Protista is an informal, polyphyletic designation for eukaryotic organisms not belonging to multicellular plants, fungi, or animals, and thus cover free-living, predominantly unicellular eukaryotes that can be heterotrophic or autotrophic (Altermatt et al. 2015, for a more detailed definition see Burki et al. 2021). Algae are also considered within Protista (Gachon et al. 2013; Saxon et al. 2020). For convenience, only microalgae (e.g., euglenoids, dinoflagellates, diatoms) will be included in this section. The diversity and abundance within Protista have been largely underestimated by traditional approaches such as culturing and microscopy, due to the challenges to collect and identify species (Caron and Schnetzer 2007; Medinger et al. 2010; Foissner 2014). Hence, culture techniques should be improved to identify and characterise morphologically, genetically, and functionally the collected specimens (Geisen et al. 2018; Gooday et al. 2020). Environmental samples and molecular approaches, however, have been currently used, and it is now a standard, at least, for soil microbial diversity assessment (Geisen et al. 2018).

Altermatt et al. (2015) have provided the first steps to the workflow standardisation in protists, leading to the development of a protocol repository.

This group of organisms is relatively understudied, and interest has mainly focused on its commercial and industrial applications (Buyck et al. 2010). Microfungi and fungus-like forms are not easy to collect, as they are either very tiny or live hidden in a substrate or within hosts. In most cases substrata are collected rather than the fungus itself. Special laboratory conditions for culturing are hence required for the isolation and identification of species (Buyck et al. 2010). Specimens have to be sampled together with the substrate and may not be visible, and they should be slowly air-dried (Wu Q et al. 2004; Agarwal and Sharma 2006). If substrata are not to be dried, they should be incubated and cultured within a day after collection (Spiegel et al. 2007). After drying, samples should be stored at temperatures as close to field conditions as possible (Wrigley de Basanta and Estrada-Torres 2017). In addition, many fungus-like organisms are obligate biotrophs and thus cannot be cultured separately from their hosts (Buaya and Thines 2020).

Plant pathogenic fungi and oomycetes can be detected either by the appearance of disease symptoms or by mycelia/fruiting bodies growing on diseased or healthy plant tissue (Callan and Carris 2004). A description of symptoms and pathogens is found in Shivas et al. (2005) and Thines and Choi (2016), the latter focusing on biotrophic oomycetes. Species identification without the aid of sequence data is in most cases only possible after identifying the host plant. Both diseased and uninfected plant material should therefore be collected (Callan and Carris 2004). It is essential to heed the following recommendations (Callan and Carris 2004; Shivas et al. 2005; Buyck et al. 2010; Lane 2012; Prospero et al. 2017):

1. Do not collect in the rain, as wet samples may become colonised by other organisms.
2. The whole plant should be examined during fieldwork, starting from the youngest leaves, and working toward the stem. Ideally, samples should be taken from all symptomatic parts of the plant.
3. When sampling trees, the upper and lower side of an angled trunk or branch should be sampled, along with various strata (canopy, trunk, ground). Epiphyte flora will be different due to different moisture conditions.
4. If the plant has a poor crown, the roots should be examined.
5. Collected samples should include different aspects and symptom stages of the disease.
6. Samples should include the border between healthy and diseased tissue.
7. Rotten material and dead plants should be avoided, as pathogens may no longer be viable and saprobic fungi may have already colonised these parts.
8. All sample tools should be disinfected after use with 70% alcohol or 5% bleach (NaOCl).
9. Avoid heating samples to more than 30 °C.

To sample the appropriate parts of the plant, some previous knowledge of disease symptoms is useful (Shivas et al. 2005). The most common parasitic fungi and oomycetes found in both agricultural and natural environments include rust fungi, which may produce various types of spore-producing structures that are often orange or brown in colour and can be found in different infection stages on leaves, shoots, fruits, and woody stems of a single host. Smut fungi can be found in specific parts of the host crop plant, especially the leaves, stem, roots, and inflorescences and often produce a mass of dark brown to black teliospores. Powdery mildews can be recognised by the white and dusty conidial state found on leaves and stem surfaces. Sooty fungi can form a hyphae network or a dark crust on leaves and small twigs (Callan and Carris 2004). Downy mildews cause chlorosis of infected leaf parts and often stunting in systemic infection, and produce a whitish, violet to dark brown down, predominantly on the lower leaf surfaces, as sporangiophores usually protrude from the host through the stomata (Thines and Choi 2016). White blister pathogens produce white to off-white pustules on infected plant parts that disseminate a white spore powder after pustules rupture (Heller and Thines 2009).

When collecting, leaves should be wrapped in moisture-absorbing paper and placed in paper bags to avoid desiccation (Shivas et al. 2005; Lane 2012). If the ambient air is too humid, a small quantity of desiccant can be added to the bag (Callan and Carris 2004). Alternatively, samples may be placed in a plastic box containing moist paper, covered with a lid, and stored in a cold room at 5 °C. They can be stored for up to 3–4 days. (APS 2022). If whole plants are sampled (i.e., small herbaceous plants), they should be placed in strong plastic bags with the roots tied off at the collar and stored in a cardboard box lined with absorbent paper (Lane 2012). Do not pull plants from the soil and scrubbing should be avoided when cleaning the roots (Shivas et al. 2005). Fruits, flowers, and tubers should be collected in the early or intermediate stages of the disease (Shivas et al. 2005), individually wrapped in dry paper and placed in open or perforated plastic bags, and should be carried in a robust container until desiccated or cultured (Lane 2012; Callan and Carris 2004). Conifers and spiny plants should be stored in large paper bags. Cankers on trunks should be removed with a knife or a chisel and should be wrapped in newspaper or in a paper bag (Callan and Carris 2004). Refer to Prospero et al. (2017) for detailed sampling protocols for bark, wood, shoots, leaves, and roots.

If the collected material cannot be cultured within a few days, samples can be air-dried after pressing them between blotter sheets. Specimens can be wrapped in newspaper for transportation and stored in paper packets (Wo et al. 2004; Buyck et al. 2010). Cultures can be obtained later if the samples contain immature fruiting bodies close to sporulation (Callan and Carris 2004). Rust and smuts can be frozen at -20 °C for up to seven days after drying, for later examination (Shivas et al. 2005).

If fungal pathogens are present inside the plant tissue (e.g., leaves, petioles, roots, or branches), tissue should be washed with 70–95% ethanol, followed by immersion in 0.5–10% sodium hypochlorite (ideally with 0.1% Tween20 or a similar detergent), and rinsed with autoclaved distilled water to kill surface contaminants such as saprobic fungi (Schmit and Lodge 2005; Shivas et al. 2005; Beales 2012). Some previous experimentation should be conducted to ascertain the correct bleach solution and exposure time for specific plant or fungal associations, as some plant tissue may be too delicate, or some fungal structures may not be affected by this treatment (Lori M. Carris, pers. comm.). The outer layers of the tissue should be removed, and the inner part should be cut into small pieces (ca. 2 mm) and placed on Petri dishes containing an agar medium (Narayanasamy 2011). Adding acidified malt can restrain bacterial growth and stimulate pathogenic fungi growth instead (Callan and Carris 2004). This also applies to endophytic fungi that do not cause diseases.

Leaf washes is another sampling alternative that also works for spores of saprotrophic fungi (Schmit and Lodge 2005). Detailed protocols for isolating pathogenic fungi from leaves, stems and roots can be found in Shivas et al. (2005). Growth requirements for specific fungi are defined in Ryan et al. (2012).

Fungi and allies can be identified by examining them under a compound light microscope. However, if reproductive structures are not present, the interface between healthy and sick tissue (ca. 2-mm sections (Akinsanmi et al. 2004)), where the pathogen is most active, should be cultured as soon as possible. Do not use necrotic tissue, as this may contain already secondary colonisers. If fruiting bodies close to sporulation are seen, the substrate can be incubated in a moist chamber for one to three days to encourage sporulation (Callan and Carris 2004).

Several microfungi can be grown in pure culture except for those that are obligate parasites or biotrophs (i.e., downy mildews, rusts, black mildews, and powdery mildews). Few obligate species may grow in vitro under non-axenic conditions and only together with the living host (Callan and Carris 2004), making this technique very laborious and expensive (Ryan and Ellison 2003). Thus, drying is the best way to preserve obligate parasites (Callan and Carris 2004) and preserve them as herbarium specimens (Homolka 2014). Collection of rusts’ basidiospores and teliospores is also possible following Ryan and Ellison (2003). However, the described method does not generate sufficient spores for preservation, even though molecular identification might be possible.

Leaf surface impressions or peels, adhesive tape, transparent cellophane tape, Mellinex, or dry water-based mucilage are additional methods to examine recalcitrant pathogenic microfungi structures (Callan and Carris 2004; Narayanasamy 2011). These procedures may be suitable for DNA recovery by touching a single lesion of a leaf so that spores can attach to the tape, which should be cut into small pieces and placed in small tubes (Pilo et al. 2022). Note that the sample will probably contain other fungi, bacteria, and PCR inhibitors (Lori M. Carris, pers. comm.). Hence, the material should be treated as an eDNA sample, unless species-specific markers are to be used.

Environmental microbiomes are systems, found together in the same habitat, that include a variety of microorganisms such as bacteria, archaea, fungi, algae, and protists, together with mobile genetic elements such as viruses, phages, and relic DNA (Berg et al. 2020; Ryan et al. 2021). So far, targeted preservation of environmental microbiomes is very rare (Smith et al. 2020; see Steiger and Heuss 2020 for a list of existing projects), as culture collections are usually maintained under axenic conditions, and only freeze-tolerant organisms endure cryopreservation procedures, leading to a partial loss of the microbial community (Ryan et al. 2019). Moreover, best practices and data standardisation in microbiome research is lacking (Berg et al. 2020). The MicrobiomeSupport project has been set up in 2018 and its main aim is to provide and improve standards and protocols for microbiome studies. Further details about concepts and challenges can be found in Berg et al. (2020).

Note that a microbiome sample will only represent the temporal and spatial structure of the moment when the sample was collected.

Sample collection and duration, as well as temperature during transportation and storage are crucial factors for maintaining the microbial community composition. Samples for intact microbiome preservation should be placed in the dark at temperatures lower than 4 °C directly after sampling. At the laboratory, samples should be immediately stored in a refrigerator (Prakash et al. 2020).

Currently ongoing studies will help to develop optimal preservation methodologies that can maintain the integrity, functionality, and inter-species interactions of microbiome samples (Smith et al. 2020; Ryan et al. 2021). Microbiome resources, such the Microbiota Vault, the Alliance for Phytobiomes and the Minimal Effective Microbiome Set (MEMS) for a plant microbiome vault are being established to provide repeatability in microbiome research (Gopal and Gupta 2019; Ryan et al. 2021). In addition, the Earth HoloGenome initiative is taking the lead on the standardisation of collection and preservation practices on animal-microbiota associations.

If living material is required (e.g., propagation), a field tissue culture can be initiated by in vitro collecting (IVC). It is important to note that this procedure has to be adapted to the specific taxon and material to be used (Sarasan 2010). IVC is suitable for species that do not produce seeds, when seeds are not available at the time of collection, or for species with recalcitrant seeds (see definition below). This additionally allows avoiding problems associated with transporting large amounts of vegetative tissue, which can die before reaching the biobank (Pence et al. 2002). However, contamination remains a large challenge, as work is performed in the field and cultures may be exposed to both air-borne contaminants and endogenous microorganisms (Cruz-Cruz et al. 2013). Check Pence et al. (2002) for contamination control protocols during IVC.

Several factors are, therefore, critical and should be considered for a successful outcome: 1) type and size of tissue, 2) removal of soil remnants and pests, 3) avoidance of damaged tissue, 4) tissue sterilisation and washing, 5) nutrient media with antimicrobial additives and, 6) storage conditions (Cruz-Cruz et al. 2013).

The material to be collected depends on the species. In general, budwoods, shoots, apices, or leaves can be used to initiate IVC. It is necessary to collect stakes, pieces of budwood, tubers, or corms for vegetatively propagated species (Cruz-Cruz et al. 2013). Two methods can be applied for tissue collecting (Pence 2005):

• The leaf punch method. Leaf discs (ca. 6 mm) are punched from young leaves and placed into vials with media including fungicide and antibiotics.
• The needle collecting method. A cross-section from soft, green, young stem tissues is collected using a syringe with a gauge needle (no. 21) to obtain a tissue cylinder which is placed into a vial with medium. Very small stems can be sliced using scissors.

Collected material can also be stored in Ziploc bags containing moist paper towels and processed indoors within a few hours of collecting if outdoor processing is not performed (Pence 2005). For culture conditions, refer to Pence (2005). Vials should be carried in opaque cloth bags and direct sunlight should be avoided. Samples can be left at ambient temperature in the field (Pence 2005). Protocols for various crops, tropical rainforest trees and endangered species can be found in Pence et al. (2002).

Collection of cuttings is preferable over tissue sampling for succulents and woody species propagation. Cuttings should be clean and neither too woody nor too soft. Cuts should be made just above a node or bud from one plant using secateurs. Secateurs or pruners should be decontaminated with 70% alcohol between plant collections. All cuttings should be free of flowers, flower buds, and fruits to minimise moisture loss. Cuttings should be stored moist and cool (3–5 °C), loosely wrapped in newspaper and placed into plastic bags. Cuttings that have been wrapped for more than 24 h should be exposed to release ethylene, which can lead to tissue damage after removing the material from the parent plant (Australian National Herbarium, updated 2015; Martyn Yenson et al. 2021). Types of cuttings, preparation and growing conditions for cuttings are available in Martyn Yenson et al. (2021).

The selected tissue preservation method will depend on the species and the duration of the field trip. Preservation methods include the use of RNAlater or equivalents, desiccation with salt buffers (e.g., NaCl-CTAB buffer), 96% ethanol (Bressan et al. 2014; Johnson 2021) or silica gel, which is the most common plant material preservation method used during fieldwork (Chase and Hills 1991).

Silica gel should ideally have a 28–200 mesh size grade and include a proportion of beads that contain a moisture indicator dye that changes from orange to colourless as gel saturation levels increase (Chase and Hills 1991). Blue dye indicator should be avoided, as it contains cobalt chloride, which is carcinogenic (Funk et al. 2017; Maurin et al. 2017). Relative humidity indicator cards provide an alternative to indicating gel (Funk et al. 2017).

Placing samples straight into Ziploc bags containing silica gel makes the replacement of saturated gel with dry gel, and any reuse of the silica gel, time consuming due to the physical contact between the plant material and the gel (Wilkie et al. 2013; Funk et al. 2017). Instead, the plant tissue can be placed in porous paper coin envelopes or tea bags/coffee filters, sealed by folding over the top and pinning with archival paper clips to ensure that the plant material does not escape (Wilkie et al. 2013; Maurin, et al. 2017). Such sample bags can be placed either into Ziploc bags or into an airtight plastic container filled with silica gel for drying. The amount of silica gel will depend on the tissue water content, but 10–15 times as much silica gel as tissue should generally be used (Chase and Hills 1991). Although coin envelopes are not usually archival quality paper, and tea bags are less robust, they can both be used for long term-storage, minimising laborious post-collecting processing work (Gemeinholzer et al. 2010; Funk et al. 2017, Maurin et al. 2017). Samples should be kept in a cool, ambient-temperature environment, and the silica gel should be changed within 12 h if the tissue is not completely dry (Davis 2011). Ziploc bags and plastic containers have to remain closed with an airtight seal to ensure that the silica gel maintains its desiccating properties (Maurin et al. 2017). Once the material is completely dried (after 24–48 h), the envelopes/tea bags/filters can be transferred into individual Ziploc bags with a small amount of fresh silica gel (Funk et al. 2017; Maurin et al. 2017; Duque-Thüs and Fulcher 2018), kept in plastic lunchboxes, or stored in low humidity cabinets (Plaisier 2019: Botanics stories). Samples can be stored at -20 °C or -80 °C (Funk et al. 2017), or if no sub-zero facilities are available, they can be kept at room temperature, if stored in hermetically sealed containers (Hodkinson et al. 2007; Tim Fulcher, pers. comm.).

Another alternative is to use FTA filter cards to preserve leaf material. Leaves should be carefully crushed onto the card, avoiding cross-contamination. Between 50–100 µl of plant homogenate can fit on one card. Cards should be placed into multi-barrier pouches containing desiccant packets, to ensure that cards remain dry during storage and transportation (Gemeinholzer et al. 2010).

For a comprehensive publication on plant tissue collection for DNA banking, see Funk et al. (2017). This paper provides recommendations for all the tasks required for voucher preparation, sample collection, LN2 preservation, and storage. The Queensland Herbarium (2016) provides collecting protocols especially for difficult plant groups such as bamboos, palms, and cacti. Barber and Galloway (2014) compiled methods for the collection of seeds, fruits, spores, underground organs, and cuttings and their further processing during fieldwork.

The following section provides guidance on collecting in the field for each plant group.

Pollen banking has had a limited use in plant germplasm conservation, but it may be a valuable method for species with recalcitrant seeds, and for different horticultural plant species (Theilade and Petri 2003). Most methods described below have been mainly developed for crops and commercial species.

Mature pollen collection must occur in the morning soon after flowers are fully opened (anthesis) (Prendini et al. 2002) by shaking the flowers over a sheet of butter paper or by using a sieve. Usually, anthers are collected when species do not shed vast amounts of pollen. Pollen should not be collected from sick or damaged flowers. Both pollen and anthers should be processed soon after collection to guarantee maximum potential longevity (Volk 2011). Be aware that large amounts of pollen are required not only for storage, but also for moisture measurements, viability testing and subsequent breeding programs and crop improvement.

