Standardised protocols for soil fauna extraction and a call for cross-lab implementation

Understanding the status and global trends of soil invertebrate diversity requires accurate and comparable data across geographical regions. However, soil animal extraction approaches still vary among laboratories, and no commonly accepted, openly available and well-documented protocols exist across taxa. Here, we present harmonised methodologies, assembled by an international group of experts, for extracting soil- and litter-inhabiting nematodes, enchytraeids, microarthropods, and larger invertebrates. The protocols are illustrated with images and videos, and include advice for overcoming the most frequent issues (‘expert tips’) for increasing extraction efficiency and reproducibility. In addition, we provide results from two pilot experiments on nematode and large invertebrate extractions. First, we show that using two layers of milk filters instead of one in wet extraction of nematodes yields very similar extraction efficiency and has little effect on the sample cleanliness. Second, we demonstrate that on average approximately 31.3% of large soil animals (3 mm and more in body length) are overlooked during hand sorting but were then captured in heat extraction of the sorted samples across three geographically distinct laboratories. The extent of bias varied by collection team and taxon, with particularly high collection bias for Pseudoscorpiones (64.4% overlooked), Diptera (62.9%) and Coleoptera larvae (57.1%). Observed differences among the laboratories call for standardized ring tests of extraction efficiency across regions and research groups. Overall, we provide openly available protocols for assessing soil animal taxa in ecological studies worldwide, facilitating comparisons for a better understanding of the dynamics of global soil biodiversity.

Soil animals including micro-, meso-, and macrofauna play key roles in ecosystem functioning and securing food provision, water cleaning and climate regulation (Lavelle and Spain 2002). Current changes in climate, land use, and pollution threaten soil animal diversity and related ecosystem functions (FAO, 2020; Geisen et al., 2019). Therefore, monitoring soil animals is crucial to detect potential changes. Recently, several initiatives to study the distribution and abundance of soil animals at global scale have been accomplished (Fierer et al., 2009; Phillips et al., 2019; van den Hoogen et al., 2019; Lavelle et al., 2022; Potapov et al., 2023). Despite important insights, these studies have also drawbacks, namely: (1) they consider different groups of organisms independently, (2) they provide snapshot data with very few temporal series available, and (3) almost all of them are based on variable sampling and extraction methods (Eisenhauer et al., 2020). The lack of unified knowledge on the dynamics of soil animal diversity and abundance restrains the drafting of legislation on their protection (Guerra et al., 2021a). As a result, the main challenge remains to include multiple soil animal groups in a regular global monitoring and to use one harmonized methodology covering different types of habitats including less studied areas of the world such as tropical regions (Cameron et al., 2019; Guerra et al., 2020). Recently established, the Soil BON Foodweb (SBF) is a global monitoring initiative collecting data on soil micro-, meso-, and macrofauna communities at a global scale using a unified methodology (Potapov et al., 2022, www.soilbonfoodweb.org). The main goals of the initiative are to explore drivers and functions of soil animal diversity, including interactions in soil food webs, and to assess the efficiency of current conservation measures for protection of soil animals at a global scale by following the Soil BON sampling design (Guerra et al., 2021b). To ensure comparability of data coming from different laboratories, an expert team was established to summarize existing methodologies that would be feasible to apply across different countries, from polar regions to the tropics. The following considerations have been decisive in the selection of extraction methods: (i) cheap or easily self-producible and easy-in-use equipment; (ii) technical feasibility of implementation across main soil types and climatic zones including remote sites; (iii) possibility of biomass-per-area assessment (quantitative) and (iv) possibility for subsequent evaluation of diversity from collected animals with molecular methods. The expert team deliberately focused on traditional animal extraction methods instead of environmental DNA (eDNA) sampling, with the former allowing for biomass quantification and avoiding cross-border transfer of soils for centralized analyses (therefore relying on local expertise and developing local capacity). Furthermore, recent EU-scale assessments highlighted potentially strong methodological biases of eDNA methods which apparently reflect very different information on biodiversity than traditional ones (Köninger et al., 2025). While this supports the use of traditional methods, these also vary in their efficiency among taxa (McSorley and Walter, 1991). Therefore, four complementary extraction approaches, including wet and dry funnel extractions and hand sorting, were adopted depending on the target animal groups. Here, we briefly review existing extraction alternatives and then focus on the protocols, including video descriptions, of selected extraction methods for nematodes, enchytraeids, microarthropods and larger invertebrates. We explain the rationale for selecting a specific method by the expert team, indicate its pros and cons and describe steps critical to maximize extraction efficiency based on expert knowledge. Finally, we provide results from two experiments testing variations of extraction methods or evaluating extraction efficiency to substantiate some of our points. We finish with perspectives for further testing and development using the methods introduced here. Our primary goal was to deliver openly available protocols maximizing reproducibility of soil animal extractions across the global monitoring network and other soil fauna assessments. Nematode extraction Free-living soil nematodes are worm-like metazoans with body lengths ranging from 0.15 to 5 mm and body widths from 2 to 100 µm, which places them into the category of microfauna (Orgiazzi, 2016; Orgiazzi et al., 2015; Swift et al., 1979). Many different methods are available to extract nematodes from soils and have been comprehensively reviewed (Nijs, 2013). The main differences