and peak height measured by a
computer.
The ”flow injection/diffusion cell“ analyzer will be turned on by the teachers. Start
analyzing a 0 mM (distilled water) standard and continue with 1, 2 and 3 mM standards. Fill a
1-ml syringe with the chosen standard solution (one syringe is dedicated and labeled for each
concentration). Turn the injection port to the ”load“ position. Insert the syringe needle into the
sampling port and inject at least 0.2 ml to fill the 0.1 ml sample loop and wait until the
baseline on the computer screen is stable. Turn the injection port to the ”inject” position, wait
5 seconds and press ”R“ on the computer keyboard. When the peak has passed the maximum,
press ”S” on the computer keyboard. Now a number representing the peak height will be
shown on the screen. Write the number down. This procedure should be repeated until the
peak heights of two replicates are within 1 % of each other. After finishing the initial standard
curve, the samples are analyzed similarly (use the syringe labeled ”sample“). For every 4
samples, one 2 mM standard should be analyzed to assure that the standard curve still is valid.
The concentration of TCO2 is calculated manually based on the standard curve.
F. DIN analysis
Analysis of NH4+, NO2- + NO3- is done spectrophotometrically on a LACHAT Flow Injection
Analyzer (the exact procedure will be explained during the exercise). NH4+ is analyzed
according to the Berthelot-reaction (Bower & Holm-Hansen 1980). In a weak basic solution
15
Exercise 2
(pH 9.5-11), NH3 reacts with hypochlorite to produce monochloramine, which in turn
provides indophenol blue in the presence of salicylate, nitroprusside-ions (catalyst) and
surplus of hypochlorite. The automated NO2- analysis is based on a reaction between the NO2ion and sulfanilamide under acid conditions. This produces a diazo-compound which is
coupled with N-(1-naphthyl)-ethylene-diamine to provide a purple azo-dye (Armstrong et al.
1967). The absorption of the color reaction is determined at 550 nm. The analysis of NO3- is
based on a reduction of NO3- to NO2- by a copper-cadmium column followed by the analysis
as described above for NO2-. Procedure: Transfer samples (in the appropriate dilution) to the
small autosampler cups and place them in the autosampler. From here, they automatically will
be mixed with reagents and measured in a spectrophotometer one by one. The concentrations
(in μM) are calculated from a relationship based on the peak areas of 5 known standards
(NH4+: 0, 10, 20, 30, 40 μM; NO3-: 0, 5.0, 10.0, 20.0 μM). NO2- and NO3- will not be
determined separately here because the concentration of NO2- is very low. Results should
therefore be presented as NO2- + NO3Calculation of fluxes: Oxygen exchange per m2 sediment surface per day (JO2, mmol
m-2 d-1) is determined from the slope (α) of a linear regression of O2 against time (days)
according to:
Flux
=
JO2
=
-2 -1
mmol m d =
slope
α
mmol m-3 d-1
⋅
⋅
⋅
water volume / surface area
π r2 h
/
π r2
m3
/
m2
This can be reduced to:
JO2 = α ⋅ h
where the slope α has the units of μM d-1 (μmol
dm-3 d-1 = mmol m-3 d-1) and h is the height of the water column above the sediment in m (or
cm/100)
Carbon dioxide (JCO2) and DIN exchange (JDIN) per m2 sediment surface per day
(mmol m-2 d-1) are calculated according to:
Flux
= change in concentration ⋅ water volume / surface area / time
=
(Ce – Cs)
⋅
π r2 h
/
π r2
/ t
Jx
-2 -1
-3
3
2
mmol m d =
mmol m
⋅
m
/
m
/ d
(values)
(units)
This can be reduced to:
Jx = (Ce – Cs) ⋅ h / t where Ce and Cs are end and start
concentration in μM (μmol dm-3 = mmol m-3), h is the height of the water column above the
sediment in m (or cm/100) and t is incubation time in days (or hours/24).
III. Questions, which should be considered in the report
1. Why is it necessary with stirring during flux incubations?
2. What can be the cause for observed differences in O2 and CO2 exchange among the 3
cores?
16
Exercise 2
3. Calculate RQ (respiratory quotient) and PQ (production quotient) and explain what
they tell us.
4. Calculate the CO2/DIN flux ratio and explain what it tells us..
5. Which processes within the sediment determine the net flux of DIN?
6. What may influence the ratio of dark CO2 production to DIN flux? Would you expect
this to be the same as the C:N ratio of the organic matter actually being decomposed?
7. How can benthic invertebrates influence the fluxes?
8. Give possible sources of errors.
17
Exercise 3
EXERCISE 3: Determination of O2 microdistribution and diffusive O2 exchange in
surface sediments
Photic surface sediments are characterized by very steep O2 gradients reflecting concurrent
production and consumption of O2. Fast microsensors are required to resolve the distribution
and the temporal dynamic of O2 in such environments – in this exercise we use Clark-type O2
microelectrodes (Revsbech & Ward 1983). However, optimized microoptodes (see also
Exercise 2) represent a viable alternative (e.g. Klimant et al. 1997) – the technique will be
demonstrated in the laboratory.
The microprofile approach makes it possible to quantify O2 exchange and penetration
in darkness and light, in order to calculate the net benthic photosynthesis. It is also possible to
measure the gross photosynthesis by the so-called light-dark shift technique (Glud et al.
1992), but that will not be done in the current exercise.
I. Materials:
2 sediment cores (can be reused from exercise 2)
Complete automated microelectrode measuring set up
Adjustable light source
Incubation facilities ensuring thermoregulation and good water flow
Light logger
Diagram for diurnal down-welling irradiance
Air-pump with tubing and air-stone
Software for O2 solubility
Table for diffusion coefficients
II. Procedures:
A. Dark O2 penetration and diffusive mediated exchange:
Experimental: The microelectrode tip is placed 1-2 mm above the sediment surface.
Subsequently the sensor is lowered in steps of 50 µm (0.05 mm), while the O2 concentration
at each depth is recorded and stored by the computerized set-up. Note when the sensor enters
the sediment. Repeat the measurements until you record low stable signals (i.e 0-3 mV) that
remain constant when you move the sensor - this is defined as the anoxic zone. The sensor is
moved back to the start position, moved 3-5 mm horizontally and the measuring procedure is
repeated. You should measure a minimum of three profiles – but are welcome to measure
more.
Calculations and theoretical work: The recorded text-files are transferred into Excel and here
the first step is to calibrate the recorded signals into µmol L-1. Oxygen sensors have a linear
response and as we know the O2 concentration in the overlying water phase at the given
temperature and salinity (see the provided computer program) and the signal size in the
anoxic (zero oxygen) sediment layers, it is simple to calibrate each of the recorded data points
18
Exercise 3
according to:
Calibrated signal (µmol L-1) = Sr (Cw /(Sw-Sa)) + Sa(Cw /(Sw-Sa))
Where Sr is the recorded signal (in mV), Sw the signal in the overlying water phase, Sa the
signal in anoxic sediment and Cw is the O2 concentration in the overlying water phase. Define
the relative position of the sediment surface. Assign this position to depth “0” and use
negative prefix for recordings at any depths below this fixed point (see Fig 3). Plot the
respective microprofiles with “Depth” (mm) on the ordinate and O2 concentration µmol L-1)
on the abscissa. Include one typical profile in the report.
Determine the average O2 penetration (P) (unit: mm) and calculate the average
Diffusive O2 Exchange (DOE – Jdark,up) from:
DOE (mmol m-2 d-1) = -Do (dC/dZ)*8640
Where Do is the molecular diffusion coefficient (cm2 s-1) at the given conditions (see Table 1)
and (dC/dZ) is the slope of the linear O2 concentration gradient in the Diffusive Boundary
Layer (DBL) (µmol L-1 mm-1) . Provide Standard Deviation for P and DOE.
Figure 3. Two microprofiles measured in sediment cores recovered from a water
depth of 1.2 m in Helsingør Harbour, Denmark. The profiles were measured at the
exact same spot in darkness and at a irradiance of 600 µmol photons m-2 s-1. The
white horizontal bars indicate the gross photosynthetic rates at each depth measured
by the light-dark-shift approach (not measured in this exercise). The temperature was
1 ºC and the salinity 12.
19
Exercise 3
B. O2 penetration and diffusive mediated exchange as a function of light:
The experimental procedure in II.A is repeated three times at 5 different irradiances (i.e. 10,
50, 100. 300, 500 µmol photons m-2 s-1). Start with the lowest irradiance. Calibrate and plot
the respective profiles as above. Include one set of light profiles - plotted in the same
diagram - in the report. Determine the average O2 penetration (P) (mm) at the respective
light levels - include the SD - and plot P against the irradiance.
Calculate net photosynthesis (Net-P, mmol m-2 d-1) in the photic zone for each light
level using a modified version of the DOE equation above (see also Fig 3B):
Net-P = DOE (Jlight,up) + DOEdo (Jlight,down) = ((Do (dC/dZ)) + (Ds ϕ (dC2/dZ2))) *8640
Where Ds is the sediment diffusion coefficient which can be approximated as Ds = ϕDo
(Ullman & Aller 1982) and (dC2/dZ2) is the downward gradient (see Fig 3B). ϕ is the surface
porosity as measured in exercise 1. DOE (Jlight,up) represents the diffusive upward flux in the
boundary layer above the sediment and DOEdo (Jlight,down) the diffusive downward flux in the
layer immediately below the sediment-water interface
20
Exercise 3
III. Questions to be answered in the report:
1. How does DOE measured in darkness compare to the total oxygen uptake (TOU)
measured during intact core incubations? Discuss potential reasons for any
differences.
2. Is the sediment net heterotrophic or autotrophic when integrated over a normal 24h
day/light cycle?
3. Discuss if the resolved benthic primary production is important on system level.
4. Compare the O2 penetration depth to the distribution of solutes and processes
measured in the other exercises.
5. How does benthic primary production and respiration obtained from exercise 3
compare with those obtained from exercise 2?
21
Exercise 4
EXERCISE 4: Measurement of sulfate reduction by the 35S technique
The purpose of this exercise is to determine the rate of sulfate reduction in the sediment.
Reduction of sulfate to sulfide is followed by the use of the radioactive isotope 35S. Labeled
sulfate in the form of 35S-SO42- is injected into the sediment (Jørgensen 1978). The rate of
sulfate reduction can be determined when the ratio between the added amount of labeled
sulfate and the amount of sulfur recovered as labeled sulfide is related to the total
concentration of sulfate in the sediment.