Depending on the species, pollen has to be separated from anthers prior to desiccation. Otherwise, the whole anther can be dried and gently crushed (Towill and Walters 2000). Sticky pollen should be treated with Lycopodium powder or talc to avoid clump formation (Sidhu 2019). Samples can be air-dried on clean Petri dishes if the ambient humidity is low (Anushma et al. 2018). Otherwise, they should be desiccated using LiCl, MgCl₂, NaOH, or silica gel to reduce moisture below the species-critical level (Towill and Walters 2000; Sidhu 2019). Most pollen can be dried to less than 5% moisture content (Theilade and Petri 2003). However, some desiccation-sensitive pollen requires controlled conditions for drying and cooling, as well as the use of cryoprotectants for successful cryopreservation (Nebot et al. 2021).

Seed banking is the most widely used ex situ conservation method for crops and wild plants. In general, standardised protocols exist for the storage and preservation of crop species, but not specific ones for wild species, because the amount of collected seeds may limit the number of seeds available for testing protocols (Hay and Probert 2013). Seed banks should ideally have a base collection for long-term conservation, which should never be distributed for use, and an active collection for propagation, multiplication, and distribution (Cromarty et al. 1990; Engels and Visser 2003).

All following described procedures are dependent on the type of seed. Seeds are classified into three groups depending on their storage conditions:

• Orthodox seeds can be dried and stored at low temperatures without losing their viability; and, importantly, their longevity in storage is increased in predictable ways by reductions in moisture content and temperature (Roberts 1973). Most crops produce this type of seed.
• Intermediate seeds cannot survive when exposed to both low humidity and low temperatures simultaneously (Ballesteros 2011). They are often chill-sensitive but can be desiccated until a certain threshold (Theilade and Petri 2003). Citrus, coffee, and rubber are economically important examples of species belonging to this category.
• Recalcitrant seeds are shed at high moisture content and do not tolerate drying. Most species bearing recalcitrant seeds are woody; and it is predicted that most tropical tree species bear this type of seed. Likewise, they do not have dormancy, or are resistant to dormancy induction procedures. They usually do not remain viable for long after falling from the tree if germination does not occur. (e.g., Wyse and Dickie 2018).

Martyn Yenson et al. (2021) have developed guidelines for the maintenance of seed collections, including identification of seeds, seed germination, dormancy, and plant nursery.

The physical quality of the seeds should be assessed, prior to collection, by using the cut-test technique. Seeds are cut along both axes to check whether they are infested, immature or empty (MSBP technical information sheet No. 4). If the proportion of damaged seeds is higher than 30%, an increased number of seeds should be collected to compensate for the non-viable ones or another population should be sampled (MSBP technical information sheet No. 2). Ideally, between 3000–5000 seeds from > 50 mother plants per accession should be randomly collected to guarantee that there is enough material for germination assays, long-term conservation, and propagation (CPC 2019). Obtaining this number of seeds for tropical species and many tree species is generally not possible. However, depending on the aim of the collection, seeds can be pooled across the population rather than keeping them separately per mother plant (Filip Vandelook, pers. comm.). No more than 20% of the seeds found in the population should be collected to avoid compromising its in-situ viability (Gold et al. 2004; CPC 2019).

Seeds have to be collected at the time of optimum ripeness, as this is a precondition to secure high seed quality (Schmidt 2000). Unripe seeds have a high RH, which may cause reduced storability and vigour (Theilade and Petri 2003), whereas seeds that have imbibed moisture and initiated germination will not tolerate re-drying (Schmidt 2000).

Many methods for seed collection have been developed, and nearly all of them can be equally applied to shrubs, herbaceous species, and trees. The choice will depend on the kind of plant to be sampled and the location. Ground collection is the easiest and most common way to obtain large seeds. However, the following issues should be considered when choosing this method: 1) the mother plant may be difficult to confirm, or a seed mixture is involved, especially in high-density stands, 2) seeds may be infested, 3) seeds can get lost on the ground, 4) seeds may germinate or deteriorate right after falling, 5) seeds from closely related species may be also sampled and, 6) debris, damaged seeds and soil-borne pathogens may be collected when raking the ground or using mechanical equipment (e.g., vacuums, rotating brushes) (Schmidt 2011). An alternative for collecting small seeds is using tarpaulins or sheet funnels under tree crowns. Seeds from grasses can be collected by stripping entire seed-heads (MSBP technical information sheet No. 3). Explosive fruits can be collected by securing cloth bags over the seed head or fruit. Flowers from aquatic plants can be wrapped in nets to capture future fruits (ENSCONET 2009). Taller trees with seed-bearing branches can be reached by using long-handled tools (e.g., pruners, telescopic poles) or by shaking. Climbing using spurs, braided ropes, or tree bicycles is another alternative, but these techniques should only be used by trained people (Schmidt 2000).

Large fruits/seeds should be placed into clean baskets or buckets, non-glossy paper bags, or woven hessian sacks. Small dry fruits/seeds should be placed on canvas sheets that can be folded and tied (Schmidt 2000). Fleshy fruits can be placed into open plastic bags for a short period of time. Do not use large containers, as ventilation cannot be assured. All container types should be tag-labelled inside and outside. A compilation of seed collection details and procedures can be found in Schmidt (2000), ENSCONET (2009), and SOS (2018).

If space is limited or field duration is long, a manual pre-cleaning from debris and plant remnants can be carried out. If the outside RH—determined by a hygrometer—is greater than 40–50%, plastic boxes containing small amounts of silica gel, charcoal or dried rice can be used for partial drying of orthodox seeds (ENSCONET 2009; Hay and Probert 2013). Otherwise, seeds can be placed in empty cloth bags for pre-drying (Engels and Visser 2003). Recalcitrant seeds lose water gradually; therefore, it is crucial to maintain the water level content at shedding levels (FAO 2014) by, for instance, storing them in the dark in closed bags to avoid drying (Ballesteros et al. 2021). Note that RH rises as temperature falls, especially overnight. Avoid extreme temperatures because they can damage the collection. Orthodox seeds can be exposed to direct sunlight after moisture reduction, whereas recalcitrant seeds should never be sun-dried (Schmidt 2000). Seeds should be stored in a cool dry location.

Seed extraction procedures are dependent on fruit and seed type. Seeds can be extracted from fruits (e.g., by drying, sifting, shaking, or flailing) in the field, unless high temperatures or threshing are required. Fruit pulp should also be removed, otherwise it will ferment, heat will be produced, and seed viability will thus be reduced (Schmidt 2000). Fruit pulp is sometimes removed upon returning to the laboratory, but it should remain fresh, as dry pulp is harder to eliminate (Filip Vandelook, pers. comm.). Some seeds cannot be separated from the fruit (e.g., nuts), as this will cause damage to the seed (Schmidt 2011). Follow Schmidt (2000) for seed extraction, depulping procedures, and temporary storage. A summary of post-harvesting recommendations can be found in the MSBP technical information sheet No. 4.

During transportation, cloth sacks containing orthodox seeds should be placed into cardboard boxes and covered with waterproof materials to protect them from moisture. Ideally, recalcitrant seeds should be stored in sacks/paper bags containing sawdust to prevent dehydration (Schmidt 2000; Schmidt 2011; MSBP technical information sheet No.3).

At the biobank facilities, seeds should be manually cleaned thoroughly or by using sieves of different sizes, rubber bungs, or aspirators to remove further debris under an extraction hood containing dust filters (ENSCONET 2009). For detailed cleaning procedures, see the MSBP technical information sheet No. 14. Seeds should be subsequently inspected for damage with a stereomicroscope and by using the cut-test (see above), and purity should be assessed using X-rays (MSBP 2015). Seeds that are empty, poorly developed, or insect-infested have to be discarded (ENSCONET 2009). If more than 50% of the tested seeds are hollow, this implies low viability, and another collection should therefore be considered (CPC 2019).

Collected seeds should be dried in a climate chamber, or in a desiccator containing either silica gel or salt solutions (Engels and Visser 2003). Regardless of the desiccation method, moisture content has to be regularly controlled, or seeds will degrade (Zimkus and Ford 2014). If immature seeds were also collected, they should be ripened before drying (MSBP 2019). Moisture content of orthodox seeds should be reduced as much as possible, and at least to the lowest possible level for recalcitrant seeds, which varies between species and within species, to avoid germination (Schmidt 2000, 2011). Consult the “culture preservation and storage” chapter of this handbook for further details on seed storage. In general, the drying process of orthodox seeds will take between one and four weeks, depending on the drying temperature and the RH. If drying at 25 °C, RH should be ca. 25–35%, whereas RH should be between 10% and 25% when drying at 5 °C (CPC 2019).

Immediately after drying and inspection, orthodox seeds should be packed in moisture-proof containers such as vacuum-sealed laminated aluminium foil bags, or air-tight containers (e.g., Kilner glasses) (Engels and Visser 2003; FAO 2014). Seed banking should occur within six months of collection or within two weeks after drying for short-lived and microscopic seeds (e.g., orchids) (MSBP 2019).

Livestock species play an important role in food production and agriculture. Following domestication, selection based on human requirements led to the development of breeds, bloodlines and landraces carrying distinct genetic profiles. The term ‘breed’ also describes the unit of conservation that will be considered for conservation management plans (CGRFA 2012).

Biobanking of livestock resources helps to optimise management decisions, breeding strategies, development of new crosses and breed-types, genetic rescue translocations, and food security (FAO 2012). However, maintaining an updated collection is a crucial key point of livestock biobanking, as genetic differences between the ex situ collection and the in situ breeding population will emerge over time (FAO 2012; Blackburn 2018). Furthermore, populations from the same breed from different geographical areas should also be sampled to uphold most of the genetic variability (Blackburn 2018). Samples must come from animals that are as unrelated as possible, as they will eventually be used for breeding purposes.

Collection and processing procedures depend on the sample type and the species. Field collection should follow strict protocols and staff should be trained for off-site collection, as disease transmission can occur when visiting different farms (FAO 2012). Consider the conservation and risk status of the breeds before sampling.

The FAO (2012) and the former IMAGE project (2020) have developed the following rules that should be applied during field collection:

• Disease testing should be accomplished before and after sampling collection.
• Ideally, samples should be collected at approved collection centres. If not the case, the sanitary status of the farm/herd/donor should be recorded as completely as possible.
• Samples from different species and with different sanitary statuses have to be placed in separate dry shippers, but may be stored in the same room, providing the tanks are clearly marked and no cooling agent can pass from one tank to the other.
• To minimise disease transmission, supplies, shipping boxes or any other instrument should not be used at other sites. These can be reused after cleaning and disinfection, but only at the same collection site.
• LN2 tanks or dry shippers used in the field should be warmed to room temperature and then decontaminated with a 10% bleach solution.
• Vehicle tires and undercarriage as well as boots have to be washed and disinfected after each farm visit. Boot covers can also be used and changed after each visit. Clothing should also be changed between collection sites or disposable suits should be worn instead. Polyvinyl or nitrile gloves should be worn and changed after handling each animal.
• Backup samples of non-germplasm material (e.g., blood) should also be collected.

Sperm collection. Semen collection is a common practice in several countries. Generally, collection procedures in mammals include electro-ejaculation, gloved-hand techniques, and the use of an artificial vagina, teaser female or a decoy animal. Abdominal stroking is used in poultry (FAO 2012). An inner liner should be placed in a collection bottle to trap the semen. For stallions, an extra non-woven filter pouch should be used to prevent the gel fraction from encountering the first sperm-rich fractions. Samples should be free of urine and other contaminants (USDA 2020). After collection, samples should be transported to the laboratory under a temperature/light-controlled environment, diluted and cryopreserved no later than 24 h upon arrival. Morphology and motility should be assessed before freezing. Species-specific collection protocols are found in FAO (2012) and USDA and can be adapted for field conditions.

When animals cannot be trained to undergo this procedure or are kept free range, it may be difficult to obtain sperm. An epididymal sperm post-mortem collection should be considered instead (ERFP 2003; FAO 2012). Preservation of epididymal sperm harvested from rats and livestock can be found in James (2004). Note that this type of sample will only preserve a single complement of chromosomes and paternal mitochondrial DNA will be lost, after fertilisation (Wolff and Gemmell 2008).

Oocyte collection. The most common method to collect oocytes involves removing ovaries from slaughtered donors. Ovaries should be kept warm in Ziploc bags during transport to the laboratory. If maintained cool, the in vitro fertilisation (IVF) embryo production rate will decrease dramatically (FAO 2012). Oocytes should be aspirated following the method described in FAO (2012, appendix H). Oocyte quality should be assessed before oocyte in vitro maturation and IVF (FAO 2012, appendix I, J). Although these methods are described for cattle, they can also be applied to other livestock species (USDA).

Oocytes can also be harvested by implementing surgical procedures such laparotomy, endoscopy, and the transvaginal ultrasound-guided method (TUGA) (FAO 2012). The latter one is commonly used in cows, goats, mares, sow, llamas, deer, and antelope. For detailed methodology, see FAO (2012). Ovariohysterectomy has also been conducted in cats to obtain oocytes (Hobbs et al. 2012).

Note that freezing and thawing techniques are not as developed as those used for sperm, and further research is needed in this field (ERFP 2003). For instance, avian and fish oocyte cryopreservation has not been successful, due to their high lipid content and polar organisation (vegetal and animal pole). The alternative option would be to use fresh oocytes and frozen/fresh semen and freeze the resulting embryos instead (FAO 2012).

Embryo collection. Embryos allow the recovery of entire genomes without back-crossing. The first step to obtain embryos is to induce superovulation in donor females by injecting hormone agents. Subsequently, in vivo fertilisation takes place. A body flush procedure should be performed to make the embryos flow out of the uterus using a physiological flushing medium. Laparoscopic surgery is required in pigs, sheep, and goats. Ultrasonography should be used before embryo collection to evaluate the potential number of embryos that can be found. As soon as the embryos are collected, they should be placed into a hypertonic solution containing a cryoprotective agent, and cryopreserved. Freezing should take place when the embryos are at blastocyst stage, which is reached by five to nine days after fertilisation, depending on the species. Follow the FAO (2012) protocols for precise methods and recommendations.

Gonadic tissues / whole ovaries collection. Ovaries are obtained by laparoscopy or immediately after the donor female dies (USDA). The ovarian tissue is mainly used to restore an animal´s fertility by transplanting it into a recipient animal. The offspring of the recipient female will then carry the donor´s genotype. This method requires surgical expertise and special grafting facilities (FAO 2012).

Although avian oocyte cryopreservation is problematic, it is still possible to freeze ovaries from newly hatched chicks within the first 24 h, because at this age, their structure is different from that of the adult ovaries. Grafting of this tissue is feasible into one-day-old chick recipients (IMAGE 2020).

Primordial germ cells (PGC) collection. PGCs are embryonic diploid stem cells that are early precursors of gametes. So far, they have only been successfully reported in fish and birds. In chickens, PGC can be collected from embryonic blood, as the PGCs migrate to the developing gonads between the fourth and sixth day of incubation. PGC can then be propagated in vitro to increase their number (Blackburn 2006; FAO 2012; IMAGE 2020). A protocol can be found in USDA.

Somatic cell collection. A piece of tissue (e.g., whole or part of an ear) should be collected and immediately frozen to obtain somatic cells that can eventually be used for cloning. This technology has been used in many domestic species, but it requires a complex approach, including reprogramming the nuclei, enucleation of the oocytes, transfer of the somatic nucleus to an enucleated oocyte, culture of the resulting embryos, and transfer into recipients of the same species (FAO 2012).

Blood or serum collection. Blood is usually taken for veterinary diagnosis and evaluation from both living and freshly dead animals. Two vials of blood (in total 10–14 ml) should be collected from the jugular or caudal vein with a needle and vacutainer tube in mammals; and from the wing veins in poultry. Vials should be frozen in LN2 vapour phase and stored in LN2 (FAO 2012).

Tissue collection for DNA/RNA extractions or pathological examinations. Organ collection should take place at slaughtering houses and should be carried out by a necropsy technician or veterinarian in a predetermined order (Leibniz Institute for Farm Animal Biology 2016). Pictures should be taken during the entire procedure. Organs should be rinsed with sterile PBS, and 1–2 cm² slices should be sampled and placed in 1 oz. Whirl-Pak bags, or 50 ml tubes to be immediately flash-frozen in LN2. Samples should then be stored on dry ice in coolers, which should be cleaned with 70% ethanol and transported in a well-ventilated vehicle to the laboratory. At the laboratory, they should be stored at -80 °C. Detailed procedures for different organs can be found in the FAANG portal: e.g., Tixier-Boichard and Fabre (2016), Burns (2018) and Jimenez and Plastow (2021).

Frozen samples (80–150 mg) should be crushed immediately for RNA extraction. DNA can be extracted using commercial kits. DNA is ideally stored in aliquots of 50 μl, with a concentration of 200 μg/ml. It can be stored at 4 °C for up to two months, otherwise it should be stored at -20 °C (for medium-term use) or lower (-80 °C or in LN2).

Conservation efforts for wild animals have so far included in situ support (e.g., habitat conservation), breeding programs at zoos and aquaria and DNA banking. The cryopreservation of blood/haemolymph, nucleotides, tissue, living cell lines, gametes and embryos has played a crucial role, among others, in assisted reproduction, animal and human medicine, evolutionary biology, systematics and conservation (Ryder et al. 2000; Bartels and Kotze 2006), as well as in biotechnology development. Animal biobanks can help to identify and prioritise which species are at risk of extinction and develop the most effective techniques for collecting, preserving, and storing genetic material (Breithoff and Harrison 2018).

Samples can be obtained during field trips or from zoological parks / breeding programs. There is no methodological difference when collecting samples from wild or captive animals. Nevertheless, the same field collection hygienic rules for livestock should be followed when sampling captive animals at zoos and aquaria (see livestock sample collection section above). Photographs of live animals, carcasses, and necropsies should also be taken from multiple angles before sampling.

The selected tissue preservation methods will depend on logistical factors, such as the duration of the field trip, convenient facilities, and taxonomic focus (Prendini et al. 2002). Wong et al. (2012) suggested a tissue sampling standardisation, ranging from one to four stars, to obtain the highest DNA/RNA quality, depending on the goal of the collection (e.g., cell line establishment, transcriptomics, whole-genome sequencing) (Table 2).