are in the extraction principle, such as live extraction based on their active movement (e.g., Baermann funnel), different sedimentation rate due to differences in density of soil particles versus nematodes (e.g., Oostenbrink elutriator) or separation based on the size of the nematode (e.g., Cobb sieves). None of these methods is perfect, and there are trade-offs between efficiency, costs, sample size, duration, and cleanliness of the sample (Nils, 2013). A global and extensive survey of nematode diversity requires avoiding complex devices, e.g., the Oostenbrink elutriator, and instead prioritizing simple methods that use inexpensive and widely available materials. The Baermann Funnel method (Baermann, 1917) with several adaptations (Cesarz et al., 2019; Verschoor and de Goede, 2000) is a simple method and the choice of the expert team. Soil with nematodes is spread on a milk filter on a coarse sieve, and is then placed in an empty funnel, which is then cautiously filled with water. Within the next 72h, the nematodes move actively downward through the soil, and then sink down passively in the free water body of the funnel. The nematodes are collected in a tube attached to the bottom of the funnel. Our recommended extraction time is three consecutive days (72 h) to include slow-moving nematodes. To counteract negative effects of possible losses due to predation, nematodes are collected and fixed in 24 h intervals by exchanging the vial with a fresh one and cautiously adding some water from above (Supplementary Materials 1: Wet cold extractors). The method is biased against certain nematode species that migrate upward rather than downward, e.g. Bursaphelenchus cocophilus and some insect parasites (Van Bezooijen, 2006). Furthermore, this method selects against very slow or inactive nematodes (dauerlarvae and cryptobiotic state) and nematode eggs. Compared to other methods, the Baermann Funnel technique is efficient in isolating nematodes from soils with high clay and organic matter content, which gives a relatively clean sample. The cleanliness of the sample, i.e. minimal amount of soil particles, is an important criterion in the Soil BON Foodweb, since it allows for efficient implementation of image analysis techniques (Potapov et al., 2022). Large amounts of soil particles make it difficult for morphological identification, counting and DNA extraction. Therefore, the use of a milk filter or a cheese cloth placed between soil sample and sieve is mandatory (Cesarz et al., 2019). Both materials are not always easy to obtain and differ in their characteristics depending on the supplier, but based on cost consideration, milk filters were selected by the Soil BON Foodweb Team. The only disadvantage is that species with a highly excrescent cuticle (e.g., Criconematidae) are more likely to get stuck on the filter materials (Ferris et al., 2004; Olson et al., 2017). The use of commercial paper towels is not recommended as it may reduce the extraction efficiency by more than 50% (Cesarz et al., 2019). A prerequisite for all sorts of filters is, however, the careful handling of the soil sample so that particles do not fall through during the sample installation or watering. To further improve cleanliness of the extraction, we tested the extraction efficiency using two layers of milk filters to decrease the amount of soil particles in the collection vial (see section Pilot Experiment 1 ). Another aspect to consider is that the thickness of the soil layer (i.e., the height of soil in the funnel) and its aggregation significantly influences nematode extraction efficiency (Cesarz et al., 2019). Due to their small size, nematodes can only move small distances, so large amounts of soil, and more specifically the soil height on the sieve, has a negative effect on the extraction efficiency (Cesarz et al., 2019; Nijs, 2013; Van Bezooijen, 2006). Cesarz et al., (2019) recommended using a maximum of about 1 cm of soil height during the extraction process. Mobility can be increased significantly when soil aggregates are broken up by gentle sieving at 5 mm or even 2 mm (Cesarz et al., 2019). In the Soil BON Foodweb protocol, we opted for gentle homogenization of the sample by hand to minimise processing time and soil disturbance. Since the extraction apparatus is filled with water and comes into contact with living organisms, we recommend using only non-demineralised water of drinking quality. Nematodes are killed by heat shock, with a sudden rise of water temperature from 20°C to 60°C by holding the rack with vials for 60s into a pot with almost boiling water or by adding boiling water in a volume ratio of 1:1 to each vial. For long-term storage and morphological identification, 4% formalin (formaldehyde) or 1-2 drops of TAF [14.9% formalin stock (35%), 1.5% triethanolamine, 83.6% distilled water] is used, but it restricts DNA extraction which is a goal of the Soil BON Foodweb. Therefore, the use of 96% ethanol is recommended instead, despite the fact that alcohol dehydrates the tissues to such an extent that morphological structures are no longer preserved and make the morphological identification impossible. The storage solution (formalin/TAF for morphological identification or ethanol for molecular studies) should be chosen depending on the goals of the individual project. Critical steps and tips 1. Thoroughly clean all parts of the extractor after each extraction to prevent contamination of the samples by soil and animals from previous extractions (Figure 1). This is important because nematodes can tolerate drought by adopting a resistant stage and revive after rehydration. 2. The height of the sample on the sieve (soil thickness) should not exceed 1 cm. 3. Set the sieve with milk filter and soil in the funnel BEFORE you pour in the water. Water should be added slowly from the side between sieve and funnel. Do not wet the soil directly from above as this will lead to a dirty sample. 4. The sample should touch the water column, but it should neither be soaked nor fully submerged during the extraction to avoid anoxic conditions. Stop adding water as soon as the soil begins to change color or “twitches”, indicating contact with water from below. Wait a few minutes, then check the sample again, adding more water only if the soil surface still appears dry. 5. The extractors in different laboratories should have a comparable length of extraction path (i.e. the distance that animals have to overcome during extraction). We recommend using a kitchen funnel of the total length of 16 cm and 5 cm long connecting hoses (up to c. 20 cm of travel distance). 