Berlese-Tullgren Arthropod Extraction From Soil and Litter
Berlese-Tullgren extraction is used to separate arthropods from litter and soil samples. Arthropods can be extracted from almost any moist substrate (nest material, decaying wood, fungi, etc.) using a Berlese-Tullgren or similar type of apparatus. Unlike many of the arthropod sampling protocols presented here, the Berlese-Tullgren apparatus extracts arthropods from a standard volume of substrate. The quantitative data (number of specimens) collected are scalable abundance measures for each species extracted from the sample.
A modified Tullgren apparatus, based on the Berlese funnel (thus often called Berlese-Tullgren funnel), and its various modifications, is the most commonly used method for separating arthropods from soil and litter. The essential component of these extractors is a sample container with wire mesh or screening on the bottom, a metal or plastic funnel in which, or over which, the sample container is placed and a collecting vessel below the funnel which usually contains a liquid preservative, generally 70-80% ethanol. A source of heat and desiccation (light bulb) is placed above the sample. The objective is to create a steep gradient of temperature and moisture throughout the sample. Arthropods react to the heat and desiccation by moving downward (away from the heat) and eventually fall through the screen at the bottom into the preservative. Cheesecloth below the sample and/or a baffle in the funnel can reduce debris from the sample falling into the preservative as the sample dries out or is agitated by the movement of larger organisms. The wattage of light bulb used in the Berlese-Tullgren funnel depends on the size and water content of the sample and on the distance of the bulb from the sample surface.
Commonly used modifications of the Berlese-Tullgren funnel in which the extraction efficiency is improved by enhancing humidity and temperature gradients include the Macfadyen high-gradient funnel, the Kempson apparatus and the Merchant-Crossley extractor. For qualitative sampling of large volumes of litter a light-weight, collapsible and easily transportable Berlese-Tullgren funnel is available.
Behavioural extractors extract only the active stages of arthropods. Eggs, prelarva, and quiescent pre-molt stages remain in the sample. In arid soils, deep soils, and for arboreal habitats such as twigs, behavioural extractors are inefficient. Processing using behavioural extractors should be carried out as soon as possible after field collection of samples. At a minimum, any study should incorporate behavioural extraction of soil arthropods. We recommend the following standards for sampling microarthropods in ecozones of Canada.
Equipment
Sifters (litter sampling)
pillowcases (litter sampling)
garden hand rake, small 30 cm length (litter sampling)
soil corer 5x15 cm (soil sampling)
gloves
scissors or knife
Berlese funnels (12-15)
supports for funnels
light bulbs - 25 watt
cheesecloth
alcohol
specimen bags (i.e., plastic whirlpak type)
label paper
pencil
Methods
Litter Sampling
Litter is the decaying plant and animal material that accumulates on the soil surface. The quantity and quality of litter and the associated arthropod species assemblages vary with the plant community, time of year, precipitation, temperature, and humidity.
- The litter is raked to the level of the soil surface and put through the sifter to remove large fragments of debris and to concentrate the arthropods until 6 litres of sifted litter are obtained.
- The sifted litter is placed in a pillowcase, labelled and tied.
- Four such samples are taken for each selected sampling period in that habitat or microhabitat.
- Samples can be kept in the pillowcases in the laboratory for up to 4-5 days as long as they are turned frequently and not exposed to excess moisture or extreme temperatures.
- Samples are processed by dividing each 6 litre sample into 3 parts of approximately 2 litres each.
- Each 2 litre sample is placed in a separate Berlese-Tullgren funnel and the arthropods extracted according to the instructions given under Berlese- Tullgren extraction below.
- The specimens extracted from each 2 litre partition of a sample (3 funnels) represent a single sample and can be recombined for ease of storage and sample tracking.
- General litter samples should be taken about every 2 months.
Soil Sampling
The sample unit can be a core or cube, ranging from 2.5 cm to 10 cm in diameter, or can be a specific volume of substrate. The most common core size used in recent publications on quantitative sampling for soil arthropods in agroecosystems is 5 cm diameter, to a depth of 15 cm in soil. However, the core size used will depend on the habitat, for instance, in humic soils and moss a 10 cm corer is preferable to reduce compaction, but these samples would require a larger extractor. The corer is constructed from case-hardened steel pipe with 5.0 cm internal diameter and is fitted with an anti-compression ring in the coring pipe. A pipe connects the corer with a tee handle and has a cross foot pedal at the top of the corer.
- If necessary, gently remove growing vegetation from sample plot using scissors or knife.
- Use the standard 5 cm diameter corer (bulk density corer, or equivalent) to remove soil sample to depth of A horizon (usually 5 to 20 cm depth).
- Divide samples into layers of 5 cm depth and bag (brown wax paper bags, cloth bags, or if using plastic leave top open) and label separately.
- Keep samples cool until arthropods can be extracted - preferably the same day as collected. If samples cannot be extracted on same day they should be maintained at 4-10° C. Arthropod extraction techniques follow under Sample Processing.
- Three to 5 replicates should be taken from each habitat within a site, and from each plot in a management regime.
- Samples should be collected a minimum of 3 times in the growing season (more frequent in the early characterization stage; twice yearly when species richness and population trends are established).
Sample Processing
Use modified Berlese-Tullgren, such as Macfadyen High Gradient, or Kempson, Lloyd & Gelhardi modifications to behaviourally extract microarthropods from soil or litter samples. The most effective extractor for microarthropods is the Macfadyen High Gradient extractor or its refinements. However, variations between the different modifications of the Macfadyen and modified Berlese-Tullgren are quantitative rather than qualitative. For this reason, any of the above extractors is acceptable if the following protocols are adhered to:
- Record wet weight of sample.
- Unless sample is very compact, place layer of cheese-cloth or fine screen over screen of extractor to prevent soil particles being dislodged into the extractant.
- Invert 5 cm soil sub-samples over screen of extractor, i.e., the top of the sample should be closest to the screen.
- Ensure an air-space between sample and collecting funnel, to avoid condensation in the funnel.
- Extract into 75% ethyl alcohol in jar or plastic bag at bottom of funnel.
- Use a dimmer (rheostat) attached to the heating source to establish gradient between top and bottom of sample; this gradient should reach about 30° C by the end of the extraction period. NOTE: dimmers should be temperature calibrated for several days to ensure uniformity between funnels. Alternatively, place extractor in refrigerated cold room (less than 15° C), and gradient will be established.
- Extract for 4 to 7 days depending on moisture and organic matter content of sample; if in doubt, extract for 7 days.
- Record dry weight of sample.
- Retain sample for determination of soil abiotic characteristics.
Sample labels must be placed inside the bag with the specimens. Change the alcohol in about 7 days, and if possible, refrigerate or freeze the samples to await further processing. Samples in storage should be double bagged.
Further Reading
Anderson (1996), Behan Pelletier et al (1996), Crossley and Blair (1991), Edwards (1991), Kempson et al. (1963), Kethley, (1991), Krantz (1978), Macfadyen (1962), Marshall et al.(1994), Martin, (1977), Merchant and Crossley (1970), Norton (1986), Norton and Kethley (1988), Seastedt & Crossley (1978), Steyskal et ai. (1986), Walter et al. (1987), Wheeler and McHugh (1987)
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