Ideally, tissues should be flash-frozen into dry shippers containing vapour-phase LN2 immediately after collection. However, LN2 tanks have to be taken to the field, if no sources are available at regular intervals, increasing costs and causing logistical obstacles that could compromise the expedition (Bennett 1999; Prendini et al. 2002; Gemeinholzer et al. 2010). This preservation method may be used when sampling captive animals or for short-term field trips. Tissues can be cut into small pieces and stored flash-frozen without preservatives, though adding preservatives prior to freezing may produce better long-term results (Mulcahy et al. 2016). They should be placed into a -80 °C freezer or LN2 tank as soon as they are removed from the dry shipper.

RNAlater, DNAgard Tissue, or AllProtect are non-hazardous stabilisation reagents that can be used as preservation solutions for several months at room temperature, but they are expensive (Zimkus et al. 2018). The most common preservation method is 95–96% ethanol, which can preserve DNA for long periods at room temperature and above (Zimkus 2018), although such temperatures will make high molecular weight (HMW) DNA applications very difficult. The use of denatured ethanol containing e.g., traces of benzene, or of lower concentrations (containing excessive amounts of water) or higher concentrations (potentially retaining chemical drying agents) of ethanol can degrade DNA (Neumann 2010). The use of DMSO/EDTA salt-saturated (DESS) solutions may be better for preserving HMW DNA in vertebrate tissues, whereas alcohol may be preferred for invertebrate tissues, although further research is needed (Mulcahy et al. 2016; Osting et al. 2020). Terrestrial invertebrates can also be trapped/preserved in food-grade propylene glycol or in ethylene glycol, although the latter poses a health hazard (Höfer et al. 2015; Liu et al. 2020). Vials should be totally filled to avoid specimen damage during transport. A minimum of a 3:1 volume ratio of ethanol to tissue is used for storage and efficient preservation. Note that to prevent watering down (due to the sample’s body fluids), ethanol should be exchanged at least once shortly after sampling, e.g., on the next morning, otherwise (or on top of the exchange) a 10:1 ratio is recommended. When using alcohol, vials should be inspected the following few days after preservation and discoloured (typically yellow) alcohol should be discarded and replaced with fresh alcohol. Tissues should be cut into small pieces to allow rapid penetration of the preservatives (Prendini et al. 2002).

Transparent plastic vials with screw-caps are used to avoid ethanol evaporation. Stripes of Parafilm can be used to seal the cap, which might become loose during transport. Be aware that some types of polymers corrode when in contact with certain insect killing agents (Krogmann and Holstein 2010). Cryotubes are sometimes used for short-term storage during fieldwork in hot climates. However, they may not be the best choice, as they are designed for contraction during cryo-processes and high temperatures may allow for high ethanol evaporation losses (Neumann 2010) in some builds.

Marine invertebrates can be collected by hand in the field in the intertidal zone or by using scuba diving (Lessios et al. 2013; Adams et al. 2019), or can be obtained from mariculture farms (i.e., commercial species). Ideally, gravid organisms should be collected, but it may be challenging to detect them. Although maturing organisms in the laboratory is possible, it is often a time consuming and costly process. Hence, it may be necessary to sample several times during the breeding season (Adams et al. 2019), due to slight fluctuations in temperature or food availability, among other factors. Specimens have to be transported alive at a constant temperature, and carefully carried, as stress (e.g., vibrations, temperature, desiccation) can make them release gametes. Individuals can be maintained together in husbandry under laboratory conditions at the highest standard (van der Horst et al. 2018; Adams et al. 2019; Zuchowicz et al. 2021) or separated by sex whenever possible (Paredes and Costas 2020). When housed together, spontaneous spawning can occur. If so, as soon as spawning is observed, the individual should be immediately put aside into an independent container, not only for gamete collection, but also to evade simultaneous spawning. Gametes can also be collected by dissecting the animals (i.e., ideal for concentrated non-activated sperm), or by inducing spawning either chemically (e.g., hormones or concentrated salt solutions) or via stress (e.g., temperature changes, vibration) (Williams and Bentley 2002; Dale 2018; van der Horst et al. 2018). Spawning and gamete collection protocols for mussels, sea urchins and sea cucumbers can be found in Paredes (2020). In crustaceans, it is possible to apply pressure on the body to obtain gametes or use electroejaculation (Aquino et al. 2022). Note that the choice of procedure will depend on the species, because the method will affect the viability of the cells. Pooling of sperm or eggs is possible for cryopreservation purposes.

Collection and maintenance methods for polychaetes can be found in Watson (1997), Williams and Bentley (2002) and Caldwell et al. (2011a); for echinoderms in Caldwell et al. (2011b), van der Horst et al. (2018) and Adams et al. (2019); for marine molluscs in Fitzpatrick et al. (2012), van der Horst et al. (2018) and Beirão et al. (2019); for crustaceans in Rendón-Rodríguez et al. (2007), Beirão et al. (2019) and Aquino et al. (2022); for nemerteans in Stricker et al. (2012); for ascidians in Yoshida et al. (2003); and for corals in Zuchowicz et al. (2021). Research on other invertebrate phyla is still scarce or non-existent (Poccia 2013; Zuchowicz et al. 2021). So far, only culturing and gamete morphology observations have been accomplished for tardigrades (Sugiura et al. 2022 and references therein).

A special issue, focused on collection, handling, and storage of gametes and embryos from marine invertebrates, has been published in the journal Animals. Further information can be found in Lewis and Ford (2012).

Insects can be reared in colonies or in small stock groups where mixed adults are free to mate to obtain eggs and embryos (Vasudeva et al. 2019; Zhan, et al. 2021). Sperm from ants and bees can be collected from freshly killed mature males by squeezing the abdomen until semen appears on the endophallus (Pearcy et al. 2014; Hayashi and Satoh 2019). Guidelines for mosquito colony maintenance can be found in FAO/IAEA (2017). Spiders should be maintained in individual tubes for some days to ensure that palps are recharged. Pedipalps should be removed, and the genital bulb separated from it to obtain the sperm (Gabel and Uhl 2013).

Parasite collections are underrepresented in biodiversity biobanks, in contrast with other taxa such as mammals or insects (Bell et al. 2018). The most effective method to collect parasites is as soon as the host animals are euthanised and processed in the field (Gardner 1996), otherwise they should be kept frozen until further analyses (Galbreath et al. 2019). Make sure to follow animal welfare regulations and approved euthanasia/necropsy protocols depending on taxon. Parasites can also be collected from fur harvest carcasses, game, and road kills, as well as from fish markets and fresh faeces (Julca et al. 2014, Kutz et al. 2014; Caira et al. 2017; Schwartz et al. 2020).

It is also possible to obtain certain parasites from live hosts subdued by light-anaesthetising or by holding (Gardner and Jiménez-Ruíz 2009). Note that mainly ectoparasites will be collected by this method.

Faeces from small mammals should be taken from the rectum and placed in a Wheaton snap-cap vial half-filled with 2% potassium dichromate K₂Cr₂O₇ to obtain coccidian parasites, helminth eggs or protozoan oocysts (Gardner and Jiménez-Ruíz 2009). Fresh faecal pellets from large mammals (e.g., ungulates) should be stored in plastic bags with silica gel. Faecal material can also be preserved in 96% ethanol or frozen until processed (Galbreath et al. 2019). Nematodes can be collected from faeces using the Baermann beaker method (Forrester and Lankester 1997; Western College of Veterinary Medicine, University of Saskatchewan) or by following a modified version by Galbreath et al. (2019).

Blood-borne parasites can be examined either by preparing blood smears in the field or by collecting blood with FTA paper, Nobuto filter strips, or hematocrit tubes, which can be sealed with Critoseal (Gardner and Jiménez-Ruíz 2009; Galbreath et al. 2019).

Crops and grasses can be parasitised by nematodes of the family Heteroderidae. When looking for nematodes, the upper soil layer (3–6 cm) should be removed before sampling (van Bezooijen 2006). Do not collect soil that is too dry, as dormant nematodes may already be damaged (Shivas et al. 2005). Soil core samples should be collected after harvest at a depth of 15–30 cm from several spots around the field in a zigzag pattern and taken from around the plant root. Entire plants that show symptoms of nematode infection can be also sampled using the “Stretcher” method (van Bezooijen 2006).

Samples should be placed in individual plastic bags or paper bags coated with paraffin to retain moisture. Samples should be kept cool and out of direct sunlight, which can cause nematodes to die from shock. Samples should be transferred to the laboratory to be processed as soon as possible (van Bezooijen 2006; Jagdale and Arnold-Smith 2011; Smiley et al. 2017). Several techniques are used for extracting nematodes from both plant material and soil using either a Whitehead tray or a Baermann funnel, or sieving, decanting, and flotation methods (Shivas et al. 2005). Refer to van Bezooijen (2006) and Shivas et al. (2005) for protocol details.

Preservation methods are identical to the ones mentioned for nematodes parasitising animals.

No general guidelines exist for sample volume, depths, and water quantities. However, 500 ml – 2 l is the usual standard sampling volume from streams, rivers, lagoons, and seawater (Shu et al. 2020). Common sampling containers are 1-l HDPE bottles (Nalgene) or 5-l buckets, which should be decontaminated by soaking them in 10% bleach (sodium hypochlorite, NaOCl) for at least 30 min to remove all traces of DNA. Containers are then rinsed with distilled water to remove residual bleach (Eichmiller et al. 2014; Spens et al. 2017).

Surface water samples are the most widely used sample in eDNA and are taken by partially submerging a sample bottle or a sterile 60 ml Luer lock syringe to collect water from the top few cm (~10 cm) to 6 m. Sub-surface samples (~50 m) can be taken using a stainless steel Van Dorn sampler (Wildlife Supply Company) (Eichmiller et al. 2014) or automated samplers (Formel et al. 2021), whereas a rosette sampler or 5 l- Niskin bottles attached to remotely operated vehicles are used for deeper sampling (>1,000 m) (Everett and Park 2018; Laroche et al. 2020; Canals et al. 2021).

Water samples should ideally be processed on-site, but if this is not possible, they have to be placed on ice and remain refrigerated (4 °C) to avoid DNA degradation. They should be processed within 24 h of reaching the laboratory (Hinlo et al. 2017; Harper et al. 2019; Shu et al. 2020; Bruce et al. 2021), otherwise, preservation buffers should be added immediately after collection (Mitchell and Takacs-Vesbach 2008; Williams et al. 2016; Kumar et al. 2020). Small volumes (<100 ml) collected with syringes can be stored in sterile Whirl-Pak until processed (Torresdal et al. 2017). The addition of 0.01% benzalkonium chloride can also mitigate microbial growth in the water samples (Yamanaka et al. 2017; Kumar et al. 2020).

Samples from the same waterbody with no stratification can be pooled and processed (preservation, eDNA capture, analysis) as one single sample or they can be treated as independent replicates. The latter should be preferred to increase species detection or if the aim of the project is to establish species distribution and habitat use (Harper et al. 2019; Shu et al. 2020).

Filtration and ethanol precipitation are the most used methods for eDNA capture (Hinlo et al. 2017; Harper et al. 2019). The precipitation method uses ethanol (twice the sample volume) or isopropanol (same volume as the samples with sodium acetate to precipitate nucleic acids in the water sample, but it is only feasible for a small water volume (<100 ml)) (Kumar et al. 2019). Samples can be kept at room temperature for up to seven days (Harper et al. 2019; Wang et al. 2021). Otherwise, they can be kept at -20 °C until DNA extraction (Egeter et al. 2018).

Filtration is a more effective method, as it can process larger volumes of water (0.5–2 l), and thus produce higher amounts of eDNA (Hinlo et al. 2017; Egeter et al. 2018). Filters are made of diverse materials, such as glass fibre, polycarbonate, and cellulose nitrate, and have diverse pore sizes, ranging from 0.22 μm to <10 μm, with 0.45 μm being the most used pore size (Egeter et al. 2018).

Two types of filters can be used: fully capsule-enclosed filters (e.g., Sterivex-GP) or open filters (Bruce et al. 2021). Filters enclosed in capsules reduce handling and contamination risk, as capture, storage and extraction take place within the capsule (Spens et al. 2017; Shu et al. 2020). Sterile syringes (e.g., 50 ml Luer-lock syringe) are used to slowly push water through the filter without tearing it (Holman et al. 2019). Any remaining water in the capsule should be removed with the syringe by pushing air through the filter. Preservation buffers (1.5–2 ml) can be added to the inlet end, and both the outlet and inlet ends should be closed with caps and sealed with parafilm. Capsules should be inverted vigorously and can be placed in -20 °C until extraction with or without preservation buffer (Spens et al. 2017; Holman et al. 2019). Another alternative to fully encapsulated filters is the use of self-preserving filters, which are compatible with suction pumps (e.g., Smith-root) and capture eDNA via desiccation at ambient temperature (Thomas et al. 2019).

Open filters require handling; thus, water samples should be filtered using sterile disposable filter funnels, and a vacuum or peristaltic pump (e.g., Nalgene 250 ml) (Spens et al. 2017). Filters should be removed with sterilised forceps and then folded in quarters with the filtrated side facing inwards (Carim et al. 2016). Filters can be placed dried or with preservation buffers in 5 ml - microcentrifuge tubes (e.g., DNA LoBind) or in Petri dishes, and frozen immediately at -20 °C (Spens et al. 2017). Alternatively, filters can be placed into clean plastic bags or tubes containing silica beads, where they can remain for several weeks. However, samples should ideally be processed within the first two weeks after collection (Carim et al. 2016; Sales et al. 2019). Once collected, eDNA samples should be kept cool, dry, and in the dark (Spens et al. 2017).

The use of sponges (Porifera) as natural samplers can also be a powerful and attractive method to detect eDNA from aquatic biodiversity. With further optimisation and validation, it will have the potential to filter more water and hence, to recover eDNA more efficiently than traditional methods (Mariani et al. 2019; Turon et al. 2020; Cai 2022; Harper et al. 2022).

Ponds and lakes. Lentic waters are basically motionless, allowing eDNA to accumulate over time (Harper et al. 2019), yet eDNA has a patchy distribution in such waterbodies (Eichmiller et al. 2014). Moreover, the presence of fallen trees and dense aquatic vegetation can be barriers to eDNA dispersion. Hence, samples should be collected at different spots, including underneath and around barriers, as well as at different depths to increase species detection (Harper et al. 2019). Filters can become blocked, when working in turbid waters, because of high levels of suspended solids and organic debris. Centrifugation, increased filter pore size, or pre-filtering will then be required (Harper et al. 2019; Bruce et al. 2021). An alternative is to use the precipitation method (Egeter et al. 2018).

Although sampling should be done at many different locations, limited accessibility (e.g., dense vegetation, or high steep banks) can hinder its optimisation. Sampling poles and boats can be used to enable water sample collection, but cross-contamination may occur between ponds. Several parameters should be recorded for better assurance and resolution of eDNA detection: 1) The total size of the pond perimeter, 2) the pond proportion that was accessible, 3) the water volume sampled and, 4) the number of samples and the distance at which these were taken (Harper et al. 2019).

It is important to note that terrestrial animals might transfer eDNA from one waterbody to another, causing false positive detections. (Harper et al. 2019).

Streams. eDNA is distributed more evenly in rivers due to the turbulent flow of streams, hence surface samples should be sufficient for eDNA detection (Shu et al. 2020). Consider sampling either downstream or upstream of tributaries, as well as across the river breadth. Note that eDNA may be diluted while moving downstream. Eddies or splashes should be avoided, as the water sample can get contaminated (Sales et al. 2019).

Carim et al. (2016), Lehmann (2016), and Pawlowski et al. (2020) developed sampling protocols for small creeks and rivers. The latter also includes step-by-step photographs of the whole sampling procedure. Kumar et al. (2022) provide protocols for both turbid and clear-water environments.

Oceans. eDNA concentration is usually low due to dilution (Bruce et al. 2021), plus eDNA can only be detected at the depth where it was released or deeper. Hence, vertical profiles play an important role in oceanic diversity detection (Canals et al. 2021). Furthermore, recovered eDNA will mainly represent local fauna, as eDNA does not exhibit a long-distance dispersal (Thomsen et al. 2016). Water samples obtained using Niskin bottles should be transferred to sterile 1 l bottles and stored at -20 °C until filtering is done at the laboratory (Thomsen et al. 2016).

It is important to note that current databases have a low representation of deep-sea organisms, hampering the analyses and further implementation of conservation measures (Laroche et al. 2020). However, attempts have been made to centralise genomic data from marine organisms (i.e., http://reefgenomics.org) (Liew et al. 2016).

Throughfall in forest ecosystems and rainwater sampling can be revised in Šantl-Temkiv et al. (2018) and Ladin et al. (2021).

For protocols for other types of freshwater bodies (e.g., subterranean water, anchialine habitats, temporary, small water bodies) refer to Valdecasas et al. (2010).

Sediment samples offer two advantages over other environmental sample types: first, eDNA can attach to particles in the sediment, providing a higher concentration that makes it easier to detect. Second, as eDNA can persist longer in this substrate, it is ideal to study past and current events, along with DNA transport and removal. However, this can also be a source of false positives, if past species are considered present in current times (Sales et al. 2019). A possible solution is to use radiocarbon dating techniques to confirm the time origin (Epp et al. 2012). eDNA in sediment samples is also sensitive enough to distinguish diverse communities that are geographically close (Laroche et al. 2020).

eDNA does not mix well in sediments, therefore it is necessary to take several subsamples that cover most of the sampling area. In general, subsamples between 0.25–1 g are collected for microorganisms, whereas 10–20 g are required for small invertebrates and macrofauna (Bruce et al. 2021). Samples are generally taken either with a sterile spatula (i.e., for shallow layers), a petite ponar grab sampler (Wildlife Supply Company) (i.e., for hard sediment samples), an epibenthic sledge (Glover et al. 2016), or a corer (Eichmiller et al. 2014; Inagaki et al. 2015; Glover et al. 2016; Bruce et al. 2021). Cores can be horizontally divided for further subsampling using single-use sterile syringes (60 ml) to extract minicores (Laroche et al. 2020; Capo et al. 2021). Clean tools between cores with 10% bleach and rinse them with double-distilled water (Laroche et al. 2020). Samples should be stored in Whirl-Pak bags on ice for up to eight hours and filtered within 24 h (Eichmiller et al. 2014). Samples should be preserved frozen at -20 °C, or -80 °C, or in some preservation solutions (e.g., ethanol for macrofauna, Qiagen’s LifeGuard for microorganisms and meiofauna, and sucrose lysis buffer for extreme environments) (Mitchell and Takacs-Vesbach 2008; Bruce et al. 2021; Pawlowski et al. (2022). DESS can also be added for meiofaunal samples Pawlowski et al. (2022).