6. Do not move or touch the funnels during extraction to prevent soil particles falling in the nematode sample. 7. Do not let all the water from the funnel into your sample and do not rinse the funnel walls, because the lighter soil particles on the funnel walls will then end up in the samples. Collect only the bottom of the extract (volume in vial or if the funnel hose ends in a clamp: roughly 2 cm above the clamp; also applies to enchytraeid extraction; see Figure 2). 8. It is important to kill the nematodes soon after the end of the extraction, so that (1) predatory nematodes or other nematode predators (e.g. tardigrades) cannot further injure or consume the others, by this reducing nematode counts drastically (Hohberg and Traunspurger, 2005). 9. Heat shock killing is recommended so a flat and typical posture can be preserved in nematodes. This is a prerequisite for morphological identification and beneficial for body size measurements. This can be done by adding boiling water 1:1 to the extracted nematodes, leading to a sudden rise in temperature to 60°C. Alternatively, if your extractor uses a vial inserted into the extraction hose, you can remove and hold the vial in a water bath (initially 100°C) for one minute to have the same effect. Figure 1 . Schematic representation of a Baermann funnel and practical tips from the Soil BON Foodweb expert team for achieving comparable and clean nematode extractions (see video guide on nematode extraction: https://www.youtube.com/watch?v=aKYANMOTBUA). Enchytraeidae extraction Enchytraeids are small whitish relatives of earthworms, ranging from 1 to 30 mm in length and with a body width of less than 2 mm, which places them into mesofauna (0.1-2 mm body width; Swift et al., 1979). Due to their soft and non-sclerotized bodies, enchytraeids must be extracted from intact soil samples (soil cores) to prevent damage during processing. Although several methods can be employed, the most widely used is the wet extraction based on migration from the heated soil sample through a fine mesh into collection vessels (O’Connor, 1955). Alternative methods include (i) flotation with Ludox (colloidal silica) followed by gentle mixing and finally counting the animals when they float at the surface of the solution, (ii) dry sieving of leaf litter, and (iii) elutriation methods that use the principle that these soil invertebrates are lighter than soil particles and hence sink less rapidly in the upward flow of water. Flotation with Ludox was suggested by Phillips et al. (1999) as a method superior to the O’Connor method, but specimens die shortly after extraction, and Ludox requires health cautionary measures (Adl, 2007). This method may be useful for a quick onsite-evaluation of enchytraeid population densities (Phillips et al., 1999) but has never been used in field studies. Dry sieving of leaf litter can render high enchytraeid numbers, but many small specimens will be overlooked (Healy and Rota, 1992). The elutriation method, never used in enchytraeid field studies, involves elaborate procedures that are cumbersome and time-consuming, and usually are based on expensive equipment (Edwards, 1991). The wet extraction has been shown to extract more than 95% of the total number of enchytraeids from organic soils in 3 h and requires easy-to-get equipment (O’Connor, 1955, 1967). Therefore, the wet extraction was the final choice for extracting enchytraeids within the Soil BON Foodweb (Supplementary Materials 2: Wet hot extractors). There are some variations to this basic funnel technique, which merely diverge in terms of heating temperature (i.e., heat vs cold extraction), duration of the extraction time and water level for soaking the sample. Because warming reduces oxygen content of water, oxygen limitation could kill enchytraeids before escaping the soil. To avoid this, a modified wet extraction method was developed in which heating is omitted and the extraction time is extended to several days for organic soils, and up to two weeks for mineral soils (Graefe, 1984; Schauermann, 1983). In a ringtest comparing methods, the cold method came out as best in terms of efficiency (Didden et al., 1995) and has become the protocol (but with the extraction time set to 24 h) used by the International Organization for Standardization (ISO) as ISO 23611-3 (ISO, 2019). However, another comparison of methods (Kobetičová & Schlaghamerský, 2003) did not find statistically significant differences of extraction efficiency. Also, the long extraction time of the “cold and wet extraction method” can lead to more damaged and dead animals in the sample. Extraction efficiency can even be higher with heat in some soil types, e.g., in the case of peat soils from British moorlands (35 °C surface water temperature over 6 h; Kwon et al., 2000). Therefore, the Soil BON Foodweb Team opted for a combination of 2 h cold extraction followed by 4 h heat extraction (e.g., heat lamp) to reduce processing time when dealing with a large number of samples, while yielding high efficiencies from samples collected in organic-rich soils (Briones et al., 1997). Critical steps and tips 1. Use deionised or mineral water with low mineral content, if possible. Tap water can also be used, but it should be tested before: it may contain chlorine, copper or other compounds toxic for enchytraeids (Figure 2). (Chlorine will evaporate if the water is left in open vessels for 2-3 days. Copper, released from the water pipes, can be avoided by letting the water run for some minutes before use.) When acid soils are extracted, water pH should be adjusted to 4.5 or 5 with drops of HCl, because some acidophilous species die in pH-neutral water. Water quality is appropriate when most of the extracted enchytraeids are complete and alive and stay so (in the fridge) for more than a day. 2. The samples should be fully submerged in water. 3. Best extraction results are achieved if the sample height does not exceed 2.5 cm. When soil cores with diameter <5 cm are used, a greater height (up to 5 cm) can be chosen. Compact samples may be broken carefully into smaller pieces. 