Radioactive sulfide is incorporated into various reduced sulfur pools in sediments. In order to
release these pools, we will distill the sediment according to the TRIS technique (Total
Reduced Inorganic Sulfur). By this technique, both the acid volatile sulfides and the
chromium reducible sulfur will be released simultaneously (Fossing & Jørgensen 1989). Acid
volatile sulfides (FeS) are converted to H2S by the addition of concentrated hydrochloric acid
(HCl). To release the chromium reducible sulfur (FeS2 and So) in the form of H2S, on the
other hand, a much stronger reducing agent (chromium) is needed. The following process
describes pyrite (FeS2) reduction: FeS2 + 2 Cr2+ + 4 H+ → Fe2+ + 2 H2S + 2 Cr3+
I. Materials:
2 sediment cores in tubes with 1 injection port per cm. (2.6 cm i.d.)
Stand + holder for tubes
Cutting plate + ruler + piston
2x6 plast centrifuge tubes (50 ml)
Centrifuge
Balance
Whirli mixer
Spectrophotometer
Cuvettes (1 cm)
Gloves
Scintillation counter
2 x 6 reaction flasks + various for distillation
Beaker for dist. H2O.
Long test tubes
20 ml scintillation vials.
7 ml scintillation vials.
Vial holders
10 ml plast centrifuge tubes
Eppendorf tubes
Waste container
Plast sheet for covering table
60 ml syringe
5 ml Finnpipette + tips
1 ml Finnpipette + tips
22
Exercise 4
200 μl Finnpipette + tips
50 μl Hamilton syringe
Long and short pasteur pipettes
Radiotracer, 35S-SO42-.
20 % og 5% (w/v) Zinc acetate.
6 M HCl
Chromium solution (1 M Cr2+ in 2 M HCl).
Scintillation liquid
Cline-reagent (low Cline).
Silicone oil (anti foam)
II. Procedures (This exercise is done in a marked and approved area. Only persons
participating in the exercise are allowed in this area. Lab coats will be handed out and gloves
must be worn)
A. Incubation (Before you start read the radiosafety instruction sheet - Appendix 3)
Two sediment cores have been sampled using core liners (2.6 cm i.d.) equipped with silicone
sealed injection ports for every cm. Mount the cores on the lab-stand and remove the
overlying water. Inject 5 μl of a 35S-SO42- solution (60 kBq) in every injection port down to
10 cm depth. Allow the cores to incubate for 4-5 hours in darkness at the same temperature as
the flux cores (Exercise 2). Note the time of start and temperature.
B. Core sectioning
Label and pre-weigh (with lids, p. 33, (1)) 2x6 plast centrifuge tubes (50-ml). Add 5.0 ml (1
cm slices) or 10.0 ml (2 cm slices) of 20% ZnAc (density 1.03 g/ml) and weigh the tubes.
Mount the incubated cores on the lab-stand. Cut 6 sections (0-1, 1-2, 2-4, 4-6, 6-8, 8-10 cm)
as described in Exercise 1 and transfer them to the centrifuge tubes. Clean the cutting plate
with paper towels between each slice. Seal the tubes immediately by the screw caps and mix
the content on by shaking before weighing the tubes. Finally, all samples are stored frozen.
Transfer the remaining sediment to the plastic bag labeled radioactive waste.
C. Distillation
Thaw the frozen samples in the centrifuge tubes and centrifuge them at 3000 rpm for 10
minutes. Transfer 200 μl of the supernatant into a 7-ml scintillation vial (label it on the lid!)
for determination of 35S-SO42- activity. Add 800 μl of distilled water. Scintillation liquid will
be added later (see below). Transfer an additional 1 ml supernatant to 1.5 ml eppendorf tube
(a spare sample in case of problems with the original) before decanting the remaining
supernatant to the waste container. Weigh the centrifuge tubes with sediment again after
removing the supernatant.
While the thawing proceeds, the ZnAc traps for retaining H2S are prepared (H2S is
precipitated as ZnS). Add 10 ml of 5% ZnAc into 6 long test tubes together with one small
drop of silicone oil (prevents “foam” formation). Place the traps in the distillation unit.
23
Exercise 4
Mix the sediment in the plast centrifuge tubes (after centrifugation and decanting the
supernatant) and transfer about 4 g of sediment (msub) into a reaction flask . Transfer then 15
ml of distilled water into each reaction flask and seal all the stoppers. When all reaction flasks
are placed on the heating plates, the entire distillation unit is assembled by sealing the ”tops”
equipped with long Pasteur pipettes. Open for the cooling water!!! Remember to check the
water flow. Open for the N2 flow using the small valves above the distillation unit. The flow
is appropriate when there is a calm and steady bubble development in the traps. Let the
reaction flasks stand with bubbling for 5-10 minutes before adding 8 ml of 6 M HCl (use a
plastic syringe) and 16 ml of a chromium solution (1 M Cr2+ into 0.5 M HCl).
Turn on the heating plates (level 3). Let the distillation proceed for 45 minutes after
boil. Keep an eye on the N2 bubble pattern and the flow of cooling water. The reaction
mixture has to boil vigorously.
Terminate the distillation by dismantling the unit (disconnect the connection to the
trap) and close the N2 flow. Mix the traps with an automatic pipet and transfer immediately 5
ml into a 20-ml scintillation vial (only labels on the lid). The remainder is transferred into 20ml scintillation vials and stored for later use.
Add 2 ml scintillation liquid to all 5-ml vials containing supernatant and 10 ml
scintillation liquid to the 20-ml vials containing trap material, and mix the solution for about 1
min. Make a blank from 1 ml H2O and 2 ml scintillation liquid. The samples are now ready
for counting. Ask the teachers about the procedure. More information about scintillation
counting can be found at http://encyclopedia2.thefreedictionary.com/scintillation+counter
The reaction mixture from the reaction flasks is transferred into a waste container. The
reaction flasks are rinsed thoroughly with distilled water before the start of a new distillation.
Warning. Bottles with N2 are handled by staff members only. Working with open
samples must occur on the covered tables. All handling of open samples must be done
standing up, partly in order to avoid inhalation of aerosols, partly in order to minimize the risk
of person contamination. Never use a whirli-mixer for samples in open containers, instead you
must mix with an automatic pipette. 35S might be mutagenic. Waste must be collected in a
plastic container marked “waste (affald in Danish)”. A solution of 6 M HCl is very
corrosive. Vapours are very easily absorbed in the mouth, throat and lungs, where it can cause
corrosion and fluid diffusion (dys-pnoea, difficulty in breathing). Chrome chloride may be
mutagenic. Waste is collected in a plastic container marked “Cr”. Scintillation fluid is a
mixture of organic solvents, and inhalation may cause brain damage. Use the fume hood! All
other waste must be collected in the buckets with radioactive marks.
D. Calculation of sulfate reduction rate
The rate of sulfate reduction (SRR) is calculated according to:
[SO42-]s ⋅ a ⋅ 24 ⋅ 1.06
SRR = ────────────── (nmol SO42- cm-3 d-1)
(A + a) ⋅ h
24
Exercise 4
where [SO42-]s is sulfate concentration in nmol cm-3 sediment = [SO42-] (mM) ⋅ φ ⋅ 1000
(determined from porewater profiles, see Exercise 6); a is the total radioactivity in the traps
(H235S - remember that only a subsample of the traps has been counted); A is the total activity
of 35SO42- (from the supernatant); h is incubation time in hours; 1.06 is a correction factor for
bacterial isotopic fractionation. Note that A and a have to be transformed to dpm cm-3, based
on the sediment weight and porosity (from Exercise 1).
Calculation of a:
K ⋅ (dpma - dpmB) ⋅ msedc ⋅ d
a = ──────────────────── (dpm cm-3)
msub ⋅ msed
where: K
= correction factor to total trap volume.
msed = weight of sliced sediment before centrifugation (g).
msedc = weight of sediment after centrifugation and decantation of ZnAc (g).
msub = weight of sediment subsample (g).
dpma = radioactivity of H235S (dpm).
dpmB = background radioactivity (dpm).
d
= sediment density (g cm-3). - from Exercise 1
Calculation of A:
(VZnAc + msed ⋅ ß/100) ⋅ (dpmA - dpmB) ⋅ d
A = ──────────────────────────── (dpm cm-3)
VA ⋅ msed
where: VZnAc
ß
dpmA
VA
= volume of ZnAc (5.0 or 10.0 ml).
= water content (%). - from Exercise 1
= radioactivity of 35SO42- (dpm).
= volume of sampled supernatant (0.2 ml).
Plot SRR as a function of depth.
E. Determination of the reduced sulfide pool
Besides the labeled sulfide, the distillation procedure has also released and trapped the total
pool of reduced inorganic sulfur in the sediment. The size of this pool can be determined by a
spectrophotometric technique. For this purpose, we will use the excess trap material, which
was stored in 20-ml scintillation vials. Mix the trap material thoroughly on a Whirli-mixer.
Transfer 50 µl (KF) or 200 μl (FS) of each sample (+ a blank consisting of 5% ZnAc) into
centrifuge tubes and dilute with 10 ml of distilled water. Add 800 μl of Cline reagent (Cline
1969) to the centrifuge tubes, close lids immediately, shake the tubes and let them rest for 20
minutes. Transfer ca. 3 ml of each sample to a cuvette and read the absorption at 670 nm on a
standard spectrophotometer. The absorbance must be below 0.8. If it is higher, the entire
procedure is repeated using less trap material.
25
Exercise 4
Warning. Cline reagent contains 6 M HCl and is very corrosive. Take good care and
use paper cover on the table. The reagent also contains dimethylphenylendiamine (0.4%),
which is very easily absorbed through the skin and methaemoglobin is formed in the blood.
Dimethylphenylendiamine may have mutagenic and other dangerous effects (not yet
completely known). By reaction with sulfide, methylene blue is created, which can be
mutagenic. All Cline-work must be done in the fume hood. Gloves and lab coat must be
worn and waste must be collected in a plastic container marked “Cline”.
The concentration is calculated using a constant factor (μM/abs) based on the
absorption coefficient for the actual Cline reagent used here (will be handed out by the
teachers). The pool of Total Reduced Inorganic Sulfur (TRIS) can be obtained according to:
abss ⋅ F ⋅ b ⋅ msedc ⋅ d
[TRIS] = ──────────────── (μmol cm-3)
k ⋅ 1000 ⋅ msub ⋅ msed
where:
abss
k
F
b
= absorbance of trap sample.
= absorption coefficient (abs/μM).
= dilution factor.
= trap volume (10 ml).
Plot TRIS as a function of depth in the sediment.
III. Questions, to be considered in writing the report
1.
2.
3.
4.
5.
What are the advantages and disadvantages by incubations using intact cores?