Protocols and laboratory pipelines for marine sediment can be found in the IODP/CDEX/KCC (2012) guideline and Glover et al. (2016). Furthermore, some of the field methodology aspects in Glover et al. (2016) are explained in a video (https://youtu.be/Io7NlFKUmYI). Pawlowski et al. (2020) have also developed sampling protocols for sediments, including step-by-step photographs of the whole sampling procedure. Pawlowski et al. (2022) provide a review regarding sediment sampling and DNA extraction. A guideline to choose the correct sampling and preservation method can also be found in Pawlowski et al. (2022: supplementary material). Capo et al. (2021) provide an overview and recommendations concerning sampling, extractions and THS technologies applied to sediments.

DNA can be extracted directly from the sediment sample, or the organisms (i.e., macrofauna) found in the sediment can be set apart by sieving and processed as a bulk sample (Bruce et al. 2021; Pawlowski et al. 2022).

Soil samples have been used for a long time to study taxonomic diversity in plants, fungi, and invertebrates (Thomsen and Willerslev 2015), as well as for characterising bacterial communities (Fløjgaard et al. 2019). Still two issues remain to be solved: the time persistence of eDNA in the soil matrix (Foucher et al. 2020), and the low representation of soil invertebrates in databases (i.e., BOLD) (Rota et al. 2020).

Ideally, two samples should be collected randomly or on a regular grid within each plot in the sampling location (Bienert et al. 2012; Taberlet et al. 2012). To account for soil heterogeneity, each sample has to comprise a pool of several (4–80) subsamples (Taberlet et al. 2012). The required amount of soil per subsample is usually 20 g, with a final volume ranging between 500 g to 4 kg (Taberlet et al. 2012; Clasen et al. 2020). Soil can be collected from the surface (a depth of 0–20 cm) and the sub-surface (a depth of 20–40 cm) using a soil corer, a disposable shovel, a thistle remover, a trowel, or by hammering a metal cylinder (Janzen et al. 2007; Bienert et al. 2012; Epp et al. 2012; Weinfurtner and Kördel 2012; Foucher et al. 2020). All tools should be sterilised with a high-temperature flame to avoid cross-contamination between soil samples (Bienert et al. 2012). Note that subsamples from different soil layers can be analysed independently to allow for comparisons (Rota et al. 2020) or can be pooled as suggested by Taberlet et al. (2012). Do not forget to remove the litter layer, large roots, and stones before sampling (Weinfurtner and Kördel 2012; Fløjgaard et al. 2019; Clasen et al. 2020). Soil samples can be placed into wide-neck barrels, Nalgene tubes or Whirl-Paks, and kept dark and cool at 4–5 °C until reaching the laboratory (Meyer et al. 2019; Clasen et al. 2020).

Samples can be sieved (3 mm) to further remove stones and enable upcoming soil homogenisation (Clasen et al. 2020). Soil samples are generally processed immediately after collection (Taberlet 2012; Rota et al. 2020). If this is not possible, samples should be immediately freeze-dried, which is the best alternative for homogenising soils prior DNA extraction (Rüdel et al. 2011; Clasen et al. 2020). If impractical, samples can be placed in Ziploc bags containing silica gel bags for drying (Foucher et al. 2020), or they can be air dried at room temperature or using an unheated air source like a fan. Alternatively, DNA/RNA Shield (1 sample:9 buffer) or DESS solution (1 sample:3 buffer) can be added to the homogenised samples but note that these preservation solutions may induce some variation in the microbial taxonomic structure (Pavlovska et al. 2021). Do not pre-freeze samples, because of the risk of thawing, which can lead to spoilage and fungal growth (Clasen et al. 2020). Samples should then be stored at -20 °C or -80 °C for up to 13 months or until DNA extraction (Rhymes et al. 2021).

Airborne eDNA studies have mainly focused on bacteria, fungal spores, algae, and pollen, with the key aim of monitoring airborne pathogens in relation to human health (Johnson et al. 2019; de Groot et al. 2021; Pumkaeo et al. 2021). Airborne eDNA methods have recently been applied to terrestrial ecosystems, proving its potential for population monitoring and biodiversity conservation (Johnson et al. 2019, 2021; Howell et al. 2021; Pumkaeo et al. 2021; Bohmann and Lynggaard 2022; Clare et al. 2022; Lynggaard et al. 2022a, 2022b). Both microorganisms and eDNA are scattered in the atmosphere, making their concentrations lower than in other environments (Banchi et al. 2020a). Sampling efficiency is, therefore, influenced by air volume, flow rate, collection medium, along with particle size and density (Behzad et al. 2015; Mainelis 2020). Several sampling approaches are used to collect airborne biological particles using either adhesive tape, air filters, or dust traps (Lynggaard et al. 2022a; see Mainelis 2020 for more details). Selecting the most fitting method depends on the studied taxon. Note that the sampling procedure, i.e., type of chosen trap, and recovery procedure, and duration can overall affect the results, especially concerning the low represented taxa (Aguayo et al. 2018).

Air sampling protocols are not standardised, which makes it difficult to compare results from different studies to choose a method. Nonetheless, knowing the differences between the methods can help to choose the best option for your sampling design. Further research is also needed to improve bioaerosol sampling along with DNA extraction methods across taxonomic groups (Bohmann and Lynggaard 2022).

Gravity and passive sampling. This method relies on the ambient wind conditions at the study site (Johnson et al. 2019). It usually involves exposing Petri dishes containing agar (Mainelis 2020), or a mix of vaseline and paraffin wax to the outer atmosphere (Aguayo et al. 2018), or using Whatman paper filter (grade 1 or 3) (Mainelis 2020). The use of Tangle-Trap sticky coating on paper filters is also possible (Aguayo et al. 2018). These traps can be secured on blocks placed 1 m above ground for fungal spores and pollen collection (Aguayo et al. 2018). Note that this type of sampling may be affected by particle size and shape, preferring larger and heavier spores and pollen sorts (Behzad et al. 2015). There are four possible methods for recovering spores from paper filters: 1) by placing them in Petri dishes containing TE buffer, 2) rubbing, 3) grinding, or 4) shaking. Methodology details can be found in Aguayo et al. (2018). It is important to note that such sampling is the most sensitive to climatic conditions and hence, biological material may get lost (Aguayo et al. 2018).

Alternatively, Big Spring Number Eight (BSNE) dust traps can be used, mainly for plant community surveys (Johnson et al. 2019). To collect samples, each tray should be rinsed with 500 ml of deionized water, which should be pooled and collected into a single 1 l sterile bottle. Water samples should be labelled and transported dark and cool at 4 °C to the laboratory and filtered following the aforementioned water filtration method. Filters should be stored at -20 °C until DNA extraction (Johnson et al. 2019; 2021).

Active sampling. Vacuum or peristaltic pumps are utilised to actively collect particles, such as bacteria, spores, pollen, whole tiny arthopods and eDNA from animals (Ovaskainen et al. 2020; Clare et al. 2021; Serrao, et al. 2021). Sampling usually takes ca. 10 min. Samples can be collected either by using an adhesive coated tape (Melinex) placed on the sampler (Banchi et al. 2020b), or by directing the aerial particles into 1.5 ml sterile centrifuge tubes (Ovaskainen et al. 2020). After sampling, the coated tape can be stored at room temperature, and before processing, it should be cut into segments that can be rolled into 1.5 ml sterile centrifuge tubes, with the sticky side facing the internal part of the tube (Banchi et al. 2020b). On the other hand, the centrifuge tubes should be removed from the sampler and transported at room temperature. Final storage should be at -20 °C until further processing (Ovaskainen et al. 2020). If insects are found in the tubes, they should be rinsed with sterile water into the sampling tube and removed with sterile tweezers (Ovaskainen et al. 2020).

Active sampling also includes impaction sampling and filtration sampling.

Impaction sampling. Particles in the air are impacted onto a wet collection medium using either liquid impinger-based samplers (Behzad et al. 2015) or Andersen samplers (Mainelis 2020). Particles are collected into sterilised deionized water, NaCl solution or PBS solution with or without surfactants (e.g., 0–25% manucol), or onto Petri dishes with agar. Note that liquid media can evaporate within two hours, damaging the sample (Behzad et al. 2015). See Šantl-Temkiv et al. (2018) for detailed methodology. Samples should be kept at 4 °C.

Once a sample (mainly bacteria and fungi) is collected, the agar plates can be transferred directly to an incubator without intermediate steps, or they can be placed in Ziploc bags and placed into a Styrofoam cooler with cold packs, to be shipped off as soon as possible. Note that culturing methods, if considered, will leave out a considerable viable fraction of the total microorganisms that is not culturable.

Filtration sampling. Filtration-based air samplers work by drawing air through filters of varying pore sizes, shapes, and compositions (Mainelis 2020). Capsule-enclosed filters with a pore size of 0.22 μm and 0.45 μm are commonly used (Clare et al. 2022). Once the particles are collected onto a filter, they can be eluted into liquid for subsequent analysis (Lynggaard et al. 2022a). Filters can also be placed dried in Ziploc bags and kept frozen until DNA extraction (Clare et al. 2022). Filter type, sampler type and speed will determine sampling duration. Sampling time ranges between 30 min and 30 h. This method has been successfully applied to terrestrial vertebrates (Clare et al. 2022; Lynggaard et al. 2022a).

Filters (e.g., F8 or F7 pleated fibrous particulate or HEPA), which are used for heating and ventilation systems, may also have the potential to collect eDNA as a byproduct of regular operation (Clare et al. 2022; Lynggaard et al. 2022a 2022b).

Permafrost-preserved bones, teeth/tusks, horns, skulls, and even large carcass remnants, like skeletons, and tissue belonging to extinct megafauna (e.g., steppe bison, woolly mammoth, woolly rhinoceros, horses, bears) are among the various fossils found in Eurasia and North America (Funck et al. 2020). Even logs and tree trunks have been found in Pleistocene peaty sediments (Martinez de la Torre et al. 2019). Usually, these fossils emerge during gold mining operations (Shapiro and Cooper 2003), construction activities (Zazula et al. 2017), or have been exposed and excavated from eroding riverbanks (Funck et al. 2020). Hence, sampling is considered opportunistic, and no excavation protocols are needed.

Gold miners regularly uncover fossils while thawing out frozen sediments to expose bedrock using high-pressure water and steam (Shapiro and Cooper 2003). Once the fossil has been removed from the permafrost, it will quickly deteriorate due to temperature and humidity fluctuations (Florida Museum). Fossil samples should not be kept frozen, unless they were recovered in that condition, because several freeze/thaw cycles damage DNA (Fulton and Shapiro 2019). Ideally, specimens should be dried slowly to avoid cracking and delamination (Florida Museum) and should be stored in a cool dry environment at a stable temperature (Fulton and Shapiro 2019).

A salvage excavation should begin when remains are found unexpectedly in a specific location (e.g., a construction site), to reveal the site stratigraphy and search for further remains. A backhoe should be employed to excavate slowly until a bone bed is detected, followed by a manual excavation using shovel and trowel (Zazula et al. 2017).

When permafrost carcasses are relatively complete, sediments within the body cavity should be discarded with a trench shovel before specimen removal from the site. Skin and any possible soft tissue should be removed from the bones by metal knives and scalpels. Hair, preserved soft tissue, and associated sediment should be individually stored and kept frozen (Funck et al. 2020). Samples for DNA studies should be taken before carcass chemical preservation (Maschenko et al. 2017).

DNA preservation depends on the environment (Fulton and Shapiro 2019). Samples from mid-continental temperate areas tend to be poorer than those from high latitudes, as warm climates and humidity accelerate post-mortem degradation of DNA (Chang et al. 2017; Kehlmaier et al. 2017; Meyer et al. 2017). However, temperate, and humid tropical caves and water-filled sinkholes have been found to contain a high diversity of faunal remains where microclimate has been stable, and no harsh environmental changes have occurred since the Late Pleistocene (Slon et al. 2022). These environments tend to be dark, cool, and wet, as well as anoxic and thermally buffered in the case of sinkholes, enhancing DNA survival (Dabney et al. 2013; Gutierrez-Garcia et al. 2014; Kehlmaier et al. 2017; Sheng et al. 2018). DNA has also been successfully retrieved from tropical lacustrine, swamp, and marine sediments (Kehlmaier et al. 2017; Dommain et al. 2020). Furthermore, dry sediments also seem to preserve DNA, although contamination and DNA leaching within stratigraphic layers may occur due to vertical transport (Dommain et al. 2020). Challenges of working with ancient samples from warm environments are described in Letts and Shapiro (2012).

New methods for retrieving DNA from sediments are being developed to overcome the dependency on fossil remains, which are a limited and scarce genetic resource (Vernot et al. 2021). sedaDNA is absorbed by clay minerals, apatite, silica, or other sedimentary materials (Dommain et al. 2020). Further information on sedaDNA, including improvements and challenges, can be found in Capo et al. (2022).

The following procedure applies both to permafrost and temperate samples. Cores should be collected under controlled conditions and contaminations should be monitored (Willerslev et al. 2003). Before collection, core equipment should be washed with boiling water to reduce DNA contamination. Cores should be wrapped in plastic wrap and placed in acrylonitrile butadiene styrene (ABS) tubes for transport and permanent cold storage. Cores should be divided in two longitudinal parts. Ideally, one section should be kept as an archive sample, and the other one used for destructive subsampling. Sediment from the core surface should not be sampled to avoid any contamination (Dommain et al. 2020). Around 2 g of sediment should be taken from the core´s centre with sterile tools and stored frozen at -20 °C in falcon tubes until further analyses (Willerslev et al. 2003; Boessenkool et al. 2014).

Samples can also be collected either with a clean scalpel blade (Slon et al. 2022), a trowel, by inserting 50 ml Falcon tubes, or by using 5 ml screw cap tubes in a drill-like manner to excavate a few grams of sediment from the middle of each stratigraphic layer or from many points sampled every few centimetres of a pit sampling column. Usually, 1–2 mm of the surface material is removed to minimise contamination (Slon et al. 2022). Bulk-bone samples can be collected by excavating sediment and dry-sieving through 1.5–3 mm sieves (Seersholm et al. 2020). Samples should be placed in individual Ziploc plastic bags and stored in a refrigerator until further processing (Vernot et al. 2021). Sterile masks and gloves should be worn during sample collection to avoid contamination by modern DNA.

As aDNA concentration is lower in deeper strata, samples from cores or pit columns should be sampled from bottom to top. Note that the mentioned core subsampling procedure has to take place at a core storage facility rather than at an aDNA laboratory (Epp et al. 2019).

More recommendations regarding subsampling and decontamination of permafrost core samples can be found in Saidi-Mehrabad et al. (2020). For optimised sedaDNA extraction protocols from both frozen and non-frozen cores, refer to Haile (2012), Epp et al. (2019) and Vernot et al. (2021).

Nonmineralized dung remains are called palaeofaeces and are usually found in caves and in dry areas. Palaeofaeces contain the DNA of both the defecator and the ingested organisms. Faeces should be cut into small pieces and added to the extraction buffer. Addition PTB, a thiazolium salt, into the buffer may help to improve the total DNA yield (Kuch and Poinar 2012).

Archaeobotanical remains, mainly domesticated plant species suitable for DNA analyses, are scarce, not only because plant material tends to decompose rapidly, but also because they are restricted to caves, arid areas, and waterlogged grounds. Most plant fossils are preserved by charring, and DNA basically cannot survive under such conditions (Nistelberger et al. 2016; Przelomska et al. 2020). Bunning et al. (2012) produced sequences from old, charred cereal grains. However, great caution should be taken when interpreting these data, as the obtained sequences might be artefacts of spurious mapping (Nistelberger et al. 2016). An assessment of carbon and nitrogen isotope values of archaeobotanical remains should be performed to determine the degree of charring before proceeding with DNA research (Nistelberger et al. 2016).

For protocols that can be used for non-charred desiccated and waterlogged plant remains see Gilbert et al. (2004), Wales et al. (2014) and Wales and Kistler (2019).

Natural history museum collections function as banks for ancient and archival DNA. Museum specimens allow studying organisms for which fresh tissue samples are impossible to obtain, either because they are already extinct, or because they are only found in inaccessible regions. Furthermore, museum collections allow temporal comparisons between historical and modern populations. Obtaining endogenous DNA from museum specimens is feasible despite their unfavourable preservation conditions. A complete review regarding the advances and challenges in museomics is provided in Card et al. (2021) and Raxworthy and Smith (2021). Experimental and computational protocols to obtain genomic data from historical specimens can be found in Cong et al. (2021).

Museum and herbarium material also require special handling and processing comparable to palaeontological remains. The principles for aDNA should therefore be followed (see DNA chapter), as many museum samples yield low amounts of endogenous DNA (Kistler et al. 2020).

Be aware that the following aDNA methods apply both to palaeontological / archaeological material and museum material.

Bones and teeth. Freshly excavated and untreated bones are preferred over fossil samples from natural history museums, as their preservation and storage conditions can be detrimental for DNA preservation. Fossils at museums are normally kept in cardboard boxes in rooms where both temperature and humidity are not stable. Hence, Pruvost (2007) suggested that fossils should be stored in a cold room, and ideally in a cryobank in small aliquots to avoid freezing and thawing cycles. Protocols at museums should be revised and assessed to improve preservation and retrieval of genetic data from archaeological/paleontological samples (Pruvost 2007; Seersholm et al. 2020).

