Biobasis Nuuk 2009 protocol. Microarthropods and decomposition
Fig. 1. Situation of plots at
Kobbefjord for microarthropod and litterbags: Salix glauca
; Silene
acaulis 
; Empetrum
nigrum 
; Loiseleuria
procumbens 
.
Show the Kobbefjord plots on Google maps.
Microarthropods
Species to be monitored
All microarthropods: Collembola at species level and mites at order level.
Frequency of sampling
Sampling is performed three times during the season corresponding to: spring (after snow melt), summer and autumn (before the snow appears). To avoid having very wet samples, for which extraction will be very slow, samples can be collected after water content is reduced.
Fig. 2. Microarthropods and litterbag sampling grid in 10x10 m plots with grid size: 0.5 x 0.5 m.
Equipment to be used
- Map/GPS with positions of plots
- Soil auger
- 64 microcosms tubes made of Plexiglas (height 5.5 cm and diameter 6 cm)
- 128 pieces DBIdut lids (size 89B)
- Measure
- Shears
- Knife to cut roots etc.
- Pre-printed labels, incl. Date, Plot Id, plant community (Silene, Salix, Empetrum, Loiseleuria), Replicate Id., Initials
- Transportation boxes
Location and marking of study plots
The sampling programme consists of collecting microarthropod samples from:
4 habitats * 2 plots * 8 subsamples * 3 sampling occasion = 192 samples.
The sampling occasions may coincide with the three litter bag
collections, if feasible. To ensure enough undisturbed sampling points
for several years each plot is divided into a ½ meter square grid (Fig.
2).
The coordinates (x,y)=(0 m, 0 m) is the exact position of the iron
corner stick with written label. An Excel
table with random sampling points includes these coordinates, for
each subsample. The sampling points are sorted according to the
x-coordinate. The random sampling Excel table with x (column) and y
(row) coordinates include 10 subsamples to be used for the litterbags
and of those 8 are used for the microarthropod soil cores. For
practical reasons the same set of random numbers are used for all 8
plots at each sampling occasion.
Sampling method
- The soil auger including two microcosm tubes is closed and ready for use.
- The point of sampling is found using the random sampling table and a measuring tape
- The soil auger is placed vertically at the sampling point so it touches the soil surface.
- At sites with dense vegetation it may be necessary to use a knife to cut around the soil auger before pushing it down into the soil or peat. Take care not to damage the soil/peat core.
- Push the soil auger vertically 5.5 cm downwards so that the lowest tube is just filled with soil. The soil surface shall level the upper rim of the lowest tube. The soil auger is open in the top so that you can follow how the soil appears in the tube. The upper tube functions only to fix the lower tube. While pushing the soil auger down turn it from side to side thereby avoiding compressing the soil in the tube.
- Tilt the soil auger from side to side loosen the soil core at the bottom and take care when you draw the soil auger including the soil core up.
- Open the soil auger and carefully remove the tube including the soil core. Place a labelled DBIdut lid at the top immediately to avoid that organism on the soil escape.
- Turn the tube around and cut surplus soil away so the soil surface levels the bottom of the lower tube. Place a DBIdut lid in the bottom of the tube.
- Place the tubes in a box top of the sample up.
Store the samples at low temperature in a shadowed place, and avoid bumping during transportation. On arrival to the lab the samples are stored in the dark at 5ºC until extraction –not later than two days after sampling.
Laboratory work
Extraction of microarthropods
The capacity for extraction is limited so it may be necessary to run the extraction more than once. To account for differences due to longer storage etc. between two extraction batches the principle of “blocking” is followed. Thus, a fraction of sub-samples, extraction block no. 1, with e.g. half of the samples from a sampling plot are randomly selected for the first extraction and the remaining other half, block no. 2, is stored at 5 oC until extractors are ready. The blocking enables a statistically valid assessment of the possible differences between the blocks, i.e. the two extraction sessions.