Discuss the depth distribution of sulfate reduction rates in the sediment.
Compare the profiles of sulfate reduction rates with those of TRIS.
Calculated the depth-integrated sulfate reduction (mmol m-2 d-1).
Compare the depth-integrated SRR with sediment oxygen uptake (Exercise 2) - How
much of the oxygen may be used to oxidize H2S?
6. Give possible sources of errors.
26
Exercise 5
EXERCISE 5,1: Measurement of denitrification in the sediment by the 15N isotopepairing technique
Sediments typically house an active and extremely interesting nitrogen cycle. Nitrate is used
in denitrification (and occasionally nitrate reduction to ammonium) and this process can
sometimes account for a significant fraction of carbon mineralization in sediments.
Denitrifiers are facultative aerobic bacteria which can use nitrate instead of oxygen in
anaerobic environments. Nitrate comes either from diffusion from the overlying water or from
nitrification in the sediments, a chemoautotrophic process where ammonium, liberated to the
sediment through organic matter decomposition, is oxidized nitrate. This exercise will focus
on denitrification.
Our determination of denitrification rates comes from an isotope pairing technique
based on addition of nitrate labeled with the stable isotope 15N and subsequent analysis of the
labeled dinitrogen pairs formed (Nielsen 1992). Denitrification rates can in theory be
determined from the accumulation of N2 in the water column. In reality, however, this is
extremely difficult due to the high background concentrations (about 500 μM) of N2 in water.
As we will see, the source of nitrate (overlying water or nitrification) is also possible to
calculate from the data we obtain, but the accuracy of this assessment has been challenged in
the literature (Middelburg et al. 1996).
I. Materials
3 sediment cores with sediment to about 10 cm from the top of the core
10 mM stock 15NO3- solution.
Alu foil and saturated ZnCl
Stirring collars
Sediment stirring set up similar to exercise 2
Plastic disposable pipets
Ruler
Glass spatulas
50 ml glass syringe
7 cm isoversinic (gastight) tube to fit syringe
10 ml Exetainers
10 ml plastic vials for nutrient samples
0.45 µm syringe filters
10 ml syringes
1 ml pipet and tips
200 µl pipet and tips
tape
markers
calculator
rack
27
Exercise 5
II. Procedures
A. Incubation
Three 6 cm cores from your location are taken and the water level is adjusted to about 0.3 cm
from the rim. Use a 50-ml syringe with a piece of tubing and try to avoid resuspension of
sediment. Then take a water sample for nutrient analysis from the circulating water in the tank
and filter it through 0.45 µm filters into the plastic vials. These nutrient samples are needed to
determine the percentage of labeled nitrate for an accurate determination of denitrification
rates.
Before addition of 15NO3- the height of the water column is measured to calculate the
volume in which the added nitrate is diluted. Calculate the volume of your overlying water
and add enough stock solution to bring the nitrate concentration to 50 μM. Check this
calculation with the teachers before adding the 15NO3-. Mix 15NO3- into the water phase up by
gentle flushing with bubbles through a disposable pipette. Notice; 15N is not a radioisotope!
Take a subsample for NO3- analysis from each core and filter it into the plastic vials.
Remember to label the vials with team number, core number and time (before or after label
addition).
Note starting time, add a stirring magnet and close the core with a stopper without
trapping air bubbles. Do not press the stopper too hard. Leave the cores with circulation in
darkness for about 3 hours.
At the end of the incubation, note the time and stop denitrification by dripping ~0.5 ml
saturated ZnCl2 solution (a little poisonous) into the overlying water of the core with a small
plastic pipet. With a glass spatula, mix the top 6 cm of sediment and water with gentle vertical
motion to distribute the labeled N2 evenly before sampling. Avoid skimming the surface as
gas may escape. Check the new height of the water column and the height of the mixed
sediment to determine the volumes in which labeled dinitrogen will be distributed. With a 50
ml syringe with a small piece of silicone tube at the end, first take in a few ml of the
suspension and discharge to wash out any gas phase in the syringe, Now, carefully withdraw
about 30 ml sample. You will have prepared 6 Exetainers by adding on drop of ZnAc
solution. Transfer the sample to an Exetainer by introducing the tube at the bottom and then
draw up as the vial is filled to the rim. This is to prevent loss of dinitrogen to the air. Put on
the screw cap and check that no bubbles are trapped. Take two samples from each core.
Dinitrogen samples are marked with Team number and core number, location and incubation
time. They are saved for later MS analysis.
B. Analysis of dinitrogen by mass spectrometry (MS).
Bring your samples to the Mass spectrometer room. You will be directed there by one of the
instructors. Initially, with the lid still on the Exetainers, one cm of sample is replaced with 1
ml helium through hypodermic needles inserted through the rubber seal, and by vigorous
shaking, dinitrogen is equilibrated into the He headspace. From the headspace, 0.25 ml is
sampled and introduced in the MS. The content of 28N2, 29N2 and 30N2 are indicated separately
as “peak areas” by the MS, which also calculate the ratios 29N2/28N2 and 30N2/28N2. Reference
28
Exercise 5
samples of atmospheric N2 are also processed, and they represent the initial isotope
composition in the cores, i.e. the natural background.
C. NO3- analysis
NO3- will be analyzed of water samples taken from cores will be done as in Exercise 2.
D. Calculations
To help with the calculations, please use the scheme below, which outlines the recommended
sequence of input data, intermittent results and final results. First clear out the formulas, then
do all the calculations for one core, and then for the other two. Concentrations should be
expressed in μM (O2, NO3-, N2 etc.) and rates as mmol m-2 d-1.
The added 15NO3- rapidly diffuses into the sediment, and mixes with native 14NO3-. In
the denitrification zone both unlabeled, single labeled and double labeled dinitrogen pairs are
formed (28N2, 29N2 and 30N2). Only the labeled pairs formed can be detected by mass
spectrometry, but assuming random pairing the formation 28N2 can be deduced from the
formation of 29N2 and 30N2.
The rate of 29,30N2 production is in principle determined by :
The principle of the calculation scheme for denitrification
[N 2 ]
29,30

N2
N2
(
)
( start ) h
−
end
28
28
N2
 N2

P 29,30 N 2 =
t
where h (m) = water height + porosity*sediment depth, and t = time (hours) and (N2) is the
concentration of ambient dinitrogen during the incubations. In reality the masses are amplified
differently in the mass spec, and this has to be corrected for, we will therefore provide you
with the P29N2 and P30N2. You can then continue the calculations as described below.
Denitrification of 14NO3- (D14) and 15NO3- (D15) can be calculated from the production of 29N2
and 30N2 according to:
D15
D14
29,30
= rate of denitrification of 15NO3
= P29N2 + 2P30N2
= rate of denitrification of 14NO3
= D15P29N2/2P30N2
The total denitrification in the cores is defined as
equal to D14
D15 + D14 and in situ denitrification is
Calculation scheme for coupled nitrification-denitrification
The data also tells how much of the denitrified nitrate that was coupled to nitrification in the
sediment, and how much was contributed from the overlying water:
29
Exercise 5
1) 14NO3- and 15NO3- from the water column are processed similarly, and therefore the in situ
denitrification of 14NO3- from the water (Dw) is calculated as:
Dw = D15 * [14NO3-] / [15NO3-]
where [14NO3-] and [15NO3-] are the measured and calculated concentrations of
15
NO3- in the water column during incubation.
14
NO3- and
2) The calculated total in situ denitrification (D14) is (hopefully) larger than Dw and the
difference is ascribed to coupled nitrification-denitrification (Dn). Calculate that.
Questions, to be considered in writing the report
1. Why do we make a slurry out of the upper part of the sediment after incubation?
2. Compare the rates of denitrification with the flux of nitrate found in Exercise 2.
3. Compare rates of denitrification with CO2 fluxes in the dark and depth integrated
sulfate reduction rates.
4. Give possible sources of errors.
30
Exercise 5
EXERCISE 5,2: Measurement of potential nitrification (ammonium oxidation) in
sediment slurry.
The measurement of nitrification (bacterial oxidation of NH4+ to NO2- and NO2- to NO3- with
O2 as electron acceptor) on intact sediment is a very difficult task. We will instead examine
the potential nitrification. When measuring potential rates, the conditions with regard to
substrate concentration (NH4+), O2 availability and temperature are optimized for the bacterial
populations under consideration (nitrifiers). Accordingly, the potential nitrification activity
(Vmax) is a relative measure of the number of nitrifying bacteria, or rather of the amount of
active enzyme molecules present. We will measure the depth distribution of NH4+ oxidizing
bacteria in the sediment using slurries of sediment from various depth intervals (Exercise 8).
We will use ClO3- (chlorate) to inhibit NO2- oxidizing bacteria. The accumulation of NO2- in
the slurries is therefore a measure of the activity of NH4+ oxidizing bacteria.
I. Materials:
1 sediment core from the study site (5 cm i.d.)
Slicing plates + stands + rulers + pistons
Alu-trays + alu-foil
Shaking platform
Måleglas (50 ml).
8 Erlenmeyer flasks (250 ml).
Balance
Black plastic bag
Tape
Centrifuge.
15 ml centrifuge tubes
Test tubes
Spectrophotometer and cuvettes
Pipet (5 ml) and pipet tips.
2 liter in situ seawater enriched with 500 μM NH4Cl, 50 μM KH2PO4 and 10
mM NaClO3 (check if pH is 7-8 otherwise adjust)
II. Procedures:
A. Incubation
First, describe the color pattern with depth in the intact sediment core and note any presence
of animals or burrows. Section the sediment core in the following depth intervals: 0-1, 1-2, 23, 3-4, 4-5, 5-6, 6-8 and 8-10 cm (see the procedure in Exercise 1). Transfer the slices to alutrays and mix the sediment thoroughly. Weigh 10 g (KF) or 15 g (FS) sediment (note the
31
Exercise 5
precise weight) from each section in an Erlenmeyer flask. The incubation is started (t = 0) by
adding 50 ml of NH4+ and ClO3 enriched seawater to the flasks. Take a time t = 0 sample.
Incubate the slurries at room temperature in darkness (cover with black plastic) on a shaking
table. Transfer at regular intervals (e.g., every half hour - we will discuss that in detail during
the exercise) 5 ml of the slurry into a labeled centrifuge tube and centrifuge immediately
(3000 rpm for 5 min). A 3 ml subsample of the supernatant is used for the measurement of
NO2- concentration (pipet the subsample into a new pre-labeled test tube)
B. NO2- analysis
Add 100 μl nitrite-reagent (be careful - nitrite-reagent is poisonous) to the cuvette
containing 1 ml sample or 1 ml standard. Mix with pipette using the sample tip and let the
samples rest for at least 10 minutes in the dark. Read the absorbance at 543 nm. Use artificial
seawater as a zero (0 standard). Calculate the NO2- concentration in the samples from a
standard curve based on the absorbance of the chosen standards.