Solid, heavy, and dense bones with few cracks are preferred for aDNA studies (Fulton and Shapiro 2019). The inner part of the petrous bone is considered the best source of mammal aDNA, and it has proven to be successful even with material from warmer climates such as Africa, the Near East, or Oceania (Hansen et al. 2017). For morphological identification in different taxa, see Bar-Oz et al. (2019). However, some destructive sampling is required, as the petrous bone has to be removed from the skull and the vestibulo-cochlear area has to be drilled into. Other options, such as the auditory ossicle and the circumferential lamellae of long bones, have also been found to promote DNA preservation, due to the absence of bone remodelling (Ferrari et al. 2021).

Non-destructive pre-screening techniques should be applied before sampling begins to determine whether a bone is suitable for aDNA analysis (Keighley et al. 2021). The amount of organic matter, such as collagen, contained in fossil bones may indicate that DNA could also be preserved within them (Kehlmaier et al. 2017). Tripp et al. (2010, 2018) suggested a micro-computed tomography (MicroCT) method to identify microstructural features associated with collagen preservation, which may help to choose the best bone, and the best sampling location within the bone. Knowing how to select the right objects will reduce the destruction of zooarchaeological remains, helping with their long-term preservation (Ferrari et al. 2021).

Different from mammalian bones, teleost fish bones undergo bone remodelling to a lesser extent, or it is totally absent due to the lack of osteocytes (Shahar and Dean 2013). This improves the bone resistance to microbial degradation, making all archaeological bones, cranial, pectoral girdle or postcranial, suitable for aDNA studies, even if they look porous and brittle (Ferrari et al. 2021).

In general, no more than 1 cm³ of bone or a single tooth root is used to recover DNA, although DNA quantity will decrease rapidly if the specimens are exposed (uncovered) for several seasons (Shapiro and Cooper 2003). The surface of the bone should first be removed using a Dremel rotary tool with a cutting blade. Engraving cutters and drills should be used to hollow out sections of the bone to gain access to the inner material (Fulton and Shapiro 2019). Bone powder can be collected by using either a Spex freezer mill or a 20 mm drill bit at very low speed (approximately 60 rpm) (Barnes et al. 2007). Note that a high drill speed will produce heat and vibration, which will be detrimental to DNA. The quantity of bone powder used for DNA extraction ranges between 30–200 mg following the Dabney et al. (2013) standard protocol that enhances the recovery of short aDNA fragments. For a more detailed protocol see Dabney and Meyer (2019).

Tooth root cementum is equally good as the petrous bone, in case it is not poorly preserved (no cementum, brittle, “chalk-like”) (Hansen et al. 2017). Often entire tooth roots are lost when retrieving DNA because of drilling or grinding. New protocols use non-destructive sampling, and teeth can still be used for radio-dating or morphological studies. Check Hofreiter (2012), Gomes et al. (2015), and Harney et al. (2021) for further information. Tooth roots may be demineralized in 0.5 M EDTA (pH 8) in a thermomixer, set at 1000 rpm and 22 °C, for 24 h, prior to extraction (Eaton et al. 2015).

To remove extraction buffers after a non-destructive sampling method is performed, bones and teeth should be transferred to double-distilled water for 24 h at room temperature and the previous step should be repeated for another few hours. Samples should be removed from the tube and air-dried at room temperature (Hofreiter 2012).

Damgaard et al. (2015) recommended the following points when extracting aDNA from ancient bone or teeth:

• A brief predigestion step (15–30 min) should be applied, but only if more than 50 mg of the sample is available.
• the pre-digest supernatant should not be discarded at first, as it contains endogenous DNA. This also applies to undigested bone pellets post-24 h digestion.
• Several extractions should be performed if sufficient material is available.
• Tooth roots should be sampled from the surface rather than the inner dentine. The outermost surface layer should be removed before sampling to minimise the risk of contamination.
• The proportion of endogenous DNA will also be maximised if genome capture, single-stranded sequencing libraries, or damage-enriched single-stranded sequencing libraries are performed.

Note that a predigestion step with or without a bleach wash may not increase sequencing efficiency (Dehasque et al. 2022). Dehasque et al. (2022) recommend the use of MinElute columns over QIAquick columns for ancient DNA extractions.

Hides. Skins should be cut into small pieces (5 × 5 mm) from the initial incision made during skin preparation using a sterilised surgical blade. This procedure should cause no significant loss and should not jeopardise further morphological studies (Moraes Barros and Morgante 2007). Each tissue sample should be transferred to a sterile Eppendorf tube. To diminish contamination, hairs and the outer layer should be removed with a sterilised scalpel and skin should be cleaned and washed three times with sterile milli-Q water, three times with 70% ethanol, then rinsed three times with sterile milli-Q water, and finally cut into small pieces. Museum skins were commonly treated with gasoline and butane, hence several washes with absolute ethanol for 24 h have to be performed to remove these PCR inhibitors (JEMU 2013).

Bird toe pads. This tissue is the best source of DNA from bird skins, as toe pads do not have much contact with preservatives (Lijtmaer et al. 2012). Approximately 2 × 4 mm of the inner toe pad should be removed with a disposable razor blade, avoiding the tissue surface. Tissue should be stored at room temperature until DNA extraction. Toe pads should ideally be rinsed in 0.5 M EDTA to wash away inhibitors, and DTT should be added in the first lysis step to reduce the amount of time required to completely dissolve the tissue. Detailed protocols can be found in Fulton et al. (2012) and Irestedt et al. (2022).

Keratin and chitin material. DNA can also be retrieved from hair, nails, horns, hooves, feathers, and insect cuticles, and these are simple to decontaminate (Campos and Gilbert 2019). Two crucial steps should be considered: 1) breaking the keratin/chitin by using detergents and reducing agents such as proteinase K, EDTA, and DTT; and 2) to purifying DNA using either silica-based methods or isopropanol when extracting with organic solvents (Campos and Gilbert 2019). Samples should be washed with a 0.5% commercial bleach solution and rinsed between two and six times in DNA-free water until traces of the bleach are removed (Gilbert et al. 2007a). A protocol for DNA extraction can be found in Campos and Gilbert (2019).

Insects. Non-destructive DNA extraction methods have been applied to pinned insects. In general, whole specimens should be placed in Eppendorf Biopur tubes, fully immersed in digestion buffer, and incubated overnight at 55 °C with gentle agitation for 16–20 h. Subsequently, specimens should be placed in 100% ethanol for 2–4 h to stop further digestion, air-dried, and returned to their collections. Refer to Gilbert et al. (2007b), Tin et al. (2014), Cong et al. (2021), and Cavill et al. (2021) for detailed protocols.

Eggs. Membranes from blown eggs can be removed through the blow-hole by using fine dissecting tools such as high-precision tweezers (e.g., no. s5/45 and 7), microforceps (no. 7) and fine shaped probes (T198 TAAB). This method preserves the egg´s appearance, patterning, colour, and shape dimensions, which is important for other scientific studies. Eggs with tiny holes and cracks around the blow-hole should be avoided. Samples should be stored in microfuge tubes at room temperature (Lee and Prys-Jones 2008).

DNA from fossil eggshells can also be obtained, as DNA is protected in calcite intracrystalline depositions within the eggshell matrix. A DNA extraction protocol can be found in Oskam and Bunce (2012). Note that the outer surface has to be removed either with a Dremel tool (>0.7-mm thin eggshell) or sandpaper (<0.7-mm thin eggshell) to avoid contamination.

Alcohol-fixed material. DNA in tissue preserved in alcohol at room temperature will degrade over time. However, different protocols have been developed to obtain useable DNA from fluid-preserved specimens, reviewed by Ruane (2021) and Bernstein and Ruane (2022).

Formalin-fixed material. DNA quality from formalin-fixed or paraffin-embedded tissues is affected by temperature, duration of exposure, and pH of the fixative. Campos and Gilbert (2012) suggested a DNA protocol that includes using heat and alkali treatment to break protein-DNA cross-links. A small piece of formalin-fixed tissue or thin slices from paraffin-embedded tissue can be taken using a sterile scalpel. Remove paraffin by trimming it away with a scalpel if it is present in large amounts. As heating is involved, O-ring screw-cap tubes should be used to avoid lids from being blown open. An optimised protocol for extracting RNA from formalin-fixed material can be found in Speer et al. (2022).

Resin-embedded specimens. It is under debate whether amber or sub-fossilised resins are suitable for genetic studies (Peris et al. 2020; Modi et al. 2021). DNA preservation in these materials may be limited by time and needs to be determined (Peris et al. 2020). A standardised protocol for insects embedded in resin has been developed by Peris et al. (2020), which can only be applied to modern samples (up to six years old). Although Modi et al. (2021) have proven that it is possible to recover endogenous ancient DNA from insects embedded in copal (a predecessor of amber), reproducibility of the results was not possible.

Mummified material. Ancient DNA research on mummified material, many animal species besides humans, is challenging, not only because mummification techniques, such as desiccation and anointment comprised the use of bandages and chemicals (e.g., heated oils, resins, natron) that are unfavourable to PCR conditions, but also because sometimes it is not anymore possible to verify how the original field collection occurred (Kurushima et al. 2012).

Another type of mummified material are the so-called bog men, i.e., human remains that survived in peatlands. Several types of tissue can be sampled by qualified staff following medical methods. Samples should be placed in vials containing distilled water and maintained in waterlogged conditions at 4 °C until further analyses are conducted (Mulhall 2020).

Samples should be removed using either a flexible fibre optic endoscope with grasping forceps, sterilised surgical tools or bore drill bits for larger animal mummies (Loreille et al. 2018; Hekkala et al. 2020). To prevent contamination with modern DNA, samples should be washed with 5–20% bleach for 15 min, then 80% alcohol, followed by rinsing with sterile water (Wasef et al. 2019). Muscle tissue (ca. 0.2 g) may be placed in glycine buffer for one week to three months with regular fluid changes to hydrate the sample (Hekkala et al. 2011). If drilling is necessary, the sampled area should first be wiped with 0.5% bleach and air-dried. Bone or tissue powder should be collected on a sterile foil reservoir (Hekkala et al. 2020).

For DNA extractions, follow Schwarz et al. (2009), Loreille et al. (2010), Rohland et al. (2010; 2018), Dabney et al. (2013), or Korlevic et al. (2015).

Herbarium material. Plant tissues preserved by freezing or desiccation are preferred for aDNA studies. aDNA has been successfully retrieved from seeds, wood, pollen, and vegetative tissues. However, a pure DNA extraction may be challenging because compounds, such as humic acids and polyphenols, along with heavily lignified epidermal layers, are considered PCR inhibitors (Wales et al. 2014). Additional reagents plus incubation time and temperature modifications to commercial kit protocols and removal of lignified tissue with a sterile blade should increase PCR success (Kistler 2012). Although DNA from herbarium tissue may be degraded, it is possible to obtain reliable sequence data (Staats et al. 2011).

Kistler (2012) suggested two protocols for DNA extraction: the CTAB protocol for single seeds, except for protein-rich, oily seeds (e.g., bottle gourd, sunflower); and the PTB (N-phenacylthiazolium bromide) protocol for gourd rind-like tissues and wood samples. See Dumolin-Lapegue (1999), Asif and Cannon (2005), Deguilloux et al. (2006) or Rachmayanti et al. (2006) for more wood DNA extraction protocols. Shepard (2017) suggested a non-destructive DNA sampling protocol for leaves and stems, and Saługa (2020) for mosses. Cubero et al. (1999), Särkinen et al. (2012), Drábková (2014), Shepherd (2017) and Drábková et al. (2021) have also developed protocols for herbarium material.

A mortar and pestle or a mechanised mill can be used for grinding samples. Sterile sand can be added for tough samples. Small seeds should be ground in an incubation tube to avoid tissue loss. Consider incubating in buffer before grinding, to soften tough samples. Remember to sterilise the equipment with a bleach solution (10%) between samples (Kistler 2012).

Protocols for historical plant specimens can be found in (Marinček et al. 2022) and for historical lichen specimens and mushrooms in Dentinger et al. (2016) Forin et al. (2018), and Kistenich et al. (2019).

Serial or periodic transfers to new medium should take place every 3–6 months and cultures should be kept at 3–8 °C or at 15 °C if containing cold-sensitive strains (Shivas et al. 2005). Keep in mind that after many serial transfers, some strains may show a compromised ability to grow because of degeneration, which can ultimately lead to their loss (Heinonen-Tanski and Holopainen 1991). Fresh or herbarium-preserved mature specimens with spores or hyphae should be used for culturing and they should not have been previously refrigerated or frozen (Wrigley de Basanta and Estrada-Torres 2017). Culture flasks (e.g., McCartney) should be closed with a plastic membrane or another means (e.g., plug or cap) to minimise contamination risks. As soon as a culture is established, several subcultures can be prepared. Some of these should be used for serial transfers at temperatures ranging from 4 °C to room temperature, whereas others should be allowed to dry or cryopreserved. Serial transfers can also be performed to maintain algal and cyanobacterial cultures under suboptimal conditions, but genetic and phenotypic stability cannot be ensured (Metting 1994; Friedl and Lorenz 2012).

Preservation can also take place on agar strips using vacuum-drying cultures, which may work for some groups otherwise difficult to preserve (e.g., Pythium (Oomycetes)) (Nakasone et al. 2004; Singh et al. 2018).

Pure colonies can be obtained after isolating one single spore or a hyphal tip. The former consists of serial spore suspensions until reaching an optimal dilution, which is then transferred to a Petri dish for incubation. The latter involves the removal of a hyphal tip from a mycelium edge and subsequent incubation to obtain a pure colony (Narayanasamy 2011; Capote et al. 2012). Many cultures can be stored at -20 °C or -80 °C in sterile 15–50% glycerol solution; the optimal combination will depend on the species (Akinsamni et al. 2017).

For permanent preservation as non-living specimens, non-sporulating cultures can also be dried in a laminar flow hood for two or three days. Then they are withdrawn from the Petri dish and placed in herbarium packets. Delicate fungi can be stored in small Petri dishes. To avoid curly and brittle results, the partially dried culture can be placed on hot glycerol agar and dried completely (Shivas et al. 2005).

Culture methods for slime moulds (Mycetozoa) have been described in Haskins and Wrigley de Basanta (2008) and in the “Eumycetozoan project”. Note that it is highly difficult to keep axenic (i.e., contaminant-free) cultures of Myxomycetes, as they require bacteria as food source (Wrigley de Basanta and Estrada-Torres 2017). Mycetozoa cultures that are dried and stored at room temperature can be revived up to a year later by rewetting with sterile distilled water (Spiegel et al. 2004).

Generally, unicellular protist cultures should be established at conditions that resemble as much as possible their environment (light/darkness, temperature, oxygen, pH), and they should be maintained along the culturing process (Altermatt et al. 2015; Esteban et al. 2015). Initial cultures usually take place in microtiter plates and can be transferred to other containers such as culture tubes, Erlenmeyer jars with loose lids, or tissue-culture flasks for further subculturing. The culture medium will depend on the diet of the organisms (Esteban et al. 2015), but usually natural seawater/freshwater is used as a basis for growth media (Altermatt et al. 2015; Weber et al. 2017; Hansen et al. 2020). The use of a proteose peptone medium is recommended for axenic cultures, whereas a protozoa pellet medium is more suitable for non-axenic ones (Altermatt et al. 2015).

Organic material can also be added to the media, but this method is less standardised and unreliable (Lee and Soldo 1992; Altermatt et al. 2015), as some of the microorganisms will be unknown (agnotobiotic cultures). Nevertheless, most protozoan strains growing in liquid culture media need the addition of sterile cereal grains (CCAP 2020). Subculturing of strains should be done weekly or up to every two months, depending on the species. Note that every time a strain is subcultured, the food source (e.g., bacteria) will be also co-transferred (CCAP 2020). It is recommended to have 4–8 replicate cultures of each species stored in two separate incubators (Altermatt et al. 2015).

Original field samples of protists can also be used as culture, but they can only be maintained if the water sample contains a sufficient amount of plant material and is kept cool. However, this type of culture can only survive up to a week, at which point most species will have disappeared from the sample due to exclusion or predation (Department of Biological Sciences, George Washington University).

Protist soil samples can be cultured following the “non-flooded Petri dish method” to obtain ciliates (Foissner et al. 2002): Depending on the type of soil, ca. 150–600 g (including litter) are placed in a Petri dish, which is then over-saturated (110%) with distilled water and set aside one month. A species succession will occur, and therefore, examination should take place on predefined days (2, 7, 14, 21, and 28). Note that this method does not reactivate all cysts (Foissner et al. 2005). See Foissner et al. (2005) for a short description of other soil culture methods.

A comprehensive compendium of isolation and culture media for fungi, Oomycetes, and Myxomycetes has been produced by Bills and Foster (2004). For further preservation protocols and maintenance of fungal cultures including recommendations, refer to Singh (2017). Methods for cultivating microalgae can be found in Acreman (1994), Guillard and Morton (2004), Day et al. (2007), Garibay and Sadio (2020), and Martinez-Goss et al. (2020). Further culturing protocols for several protists can be found in Lee and Soldo (1992). Culturing protocols focusing on specific taxonomic groups are also available, i.e., for Acanthamoeba species (Axelsson-Olsson et al. 2009), choanoflagellates (King et al. 2009) and ciliates (Liu et al. 2019). Media recipes for protists can be found in Andersen (2005) and Altermatt (2015).

This method is ideal for mycelial and non-sporulating cultures that cannot undergo freezing, except Basidiomycota (Homolka 2014). Sterile mineral oil is used to cover agar cultures, thus preventing dehydration and slowing down metabolic activity and growth (Bégaud et al. 2012; Caleza et al. 2017; Singh et al. 2018). Vials can be sealed with parafilm before storage (Abd-Elsalam et al. 2010). Even though this method reduces mite infestations, vials need to be regularly inspected for air-borne spore contamination (Homolka 2014). Cultures can persist for up to 40 years, depending on the species, and ideally renewal should take place every five years (Shivas et al. 2005). This method is preferred when lyophilisation or cryopreservation is not possible (Ryan et al. 2000). Note that selection of mutants may occur due to fungi continuously growing in this preservation method (Homolka 2014). Singh et al. (2018) provide a protocol for long-term preservation of live cultures.