4. Temperature in the upper part of the extractor should reach 40 °C; however, critical is the temperature at the bottom of the sieve, it should reach 35 °C or more. The temperature at the bottom of the tube is usually not affected by the heating device above, but cooling is recommended when extraction is carried out in hot weather (ambient temperature 35 °C and more.) 5. In order to prevent excessive amounts of soil particles falling through the mesh, place coarsely woven cotton or kitchen cloth at the bottom of the sieve before placing the soil sample. In case of uncertainty about the suitabletype of cloth, hold it against the light and stretch it; mesh holes of about 1 mm diameter should be seen. Such clothes allow passing of worms while retaining most of the soil particles. However, do not move or touch the funnels during extraction, because some soil particles will fall through. 6. During extraction, worms assemble at the very bottom of the tube, which means that releasing the bottom 2-4 cm of water into a Petri dish will assemble all the extracted worms, alive and moving in the about half water-filled Petri dish. From there they can be collected or photographed. In clayey soils, fine particles make the extraction-water turbid and high-resolution photos impossible. Such extracted samples can be left in the Petri dishes in the fridge (4 °C) overnight; clay particles will settle and the animals can be found alive on the clay layer; they do not move inside. From there, they can be transferred (pipetted) into ethanol. Figure 2 . Schematic representation of funnel extractor of enchytraeids and practical tips from the Soil BON Foodweb expert team for achieving comparable and clean extractions. Microarthropod extraction Microarthropods are small arthropods with body widths ranging from 100 µm to 2 mm, included in mesofauna (Swift et al., 1979). Dominant groups of microarthropods are springtails (Collembola) and mites (Acari); microarthropods also include proturans (Protura), diplurans (Diplura), small myriapods (Symphyla, Pauropoda), and small larvae of flies (Diptera), butterflies and moths (Lepidoptera), pseudoscorpiones, and beetles (Coleoptera) (Lavelle and Spain, 2001). Most of the extraction devices for microarthropods are based on their light-, heat- and drought-avoidance behavior. An intact soil core (normally in the form of a monolith) is placed on a sieve resting on a funnel and, at the lower end of the funnel a jar with a fixative solution is placed. A heat source is placed above the funnel to ensure soil drying from the top. Microarthropods start moving down the soil through natural cavities and pores, and fall through the sieve into a jar. The currently used extraction devices are mainly modifications of the Berlese and Tullgren extractors (Berlese, 1905; Tullgren, 1918). Their extraction efficiency generally varies between 25 and 80% depending on the soil type, light bulb wattage and thermal regime in the extraction room and extracted taxa (Gongalsky, 2021; Krivolutsky et al., 1995). To increase extraction efficiency, several modifications have been proposed. The most important is cooling of the lower parts of the funnel in addition to heating from the above, which creates a marked temperature gradient (MacFadyen, 1962). Other modifications include the gradual increase of temperature during extraction (Zaitsev et al., 2002), use of infrared lamps or electric heating plates or the use of infrared lamps that illuminate the soil in short pulses and thus minimize mortality of invertebrates from overheating (Kempson et al., 1963). In addition to dynamic methods for extracting microarthropods from the soil, there are a number of mechanical methods based on the principles of flotation, substrate vibration and centrifugation (see Krivolutsky et al., 1995 for a review). However, for the Soil BON Foodweb Team such approaches were not considered, as they are very sensitive to the type of soil investigated, operator-dependent, as well as resource- and time-consuming. Therefore, the expert team opted for the basic Berlese/Tullgren extraction (Supplementary Materials 3: Dry hot extractors). This method requires building extractors, but constructing it involves relatively little monetary investment. We also provide the device schemes to make them more accessible (Supplementary Materials 4: Wet hot extractors crafting manual; 5 Dry hot extractors crafting manual). Although the general extraction principles are straightforward, we recommend paying attention to several details. First of all, since most microarthropods cannot efficiently burrow through the soil, and the largest ones typically live on the surface, it is advisable to extract the soil cores upside down and have some free space on the edge. When extracting loosely structured soils, the sample should form a layer of no more than 3-5 cm thick with some space around the substrate for large animals to move down. We also recommend using additional layers of mesh (cell size 1-2 mm) to prevent soil from falling into the collection jars and ensuring clean extraction. This is essential for both image analysis and any other subsequent visual processing of the material. Another important point is to make sure that the substrate is completely dry at the end of the extraction because some animals will not leave the soil if it is still moist. Therefore, different soil types require different extraction times (typically 3-4 days is needed for extraction from litter and 7-10 days for extraction from soil). It is advisable to check the moisture content of the sample during the extraction by inspecting the bottom of soil samples. In case of intact soil monoliths, it is recommended to assess the moisture at the center of the core after the extraction and extend the process if it remains wet. Never check the moisture of the sample directly above the extraction device as this causes a lot of substrates to fall and make the extraction dirty. A gradual increase in temperature helps prevent sample overheating, and a slower extraction process is often more efficient. The surface temperature of the samples should not exceed 50 °C. Regularly monitor both the temperature regime and ethanol level in the collection jars. Standardized lab comparisons could help to estimate potential differences in extraction efficiencies among various laboratories caused by operational differences in the procedure (see section Ring test ). Critical steps and tips 1. Do not overload the funnels with soil. If there is no free space around the edge, or the layer is too thick, animals will not find their way out (Figure 3). 2. Put the samples upside down so animals can move through larger pore space. 3. Avoid overheating samples in the first few days of the extraction. This can lead to the death of microarthropods. Slow extraction is often more efficient. 