C. Calculations:
Transform the NO2- results from μM in the slurry to nmol cm-3 sediment by the following
equation:
nmol cm-3 = μM ⋅ φ ⋅ F
where:
φ = porosity - from Exercise 1
F = dilution factor
The dilution factor is obtained according to:
F = (m ⋅ (β / 100) + 50) / (m ⋅ (β / 100)
where:
m = weight of sediment in the slurry (g)
β = water content (%) - from Exercise 1
Plot the accumulated NO2- (nmol cm-3 sediment) for each depth as a function of time (hours).
Calculate potential NH4+ oxidation (Vmax) for all depths (nmol cm-3 h-1). Then, plot Vmax as a
function of sediment depth.
III. Questions, to be considered in writing the report
1. Why is it necessary to shake the slurry during the incubation?
2. What conclusions can you make with regard to the presence and distribution of NH4+
oxidizing bacteria?
3. Which factors control the in situ activity of NH4+ oxidizing bacteria?
4. Give possible sources of errors.
32
Exercise 6
EXERCISE 6: Determination of vertical TCO2, DIN (NH4+ and NO2- + NO3-) and SO42profiles in the sediment.
During this exercise we will determine the vertical distribution of selected dissolved
compounds (TCO2, DIN and SO42-) in the sediment porewater. The profiles of these
compounds reflect the metabolic activity in the sediment, i.e., accumulation of metabolites
(such as TCO2, NH4+) and consumption of electron acceptors (such as NO3-, SO42-). The
obtained profiles can also be helpful in the interpretation of the results of the other exercises.
Furthermore, the porewater concentrations of SO42- are essential for the calculation of sulfate
reduction rates (Exercise 4).
I. Materials:
2 sediment cores from the study site (5 cm i.d.)
Core slicing equipment (see Exercise 1)
16 double centrifuge tubes
10 ml syringes
2.5 cm GF/C filters
Pincers
0.45 µm disposable syringe filters
20 ml scintillation vials.
3 ml Exetainers
Alu trays
Spoons
Balance
Centrifuge.
1 ml and 5 ml Finnpipette + tips
”flow injection/diffusion cell” analysator.
II. Procedures
A. Sediment sectioning and porewater extraction
Take one core from the storage in the cold room and finish that before taking the second to
the laboratory. First, describe the color zonation of the sediment and note any trace of animals
in each of the two cores. Transfer about 15 ml of the water phase above the sediment into a
20-ml scintillation vial (to be analyzed as the porewater samples). Section the sediment cores
into the following depth intervals: 0-1, 1-2, 2-3, 3-4, 4-5, 5-6, 6-8, 8-10 cm as described in
Exercise 1. Cut 4 slices at the time. Transfer each slice into double centrifuge tubes
(remember to put one GF/C filter in the bottom first). Weigh the centrifuge tubes with
sediment and assure that they pairwise only differ by max. 0.2 g. Place the tubes in the
centrifuge (tubes of equal weight should be opposite each other) and centrifuge them at 1500
rpm for 10 minutes. Filter the extracted porewater carefully through a 0.45 µm disposable
33
Exercise 6
syringe filter into a labeled 20-ml scintillation vial after centrifugation (one vial per depth
section per core). Transfer 1 ml of the porewater (for TCO2 and SO42- analysis) to a labeled 3ml Exetainer and freeze the remaining porewater in the 20-ml vial for later DIN analysis.
Rinse (in distilled water) and dry the centrifuge tubes when the entire procedure is terminated
for the first 4 slices and repeat the procedure with the next 4 slices (do not throw out the
centrifuge tubes after use – we re-use them).
B. TCO2 analysis
Porewater TCO2 will be analyzed on a ”flow injection/diffusion cell“ analyzer. As described
in Exercise 2.
C. DIN analysis
DIN (NH4+, NO2- and NO3-) in the porewater will be analyzed as described in Exercise 2.
Fællesstrand porewater must be diluted 1:1 with distilled water and Kærby Fed porewater
must be diluted 1:4 with distilled water.
D. SO42- analysis
Sulfate will be analyzed by ion chromatography using a Dionex ICS-2000 Ion
Chromatography system (the exact procedure will be explained during the exercise).
III. Questions, which have to be answered in the report
1. Plot the concentration of TCO2 (mM), DIN (μM) and SO42- (mM) with depth in the
sediment.
2. Discuss the distribution of NO3- in the sediment?
3. Are there any relationships between TCO2 and NH4+ or SO42-? If any, then plot the
relationship and explain why.
4. Give possible sources of errors.
34
Exercise 7
EXERCISE 7: Determination of particulate iron and redox profiles in the sediment.
In this exercise, we will compare the vertical distribution of oxidized (Fe3+) and reduced
(Fe2+) particulate iron pools with the redox potential in the sediment. The oxidation of Fe2+ to
Fe3+ by both NO3- and free O2 occurs rapidly in the oxidized zone of sediments. The process
is catalyzed either by microbial enzymes (chemolithotrophic Fe2+ oxidation) or by chemical
auto-oxidation. In the reduced zone, the opposite process occurs - also catalyzed by bacteria
(heterotrophic Fe3+ respiration) or by chemical reduction. The idea behind the present iron
measurement is to obtain the biologically available particulate iron (the pool which is
accessible for bacteria) by extracting the sediment in 0.5 M HCl. The acid also prevents any
oxidation of reduced iron. First the reduced iron (Fe2+) is measured using Ferrozin (produces
a red color complex which is stable for at least 24 hours). Subsequently, the oxidized iron
(Fe3+) is reduced to reduced iron (Fe2+) by hydroxylamine and the total iron is determined
using Ferrozin. The difference between total and reduced iron is oxidized iron.
Each specific anaerobic process in sediments usually has a demand for a particular
redox environment. By determining the redox potential, we can roughly predict the microbial
processes which may occur in different sediment depths. The redox potential is a measure of
the electron potential - the lower redox, the more available electrons. At redox potentials
below about 300 mV, the stable form of iron is Fe2+ and the stable form of nitrogen is NH4+.
Sulfate reduction usually occurs at redox potentials below -100 mV. Oxic water has a redox
potential of about 500 mV. The redox potential is measured by a platinum electrode with a
calomel electrode as a reference. The theoretical Eh values are, however, the potential relative
to a hydrogen electrode. It is therefore necessary to correct the values obtained using a
calomel electrode according to:
Eh = Emeasured + 244 mV
I. Materials:
2 sediment cores from the study site (5 cm i.d.)
Core slicing equipment (see Exercise 1)
Alu trays
Steel spatula
Balance
15 ml centrifuge tubes with stopper
Shaking platform
5 ml and 200 μl Finnpipette + tips
18 plast test tubes with stoppers
Centrifuge
Spectrophotometer and disposable cuvettes
100 ml 0.5 M HCl
100 ml 100 mM FeSO4 in 0.5 M HCl
1.5 M Hydroxylammoniumchloride (NH2OH⋅HCl) in 0.25 M HCl
0.02% Ferrozine in 50 mM HEPES, pH 7
mV-meter.
35
Exercise 7
Redox electrodes
Stop watch
II. Procedures
A. Redox measurement.
First, describe the color zonation of the sediment and note any trace of animals in each of the
two cores. Make two redox measurement on one of the cores, while the centrifuge tubes (no.
1-9 containing 5 ml of 0.5 M HCl), test tubes (no. 1H-9H) and cuvettes (no. 1-9 and 1H-9H)
are being labeled. When the redox measurements are terminated, the core is sectioned for iron
measurement. During the slicing procedure of the first core, redox is measured on the second
core. When redox is done on the second core, it is sectioned immediately for iron
measurement. If you find it appropriate, redox can be measured simultaneously on both cores.
The redox measurement is done with a needle platinum electrode fixed on a
micromanipulator. The redox electrode is together with a calomel reference electrode
connected to a mV-meter, which is switched to the 200 mV scale. Fix the sediment core to
the stand and place the tip of the platinum electrode in the water-phase 1 cm above the
sediment. The calomel electrode must be in contact with the overlying water. Push the
platinum electrode downwards and measure the redox potential in 2.5 mm steps relative to
the sediment surface, i.e.: -10, -7.5, -5, -2.5, 0, 2.5, 5, 7.5, 10 etc (negative values represent
measurements above the sediment-water interface and positive below). Each reading is made
1 minute after moving the electrode to a new depth. Clean the tip of the redox electrode with
scouring powder between measurements (the electrode is usually poisoned – reduced – by
sulfide when inserted into anoxic sediment). Be careful not bending the needle during
handling. Rinse the electrode with distilled water after cleaning.
Figure 3. Volumes of sample transfer into various vials with their labeling indicated
36
Exercise 7
B. Determination of iron pools (Stookey 1970, Lovley & Phillips 1987)
The particulate iron pools are determined by the Ferrozin method after extraction with HCl.
Section the sediment in the chosen depth intervals: 0-0.5, 0.5-1, 1-1.5, 1.5-2, 2-2.5, 2.5-3, 3-4,
4-5, 5-6 cm according to the technique described in Exercise 1. Finish the handling of each
slice before cutting the next - it is important to work fast because Fe2+ will be rapidly
oxidized to Fe3+ when the sediment is exposed to O2. Transfer the slice to an alu-tray and mix
it fast to a homogenous paste. Transfer about 300 mg (KF) or 600 mg (FS) of the
homogenized sediment to the centrifuge tube containing 5 ml of 0.5 M HCl (place the
centrifuge tube on a balance and zero the balance before transferring sediment). The weight
of the transferred sediment is determined in the centrifuge tube - note the weight. Place all
centrifuge tubes horizontally on a shaking table for 30 minutes when subsamples of all slices
from one core have been transferred to the tubes. Pipet 0.2 ml hydroxylamine into the 9 test
tubes (1H-9H), which are stoppered, and 2 ml of 0.02% Ferrozin to all 18 cuvettes (Fig. 3).