Young and vigorous colonies, including spores, hyphae, or yeasts, on agar are cut in blocks and placed in McCartney bottles, containing sterile distilled water. The bottle lids should be tightly screwed down. Storage is recommended at room temperature (Agarwal and Sharma 2006) or kept cool in a refrigerator (Abd-Elsalam et al. 2010). Water and inoculum blocks should follow a 40:1 volume ratio. If excessive amounts of inoculum and medium are added, the fungus will not endure (Caleza et al. 2017).

This method works very well for early-diverging fungi, filamentous fungi, moulds, and yeasts which can survive more than a decade, including plant pathogens (e.g., wood-decomposing Basidiomycota and oomycetes such as Pythium, Phytophthora spp.), and ectomycorrhizal fungi (Heinonen-Tanski and Holopainen 1991; Burdsall and Dorwoth 1994; Bégaud et al. 2012; Homolka 2013). This method is not recommended for taxa previously assigned to Zygomycota since these are less stable in this type of preservation (Singh et al. 2018).

This method is ideal for microorganisms that can produce conidia (asexual spores) or resting structures (Jong and Mirmingham 2001; Bégaud et al. 2012). A spore suspension is added to sterilised sieved soil and incubated at room temperature for 5–10 days depending on the fungus growth rate. Then it should be stored at 4–7 °C (Shivas et al. 2005; Agarwal and Sharma 2006). A detailed protocol can be found in Singh et al. (2018). Soil-borne pathogens (e.g., Fusarium) can be preserved by this method (Abd-Elsalam et al. 2010); however, soil preservation is not recommended for ectomycorrhizal fungi (Heinonen-Tanski and Holopainen 1991).

Silica gel can prevent fungal growth and metabolic activity, minimising the risks of genetic and morphological changes (Abd-Elsalam et al. 2010). Some fungi can be stored for up to 18 years in this way (Bégaud et al. 2012). Typically, glass flasks are filled to one quarter with non-indicating silica gel and sterilised in the oven at 180 °C for 3h. Bottles should be placed in a water tray and frozen in a deep freezer. Spore suspensions are added to the bottles and incubated at 25 °C for 10–14 days. Bottles are then stored at 4 °C in airtight containers with indicator silica gel. Silica gel is suitable for organisms that produce sclerotia or chlamydospores (Shivas et al. 2005). However, this method is not ideal for thin-walled spores, spores with appendages and mycelia, or for ectomycorrhizal fungi (Heinonen-Tanski and Holopainen 1991; Agarwal and Sharma 2006). Two protocols are provided in Nakasone et al. (2004).

This method has been designed specifically for wood-inhabiting fungi (e.g., Basidiomycota and Ascomycota), but some pathogenic fungus-like protists can also be maintained with this preservation method (Abd-Elsalam et al. 2010). Around 50 small pieces of wood should be collected into a flask, which should be sterilised at 121 °C for 20 min twice. Fifteen chips should be placed on pre-grown colony Petri dishes and incubated for 10–15 days. Colonised wood chips are transferred to sterile test tubes, which are stored at 4 °C (Nakasone et al. 2004; Singh et al. 2018).

Collected pieces of rotting wood can also be placed in standing water in a Petri dish or onto agar plates to grow protists such as protostelids. Cultures should be kept in moist chambers and inspected daily for up to two weeks with a microscope (Spiegel et al. 2004).

This method uses sterile cereal grains (e.g., rye, barley, millet) to grow both microfungi (including pathogenic species, e.g., Rhizoctonia), and mushrooms (see macrofungi section below for more details). After inoculation, microfungi can be stored at -20 °C to 5 °C, depending on the species. It should be used for storage times of less than one year (Singh et al. 2018). One advantage of this method is that subculturing is not needed (Singh 2017). Cultures can be dried in a desiccation chamber and kept at -25 °C. Alternatively, colonised grains can be transferred to a sterile Eppendorf tube and stored at -80 °C (Abd-Elsalam et al. 2010).

Agar plugs are placed on the filter paper (e.g., Whatman No. 1) and stored in Petri dishes at room temperature until the paper is fully colonised by fungi. Filter paper should be completely dried, cut in small pieces and stored in air-tight vials at 4 °C (Shivas et al. 2005). A protocol can be found in Singh et al. (2018). Mycorrhizal fungi and saprotrophic fungi can be preserved on charcoal filter paper strips for subsequent cryopreservation (Stielow et al. 2012). Ciliate dried cysts can also be preserved using filter paper (McGrath et al. 1977).

Microalgae can be immobilised and preserved in pieces of cotton cloth at 4 °C in the dark. Refer to Prasad et al. (2016) for details.

Note ectomycorrhizal fungi and obligate pathogens can only be maintained with their host plant either in diaxenic or semi-aseptic cultures.

A substantial number of protocols is available for pure culture preservation and storage of microorganisms, but techniques are still scarce for intact (unhomogenised) samples or microbiome preservation. Some techniques developed in the food industry and therapeutics can be applied to the long-term preservation of microbiomes, such as CAS (cell alive system) freezing under an alternating magnetic field (Morono et al. 2015) for up to six months, or high-voltage microencapsulation techniques (e.g., electro-spraying and electro-spinning) (Lim 2015; Rodríguez-Tobías et al. 2019; Stojanov and Berlec 2020). Prakash et al. (2020) provide an overview of current preservation methods of intact samples, mixed cultures and microbiomes.

Pollen. Comparable to seeds, pollen can also be tolerant or sensitive to desiccation. Most pollen grains are desiccation-tolerant and have moisture contents below 30% (fresh weight basis) at dispersal. They are also termed partially dehydrated or orthodox and are characterised by small size, often between 30 μm and 100 μm (Pacini and Franchi 2020) and a storability at low moisture contents (< 15%) and low temperatures (Dinato et al. 2020). Pollen grains with a moisture content of above 30% at dispersal are often desiccation-sensitive and are also termed partially hydrated or recalcitrant. They often measure between 15 μm and 150 μm (Pacini and Franchi 2020) and commonly do not tolerate water loss below 30% (Nebot et al. 2021). However, pollen of species producing orthodox seeds are not necessarily desiccation-tolerant (Franchi et al. 2011).

Orthodox pollen can be stored dry in Eppendorf tubes, cryovials, glassine bags or gelatine capsules sealed airtight in aluminium pouches at -20 °C, -80 °C or -196 °C in LN2 (Anushma et al. 2018; Sidhu 2019). Drying is recommended at RH near 30% (Nebot et al. 2021). Alternatively, pollen may be stored dry at 4 °C, but viability will decrease after a couple of days or weeks, depending on the species (Sidhu 2019). Pollen is generally very short-lived, and viability can eventually decrease even during LN2 storage (Pence et al. 2020), for yet unknown reasons. Therefore, viability testing procedures should be standardised and carried out periodically (Towill and Walters 2000).

Recalcitrant pollen requires cryopreservation (see cryopreservation chapter). Note that the moisture content has to be chosen carefully. It needs to be above a level at which desiccation damage occurs and below or close to the limit at which water can freeze (Nebot et al. 2021).

Seeds. More than 90% of angiosperm species are known to produce orthodox seeds that can tolerate water contents of less than 10% (Alpert 2005; Hoekstra 2005). Hence, they can be maintained for extended periods if storage conditions are optimal, namely dry and cool (FAO 2014). By contrast, less than 10% of the species are known to produce recalcitrant or intermediate seeds that do not survive drying or are sensitive to drying, respectively (Wyse et al. 2018). However, so far, many seeds of tropical species have not been classified yet and might belong to the last two categories. In any case, seed storability is dependent on seed moisture content, storage temperature and atmosphere (Roberts 1961, 1972; Theilade and Petri 2003). Anoxic conditions may prolong seed longevity during storage (Groot et al. 2015). A compendium of storage behaviour of different species and protocols to determine seed storage behaviour can be found in Hong et al. (1996), Royal Botanic Gardens Kew (2017), and MSBP-Kew technical information sheet 10. Further information about factors that affect long-term conservation of seeds in genebanks are provided by Kameswara Rao et al. (2017), and suggestions to improve seed longevity can be found in Ballesteros et al. (2020). Comprehensive overviews regarding seed bank storage can be found in Smith et al. (2003), Hay and Probert (2013), Solberg et al. (2020) and Martyn Yenson et al. (2021).

Orthodox seed storage. Typically, dry mature seeds are further dried to an equilibrium moisture content in a controlled environment of 5–20 °C and 10–25% RH. Long-term storage collections are mainly kept at -18±3 °C (FAO 2014; MSBP 2015). Note that seeds of wild relatives may behave differently from seeds of domesticates, hence optimal storage conditions should be individually defined (Engels and Visser 2003). For optimal short-term storage, the seeds should be stored at the same temperature at which they were dried, which is usually 15 °C and 15% RH (ENSCONET 2009; FAO 2014).

Silica gel can be useful for drying seed collections to recommended moisture contents in the laboratory, in the field, or at other situations where a permanent drying facility is not available. However, it is advisable to monitor the seed moisture content when seeds are kept in closed containers with indicating (colour-changing) silica gel for extended periods. There is a lower limit for the effectiveness of moisture content reduction in extending storage longevity and for some otherwise orthodox seeded species (e.g., Salix spp.), over-drying can be harmful (Walters and Engels 1998; John Dickie pers. comm.).

Some species produce orthodox seeds that are short-lived, such as seeds from Allium species or lettuce (Nagel and Börner 2010), or various wild species. These species are prepared for storage rapidly after maturation drying or may be stored in the vapour phase of LN2 (see cryopreservation chapter).

Alternatives to silica gel drying are ultra-dry storage (Hong et al. 2005), sun/shade drying (Hay and Probert 1995, 2013), and vacuum/freeze drying that allow storage at room temperature (Theilade and Petri 2003). The use of zeolite beads is common in horticulture, allowing seeds to be kept at room temperature (Hay et al. 2012; University of California, Davis). The protocol for seed drying and storage using zeolite beads has been developed at the University of California Davis.

Recalcitrant and intermediate seed storage. Unlike orthodox seeds, recalcitrant seeds are not desiccation-tolerant and die when exposed to low RH and lower temperatures, whereas intermediate seeds usually have short lifespans in dehydrated or/and low-temperature conditions (FAO 2014; Ballesteros et al. 2021). For short- or medium-term storage (i.e., in the case of tropical species or in some temperate species), recalcitrant seeds are maintained in hydrated storage to avoid water loss, which can lead to germination or poor viability and vigour (FAO 2014). Seeds should be placed into paper bags within polyethylene bags, or they can be placed in sterilised plastic buckets with sealing lids. In general, seeds should be stored at the lowest temperature they can tolerate, and it is critical that temperature remain constant (i.e., temperate seeds 4 ± 4 °C, tropical seeds 16 ± 2 °C). Containers should be regularly ventilated to evade anoxic conditions (FAO 2014).

Plant tissue can be stored in various ways: 1) boxes containing small amounts of silica gel and a relative humidity indicator card, 2) into vials containing RNAlater or a similar product, 3) in CTAB buffer at -20 °C, or -80 °C, or 4) they can be flash-frozen in liquid nitrogen, particularly if high molecular weight DNA is required (CPC 2019; Forrest et al. unpublished)

In vitro techniques are important tools for conservation, especially for the propagation, regeneration, multiplication, and restoration of plant genetic resources that cannot be efficiently maintained and regenerated via seeds (Rowntree 2006). Vegetatively conserved species are mainly those that rarely or never produce seeds (e.g., garlic, banana), species with recalcitrant or intermediate seeds, (e.g., mango, coffee, papaya), or species that do not breed true and produce heterozygous seeds (e.g., potato, apple, strawberry) (Reed et al. 2013). Commonly, these species are immediately introduced into tissue culture (Pence et al. 2002) and preserved under so-called ‘slow-growth conditions’ for short- and medium-term storage (Chauhan et al. 2019; Panis et al. 2020). Benefits and risks of plant tissue culturing are described in Sommerville et al. (2021).

It is important to note that in vitro tissue culture should be initiated immediately after plant material collection, either in the field (see plant collection section) or more commonly in the laboratory. For introduction to tissue culture and propagation of in vitro plantlets, young or growing tissues, preferably meristematic tissues, are used as explants. This often includes shoot tips, nodal and floral segments, embryos, and embryonic axes (Paunescu 2009). Explants are commonly immersed in sodium dichloroisocyanurate (NaDCC or SDICN) or sodium hypochlorite to limit endophytic contamination prior to placing them onto an axenic culture (Engelmann 1997; Sarasan et al. 2006; Senula and Nagel 2021). Possible solutions to overcome critical obstacles (e.g., tissue and medium browning or necrosis) to successful culture initiation include the addition of activated charcoal, DMSO or PVP, or transferring to fresh media cultures (Sarasan et al. 2006). A detailed review regarding contamination control during in vitro initiation can be found in Pence et al. (2002) and Reed et al. (2004).

In general, tissue culture protocols and media composition should be developed for each plant group, species, or in some cases even for specific genotypes, as they can behave differently in similar culture conditions (Ashmore 1997). Before culture initiation, it should be determined whether different media for initiation, proliferation and storage are necessary (Reed et al. 2004). In addition, growth conditions (e.g., light quality, intensity, and exposition time, temperature, O₂ and CO₂ concentration, availability of space, and addition of mycorrhiza) should be carefully adjusted and controlled for different species (Kubota 2001; Sarasan 2010), especially for those that require particular treatments to grow in vitro (e.g., immature seeds from temperate orchids can only germinate using a mycorrhiza-assisted germination technique (Diantina et al. 2020)). Depending on the species, cultures may be maintained in liquid or solid media and can contain macro- and micronutrients, sugars, supplements such as phloroglucinol or difluoromethylornithine (DFMO), and phytohormones to promote regeneration (Sarasan et al. 2006; Chauhan et al. 2019; Panis. et al. 2020).

In recent decades, different techniques have been widely implemented for conservation, propagation, and multiplication of plant material, mainly of Angiosperms (flowering plants) and Gymnosperms (conifers, cycads). In vitro techniques are dependent on the plant species, the intended use and the targeted organ (Reed et al. 2004; Sarasan et al. 2006; Cruz-Cruz et al. 2013; Sommerville et al. 2021):

• Micropropagation of nodal segments, axillary or terminal vegetative buds, or excised meristems. Generally, plant segments containing meristematic tissue are used for vegetative propagation (Engelmann 2010). There are a number of protocols established for root and tuber crops, fruit trees, ornamental and medicinal plants, and wild species among others, from temperate and tropical origin (Pence et al. 2002). For root cuttings of woody species, vermiculite, coir, perlite, or paper pulp can support efficient root development (Sarasan 2010).
• Callus culture. Callus cells are considered unorganised, non-differentiated meristematic cells that can develop into a differentiated plant under appropriate conditions (Efferth 2019). To reduce the risk of mutations, callus cultures are usually not used for the conservation of plant genetic resources (Panis et al. 2020). However, it has been applied efficiently for mass production of plantlets (i.e., in tree species such as Neolamarckia cadamba) (Huang et al. 2020). Different ratios of plant growth regulators of auxins, cytokinins and gibberellins are used to stimulate callus cultures or root and shoot formation (Efferth 2019).
• Suspension cell culture. Loose clumps of callus are usually placed on a liquid culture medium, where they break into small fragments to form a cell suspension. Usually, a sample of this solution is transferred to other types of medium to induce the production of more callus, somatic embryos, or adventitious shoots (Sommerville et al. 2021).
• Somatic embryogenesis. In somatic embryogenesis, the formation of plant embryos from somatic cells is initiated. This can occur in nature under stressful environmental conditions. In tissue culture, it is controlled by plant growth regulators and includes induction, embryo development, maturation, and germination (Méndez-Hernández et al. 2019). The composition of the media and environmental conditions are species-dependent, and require precise adaptation (Bhojwani and Dantu 2013).
• Protocorm-like bodies (PLBs) formation. PLBs are a specific type of somatic embryos that occur only in orchids and are highly used for micropropagation purposes (Cardoso et al. 2020). Reviews, including optimised parameters for the development or improvement of protocols, can be found in Teixeira da Silva et al. (2015), Zanello and Cardoso (2019) and Cardoso et al. (2020).
• Zygotic embryo culture. Zygotic embryo culture is often implemented after so-called “embryo rescue”, which implies the excision of embryos from seeds to assist the development of a vigorous plant from dormant, weak, immature, or hybrid seeds (Raghavan 2003). Culture methods are reviewed in Winkelmann et al. (2010) and Thorpe and Yeung (2011).
• In vitro seed germination. To support seed germination, seeds are disinfected and grown under optimum conditions on agar. After normal seedlings develop, they are commonly used for molecular studies or transferred and planted into soil.

For bryophyte tissue growth, cultures are commonly started from spores but sporophytic and gametophytic tissues can also be used to initiate cultures. Plant material should be sterilised using NaDCC (sodium dichloroisocyanurate) without further detergents, as they can have detrimental effects on the cultures. Ideally a pre-culture step should take place to increase the amount of starting material before culture initiation. Two protocols can be found in Rowntree (2006).

For fern tissue growth, spores are sterilised to generate in vitro gametophyte cultures and sporophyte culture lines. Sporophyte cultures can also be initiated using other fern tissues such as rhizomes, shoot tips or bud scales. However, the culture growth often takes weeks to years, until a sporophyte can be obtained. Refer to Ballesteros and Pence (2018), Barnicoat et al. (2011) and Menéndez et al. (2011) for detailed protocols and suggested media.