4. If possible, extraction jars with wide openings should not be tightly connected to funnels because intense alcohol vapor from the jars may reduce extraction efficiency (A. Zaitsev, pers. obs.). For extractors, where funnels are tightly connected with tubes, extraction into low percentage ethanol solution (60-70%) or 50% ethylene glycol followed by further transferring extracted animals into high-percentage ethanol is recommended. Using small diameter vials is also beneficial to minimize evaporation. 5. Avoid shaking or any kind of vibration in the room during extraction, including general cleaning activities. When unloading your extractors, first carefully remove jars with animals and then the sieves with soil to prevent additional soil particles falling into the collection jars. Never do any manipulations with soil above the extractors. 6. No matter whether you have a computer-controlled or a manually controlled extractor, check your samples regularly for the temperature regime (≤50 °C) and levels of ethanol in the jars. Figure 3 . Schematic representation of dry hot extractors and practical tips from Soil BON Foodweb expert team for achieving comparable and clean extractions. Large invertebrate extraction We define large invertebrates as all invertebrates with body width > 2 mm and body length > 3 mm. We on purpose do not use the term macrofauna (invertebrates with body width > 2 mm; Swift et al., 1979), because in many studies macrofauna is defined by taxonomic groups and not solely body size (Gongalsky, 2021). Here we follow body size criteria because large microarthropod groups (e.g. large springtails and mites) cannot be efficiently collected from small soil cores because of their high mobility. To note, large ground-dwelling animals are most effectively collected with Barber pitfall traps - jars dug into the soil with the rim being at the same level as the soil surface (Spence and Niemelä, 1994). However, this approach estimates invertebrate activity rather than density (abundance per area), which is one of the criteria in the Soil BON Foodweb Team methods, so we excluded pitfall traps as well as other non-quantitative collection methods from consideration below. Quantitative methods for soil large invertebrate assessment are based on the extraction of animals from soil samples of various area sizes, usually spanning from 10 to 50 cm in diameter or side length, or even larger in the case of litter samples. The oldest and most efficient method is the direct counting of animals in the soil under a microscope without any extraction (André et al., 2002), but this method is extremely laborious and thus rarely used. Probably, the most common method is hand sorting (i.e., carefully breaking up the soil and manually collecting the animals) of litter and soil samples, either in the field or in the laboratory (Gilyarov, 1941). This method is laborious, but can be applied with minimum equipment (a spade and a plastic tray) and requires little training. Hand sorting is the standard method in the Tropical Soil Biology and Fertility (TSBF) protocol (Anderson and Ingram, 1993; Römbke et al., 2006), and produced data summarized in the recent global databases of soil macrofauna (Lavelle et al., 2022; Mathieu et al., 2022). The time that can be saved and the resultant reductions in extraction efficiencies when sorting is time-limited has been quantified for earthworms (Schmidt, 2001a). However, to give universal recommendations on time-limited sorting, a ring-test covering diverse soils and settings would be required. Heat extraction (see above) can also be used for extraction of large invertebrates from litter and soil samples, but this requires large equipment and may lead to a different representation of taxa compared to hand sorting (see the results from our Pilot Experiment 2). Published comparisons suggest that heat extraction underestimates the abundance of earthworms (mostly endogeic and anecic earthworms undersampled) and social insects (ants and termites), while hand sorting underestimates the abundance of small taxa, e.g. Thysanoptera (Gongalsky, 2021). As a variation of heat extraction, the Winkler extractor is often applied in the tropics to extract invertebrates from litter. This extractor consists of a coarse-mesh bag with substrate, which is placed inside a cloth container suspended over a collection jar at the bottom (Sabu et al., 2011). However, social insects are often poorly recovered by Winkler extractors as well, so alternative methods such as counting the number of nests at large spatial scales, e.g. per hectare (Delabie et al., 2021), have been proposed. Additionally, a comprehensive assessment of earthworm communities may require, in some cases, sampling deep soil down to 1 m. As alternative approaches (or in combination with hand sorting), chemical extraction using mustard oil or allyl isothiocyanate as well as electrical octet method have been suggested to collect earthworms (Čoja et al., 2008; Schmidt, 2001b). In the expert team, we opted for hand sorting of a quadrat of 25 x 25 cm (the standard TSBF area) and 10 cm depth (a compromise solution in comparison to 20 cm in TSBF to reduce labor) to extract large invertebrates. Our main motivation was feasibility of this method under diverse climatic conditions and soil types due to low requirements of extraction equipment and often low transportation costs. In addition, hand sorting ensures better comparability of collected data with the existing global-scale macrofauna initiative (Mathieu et al., 2022). Although it can underestimate some smaller or deep-burrowing taxa, it was proven to render better results than chemical extraction for earthworms (Briones and Schmidt, 2017). Moreover, hand sorting also provides good estimates of less mobile animals like gastropods and some insect larvae, which might be underestimated by heat extraction. However, the method is also very prone to human biases because every person will have different diligence and detection abilities. To assess the accuracy of hand sorting, we conducted pilot Experiment 2 comparing the number of heat-extracted animals from the soil samples that were hand-sorted prior to the heat extraction. Our aim was to see which taxa are usually omitted using hand sorting (see section Pilot Experiment 2 ). Critical steps and tips 1. Hand-sort leaf litter layers initially. Dig quickly to reduce animals escaping. Cut straight and clean along the edges, lift out intact soil blocks. Do not cut inside the block to reduce damage to animals. 2. Hand sorting should be done under good ergonomic and light conditions. When hand sorting is done in the field, use headlights in case of shady conditions (Figure 4). Use white background to detect animals easily. 