When the shaking process is terminated, the centrifuge tubes are centrifuged at 3000 rpm for
5 minutes. Pipet from the supernatant (be careful not to resuspend the precipitate): A. 40 μl
to 9 cuvettes (1-9) containing Ferrozin (mix with the pipette); B. 1 ml to each of the 9 test
tubes (1H-9H) containing hydroxylamine (Fig. 3) - shake these frequently for the next 15
minutes before transferring 40 μl to the remaining 9 cuvettes (1H-9H) containing Ferrozin
(mix with the pipette). The samples (in the cuvettes) are now ready to be read at 562 nm on
the spectrophotometer. Remember to make a standard curve. Use FeSO4 to make the
following standards: 0, 50, 100, 200, 400, 800 µM Fe2+ in 0.5 M HCl. Handle the standards a
mentioned under A above. Zero the spectrophotometer using a solution of 2 ml Ferrozin
reagent + 40μl of 0.5 M HCl.
Warning. Hydroxylamine-hydrochloride is poisonous. By breathing in aerosols, or by
absorbing it through the skin, methaemoglobin is formed in the blood. Hydroxylaminhydrochlorid can be mutagenic and 0.5M HCl can be corrosive. Gloves and lab coat must be
worn and waste must be collected in a plastic container marked “Fe”.
Calculation of iron pools: (unit: μmol g ww-1)
Abs562 ⋅ A ⋅ (5 + (m ⋅ ß ⋅ 10-2))10-3
[Fe2+] = ─────────────────────────
M
Abs562 ⋅ A ⋅ 1.2 ⋅ (5 + (m ⋅ ß ⋅ 10-2))10-3
[Fe ] = ──────────────────────────── - [Fe2+]
m
3+
where:
Abs562
A
m
ß
= absorbance at 562 nm
= absorption coefficient (obtained from standard curve)
= weight of sediment (g)
= water content (%)
- from Exercise 1
37
Exercise 7
III. Questions, to be considered in the report
1. Describe the distribution of oxidized and reduced iron in the sediment. Is oxidized Fe
found deep in the sediment? If so, why?
2. Did you find any relationship between redox potential, iron pools and possible color
changes in the sediment?
3. Are there any relationships between Eh and the rates of sulfate reduction (from
Exercise 4)?
4. Explain the cause for the redox discontinuity - base the answer on the results from the
other exercises.
5. Give possible sources of errors.
38
Exercise 8
Exercise 8: Molecular analysis of microbial community structure and function.
Microbial ecology has been revolutionized by the introduction of molecular techniques.
These new methods target total community (metagenomic) DNA that has been directly
isolated from environmental samples. In contrast to the isolation and cultivation approach in
classic microbiology, molecular ecological methods are used to study not yet cultivated
microorganisms from the environment. It has been estimated that less than 1% of all
microorganisms have been cultivated to date, showing the importance of molecular methods
for our understanding of microbial ecology.
Molecular microbial ecology techniques are used to answer the following questions:
1. Who is there?
2. How abundant are the different microorganisms present?
3. What are they doing?
4. How active are they?
Here we will demonstrate two of several possible approaches using molecular techniques:
Community Fingerprinting of bacterial communities using Terminal Restriction
Fragment Length Polymorphism (T-RFLP)
Molecular community fingerprinting allows us to obtain comparative information about the
composition of microbial communities. The basic idea is to amplify all the copies of a
particular gene present in a microbial community using polymerase chain reaction (PCR) and
analyze the composition of the resulting mixture of PCR products. The most commonly
targeted gene is the one encoding 16S ribosomal RNA as it is present in all prokaryotic cells
and a comprehensive, continuously growing database for it is available. Usually a "general
primer" set targeting the majority of Bacteria is used, but some studies also use groupspecific primers. Alternatively, the PCR may target a functional gene coding for a protein
characteristic of a particular physiological group of organisms. The PCR products will
typically have similar size but differ in sequence as the result of evolutionary changes.
In the exercise we will use an approach known as Terminal Restriction Fragment Length
Polymorphism (T-RFLP). Here, different phylogenetic groups (or operational taxonomic
units, OTUs) are distinguished after treating the PCR products with a restriction enzyme,
which will produce a mixture of DNA fragments of different lengths. If one of the two PCR
primers is labeled with a fluorescent group, each characteristic sequence will produce one
end-labeled DNA molecule. The mixture of products can be separated by capillary
electrophoresis using an apparatus similar to that used for DNA sequencing. It is capable of
separating DNA molecules of 600 bp or more, resolving differences of only 1 base pair. Using
a standard mixture of DNA molecules of known sizes (size standard) labeled with a second
fluorophore it is possible to determine the absolute length of the products, facilitating
39
Exercise 8
comparisons between samples and allowing comparison with predicted restriction product
sizes calculated from sequence information in databases.
In this exercise we will not generally attempt to identify the individual peaks. We will rather
use the abundances of the different peaks (representing the OTUs) to obtain an overall pattern
as a community fingerprint to compare samples from different sampling sites and sediment
depths in order to answer question 2: How abundant are the different microorganisms
present?
Quantification of functional genes with quantitative PCR
Quantitative PCR (qPCR) is a method often used in molecular microbial ecology to
determine either how abundant microorganisms are or how active they are. Measurements of
activity are often performed by extracting mRNA from environmental samples and
determining the copy numbers of marker genes for metabolic pathways (also called
functional genes) by qPCR. In order to compare how many microorganisms in a sample
harbor a certain metabolic pathway, the metagenomic DNA can be analyzed with qPCR as
well. By using primers targeting the 16S rRNA gene of a specific phylogenetic group of
microorganisms in parallel to primers for marker genes for a certain metabolic pathways in a
qPCR analysis, the results allow the estimation of the metabolic potential of the group of
organisms in question.
Quantitative PCR is fairly similar to a normal PCR. However, by using a method to quantify
the amount of DNA in the PCR reaction during each step of the amplification allows to
calculate back to the initial amount of target DNA molecules in the sample (the so-called
copy number). We use a method utilizing SYBR Green, a fluorescent dye that binds to
double stranded DNA, for quantification, which is detected by an optical system in the qPCR
machine.
In this exercise we will quantify the marker gene for nitrification, the ammonia
monooxygenase (amoA) gene. The process of nitrification can be carried out by two different
groups of organisms, ammonia oxidizing bacteria (AOB) or ammonia oxidizing archaea
(AOA), also known as Thaumarchaeota. From the measurement of the potential nitrification
activity in exercise 5,2 we already know if the sediment harbors organisms that can nitrify.
Here we try to confirm the results from the biogeochemistry but take it one step further and
also determine which of the two groups of nitrifiers is present.
Overview of the laboratory exercises
We will analyze sediment samples from four different depths from the same sites you are also
investigating during the biogeochemical part of the course (Fællesstrand, Kærby Fed).
Sediments are difficult to work with due to heterogeneity (resulting for example from animal
activities) and the common presence of PCR inhibitors like humic substances. We will use a
40
Exercise 8
special method to purify metagenomic DNA so that we can use PCR and qPCR methods
downstream in the experiment.
Overview scheme of exercise:
Precautions to be taken when working with samples for molecular work
All reagents are autoclaved or sterile-filtered
Use only autoclaved pipette tips; keep the box closed when not in use
Use forceps to remove Eppendorf tubes to avoid touching the insides of the lids
Wear gloves to prevent contamination from the skin
Treat PCR tubes gently since they have thin walls
Ensure that the liquid is in the bottom of the tube; give a brief centrifugation if needed
Label tubes on the lids with fine-point markers
41
Exercise 8
I. Materials:
1 sediment core from the study site (5 cm i.d.)
Core slicing equipment (see Exercise 1)
aluminum trays (sterile)
70% ethanol
Paper towels
2 ml Eppendorf tubes (sterile)
Metal spoon
Metal spatula
Mo Bio PowerLyzer PowerSoil DNA Isolation Kit
Set of pipettes
Sterile pipette tips
Balance
Eppendorf tube rack
FastPrep instrument
Eppendorf centrifuge
DNAs extracted from the sediment
Positive control DNA
Forward primer 27F FAM (20 pmol/µL)
Reverse primer 519R (20 pmol/µL)
PCR reaction components
PCR machine
PCR tubes
Gel electrophoresis equipment
PCR products
Buffer R (10 x concentrated)
Restriction enzyme BsuRI (10 U/µL)
37 °C incubator or PCR machine
Restriction enzyme treated PCR products
Thermo Scientific GeneJET PCR Purification Kit
Sterile 1.5 ml Eppendorf tubes
PCR water
qPCR cycler
qPCR tubes and lids
RealQ Plus 2 Master Mix (Ampliqon)
Forward primers AOA-amoA-f and AOB-amoA-1F
Reverse primers AOA-amoA-r and AOB-amoA-2R
42
Exercise 8
II. Procedures
A. Slicing of cores
1. Wipe metal plates used for core slicing with 70% ethanol
2. Slice core and save slices from 0-1 cm, 2-3 cm, 4-5 cm, and 6-8 cm on separate sterile
aluminum trays
3. Wipe the metal spoon with 70% ethanol to disinfect it and homogenize the sediment
on the tray
4. Wipe spoon with ethanol and repeat with the other slices
5. Wipe the spatula with ethanol and use it to transfer ~1 g of the 0-1 cm homogenized
sediment slice into a 2 ml Eppendorf tube
6. Fill two additional 2 ml Eppendorf tubes with 1 g sediment each from the same
sediment slice
7. Wipe spatula with ethanol and repeat with the other homogenized sediment slices
In total you have taken 12 sediment samples now, 3 from each depth. Put one sample from
each depth on ice for further processing and freeze the others in liquid nitrogen. We will store
them at -80 °C in case we might need them.
B. DNA extraction with Mo Bio PowerLyzer PowerSoil DNA Isolation Kit
For the DNA extraction we will follow the protocol of the kit. For further information please
refer to it. Here is the excerpt from the manual (with modifications):
1.
Properly label each Glass Bead Tube on both the cap and on the side
2.
To the PowerLyzer Glass Bead Tube (0.1 mm provided) add approx. 0.25 g of
sediment sample. (IMPORTANT: please determine the exact weight of the sediment
sample you are extracting; you will need this information later!)
3.
Add 750 µl of Bead Solution to the Glass Bead Tube. Gently vortex to mix.
4.
Check Solution C1. If Solution C1 is precipitated, heat solution to 60 °C until
dissolved before use.
5.
Add 60 µl of Solution C1 and invert several times or vortex briefly.
6.
Bead Beating: place the Glass Bead Tubes into the Tube Holder for the FastPrep
instrument. The Glass Bead Tubes must be balanced (evenly spaced) on the Tube
Holder. Run the samples on setting “6” for 45 seconds.
7.
Make sure the Glass Bead Tubes rotate freely in your centrifuge without rubbing.
Centrifuge Bead Tubes at 10,000 x g for 30 seconds at room temperature. CAUTION:
Be sure not to exceed 10,000 x g or tubes may break. If the sediment is not completely
pelleted after 30 seconds, centrifuge again for 3 minutes at 10,000 x g.