Slow-growth preservation procedures are suitable for short-term and medium-term storage (Chauhan et al. 2019). This approach reduces staff time because subculture intervals can be prolonged to up to four years (Ashmore 1997). Various factors should be considered when using slow-growth procedures and a combination of these is commonly used for several species (Ashmore 1997; Reed et al. 2004; Sarasan et al. 2006; Keller et al. 2011, 2012; Priyanka et al. 2021; Senula and Nagel 2021; Sommerville et al. 2021):

• Low temperature: plant material is kept at temperatures ranging from 1° C to 10° C for cold-tolerant species, and from 15°C to 28°C for tropical species.
• Reduced light conditions or culture in the dark.
• Addition of osmotic agents (e.g., mannitol, sorbitol, PEG) or growth retardants (e.g., abscisic acid). Be aware that these may increase somaclonal variation. Culture media should be preferably hormone-free.
• Covering explants with mineral oil or paraffin can be useful to reduce culture growth (Cruz-Cruz et al. 2013).
• Low oxygen: the concentration of oxygen is reduced, but the atmospheric pressure is maintained by adding nitrogen. Plant tissue growth will decrease, as will the photosynthetic activity.

Plant propagules, especially somatic embryos, nodal segments, shoot tips, and somatic embryos, as well as spores are artificially encapsulated in a protective coating, forming the so-called synthetic seed (Jang et al. 2020). Seed production and storage can be accomplished either by slowly desiccating the seeds and encapsulating them in polyoxyethylene (Polyox), or by using hydrogel capsules (i.e., sodium / calcium-alginate), in case the plant species are recalcitrant. Capsules can be 1) placed directly for sowing (no need for acclimatisation), 2) placed on nutrient media until formation of plantlets, 3) stored in a laboratory fridge at 4 °C for short-term storage, or 4) stored in LN2 for long-term storage (Qahtan et al. 2019). For further details on production, techniques, protocols, and examples, see Faisal and Alatar (2019).

Different techniques are employed for culturing invertebrates (i.e., to maintain in captivity for research or teaching purposes), and they depend on the organism group. Culture and maintenance of micrometazoans, coelenterates, scyphozoans, bryozoans, polychaetes, and small crustaceans can be found in Smith and Chanley (1975). Culture of sea hares, nematodes and fruit flies can be found in Smith et al. (2011). Methods and protocols for culturing eutardigrades can be found in Degma (2018) and Roszkowska et al. (2021). Further methods for aquatic and terrestrial invertebrates can be found in Thorp (2015), and recommendations in Rogers and Thorp (2015).

In vitro cell cultures are an important tool for ex situ conservation due to their potential to maintain a renewable source of high-quality genetic material (e.g., genomic DNA) for in principle unlimited periods without affecting animal welfare (Houck et al. 2017; Ryder and Onuma 2018; Cardoso et al. 2020; Sano et al. 2022). Cell cultures have been used for genomic and evolutionary studies, conservation (e.g., genetic rescue), toxicology, host-pathogen interactions, and medical research (Ezaz et al. 2009; Bui-Marinos et al. 2022), as well as for cloning, generation of induced pluripotent stem cells (Ben-Nun et al. 2011; Borges and Pereira 2019; Iqbal et al. 2021) and even for food production (FDA 2018; Goswami et al. 2022).

Tissues to be used for the establishment of cell cultures are mostly collected post-mortem, usually during necropsy or surgical procedures (Houck et al. 2017; Bui-Marinos et al. 2022; Logan et al. unpublished), or when carcasses are discovered in the field (Calle and Ramírez 2022; Mattos et al. 2022). It is crucial that tissues are not necrotic, decomposed and do not bear signs of disease. Non-invasive and minimally invasive techniques can also be used, especially for threatened or rare animals (Ezaz et al. 2009; Moghanjoghi et al. 2018; Cardoso et al. 2020b), which depending on the size, must be anaesthetised (Praxedes et al. 2018). The collector should be a specialist in animal capture and should be familiar with laboratory-based tissue cultures, in case sample processing occurs under field conditions (Ryder and Onuma 2018). Feathers, placenta tissues at birth, gonads, skin, eye, trachea, lung, and kidney tissues are the main sources for the establishment of fibroblast cultures (Houck et al. 2017; Ryder and Onuma 2018). Blood samples can also be used, but collection may cause bleeding and stress to animals. Furthermore, it is difficult to obtain enough blood from small animals (Ezaz et al. 2009).

Before sample collection, the tissue surface should be wiped down with 70% alcohol. Collected tissue should be washed with sterile balanced salt solutions (e.g., PBS) and antibiotics, dried on cellulose filter paper, and minced into small pieces (0.5–1 mm²) using a sterile scalpel blade or fine scissors (Ezaz et al. 2009; Wong et al. 2012; Siengdee et al. 2018; Bui-Marinos et al. 2022). If skin biopsies are taken, subcutaneous tissue (e.g., connecting tissue) and the front and back of the skin can be separated (Siengdee et al. 2018).

Ideally, the sample should be cultured immediately, but if this is not possible, it can be placed in a vial containing growth medium or PBS and antibiotics, stored within 24 h post-mortem at 4 °C up to five days (Wong et al. 2012; Houck et al. 2017; Praxedes et al. 2018; Strand et al. 2020; Sano et al. 2022). During transportation, samples should be carried in a chilled carrier (e.g., styro-foam container) and must not be in direct contact with ice packs to avoid sample freezing (Siengdee et al. 2018). Note that storage time for non-mammalian vertebrate tissues may be shorter than for mammals (Wong et al. 2012). If long-term storage of primary tissues is necessary, the vial should contain a cryoprotectant in concentrations between 5% to 15% (usually 10%) covering the whole sample, and it should be first placed at -80 °C in a CoolCell, Mr Frosty, or any other freezing container (cooled at 1 °C/min) for 24 h before being transferred to LN2 (Freshney 2010; Wong et al. 2012; Strand et al. 2021; Sano et al. 2022). Initiating cultures from tissues that were previously frozen and thawed will take longer and cultures will, in general, grow slower (Ryder and Onuma 2018), as freezing may affect cell adhesion and growth rate (Sano et al. 2022). Wong et al. (2012) and Sano et al. (2022) developed sample protocols for freezing tissue biopsy samples from wild animals prior to the initiation of cell culture.

Two methods exist to establish cell cultures: the explant and the enzymatic digestion method. In the former, the explant tissue adheres to the vessel surface, so that fibroblasts can migrate out of the tissue, whereas in the latter, the tissue is disaggregated by using an enzyme such as collagenase or trypsin for 30 min or longer, depending on the type of material (Cardoso et al. 2020b). Note that explants might lead to a more short-lived cell culture, especially relevant for small samples (Wong et al. 2012). Coating mixes (e.g., FNC coating mix) can also be added to improve cell adhesion (Sano et al. 2022).

Tissue should be transferred to a cell culture vessel and incubated at 37 °C for mammals, 38–40 °C for birds (Cardoso et al. 2020b), 26–32 °C for reptiles (Fukuda et al. 2012; Moghanjoghi et al. 2018; Kulak et al. 2020; Logan et al. unpublished), 20–30 °C for amphibians (Strand et al. 2021), and at a broad temperature range (20–32 °C) for fish (Lakra et al. 2011; Goswami et al. 2022), being 28 °C the optimum for marine fishes (Yashwanth et al. 2020). Cells usually start growing within 4–5 days and cell migration and proliferation may take 1–2 months or longer depending on the tissue, the growth conditions (Bui-Marinos et al. 2022) and the group of organisms (amphibian and reptile growth rates are lower than in mammals and birds) (Camilla Di Nizo, pers. comm.). Subculturing is performed by detaching the cells from the vessels by trypsinization or mechanical means when the cells reach 70–90% confluence, or when cell growth is excessively dense (Phelan 1998; Ezaz et al. 2009; Ryder and Onuma 2018; Praxedes et al. 2021; Bui-Marinos et al. 2022; Logan et al. unpublished). The obtained cell suspension can then be split into fresh cultures (Phelan 1998). Bui-Marinos et al. (2022) provide guidance in determining when cells from a tissue explant culture should be subcultured.

Sample collection (including post-mortem biopsies), processing and cell culture protocols for different vertebrates can be found in Wong et al. (2012) and Houck et al. (2017). Primary culture methods (i.e., first culturing event) and cryopreservation methods can be found in Freshney (2010) and Ryder and Onuma (2018). Guidance on good cell culture practice and on fundamental techniques in cell culture can be found in ECACC (2018).

Note that these cell cultures are not technically immortalised, implying that after many passages they will reach a state where they undergo ageing and stop propagation (Galli and Lazzari 2021). Ideally, established finite cell cultures with no signs of senescence and in early passages (3–12 events) should be used for cryopreservation (Logan et al. unpublished; Bui-Marinos et al. 2022), because there is a lower risk of karyotype abnormalities or epigenetic alterations (Galli and Lazzari 2021). Cells to be cryopreserved should be harvested and washed by adding a salt solution (e.g., HBSS, PBS) and a dissociating solution (e.g., Gibco TrypLE, trypsin) (Strand et al. 2021). Prior to freezing, cells can be counted in a Neubauer chamber, if necessary (Cardoso et al. 2020b).

Note that most of the following cell culturing protocols developed for different taxonomic groups also include their corresponding cryopreservation protocols, and hence are not included in the cryopreservation chapter of this handbook.

In vitro cell cultures can be assessed by performing the same procedures applied after somatic cell cryopreservation recovery (see retrieval and viability chapter). Briefly, cell morphology and membrane integrity, mitochondrial dehydrogenase activity (MMT), population doubling time (PDT), and chromosomal quantification are analysed for culture characterisation (Praxedes et al. 2018).

Marine invertebrates. Fifty years ago, the development of cell cultures in marine invertebrates started, especially from sponges, cnidarians, molluscs, crustaceans, echinoderms, urochordates and cephalochordates. Until recently, permanent establishment of cell lines was not possible, (Rinkevich 2005; Cai and Zhang 2014; Conkling et al. 2019), due to cellular quiescence after 24–72 h of cell isolation (Rinkevich 2011; Potts et al. 2020). Currently, some successful cell lines are being established for sponges and cnidarians (Kawamura et al. 2021; Urban-Gedamke et al. 2021).

Sponges are used whole as tissue source, with the difference (to the procedure described above) that the sponge should be sieved to remove large aggregates of cells, right after cutting the specimen into little pieces (Conkling et al. 2019). Apical branch tips from corals, hemolymph from crustaceans and molluscs, and guts from echinoderms are mainly used as a source tissue (Cai and Zhang 2014). Although it is also possible to use other types of tissue, embryonic tissue seems to be the most promising for all phyla (except Porifera) to initiate cultures. Note that the developmental stage of the embryo will affect the effectiveness of cell culture approaches (Cai and Zhang 2014). The Cnidarian Cell Culture Consortium is currently establishing cell line protocols for the taxon (Roger et al. 2022). Cai and Zhang (2014) reviewed several protocols published until 2012 for all the phyla mentioned above, whereas Zahiri and Zahiri (2016), Rosner et al. (2021) and Ballarin et al. (2022) reviewed different studies regarding stem cells in marine invertebrates. Further details can be found in Mitsuhashi (1989).

Refer to Schippers (2013), Conkling et al. (2019) and Urban-Gedamke et al. (2021) for protocols in sponges, Barnay-Verdier et al. (2013), Ventura et al. (2018), Fricano et al. (2020) and Kawamura et al. (2021) for cnidarians; to Jayesh et al. (2012), Söderhäll (2013), and Potts et al. (2020) for crustaceans; to van der Merwe et al. (2010), Yoshino et al. (2013), Maselli et al. (2018), and Suzuki et al. (2021) for molluscs; and to Wang et al. (2020), and Rodríguez-Hernández (2020) for echinoderms.

Arthropods. Cell cultures have mainly been established to study biological control of insect pests and disease vectors (Mitsuhashi 1989). Hence, several cell lines, especially for lepidopteran species, are available (Lynn 2007a). Hemolymph from larvae or embryos can be used to initiate cultures (Guo and Weng 2020). First, eggs should be immersed in 70% alcohol and rinsed with sterile distilled water. The eggs are opened to release the embryo, which is separated from the yolk and macerated. Incubation occurs at 27 °C (Johnson et al. 2010). Specific tissues from larvae can also be used to establish cultures following Lynn (2007b). Protocols for ticks and other arthropod disease vectors (midges, sand flies) have been developed by Bell-Sakyi et al. (2019). For hymenopterans, see Hunter (2010), Goblirsch et al. (2013), Guo and Weng (2020), and Watanabe et al. (2021). Refer to Lynn (2002) for maintenance of insect cell cultures. Further culture conditions can be found in Drugmand et al. (2011) and Maramorosch and McIntosh (2017).

Other invertebrates. Protocols have been developed for the freshwater polyp Hydra (Li et al. 1963) and for the nematode Caenorhabditis (Strange et al. 2007; Zhang et al. 2011, Zhang and Kuhn 2005–2018). Refer to Mitsuhashi (1989) for protocols in land slugs and nematodes, and to Fuller-Espie et al. (2015) and Riedl et al. (2022) for earthworms. Some first attempts have also been made for tardigrades (Haldrup et al. 2015). Bayne (1998) provides recommendations and considerations for insects and worms.

Fish. Several fish cell lines have been reported (Nagpure et al. 2013). Generally, skin explants from the caudal fin are the main source for establishing cell cultures (Chenais et al. 2014), although gills, eye, heart, kidney, muscle, and liver tissues have also been used (Nagpure et al. 2013; Goswami et al. 2022). Note that the skin needs to be wiped thoroughly with sterile gauze before sampling to remove the build-up mucous, which can lead eventually to culture contamination (Wong et al. 2012). The choice of temperature for culture incubation will depend on the species, but usually it is 20–23 °C for cold-water fish, and 26–30 °C for warm-water fish (Bols et al. 2011).

Standard necropsy procedures and culture maintenance protocols can be found in NWFHS (2004) and Bols et al. (2011). Lakra et al. (2011) provide an overview regarding development, characterisation and storage of fish cell lines, as well as a list of cell lines that have been developed for different freshwater fishes.

Amphibians. Skin biopsies can be taken from the hind legs, or the posterior end of both dorsal and ventral sides, but other kinds of tissues (e.g., tongue, eye) can be used (Zimkus et al. 2018). Strand et al. (2022) evaluated and validated a protocol for different types of tissues, since most cell line protocols were developed more than 20 years ago. Bui-Marinos et al. (2022) optimised a step-by-step explant protocol for skin cell cultures that can be adapted to many frog species by adjusting the culture medium and solutions. A protocol for the establishment of cell cultures from the great crested newt has been developed by Strand et al. (2021). Recommendations for initiating, establishing, and maintaining frog cell lines can be found in Bui-Marinos et al. (2022).

Reptiles. Primary fibroblast cultures can be established from tail (5–10 mm) and toe clips (3 mm) or toe webbing, but also from blood, heart, spleen, kidney, and internal connective tissue (Ezaz et al. 2009; Logan et al. unpublished). Reptile cells can be maintained at a range of different temperatures, possibly because a reptile’s temperature adjusts to the environment (Fukuda et al. 2012; Moghanjoghi et al. 2018). A non-invasive protocol can be found in Ezaz et al. (2009) and can be applied to a wide range of reptile species. Logan et al. (unpublished) developed a universal protocol for terrestrial reptiles, whereas Moore et al. (1997) and Fukuda et al. (2012) developed protocols for marine turtles.

Birds. Blood feathers are the preferred source for initiating cell cultures, followed by blood (Kroglund et al. 2022). Blood feathers (i.e., developing feathers) from the wings or tail should be pulled out at the base and the pulp removed from the calami to establish fibroblast cultures (Houck et al. 2017). Feathers in early developmental stages are preferred because they contain larger amounts of pulp tissue than mature feathers (Cardoso et al. 2020). Refer to Sasaki et al. (1968), Romanov et al. (2009), Cardoso et al. (2020), and Kroglund et al. (2022) for protocols.

Mammals. Fibroblasts from skin and foetal fibroblasts are the most common cell types used in mammals (Galli and Lazzari 2021), although other tissue sources such as muscle, bone marrow, intestines, adipose tissue, and oral mucosa have been used (Praxedes et al. 2018; Calle and Ramírez 2022). Skin biopsies, mainly from the groin area or the abdomen, or ear notches (1–2 cm² for medium-size (e.g., agouti), 2–3 cm² for large animals) can be obtained, for instance, during veterinary exams or radio-collaring, or from hunters (Houck et al. 2017; Sano et al. 2022). Note that hair and fur are a source of contamination, so they should be removed before sampling by shaving and cleaning the skin with a gauze soaked in 70% alcohol. If shaving is not possible, the biopsy area should be washed with soap and water, dried, and subsequently cleaned with 70% alcohol for 15–20 s. Harsh disinfectants can harm cells and should be avoided (Wong et al. 2012).

Protocols, recommendations, and challenges to establish fibroblast cultures can be found in Ma et al. (2011), Siengdee et al. (2018), Praxedes et al. (2021), and Sano et al. (2022). Phelan (1998) developed protocols for human cell cultures that can be used for other mammals. Species-specific protocols have also been developed, among others, for agouti (Costa et al. 2020), rabbit (Gavin-Oagne et al. 2020), peccary (Borges et al. 2020), Iberian lynx (Leon-Quinto et al. 2009; 2011), puma (de Oliveira et al. 2021a), jaguar (de Oliveira et al. 2021b), buffalo (Selokar et al. 2018) and bats (Crameri et al. 2009; Yohe et al. 2019).

Soil samples and sediment cores can be stored in paper bags and sealed containers, respectively. Samples should preferably be kept in cold (4 °C), dark, and ideally, oxygen-free conditions to maintain the microbial community, constrain bacterial growth and decrease metabolic processes (Vandeputte et al. 2017; Capo et al. 2021; Duan and Bau 2021; Brasell et al. 2022). Depending on the type of sediment, samples should be kept for up to two (e.g., for lacustrine material) or six months to prevent loss of DNA (Brasell et al. 2022). High temperature-drying (60 °C) of soil for 12 h is also possible but microbial diversity and reliable sequencing precision may be affected (Duan and Bau 2021). Note that storage as well as sediment type (e.g., clay, organic) will impact the abundance of some organisms in the samples (Tennant et al. 2022).