3. Use a large plastic sheet or box, take small subsamples to break them up and check them, and move soil from unsorted to sorted and from one side to the other in a systematic fashion. Carefully inspect the plant roots, twigs, folded leaf litter, and the content of decaying fruits, acorns, nuts etc. 4. Use flexible (soft) tweezers that do not damage specimens during collection. 5. To catch small, quickly moving animals use entomological aspirators, paintbrushes or put a drop of ethanol on an individual before transfer. 6. When multiple people are performing the hand sorting, have each sample processed by more than one person to evenly distribute individual variation across treatments. 7. Return the hand-sorted soil into the pit if feasible, ensuring minimal disturbance to the site and reducing subsequent risk of injury to animals and people. Figure 4 . Schematic representation of hand sorting of earthworms and other large soil invertebrates and practical tips from the Soil BON Foodweb expert team for achieving comparable extractions. Pilot Experiment 1: Extraction efficiency of nematodes using one vs. two milk filters The cleanliness of the samples is crucial to develop automated recognition, counting and biomass estimation using supervised machine learning (Sys et al., 2022). Overlaps between animals and soil particles could blur the shape of the individuals and challenge their detectability by machine learning algorithms. Therefore, we tested if the use of two milk filters in the nematode extraction could provide cleaner samples and whether it could influence extraction efficiency compared to the use of only one milk filter. The extractions were performed in two separate laboratories (Finland - University of Jyväskylä and the Czech Republic - Biology Centre of the Czech Academy of Sciences). In Finland, samples were taken from two boreal coniferous forests on sandy soils, one Scots pine dominated and another Norway spruce dominated stand, while in the Czech Republic, samples were taken from a spruce forest on loamy soils. At each of three localities, we sampled 5 soil cores (5 cm inner diameter) to a depth of 10 cm. At the laboratories, all samples were first carefully mixed by hand and then two subsamples were taken for extractions (12-21 g fresh weight). One subsample was extracted using one milk filter and the other using two milk filters. The extraction procedure followed the Soil BON Foodweb Team protocol (Supplementary Materials 1: Wet cold extractors). Extracted nematodes were counted under a microscope. To evaluate differences in the numbers of nematodes extracted with one or two milk filters, we used a two-way ANOVA in R v4.5 with location and number of filters as independent factors. We found no difference between the numbers of nematodes extracted with one or two milk filters (Figure 5; F 1,25 = 1.88, p = 0.18). Against our expectations, clean samples were obtained in both one- and two-milk filter extractions. We speculate that the sample cleanliness probably depends more on the other factors such as soil treatment during extraction, soil clay content, initial soil moisture, milk filter type, and number of animals in the sample. It has been previously shown that the filter type is crucial for extraction efficiency and may differ considerably depending on the used material and soil type (Cesarz et al., 2019). Although the milk filters used in two studied laboratories were from different suppliers, similar treatment effects were observed. Overall, although their effects on community composition and sample cleanliness are to be evaluated in future studies, our results suggest that two layers of milk filters do not significantly reduce the abundance of extracted nematodes and this approach may be tested in the cases where samples are particularly ‘dirty’. A ring test could be an optimal solution to ensure data comparability (see section Ring test ). Figure 5 . Mean number of nematodes extracted from 1 g of fresh forest soil sample using either one and two milk filters at two laboratories (Finland, Czech Republic). Bars represent means and error bars indicate upper 95% confidence intervals. The dashed line shows the mean for each country. Pilot Experiment 2: The efficiency of hand sorting method for large soil invertebrates To test the efficiency of the hand sorting method, we implemented a two-step process. The samples were hand-sorted and then the same soil and litter was subjected to subsequent Kempson´s dry and heat extraction. To evaluate differences among teams working in different regions, we performed this test in three independent laboratories: the Czech Republic (CZE, Biology Centre of the Czech Academy of Sciences), Germany (GE, University of Leipzig) and Italy (IT, Eurac Research). In the Czech Republic, samples were taken from two mixed spruce- and beech-dominated forests (USDA soil texture class: loam). In Germany, samples were taken from two forests (beech- and oak-dominated, silty loam) and two meadows (loam to silty clay loam). In Italy, the samples were taken from one oak (sandy loam), two subalpine spruce and two subalpine larch forests (loam). The soil samples were obtained following the Soil BON Foodweb protocol (Potapov et al., 2022) by digging out a soil monolith of 25 x 25 cm square of soil, including the litter layer and the underlying 0-10 cm of soil. The vegetation was cut and removed before the digging. The litter was collected and hand sorted separately from the soil. The samples were transported to the laboratory and hand sorted to catch all large invertebrates (> 3 mm in body length) except enchytraeids. The screened soil and litter samples were then put separately in Kempson’s extractors (Kempson et al., 1963) and extracted for five days (or until the sample was completely dry) in order to expel all the remaining animals from the soil. In Germany, a 25% subsample of each sample was used for heat extraction due to a limited space in the extractors. Obtained invertebrates were stored in pure 96% ethanol at -20°C until identified to high-rank taxa (orders, families) under a dissecting microscope. The proportion of missed individuals for each taxon was calculated as the number of Kempson-extracted individuals divided by the total number of individuals collected (hand sorting + Kempson). To compare between laboratories and animal groups, the proportion of missed individuals were evaluated by linear-mixed effect models using R v4.5 and with laboratory, animal group and their interactions as fixed factors while plot ID as random factor. Figure 6 . The average