8.
Transfer the supernatant to a clean 2 ml Collection Tube (provided). Note: Expect
between 400 to 500 µl of supernatant. Supernatant may still contain some sediment
particles.
9.
Add 250 µl of Solution C2 and vortex for 5 seconds. Incubate at 4 °C for 5 minutes.
10. Centrifuge the tubes at room temperature for 1 minute at 10,000 x g.
43
Exercise 8
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
Avoiding the pellet, transfer up to, but no more than, 600 µl of supernatant to a clean 2
ml Collection Tube (provided).
Add 200 µl of Solution C3 and vortex briefly. Incubate at 4 °C for 5 minutes.
Centrifuge the tubes at room temperature for 1 minute at 10,000 x g.
Avoiding the pellet, transfer up to, but no more than, 750 µl of supernatant into a clean
2 ml Collection Tube (provided).
Shake to mix Solution C4 before use. Add 1200 µl of Solution C4 to the supernatant
and vortex for 5 seconds.
Load approximately 675 µl onto a Spin Filter and centrifuge at 10,000 x g for 1 minute
at room temperature. Discard the flow through and add an additional 675 µl of
supernatant to the Spin Filter and centrifuge at 10,000 x g for 1 minute at room
temperature. Load the remaining supernatant onto the Spin Filter and centrifuge at
10,000 x g for 1 minute at room temperature. Note: A total of three loads for each
sample processed are required.
Add 500 µl of Solution C5 and centrifuge at room temperature for 30 seconds at
10,000 x g.
Discard the flow through.
Centrifuge again at room temperature for 1 minute at 10,000 x g.
Carefully place spin filter in a clean 2 ml Collection Tube (provided). Avoid splashing
any of Solution C5 onto the Spin Filter.
Add 100 µl of Solution C6 to the center of the white filter membrane.
Centrifuge at room temperature for 30 seconds at 10,000 x g.
Remove the Spin Filter and put it in a fresh Eppendorf tube (not provided). The DNA
in the tube is now ready for any downstream application.
Store the DNA at -20 °C until usage (Solution C6 contains no EDTA!).
C. Determination of DNA concentration with Nanodrop
1. Pipet 2 µl of sterile Milli Q water on Nanodrop and “initiate”.
2. Wipe off and pipet 2 µl sterile Milli Q water on Nanodrop and read “blank”.
3. Wipe off and pipet 2 µl sterile Milli Q water on Nanodrop and measure as sample;
reading should be close to “0”.
4. Wipe off and start measuring samples.
D. Polymerase chain reaction of 16S rRNA genes
With PCR it is possible to amplify certain regions of genomic DNA using specific starter
oligonucleotides (called “primers”). Here we are interested in the 16S rRNA gene, a universal
phylogenetic marker for Bacteria and Archaea (in Eukaryotes the equivalent is the 18S rRNA
gene). We will amplify the first part of the 16S rRNA genes of all bacteria in the sample using
a primer set where one of the primers is labeled with a fluorophore. This will allow us to
obtain a community profile after treatment of the mixed sequences with a restriction enzyme.
44
Exercise 8
Samples:
Each group will use the samples that were purified with the Mo Bio PowerLyzer PowerSoil
DNA Isolation Kit. For a better comparison of the results we will have to dilute the DNAs to
5 ng/µL using PCR grade water before we can use it in the PCR reaction.
Each group should thus have the following “samples” to use in the PCR:
1. DNA from core slice 0-1 cm, adjusted to 5 ng/µL
2. DNA from core slice 2-3 cm, adjusted to 5 ng/µL
3. DNA from core slice 4-5 cm, adjusted to 5 ng/µL
4. DNA from core slice 6-8 cm, adjusted to 5 ng/µL
5. Positive control DNA
6. Negative control (only premix, no DNA)
PCR reaction mixture
Combine the reaction mix (total of 50 µL) in a PCR tube, containing template, the two
primers, and the Master Mix with dNTPs, polymerase and reaction buffer.
Template (purified DNA, diluted purified DNA)
2 µL
PCR-grade water
30.75 uL
dNTP (2 mM)
5 uL
MgCl2 (25 mM)
5 uL
Forward primer 27F FAM (20 pmol/µL)
1 µL
Reverse primer 519R (20 pmol/µL)
1 µL
Taq-buffer (10x)
5 µL
Taq-pol (5 U/uL)
0.25 uL
It is advantageous to prepare a “premix” of the PCR reagents including everything besides the
template for all samples that you want to amplify, e.g. if you want to amplify 6 samples
prepare a “premix” with 6.5 x the volume you would need.
PCR program
Hotlid held at 105 °C
Initial:
94 °C
Repeat 29 ×:
94 °C
54 °C
72 °C
Finish:
72 °C
4 °C
1 min
denaturation
15 sec
45 sec
45 sec
denaturation
annealing
elongation
5 min
indefinite
finishing
holding
After the reaction is complete, the products will be analyzed on a 1.5% agarose gel. If the
PCR reaction is successful we will obtain a band at about 500-550 bp, corresponding to the
45
Exercise 8
distance between the forward primer at position 27 and the reverse primer at position 519 of
the 16S rRNA gene.
E. Restriction of amplified 16S rRNA genes
In order to perform a Terminal Restriction Fragment Length Polymorphism (T-RFLP)
analysis we have to digest the PCR products with a restriction enzyme. To do this, each group
will use the PCR products received from the sediment sample DNAs, avoiding the positive
and negative control products.
Restriction reaction
The reaction mix (20 µL) in a 0.2 mL tube contains the PCR product, the restriction enzyme
of choice (BsuRI; GG/CC) and the reaction buffer.
PCR product
10.0 µL
Buffer R (10 x concentrated)
2.0 µL
Restriction enzyme BsuRI (10 U/µL)
0.5 µL
PCR water
7.5 µL
Final volume
20.0 µL
It is advantageous to prepare a “premix” of the reagents including everything except the PCR
product. The mixture is incubated at 37 °C for at least 3 hours (better over night), followed by
heat inactivation for 20 min at 80 °C.
F. Purification of restriction fragments
The restriction fragments must be separated from proteins and remaining, unused fluorescentlabeled primer before they can be run on the apparatus, which separates the DNA molecules
by size. For this clean up we will be using another “kit”, called GeneJET PCR Purification
Kit from Thermo Scientific:
1. Add a 1:1 volume of Binding Buffer to completed PCR mixture (e.g. for every 100 μl
of reaction mixture, add 100 μl of Binding Buffer). Mix thoroughly. Check the color
of the solution. A yellow color indicates an optimal pH for DNA binding. If the color
of the solution is orange or violet, add 10 μl of 3 M sodium acetate, pH 5.2 solution
and mix. The color of the mix will become yellow.
2. Transfer up to 800 μl of the solution from step 1 to the GeneJET purification column.
Centrifuge for 30-60 s at 12.000 g. Discard the flow-through.
3. Add 700 μl of Wash Buffer (diluted with the ethanol as described on p. 3) to the
GeneJET purification column. Centrifuge for 30-60 s at 12.000 g. Discard the flowthrough and place the purification column back into the collection tube.
4. Centrifuge the empty GeneJET purification column for an additional 1 min at 12.000 g
to completely remove any residual wash buffer.
46
Exercise 8
5. Transfer the GeneJET purification column to a clean 1.5 ml microcentrifuge tube (not
included). Add 50 μl of Elution Buffer to the center of the GeneJET purification
column membrane, incubate for 5 min at RT and centrifuge for 1 min at 12.000 g.
6. Discard the GeneJET purification column and store the purified DNA at -20 °C.
Determine the DNA concentrations of the resulting eluate using the Nanodrop, adjust the
concentrations to 7,5 ng/µl in a total volume of 20 µl. Pipet the adjusted DNA solution into a
well of a PCR strip or microtiter plate. Subsequently the samples will be send for T-RFLP
analysis.
G. Analysis of T-RFLP results
To be able to use the results of the T-RFLP analysis we have to perform several steps of data
processing and will use the following software packages or online resources:
•
•
•
For the initial processing of the raw chromatogram file we will use the free Peak
Scanner software (http://download.sharewarecentral.com/6/271833/peak-scannersoftware.html)
For noise filtration, aligning of peaks and the matrix output file we will use a free
web-based program called T-REX (http://trex.biohpc.org).
For the statistical analysis of the data you should use the statistics package of your
choice, e.g. PRIMER (http://www.primer-e.com), R (http://www.r-project.org), PAST
(http://nhm2.uio.no/norlex/past/download.html) or whatever program you are familiar
with.
The use of the programs will be demonstrated during the course.
H. Quantitative PCR
For the qPCR analysis we are using a BioRad CFX Connect qPCR machine. We will quantify
both, archaeal and bacterial amoA genes and run standards in parallel.
qPCR reaction mixture and PCR program
Prepare a premix containing the two primers, PCR water, and the Master Mix with dNTPs,
polymerase, reaction buffer, and SYBR Green. Calculate the amount of premix based on the
equation below. For each samples resp. standard combine 24 µl of the premix with 1 µl
template DNA in a qPCR tube and seal with qPCR lids.
multiplication factor for premix =
2 x (Number of samples + Number of Standards + Blank) + 2
AOA (Coolen et al., 2007)
Reaction mixture
47
Exercise 8
Template (purified DNA, diluted to 5-10 ng/µl)
Forward primer AOA-amoAF (20 pmol/µL)
Reverse primer AOA-amoAR (20 pmol/µL)
RealQ Plus 2 Master Mix (Ampliqon)
PCR water
1 µL
0.25 µL
0.25 µL
12.5 µL
11 µL
qPCR program
Hotlid held at 105 °C
Initial:
95 °C
15 min
denaturation
Repeat 40 x:
95 °C
30 sec
denaturation
59 °C
30 sec
annealing
72 °C
30 sec
elongation
Melt curve:
65-95 °C in 0.5 °C increments; 5 sec/step
AOB (Rotthauwe et al., 1997)
Reaction mixture
Template (purified DNA, diluted to 5-10 ng/µl)
Forward primer AOB-amoA-1F (20 pmol/µL)
Reverse primer AOA-amoA-2R (20 pmol/µL)
RealQ Plus 2 Master Mix (Ampliqon)
PCR water
1 µL
0.25 µL
0.25 µL
12.5 µL
11 µL
qPCR program
Hotlid held at 105 °C
Initial:
95 °C
15 min
denaturation
Repeat 40 x:
95 °C
30 sec
denaturation
60 °C
30 sec
annealing
72 °C
30 sec
elongation
Melt curve:
65-95 °C in 0.5 °C increments; 5 sec/step
I. Analysis of qPCR results
In order to calculate absolute copy numbers for the bacterial resp. archaeal amoA gene the
fluorescence signal of the environmental samples will be compared with a calibration series of
DNA solutions with known copy numbers. The software, which is installed to run the BioRad
CFX Connect qPCR machine, will also generate the calibration curve and calculate the copy
48
Exercise 8
numbers in each sample. The use of the software and how to extract data after the run is
finished will be explained during the exercise.