Equipment
- Extractor with temperature sensor and data logger
- Insulation foam
- X number of soil samples
- X number of meshes with a mesh size of 1x1 mm
- X number of extraction cups
- Saturated benzoic acid (14.5 g benzoic acid and approx. 1 ml detergent per 5 L)
- Manual for extractor
- Detergent
- X number of lids for extraction cups
- Incubator
- 96% alcohol (may be denatured if pure alcohol is not
available)
- Small cups for transportation of extracted organisms in extraction liquid and alcohol
Extraction procedure
1. One day before extraction: Start the refrigerator connected to the extractor as the samples may not be stored at temperatures higher than 5°C.
2. At the day of extraction: Bring the samples carefully from the storage room to the extraction room.
3. Fill all extraction cups with a saturated solution of benzoic acid + detergent (14.5 g in 5 L water + roughly 1 ml) up to 0.5 cm.
4. For each sample: Take a tube containing a soil sample. Move the label from the lid to the extraction cup. Carefully remove the upper lid and place the mesh on the tube with the sample.
5. Place a suitable cup above the soil sample unit and turn the cup with the sample around.
6. Remove the DBIdut lid from the bottom and sweep surplus soil down into the cup.
7. Place the microcosm tube with a net on an extraction cup with the benzoic acid.
8. Pour the surplus soil into the soil sample.
9. Carefully place the microcosm tube with the soil surface facing downwards into the extractor.
10. Place the insulating material around the samples when all samples are in place in the extractor. The insulation around the tubes must be placed carefully so that no soil particles will drop into the cups.
11. Connect one temperature sensor in the extractor for regulation of temperature and connect three temperature sensors to a data logger to follow the temperature during the extraction in the benzoic acid liquid, just above the mesh and on surface of the soil sample facing the heater.
12. Close the extractor.
13. Turn on the extractor and press the green start button.
The extractor will now
heat the samples according to this schedule:
- 30°C for 48 hours
- 40° C for 48 hours
- 50° C for 48 hours
- 60° C for 24 hours, terminated manually by switching off the power supply,
but it may be continued until all the samples are dry on the
down-facing surface
on the mesh.
The cooling system should ensure that the temperature of the benzoic
acid solution is minimum 4 oC and maximum 20 oC
throughout the extraction.
14. Samples with high organic matter such as peat should be divided into two horizons, e.g. the lower 3 and the upper 3 cm, and extracted independently. The samples may be divided either from the beginning of the extraction or at the temperature, e.g. 50 oC, where the upper 2 cm has become completely dry. In the latter case the upper 2-3 centimetre is cut off the sample and discarded provided they are completely dry. The sample is removed from the extractor during this operation, to ensure that no sample material will drop into the extraction beaker.
15. The extraction is stopped manually by turning the power off.
16. Check that the samples are dry on the surface facing downwards after termination of the pre-programmed extraction process. If some samples are still wet continue the extraction at 60 ºC until the samples are dry. Samples with high organic content can be divided in two, to ensure complete extraction.
17. Turn off the extractor and data logger.
18. Throw the soil away
19. Brush the nets clean. Wash the tubes.
20. Add a drop of detergent to all cups in the extractor to reduce the surface tension of the benzoic acid.
21. Take the cups up from the extractor and put lids on. If there are organisms on the sides of the cups then flush or move them into the benzoic acid with a brush.
22. Put all cups with lids on into a heating oven for 24 hours at 50 °C. The heat and the detergent ensure that all organisms sink to the bottom.
23. Pour the content from each cup into plastic cups and fill up with 96% alcohol to one part water to two parts of alcohol (resulting in approx. 70% alcohol). If necessary to obtain this proportion divide the sample into two plastic cups.
24. Store the samples with lids closed tightly until filtering at NERI or GNIR.
25. Draw a graph of the extraction in Excel and save it on the server drives. The curves are used when evaluating the results.
Decomposition
Organic material used for monitoring
Filter paper is generally used for litterbags. A batch of litterbags with Salix glauca leaves available at GINR is positioned together with the filter paper litterbags for the 2009 litterbag study only.
Frequency of sampling
Three times during the season depending on the stage of decomposition.