proportion of missed soil fauna individuals during hand sorting of soil monoliths in three laboratories (the Czech Republic n = 20, Germany n = 8, Italy n = 10 soil monoliths). The proportion is the difference between the total number of extracted individuals with Kempson’s apparatus divided by the number of total (hand-collected and extracted) individuals across all samples in each laboratory. These total numbers are shown above the columns. The dashed line represents the average proportion of missed individuals across all taxa in each laboratory. The category ‘Other insects’ contains all insect groups (e.g., Hemiptera and Lepidoptera) not belonging to the dominant groups of soil fauna. Earthworms are Lumbricina. In general, our results show that there are overlooked individuals in virtually all soil animal taxa. On average across taxa, 31.3% of individuals were missed, but the percentage of missed individuals varied strongly depending on the taxon (taxon effect F 13,276 = 6.5, p < 0.001). The most missed animal taxa across all three laboratories were Pseudoscorpiones (64.4% ± 46% missed on average across three laboratories), Diptera larvae (62.9% ± 35%), Coleoptera larvae (57.1% ± 33%) and other insects (55.2% ± 45%). Coleoptera and Diptera larvae were also the most abundant taxa across all samples. Although overlooked in some cases, the overall missing percentages were low for Araneae (29.3% ± 36%) and Lumbricina (23.0% ± 33%; Figure 6). Individual laboratories also differed in the collecting efficiency of specific taxa (taxon:laboratory interaction F 23,276 = 3.7, p < 0.001). Based on ANOVA of each animal group, significant differences were observed between laboratories in Lumbricina (CZE = 3.9%, GE = 65.0%, IT = 0%), Diptera larvae (CZE = 34.0%, GE = 81.3%, IT = 73.3%) other insects (e.g. Homoptera, Lepidoptera, CZE 17.9%, GE = 57.1%, IT = 90.7%), Araneae (CZE= 13.9% GE =3.9%, IT = 70.0%), Formicidae (CZE = 0%, GE = 7.8%, IT = 76.0%), adult Coleoptera (CZE = 13.9%, GE = 25.2%, IT = 88.1%) and Symphyla (CZE = 0%, GE = 32.6%, IT = 50.0%). On average across target animal taxa, 20.3% of all individuals were missed in Czech sites, 35.3% in German sites, and 60.5% in Italian sites. Overall, our test showed that efficiency of hand sorting depends on both individual taxa and the collecting team/ecosystem type (it was not possible to separate the latter two factors with our sampling design). Furthermore, individual taxa were overlooked to a variable extent depending on the collecting team which may likely be explained by differences in the local fauna (e.g. more obscure life forms of Araneae in one location than in another). The results highlighted two main disadvantages of the hand sorting method from a soil monolith. First, the method misses a significant proportion of large invertebrates (18-64%), both due to human bias and different local characteristics of the soil and fauna. Therefore, total densities evaluated by hand sorting are likely to be significantly underestimated. Moreover, the number of missing individuals differs significantly among animal taxa and thus hand sorting skews the information about community structure. The overlooked individuals were especially from the ecologically important and abundant groups of macrofauna such as the larval stages of Coleoptera and Diptera (> 3mm). The underestimation might be less pronounced when the animal biomass is calculated, as omitted specimens within taxonomic groups were mainly small in size (visual observations). We speculate that the larvae and isopods were missed due to often dense cover by soil particles that make them hard to recognize. In addition, earthworms were often covered by soil during the sorting, and thus probably obscured and overlooked in the German sites. Moreover, in many of these taxa, thanatosis (immobility in face of danger) could be a protective behavior that could be provoked by hand sorting (Humphreys and Ruxton, 2018). For example, ants, typically exhibiting active movement, were less likely missed. Overall, the hand sorting method captured better actively moving taxa such as spiders, adult beetles, centipedes and ants. A thorough assessment and large time investment (at least 60 minutes for one person per 25 x 25 x 10 cm monolith) were needed to collect most animals with this method, and even then small individuals were overlooked. This critical information needs to be taken into account when interpreting the results of hand sorting. The second disadvantage lies in the significant differences observed in results coming from different laboratories. The explanation can be variable searching time per sample among laboratories. The searching time through one sample was on average one hour for the four people involved in the Czech team (4 hours/sample of soil+litter), one hour for two people from the German team (2 hours/sample), and 25 minutes for 3 people in the case of the Italian team (1 hour 15 minutes/sample). This is aligned with the overall collection efficiency that descends in the order from Czech to Germany to Italy (see above). However, the searching time cannot be unified to standardize the method because effectiveness is strongly related to the soil type, moisture of the soil, weather conditions, amount of roots, aggregation of soil and type of substrate (e.g. litter vs soil) and the number of specimens present. A ring test following our pilot experiment, involving more laboratories and improved methodology, would be necessary to identify main sources of bias and potentially develop calibration coefficients for different animal taxa, ecosystems and searching time. Ring test Ring tests involve conducting identical research interventions in geographically distant scientific units following the same protocol to standardize these protocols and ensure comparable results across the units. This approach is commonly used in soil ecotoxicology to draft and quality-check international standards for research methods and facilities (see e.g., Knacker et al., 2004; Koolhaas et al., 2004; Moser and Römbke, 2009). In soil ecology this approach has received little attention up to date (but see Crossley and Blair, 1991). Results of the pilot experiment 2 presented in this paper show that even with the same protocol, different laboratories achieve variable efficiency of hand sorting. Hence, it is of vital importance to verify soil fauna extraction efficiency in global studies assessing soil communities and in large scale monitoring surveys. There are two possible ways to implement the ring test in the global assessment