With the extracted data you eventually can calculate the copy numbers of the genes in your
original samples in the unit copy numbers per mg of sediment. Therefore you need the exact
weight of the sediment samples used for the DNA extraction, the total amount of extracted
DNA and the amount of DNA used in the qPCR reaction.
III. Questions, to be considered in the report
1. What amounts of DNA did you extract from the sediment? Did they differ between
sites and depths? Do you think this could be a good measurement for prokaryotic
abundance?
2. Are there differences in the microbial communities between depths? What is with
differences between the two study sites?
3. Are there specific OTUs (operational taxonomic units) that are present in both
samples, at all depths or the opposite, that are unique for a certain depth or sampling
site?
4. How do the copy numbers of the amoA gene correlate with the measured potential
nitrification rates?
5. Which types of nitrifiers are present in the two different sites?
49
References
Reference list
Armstrong, F.A.J., Stearns, C.R., Strickland, J.D.H. (1967) The measurement of upwelling
and subsequent biological processes by means of the Technicon Autoanalyser and associated
equipment. Deep-Sea Res. 14:381–389
Bower, C. E., Holm-Hansen, T. (1980) A salicylate-hypochlorite method for determining
ammonia in seawater. Can. J. Fish. Aquat. Sci. 37: 794–798.
Castle, S.C., Morrison, C.D., Barger, N.N. (2011) Extraction of chlorophyll a from biological
soil crusts: A comparison of solvents fro spectrophotometric determination. Soil Biol.
Biochem. 43: 853-856
Cline, J.D .(1969) Spectrophotometric determination of hydrogen sulfide in natural waters.
Limnol .Oceanogr. 14: 454–458
Fossing, H., Jørgensen ,B.B. (1989) Measurement of bacterial sulfate reduction in sediments:
evaluation of a single-step chromium reduction method. Biogeochemistry 8: 205–222
Glud, R.N., Ramsing, N.B., Revsbech, N.P. (1992) Photosynthesis and photosynthesiscoupled respiration in natural bio-films quantified with microsensors. J. Phycol. 28: 51-60
Hall, P. O. J., Aller, R. C. (1992) Rapid, small-volume, flow injection analysis for ΣCO2 and
NH4+ in marine and freshwaters. Limnol. Oceanogr. 37: 1113–1119.
Jørgensen, B.B. (1978) A comparison of methods for the quantification of bacterial sulfate
reduction in coastal marine sediments. Geomicrob. J. 1: 11–27
Klimant, I., Kuhl, M., Glud, R.N., Holst, G. (1997) Optical measurements of oxygen and other
environmental parameters in microscale: Strategies and biological applications. Sensors and
Actuators B 38-39: 29-37
Kuhl, M., Polerecky, L. (2008) Functional and structural imaging of phototrophic microbial
communities and symbioses. Aquat. Microb. Ecol. 53: 99-118
Lovley, D.R., Phillips, E.J.P. (1987) Rapid assay for microbially reducible ferric iron in
aquatic sediments. Appl. Environ. Microbiol. 53: 1536–1540
Middelburg, J.J., Soetaert, K., Herman, P.M.J. (1996) Evaluation of the nitrogen isotopepairing method for measuring benthic denitrification: A simulation analysis.
Limnol. Oceanogr. 41: 1839-1844
Nielsen, L.P. (1992) Denitrification in sediment determined from nitrogen isotope pairing.
FEMS Microbiol. Ecol. 86: 357–362
Revsbech, N.P., Ward, D.M. (1983) Oxygen microelectrode that is insensitive to medium
chemical composition: Use in an acid microbial mat dominated by Cyanidium caldarium.
Appl. Environ. Microbiol. 45: 755-759
50
References
Stookey, L.L. (1970) Ferrozine - a new spectrophotometric reagent for iron. Anal. Chem. 42:
779–781
Ullman, W.J., Aller, R.C. (1982) Diffusion coefficients in near-shore marine sediments.
Limnol. Oceanogr. 27: 552-556
51
Appendix 1
Appendix 1
SCHEDULE FOR SEDIMENT COLLECTION
These will be collected on Sunday 31.07.2016. All interested in the sampling and to have a look
at the sites are welcome to join the sampling trip. Otherwise, we plan another sampling trip later.
Kærby
Ex 1: (5 cm cores)
Ex 2: (5 cm cores)
Ex 3: (5 cm cores)
Ex 4: (2.6 cm cores)
Ex 5: (5 cm cores)
Ex 6: (5 cm cores)
Ex 7: (5 cm cores)
4
6
4
4
10
4
4
52
Fællesstrand
4
6
4
4
10
4
4
Appendix 2
Appendix 2
Tables for the data obtained during the exercises
(Exercise 1, 2, 4, 6, 7 only)
53
Appendix 2
Exercise 1
Sediment characteristics
Site:___________________________
Team nr.:__________
WATER CONTENT
Alu-tray
Depth
Alu-tray weight
nr.
(cm)
(g)
+ wet sed.
+ dryt sed.
β
(g)
(g)
(%)
Core 1
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
Core 2
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
54
Appendix 2
Exercise 1
Sediment characteristics
Site:___________________________
Team nr.:__________
DENSITY and POROSITY
Depth
(cm)
Syringe
(g)
Syringe
+ 4 ml sed.
(g)
Core 1
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
Core 2
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
55
Density
(g cm-3)
Porosity
φ
Appendix 2
Exercise 1
Sediment characteristics
Site:___________________________
Team nr.:__________
ORGANIC CONTENT (loss on ignition)
Crucible
nr.
Depth
(cm)
Crucible weight
(g)
+ dry sed.
(g)
Core 1
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
Core 2
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
56
+ ash
(g)
LOI
(%)
Appendix 2
Exercise 1
Sediment characteristics
Site:___________________________
Team nr.:__________
Chlorophyll a
Core 1
Depth
(cm)
Sed weight
(g)
Abs 665o
Abs 665a
0-0.5
0.5-1
1-2
0-0.5
0.5-1
1-2
2-3
2-3
57
Abs 750o
Abs 750a
Chl. a
µg g-1
Appendix 2
Exercise 2:
Oxygen measurements
Site:___________________________
Salinity:
Core No
Team nr.:__________
Temperature:
1
2
3
4
Height
Time 1
%O2 1
Time 2
%O2 2
Time 3
%O2 3
Time 4
%O2 4
Time 5
%O2 5
Time 6
%O2 6
Time 7
%O2 7
Time 8
%O2 8
Time 9
%O2 9
Time 10
%O2 10
58
5
6
Appendix 2
Exercise 2
Flux measurements-dark
Site:___________________________
DARK
Team nr.:__________
CONCENTRATION
O2 (μM d-1)
slope
Core
nr.
NH4+ (μM)
start
end
NO2- + NO3- (μM)
start
end
1
2
3
CO2 CONCENTRATION
Standard
Conc.
Height
Core
No.
1 mM
1
2 mM
2
3 mM
3
start
Peak height
end
2mM
CO2 concentration
start
end
FLUX
Core
nr.
Height of
water phase
(cm)
Incubation
time
(hours)
O2-flux
CO2-flux
NH4+-flux
1
2
3
average
mmol m-2 d-1
59
NO2-+NO3--flux
Appendix 2
Exercise 2
Flux measurements-light
Site:___________________________
LIGHT
Core
nr.
Team nr.:__________
CONCENTRATION
O2 (μM d-1)
slope
NH4+ (μM)
start
end
NO2- + NO3- (μM)
start
end
1
2
3
CO2 CONCENTRATION
Standard
Conc.
Height
Core
No.
1 mM
1
2 mM
2
3 mM
3
start
Peak height
end
2mM
CO2 concentration
start
end
FLUX
Core
nr.
Height of
water phase
(cm)
Incubation
time
(hours)
O2-flux
CO2-flux
NH4+-flux
1
2
3
average
mmol m-2 d-1
60
NO2-+NO3--flux
Appendix 2
Exercise 4
Sulfate reduction
Site:___________________________
Time
Depth
(1)
tube+lid
(g)
cm Tube nr
core 1
start:
stop:
core 2
start:
stop:
0-1
1-2
2-4
4-6
6-8
8-10
t t
Team nr.:__________
(2)
tube+lid
+ZnAc
(g)
(3)
tube+lid
+sed+ZnAc
(g)
(4)
tube+lid
+centr sed
(g)
msed
(g)
[3-2]
msedc
(g)
[4-1]
msub
(g)
[SO42-]
nmol cm-3
SRR
nmol cm-3 d-1
[TRIS]
μmol cm-3
23
ΣSRR:
mmol m-2 d-1
ΣSRR:
mmol m-2 d-1
0-1
1-2
2-4
4-6
6-8
3717
61
Appendix 2
TCO2, DIN and SO42- profiles
Exercise 6
Site:___________________________
Team nr.:__________
TCO2 og DIN
Depth
(cm)
Core 1
TCO2
(mM)
NH4+
(μM)
NO2-+NO3(μM)
Core 2
SO42(mM)
Surface
0-1
1-2
2-3
3-4
4-5
5-6
6-8
8-10
62
TCO2
(mM)
NH4+
(μM)
NO2-+NO3(μM)
SO42(mM)
Appendix 2
Exercise 7
REDOX
Site:___________________________
Depth
(mm)
Team nr.:__________
Core 1
Eh
meas
(mV)
Eh
corr
(mV)
Core 2
Eh
meas.
(mV)
Eh
corr
(mV)
-10.0
-7.5
-5.0
-2.5
0
2.5
5.0
7.5
10
12.5
15.0
17.5
20.0
22.5
25.0
27.5
30.0
32.5
35.0
37.5
40.0
45.0
50.0
55.0
60.0
63
Eh
meas.
(mV)
Eh
corr
(mV)
Eh
meas.
(mV)
Eh
corr
(mV)
Appendix 2
Exercise 7
Particulate iron
Site:___________________________
Depth
(cm)
Sed. wt.