Equipment to be used
- Map/GPS with positions of plots according to random allocation table
- Litterbags filled with 2 g VWR filter paper corresponding to 4
round pieces of filter paper, 9 cm in diameter.
- Knife
- Preprinted labels, incl. Date, Plot Id, subsample no., x-y coordinates, plant community (Silene, Salix, Empetrum, Loiseleuria), Replicate Id., Initials
- Transportation boxes
Litterbags
VWR filter paper is used as a surrogate for indigenous litter and filled into litterbags. Each litterbag (5 mm mesh and 10 by 10 cm) is filled with 2 g of filter paper (corresponding to 4 pieces of filter paper, 9 cm in diameter). Each filterpaper is cut into 4 pieces:
Each plot holds 10 litterbags buried horizontally 3-5 cm into the soil/peat. Each bag is identified with a unique labelling embossed on a plastic tag and placed inside the closed bag including: sampling occasion (date), habitat, plot number, subsample no., x coordinate meter, y coordinate meter. The litterbags are left open in one end for ease of emptying and further processing. Marker sticks are used to locate the litterbags at the 3 sampling occasions to ease the idenfication and retrieval.
Bag design and litter
When employing a new batch of filter-paper the dry-weight (DW) is determined by taking about 5 representative samples and dry them in an oven at 50o C until constant weight. As the filterpaper take up water from the surrounding air they should be stored in an exicator if they cannot be weighed immediately after drying in the oven.
Location and marking of sampling plots
During the 2009 sampling season each of the 4 habitats will be characterised concerning pH, texture and plant communities. About 0.5 kg soil is collected and send for analysis in Foulum, Denmark according to the soil sampling procedure.
Litterbags are placed according to a random sampling scheme in the field monitoring site each autumn and the last (third) batch is collected at the same time as the placement of the next year’s set. The litter bags are placed in a manner ensuring good, natural contact with the underlying litter layer. The bags are covered by some of the surrounding litter if the habitat includes a natural litter layer. If the habitat consists of peat, the bags will be put into the peat layer at max 5 cm depth. In soil habitats they are covered by approx. 3 cm soil. In this case a slit is made with a shovel and the litterbag is slided into the slit and covered by the soil.
The 30 litter bags in each plot are placed in a ½ m square grid. A stick is used to fix the litter bag to the soil for easy retrieval.
Sampling method
Three sets of litter bags will be collected and brought to the lab
for measurement between spring and autumn. When roughly 50% of the
original plant material has disappeared from the litter bags in autumn
the 3rd set of bags are collected. If less than 30% has been decomposed
in the autumn, another season may be added to the duration of the
decomposition period to obtain a higher decomposition rate around 50%.
Each set of litter bags consists of:
4 habitats x 2 replicates x 10 litter bags (subsamples) * 3 sampling
occasions = 240 litter bags.
To check the state of decomposition in addition 10 extra bags are
placed at each habitat, i.e. totally 40, to monitor the current level
of decomposition. Only one replicate plot may be chosen for the
monitoring purpose. Three of the these monitor litterbags may be
collected corresponding to each sampling occasion and measured before a
final date for collection is decided. Decomposition should be
terminated when the remaining dry-weight is about 30%, so the rate of
decomposition of the first and second sampling occasion would be about
75% and 50%.
Laboratory work
After collection the filterpaper is oven dried in paper bags at 50oC for 24 hours or longer to ensure the mass (DW) is constant. Any mosses, lichens, fine roots, or other plant parts that have grown into the bags should be removed prior to weighing.
Input of data into database
- Labelling of the batch of filterpaper with: batch no., date
- Dry-weight of filterpaper after decomposition
- Daily temperatures and precipitation best quality data covering the exact location or GeoBasis data.
Subproject organisation
- Josephine Nymand, +299 36 12 34, and staff at Naturinstituttet, are performing the field work in Greenland and installs and analyses the litterbags from the study site for DW.
- NERI/AU technical staff is handling samples of extracted animals and does species density estimation and identification (Zdenek Gavor and Elin Jørgensen).
- Paul Henning Krogh, +45 89 20 15 88, is responsible sub-project manager, and performs data analyses and reporting.