of soil animal biomass, diversity and functioning. The first and more complex method is to extract artificial communities formed by focal taxa that are either pre-extracted or taken from laboratory cultures. The second approach is based on extraction of samples from a single natural community in different laboratories. The first option provides detailed taxon-specific information, valuable due to potential extraction efficiency variation by taxon and functional group. The second approach avoids soil animal stress due to the pre-extraction. However, there are three major constraints of the second approach. First, many more samples need to be extracted to account for local spatial variability in species distribution. Second, legal hurdles are numerous on the way to authorize the admission of non-sterilized soil across customs borders. And last, logistical efforts are necessary to prevent community changes caused by varying shipment duration and conditions. Still, the second option of ring test implementation might be the preferred one, if performed regionally. Globally, we propose two levels of ring testing: (i) calibrate devices in a few core facilities in different regions and then (ii) calibrate as many devices as necessary at the regional level using samples collected within the same country or customs-free areas. In the case of studies covering wide geographic ranges, ring tests represent a very promising yet technically and organizationally challenging opportunity to increase the quality, comparability, and reliability of the collected data. With this paper we call for including such ring tests in the agenda of existing and future large-scale projects and initiatives and during establishment of new laboratories. Future perspectives Comparable data from different locations is essential to test major (macro)ecological hypotheses related to soil fauna (Mathieu et al., 2022), assess changes of soil communities along wide climatic and disturbance gradients, correctly estimate contribution of soil animals in the global biomass stocks (Bar-On et al., 2018), and understand global trends of soil biodiversity (Guerra et al., 2021a). Adequate biomass estimates are necessary to implement soil food web and energy flux analyses, in order to describe variation in functioning of soil animal communities (Jochum et al., 2021; Potapov et al., 2022). Here, we describe the methodological basis for the global-scale monitoring of soil animal communities in the framework of the Soil BON Foodweb. Such assessments have been implemented across 30+ countries globally in 2022-2023, and will continue in 2025-2026, 2028-2029 and beyond to provide realistic assessments on the status, trends and drivers of soil animal biomass and community composition (Potapov et al., 2022). Although none of the methods and protocols are perfect to assess soil fauna comprehensively, the Soil BON Foodweb expert team has put a large collaborative effort into finding balance between labor, financial investments and representativeness of each method. Despite the methods described above are not new and are being routinely used in many laboratories, there are many inconsistencies in results. By summarizing ‘expert tips’ and providing visual materials, we aim at achieving a better standardization of transnational soil biodiversity assessments. Furthermore, we call for a global-scale ring testing of extraction methods of soil fauna as the next crucial step towards understanding of global soil animal diversity status and trends. Similar standardization has been implemented for the assessment of other biodiversity dimensions like soil microorganisms and soil microbial functions (e.g. in Soil BON; Guerra et al., 2021) via centralised laboratory facilities. However, this is not possible for most soil fauna taxa and local facilities are to be established, and expert capacity built. Our recommendations are aimed to reach scientists also beyond the Soil BON Foodweb. By small amendments of existing protocols or expanding toolbox, soil ecology laboratories are invited to join our efforts in the global standardization of soil animal assessment and monitoring. Our protocols can be used in local, national or regional assessments and monitoring schemes to contribute to the accumulating body of knowledge on soil fauna. In the UN Decade on Restoration and with important initiatives like the EU soil strategy for 2030 including the upcoming European Soil Monitoring and Resilience Directive being implemented, it is of utmost importance to understand status and trends in soil biodiversity. For this, time series analyses need to be implemented (Eisenhauer et al., 2022). Many countries are establishing their national biodiversity monitoring programs across taxa and to aid inclusion of soil biodiversity, standard, open and efficient methods need to be applied. Our methods set may serve as a basic (and potentially expandable) reference making such assessments comparable across borders.

We dedicate this paper to the memory of Dr. Diana Wall, who made significant contributions to this work before her passing on March 25, 2024. We acknowledge Helene Blasbichler, Lucie Kryštofová, Peter Čuchta, Marco Antonio Jimenez Santos, Verča Grygarová, Šárka Otáhalová for field and laboratory assistance. We also thank Svenja Meyer and Carlo De Rito for providing the animal and soil profile drawings. SC and NE acknowledge support of iDiv funded by the German Research Foundation (DFG– FZT 118, 202548816) and the DFG (Ei 862/29-1). AP and JZL are funded by the European Union (ERC CARBONWEB, 101170898). Views and opinions expressed are however those of the authors only and do not necessarily reflect those of the European Union or the European Research Council. Neither the European Union nor the granting authority can be held responsible for them. Supplementary Material

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Authors Metrics & Citations Metrics Article Usage 989views 361downloads Citations Download citation Michala Tůmová, Jing-Zhong Lu, Maria Briones, et al. Standardised protocols for soil fauna extraction and a call for cross-lab implementation. Authorea. 05 November 2025. DOI: https://doi.org/10.22541/au.176233217.71833958/v1 DOI: https://doi.org/10.22541/au.176233217.71833958/v1 If you have the appropriate software installed, you can download article citation data to the citation manager of your choice. Simply select your manager software from the list below and click Download. For more information or tips please see 'Downloading to a citation manager' in the Help menu.