(mg)
Fe2+
Abs
Team nr.:__________
Fe2+
(μmol g-1)
Core 1
0.0 - 0.5
0.5 - 1.0
1.0 - 1.5
1.5 - 2.0
2.0 - 2.5
2.5 - 3.0
3.0 - 4.0
4.0 - 5.0
5.0 - 6.0
Core 2
0.0 - 0.5
0.5 - 1.0
1.0 - 1.5
1.5 - 2.0
2.0 - 2.5
2.5 - 3.0
3.0 - 4.0
4.0 - 5.0
5.0 - 6.0
Absorption coefficient:_________________________
64
tot-Fe
Abs
tot-Fe
(μmol g-1)
Fe3+
(μmol g-1)
Appendix 3
APPENDIX 3
How to do tricky calculations
A. How to estimate partitioning of electron acceptors
In exercise 2 you measured O2 uptake, which is the sum of all oxygen consuming processes:
1. aerobic respiration (incl. worm respiration)
2. reoxidation of metabolites (H2S, NH4+)
You also measured CO2 prod, which is the sum of all heterotrophic processes:
1. fermentation
2. respiration (aerobic + anaerobic)
From these you can estimate the respiratory quotient, RQ = CO2 prod/O2 uptake. If RQ > 1,
there is a net storage of reduction equivalents (e.g. FeS) and if RQ < 1, there is a net
consumption of reduction equivalents.
We know (if fermentation is ignored) that:
CO2 prod = O2 resp + NO3 resp + Mn resp + Fe resp + SO42- resp
You have measured CO2 prod, NO3 resp and SO42- resp. We can for now ignore Mn resp. Fe
resp can be deduced from the Fe(III) content (see below). Based on these calculations, you can
estimate how much (in %) the various electron acceptors contribute to total CO2 production.
The remainder is an estimate of aerobic respiration:
O2 resp = CO2 prod - NO3 resp - Fe resp - SO42- resp
You need to do all the calculations in carbon units according to the conversions below:
O2 reduction x 1 = CO2 production
NO3 reduction x 5/4 = CO2 production
(MnO2 reduction x 1/2 = CO2 production)
FeOOH reduction x 1/4 = CO2 production
SO42- reduction x 2 = CO2 production
B. How to estimate Fe resp (FeR)
We know from above that total anaerobic C-oxidation is (Mn resp ignored):
NO3 resp + Fe resp + SO42- resp
FeR in % of anaerobic C-oxidation is then:
(Fe resp x 100)/(NO3 resp + Fe resp + SO42- resp)
From the literature we know that:
%FeR = 1 – exp(-0.056 [FeIII])
Where Fe[III] is measured reactive oxidized iron.
65
Appendix 3
C. How to estimate nitrification and denitrification from porewater profiles
1. C:N ratio of organic matter being degraded can be estimated from steady state porewater
profiles of CO2 and NH4+ because these provide the outcome from degradation (reaction rates)
and transport.
The reaction stoichiometry in diffusion dominated sediment is Rc/Rn = Dsc/Dsn x dCc/dCn,
where R is the reaction rate; dCc/dCn is the slope of a
porewater TCO2 vs. NH4+ plot; Ds is sediment
diffusion coefficient. In other words, the ratio of net
TCO2 and NH4+ production (C:N ratio of organic
matter being mineralized) is equal to the slope of the
TCO2 vs. NH4+ plot multiplied by the ratio between
transport of major ions, i.e., for molecular diffusion:
DHCO3-/DNH4+ = 0.59. In the bioturbated zone of
permeable sandy sediment there is a significant
advective component (Dsc/Dsn → 1).
This means that the C:N ratio of organic matter being
degraded is:
Non-bioturbated:
dCc/dCn x 0.59
Bioturbated:
dCc/dCn
2. Net NH4+ production (Rn) in the sediment can be estimated from measured CO2 production
(Rc = CO2 efflux) as:
Rn = Rc/( Dsc/Dsn x dCc/dCn)
3. Nitrification in the sediment can be estimated from the Rn in excess of measured NH4+ efflux
as:
Nitr = Rn - NH4+ efflux
4. Denitrification in the sediment can be estimated from the Nitr in excess of measured NO3efflux as:
Denitr = Nitr - NO3- efflux
66
Appendix 4
APPENDIX 4
PRECAUTIONS WHEN WORKING WITH RADIOACTIVE MATERIALS
Be sure that you have read and understood these instructions before you start the work and
think carefully about what you are doing.
1. The work area (i.e. table or bench) must be covered with plastic (clear plastic on
bottom, black plastic on top) which is taped to the table. Place absorbing paper on top of
the plastic cover. All items can be found in the laboratory.
2. A contained of suitable size is used for waste – place a plastic bag in the container.
3. Always wear a lab coat in the laboratory – the coat is not to be worn outside the
laboratory.
4. Always wear gloves while working with radioactive materials. Never touch
uncontaminated items with gloves that can be contaminated.
5. Persons with open wounds on their hands must not work with radioactive materials –
even if the wound is covered by a patch.
6. All work with radioactive sources must be carried out in a tray containing an2-3 layers
of absorbent material.
7. Glassware and other equipment used for radioactive materials must be kept separate
from the rest of the equipment.
8. The following items must not be brought into or used in laboratories where open
radioactive sources are being used:
a) Food and drinks
b) Cigarettes and other tobacco products
c) Handbags
d) Lipsticks and other cosmetics
e) Cutlery, cups, glasses, plates etc.
9. When the laboratory work is finished, everything must be cleaning thoroughly
a) Place all contaminated equipment in the special container for washing.
b) All waste (including absorbent paper, gloves, plastic bags, tissue remains,
sediment etc.) are packed into the plastic bag in the waste container.
c) Wash your hands meticulously.
10. All spills of radioactive materials must be reported immediately to the instructor.
67
Appendix 4
68
Appendix 4
General Health & Safety Regulations
for laboratory exercises at The Institute of Biology
Good laboratory practice
1. Plan your experimental work carefully. Read the Laboratory Manual carefully and
follow the delivery notices before starting to work. Plan your work and keep your working
bench clean. You are not allowed to bring your coat or bag into the lab.
2. Be well prepared. Even if you have prepared your experimental work, accidents may
happen. Be sure to know where to find the fire extinguishers, emergency showers and eye
wash bottles and know how to use these.
3. Wear a lab coat. When working with radioactive isotopes, you will receive a lab coat.
When working with other experiments it is recommended that you bring your own lab coat.
Used lab coats can be bought for a reasonable price at ‘Odense Kittelservice’.
4. You are not allowed to smoke, eat or drink in the laboratories.
5. Use personal protection correctly. Wear gloves when needed and always be very careful
to protect your eyes, hearing and respiration airways
6. Handle chemicals correctly. Never pour the chemicals back into the original bottle. Use
clean spoons when weighing. Keep your hands on the label when pouring. Clean the
balance if you have a spill.
7. Put labels with your name and date on your solutions.
8. Waste must be collected in the correct manner. Chemical waste must be contained in
labelled plastic jars. Organic waste, e.g. tissue and blood must be put in a plastic bag, which
will then be destroyed.
9. Wash your hands during your work in the laboratory and when you have finished your
experiments. If you wash your hands very often you may experience that your skin
becomes dry, which increases the danger of having open wounds on the skin. Therefore use
skin crème whenever you wash your hands.
10. Clean up before leaving your working place. Wipe down your workbench, all glass items
must be rinsed before being placed into the grey plastic trays next to the sink.
69
Appendix 4
Chemicals and materials
The practice data sheets of every chemical in use will be present in the laboratory. Data sheets
give information about the danger of the drug and how it is handled. At each exercise the most
important information in the data sheets are available separately. This is to give an overview and
does not imply that it is unnecessary to read the datasheets.
Gloves
If used incorrectly they can procure a false sense of safety. The gloves are only used while
working with chemicals that you need protection against. Take them off before opening a door,
answering the phone or using the computer etc. Also take them off and throw them away if you
discover that you have spilt any chemical on them. The following types will be available:
Latex: To be used when working with aqueous solutions, as they are made of natural rubber that
contains latex proteins. It is common to suffer an allergic reaction from these proteins and this
reaction is often worsened if powder is added to the glove.
Vinyl: Also to be used when working with aqueous solutions.
Nitrile: To be used with some organic solutions.
Barrier-glove: Resistant to most chemicals but because of its size it is difficult to work with.
Wear a latex glove over the barrier glove, which gives a better feeling with things.
Insurance
The working environment law covers students but the Working Risk Insurance does not cover
them. This implies that students must follow the same rules as the employees in a laboratory, but
if they are hurt, they are not entitled to compensation. Therefore you must take out your own
accident insurance. Some insurance companies may add an excess to the policy for claims made
relating to laboratory work. The service shop of the “Studenterråd” can help you with the proper
insurance policy.
70
Appendix 5
APPENDIX 5
Guidelines for writing a report
Scientific reports are usually divided into the following sections:
1.
2.
3.
4.
5.
Introduction
Materials and methods
Results
Discussion and conclusions
References
We want you to follow this structure as closely as possible in the reports of the present
course.
Re. 1. Here you state the scientific background, current status of the topic and
objectives of the present work. It is here scientific papers argue for the necessity of a
particular research area.
Re. 2. Describe first the sampling sites. Then the methods used (both in the field
and in the laboratory) are described. The tools, equipment, chemicals etc. used for the
work must be mentioned. It is not necessary to describe standard methods that have been
published. The relevant publications where the methods are described in detail can instead
be cited – with mention of possible modifications.
Re. 3. Only the treated data should be presented in the results section to make life
easier for the instructors. Raw data (if they have to be included) must be placed in
Appendices at the end of the report. Present the results in the form of easy understandable
tables and figures. Make figures with readable axis titles and use sufficiently thick lines for
curves (in case figures must be reduced in size). All tables and figures must be numbered
(cite them in the text) and supplied with an explanatory legend. Explain what the results
show in the text without interpretations and conclusions (save these for the discussion).
Re. 4. Discuss the results. What do they show? Relate the results to and cite the
available literature. Make relationships based on your own data that can explain the
phenomena and check if they are consistent with the general knowledge obtained from the
literature. Try to explain if and why the results disagree with the generally accepted rules
and laws. Unless everything is perfect, this part may account for a large part of the
discussion.
Re. 5. References must be cited in the text as (Henriksen et al., 1981) or if they are
an active part of a sentence as Henriksen et al. (1981). If more than one reference is given,
it must be like: (McCave, 1976; Henriksen et al., 1981). Make the reference list at the end
of the report according to the following format:
Henriksen, K., Hansen, J.I. & Blackburn, T.H. (1981). Rates of nitrification, distribution of
nitrifying bacteria, and nitrate fluxes in different types of sediment from Danish
waters. Mar. Biol. 61: 299-304.
McCave, I.N. (1976). The benthic boundary layer. Plenum Press. New York.
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