Many environmental specimen banks (ESBs) or other biobanks store soil samples at -80 °C rather than in LN2 for long-term storage (Clasen et al. 2020; Duan and Bau 2021; Schroeder et al. 2021). However, it is recommended to snap freeze samples – including those preserved in e.g., DNA/RNA buffers or DESS – in LN2 and store them in the LN2 gas phase (Rüdel and Winsgärtner 2008; Pavlovska et al. 2021).

Soil samples should be placed in cryovials and frozen right after collection (Rüdel and Winsgärtner 2008; Duan and Bau 2021). The cooling chain should not be interrupted during transport (Rüdel and Winsgärtner 2008). Samples can undergo a cryomilling procedure, meaning they should be ground and homogenised under cryogenic conditions (-130 °C) using, e.g., an oscillating mill or a planetary ball mill in a laminar flow hood (Rüdel et al. 2008). It is critical that the samples do not warm up during the process. Hence, mills and vessels have to be cooled to -150 °C using LN2, following LN2 safety regulations (ISBER 2019). If the sampled material has a large grain size (<1 cm), stainless-steel mortar and pestle, either pre-cooled or immersed in LN2, should be used beforehand. This procedure can also be applied to filters (Rüdel and Winsgärtner 2008; Rain-Franco et al. 2021).

Sediment cores can be cryopreserved at -80 °C, but it has to be noted that subsampling of cores should be done before freezing to avoid compromising their stratigraphic integrity (Brasell et al. 2022). Sediment subsamples should be taken immediately after opening the core and stored at -20 °C or -80 °C to reduce sample degradation due to fungal and bacterial growth, and cross-contamination (Massana et al. 2015; Capo et al. 2021; Duan and Bau 2021; Tennant et al. 2022). The CAS-freezing technology has also been used to preserve seafloor sediment samples (Braun et al. 2016; Trembath-Reichert et al. 2017).

Water samples and filters used for water and airborne samples can be stored at -20 °C until further processing (Egeter et al. 2018; Johnson et al. 2019; Ovaskainen et al. 2020; Rhymes et al. 2021). Recommendations for preservation of freshwater samples are provided in Baricevic et al. (2022).

Long-term preservation of protists is mainly achieved by subculturing and only cyst forms are stored frozen (Altermatt et al. 2015). Yet, ciliate cysts have been successfully cryopreserved, using vitrification methods (Müller et al. 2008). In principle, cryopreservation using DMSO as cryoprotectant presumably works for all protist species, including the parasitic forms (Tedeschi and De Paoli 2011). However, low survival after thawing may still occur. Therefore, the following recommendations should be followed to increase survival rates (Altermatt et al. 2015):

• Thawing and survival rates should be experimentally determined for each taxon/strain and cryopreservation method (e.g., cryoprotectant concentration between 5–12.5%) before it becomes a routine method.
• Eight cryovials should be prepared per culture to increase survival rate.
• Controlled cooling-down should precede LN2 storage.
• Frozen samples should never be kept outside LN2 storage for any longer than 1 min.
• Frozen cultures should be regularly reinitialised to assess genetic variation.

Parasitic blood protozoa, however, can be frozen without cryoprotectants and without an accurate cooling rate control (De Paoli 2005). Protocols for parasitic forms can be found in Diamond (1995), Miyake et al. (2004), Murilla et al. (2016), and Jaskiewicz et al. (2020).

In addition to the mentioned generic protocol for many protists (Altermatt et al. 2015), Liu et al. (2019) established a cryopreservation protocol for marine ciliates with a broader application to other marine protozoa. Folgueira et al. (2018) also developed a protocol specifically for marine ciliate endoparasites. Free-living amoebae protocols can be found in Seo et al. (1992). Further cryopreservation and freeze-drying protocols for specific protozoa species can be found in Lee and Soldo (1992).

Regarding microalgae, methanol and DMSO are the preferred cryoprotectants (Day et al. 2005). Yet, many microalgae with complex morphology (e.g., euglenoids), filamentous algae, and large-cell-sized algae are recalcitrant to current cryopreservation procedures (Harding et al. 2004; Day and Harding 2008; Day et al. 2010; Tessarolli et al. 2017; Kapoore et al. 2019; Paredes et al. 2021a). The presence of rigid cell walls and vacuoles usually hampers the penetration of the cryoprotectant into the cells and complicates the freezing process (Day 2005; Paredes et al. 2021a). Pre-culture conditions, cryopreservation protocols and further recommendations can be found in Mori et al. (2002), Day (2007), Day and Harding (2008), Gäbler-Schwarz et al. (2013), Day et al. (2017), Paredes et al. (2021a), and Arguelles et al. (2020). Furthermore, Day and Brand (2005) provided several detailed cryopreservation and thawing protocols that have been applied to a broad range of microalgae. Harding et al. (2008) also developed an encapsulation/dehydration method for microalgae. Cryopreservation protocols for marine microalgae have been established by Rhodes et al. (2006) and by the ASSEMBLE+ project (Paredes et al. 2020a). Step-by-step cryopreservation protocols for Euglena and related genera were developed by Harding et al. (2010) and Shah et al. (2022). Customised protocols for chlorophytes are provided in Fernandes et al. (2019) and for marine dinoflagellates in Kihika et al. (2022). A list of successful cryopreservation approaches in various microalgae species can be found in Nugroho et al. (2016) and in Paredes et al. 2020a).

Phenotypic characterisation of microalgae using a miniaturised growth measurement should also be performed before and after cryopreservation to determine the success of the procedure (Day et al. 2005).

Cryopreservation is considered the best preservation procedure for fungi and fungus-like forms (e.g., Mycetozoa, Oomycota). Yet, changes both in hyphae morphology and viability during freezing and thawing have been recorded when storing cryopreserved strains at merely -80 °C (Ryan et al. 2000). Both sporulating and nonsporulating cultures, as well as those that cannot be lyophilised or have delicate spores, can be kept at ultra-low temperatures (Nakasone et al. 2004; Agarwal and Sharma 2006).

Spores, mycelia cultures, and air-dried conidia can be cryopreserved, using cryoprotectants such as glycerol, trehalose, or DMSO to reduce injuries during the cryoprocess, although the latter is often toxic for sensitive organisms (Hubálek 2003; Ryan and Smith 2004; Homolka 2013; Singh and Baghela 2017).

Some cultures can be frozen directly after attaining suitable growth (Abd-Elsalam et al. 2010). Otherwise, mycelia agar should be cut from the margin of the colony with a sterile cork-borer or a sterilised plastic straw. The excised disks (ca. 8–10 disks with 4 mm diameter each) should be placed in cryovials containing 10% glycerol. This method applies to mycelia that grow deep into the agar or fungi that do not sporulate. Five tubes should be prepared in this way from each iso­late (Vasas et al. 1998; Nakasone et al. 2004; Overy et al. 2019). Polypropylene vials must be placed directly into the vapour phase of LN2 (Nakasone et al. 2004).

Mycelia growing in liquid culture should macerated and fragmented in a miniblender prior to pipetting. An equal part of 20% glycerol should be added to the vial (Nakasone et al. 2004). Mycelia growing on the agar´s surface can be gently scratched with a pipette and placed in a cryovial. Spore suspensions should ideally be placed in polypropylene straw ampoules together with 10% glycerol. Both mycelia and spores should be precooled at 7–4 °C, then prefrozen at -40 °C, at a rate of 1 °C/min, followed by a rate of 10 °C/min until reaching -90 °C. Ampoules and vials should be immediately submerged into LN2 (Juarros et al. 1993; Nakasone et al. 2004; Agarwal and Sharma 2006).

Most cultures are stored in mechanical freezers at -80 °C. Fungi and fungus-like forms growing on different organic substrata (e.g., cereal grains, agar strips, filter paper), and that do not sporulate excessively, should be dried before freezing (Nakasone et al. 2004; Abd-Elsalam et al. 2010). Deep-freezing at -80 °C is not recommended for ectomycorrhizal fungi (Heinonen-Tanski and Holopainen 1991). Cryopreservation protocols for oomycete cultures can be found in Kitamoto et al. (2002), Uzuhashi et al. (2020), and Eszterbauer et al. (2020).

Preservation on porous beads. This was originally developed for storing bacteria in the LN2 vapour-phase (Homolka 2014). Beads (e.g., ceramic or glass beads) work as carriers and have been used for nonspore-forming fungi (Lakshman et al. 2018), sporulating fungi and yeasts (Homolka 2013). Beads should be taken with sterile forceps out of their vial and placed in a Petri dish containing the culture for up to seven days, or until the beads have been covered with mycelia. Beads should be returned to the original vial after removal of the contained preservative solution. Preservation temperatures can be either -80 °C or -150 °C (Homolka 2013). Commercial Microbank beads should never be immersed in LN2 (Lakshman et al. 2018). Cryopreservation of air-dried or suspended conidia can also be carried out using porous or polystyrene beads as carriers (Chandler 1994).

Preservation on perlite. Perlite is a unique aluminosilicate volcanic mineral that can retain water and release it when needed (Homolka 2013). Perlite has been suggested as a favourable substitute for serial transference used with agar cultures in long-preservation of fungi (Simões 2013). This method has been successfully applied to yeasts, Ascomycota, Zygomycota, and some Basidiomycota strains that cannot survive other types of preservation (Homolka 2013). Perlite is used as a carrier of mycelia and can be added directly to the cryovials. For further details, refer to Homolka et al. (2007). The protocol is described in Homolka (2013).

Recalcitrant species such as the water mould Saprolegnia spp. and unculturable fungi such as microcylic rusts can be cryopreserved by vitrification and immobilisation or encapsulation (Ryan 2001; Eszterbauer et al. 2020). The vitrification technique has not been broadly tested on fungi, so further investigation is needed (Homolka 2014; Singh and Baghela 2017). Immobilisation is a technique involving alginate encapsulation of mycelium/spores prior to preservation (Ryan and Smith 2004; Ryan et al. 2019). Simões (2013) has used this method for preservation of recalcitrant fungal strains and several filamentous fungal species maintained in 10% glycerol. The combination of encapsulation and vitrification may be appropriate for mutualistic associations such as orchidaceous mycorrhizal fungi (Kermode et al. 2012), endophytes and lichens (Ryan and Smith 2004), as well as for obligate pathogens, which must be cryopreserved together with their hosts (Homolka 2014; Singh and Baghela 2017; Strittmatter et al. 2020). Plants used as hosts should go under a quarantine period prior to inoculation with infected plant material (Ryan and Ellison 2003).

Ryan and Ellison (2003) developed a protocol for cryopreservation of microcylic rusts. However, their pathogenicity/infection ability was not successful after retrieval, and spores did not survive after direct plunge freezing. The authors recommended the use of stem or petiole tissue, as leaf material may have moisture issues after cryopreservation.

Some institutions have specialised in specific types of fungi and have developed their own preservation protocols. For instance, the West Virginia University has developed methods for spore extraction, hyphal harvesting, culturing, voucher preservation and storage of vesicular arbuscular mycorrhizal fungi. The methods include conservation of in vivo pot cultures in sterile soils or other supporting materials, in vitro cultures with genetically modified root-organ culture of host plants, and in vitro autotrophic systems in artificial media with axenic plants (Drazhnikova and Andrianova 2020).

Further cryopreservation protocols for fungi can be found in Westerdijk fungal biodiversity Institute. Protocols for yeasts and filamentous fungi can be found in Bond (2007) and Ryan and Smith (2007), respectively.

Information regarding macrofungi can be found in Linde et al. (2018) and Ilyas and Soeka (2019). Mata and Pérez-Merlo (2003) and Mata and Savoie (2013) established protocols for preservation of macrofungal mycelia on cereal grains without using cryoprotectants. Furthermore, Sato et al. (2020) assessed a novel cryopreservation protocol that uses vermiculite grains – a soil additive – to improve the viability of ectomycorrhizal basidiomycetes (e.g., Amanita spp.) that are difficult to cryopreserve.

So far, cryopreservation is the only available method providing long-term conservation for plant genetic resources that can be propagated only vegetatively, and for recalcitrant and rare species (Engelmann 2009; Pence et al. 2020; Funnekoter et al. 2021). The physiology and ecological characteristics of each species must be considered to identify the most optimal cryopreservation technique (Sarasan et al. 2006; Sarasan 2010; Engelmann 2010; Pence et al. 2020). Various strategies have been developed to adapt cryopreservation protocols efficiently (Kim et al. 2012; Funnekotter et al. 2021):

Slow cooling procedures. Freezing temperatures are usually applied to cell suspensions, calluses, and dormant buds, as well as to cold-tolerant species. This process involves slow cooling to about -40 °C, followed by a rapid immersion in LN2. A protocol designed for plant cell suspensions can be found in Grout (2007). Jenderek et al. (2010) gives an overview on dormant bud cryopreservation.

Vitrification technique. This approach uses a highly concentrated solution of different cryoprotectants such as DMSO, glycerol, sucrose, sorbitol and/or ethylene glycol, which allows cells to dehydrate and to vitrify during freezing. Cryoprotectants can be used individually or in combination. The most commonly used plant cryoprotectant is PVS2, a combination of DMSO, glycerol, sucrose and ethylene glycol (protocol) (Panis and Lambardi 2005; Funnekotter et al. 2021). Other cryoprotectants are PVS3 with 50% sucrose and 50% glycerol (Nishizawa et al. 2008; Senula and Nagel 2021) or PVS1, PVS4 or Vitrification Solution L (VSL) (Zamecnik et al. 2021). Instead of using pure DMSO (Schäfer-Menuhr et al. 1996), potentially toxic for the cells, propylene glycol can be used as a replacement. Vitrification methods can be highly reproducible and can be applied to different types of tissue and to a broad range of species (Panis and Lambardi 2005). It has been noted that plant propagules may age during LN2 storage, potentially due to the inherently short lifespans of certain species and due to the fact that cell architecture may change after vitrification processes (Ballesteros et al. 2021). Further research on this topic is needed.

Droplet vitrification method. Tissues (e.g., apices) are treated with droplets of a vitrification solution on aluminium foil strips that are rapidly frozen in LN2. This method is the most widely used as it can be applied to a wide range of species (and genotypes) including woody and herbaceous species (Wang et al. 2021). A step-by-step guide to developing new protocols can be found in Panis et al. (2011).

Cryo-plate techniques. Calcium alginate capsules containing tissue (e.g., shoot tips) are secured on aluminium cryo-plates. The vitrification cryo-plate (V Cryo-plate) method is a combination of encapsulation-vitrification and droplet vitrification. The dehydration cryo-plate (D Cryo-plate) combines the encapsulation with the dehydration method, and usually shoot tips are air-dried. Handling becomes easier, as the cryo-plates are manipulated instead of the plant material. Both methods are promising for herbaceous and woody plant preservation after modifications of the original protocols (Yamamoto, et al. 2011; Matsumoto 2017).

Cryo-mesh technique. It is comparable to the cryo-plate techniques, but the main difference is that a stainless-steel mesh strip is used instead of a cryo-plate (Wang et al. 2021).

Desiccation technique. The plant material, mainly zygotic embryos and embryonic axes, is dehydrated using a stream of compressed air or under the laminar flow of a lab bench on silica gel and, then immersed in LN2 (Sakai et al. 2000).

Encapsulation–dehydration technique. This method is comparable to the synthetic seed technology, as the plant material (e.g., shoot tips) is also encapsulated in alginate beads. Beads are then dehydrated, either by air-drying or using silica gel, and then immersed in LN2. Some of the advantages of this technique include easier tissue manipulation, protection during dehydration, and no need for cryoprotectants (Niino and Sakai 1992).

Encapsulation-vitrification technique. Tissue samples are encapsulated in alginate beads and then submitted to freezing by vitrification. A protocol for shoot tips and meristems can be found in Benson et al. (2007).

The use of other tissue samples such as small leaf square-bearing adventitious buds (SLS-BABs), stem disc-bearing adventitious buds (SD-BABs), rhizome buds and microtubers can simplify cryopreservation protocols, as the most time- and labour-consuming step –the shoot tip excision– can be excluded (Wang et al. 2021). However, dormancy, viability and recovery of the tissue are factors to consider in decision making.

Several protocols for cryopreservation can be found in:

• Specific journals such as Acta Horticulturae, Journal of Plant Physiology, Plant Cell, Tissue, and Organ Culture (PCTOC), Frontiers in Plant Science, Methods in Molecular Biology, Plant Science, Propagation of Ornamental Plants, Physiologia Plantarum, and Journal of Horticultural Science and Biotechnology.
• Bettoni et al. (2021) provide information regarding facilities, technical skills, and plant material conditions to implement plant shoot tip cryopreservation.
• The Laboratory of Tropical Crop Improvement at KU Leuven defined cryopreservation protocols for 26 crop species such as banana, taro, and common chicory.
• Wang et al. (2021) provided a comparison of new technologies for in vitro based cryopreservation, as well as a reference list for several cryopreservation protocols.
• Cruz-Cruz et al. (2013) provided an extensive reference list for protocols in different taxa.
• Thammasiri et al. (2019) compiled cryopreservation protocols for crops.
• Reed (2008) provided step-by-step instructions for developing new cryopreservation protocols based on well-established and successful protocols. It includes protocols for the cryopreservation of both orthodox and recalcitrant seeds, pollen, dormant buds, bryophytes, ferns, de-differentiated cell cultures, embryogenic cultures, embryonic axes, crops, herbaceous plants, and trees.
• Engelmann and Takagi (2000) included papers from a workshop, describing different protocols for cryopreservation of tropical plants.
• Normah et al. (2019) suggested strategies for cryopreservation of shoot tips of recalcitrant and tropical species.
• Popova et al. (2015) reviewed different cryopreservation protocols for diverse plant species. They also provided strategies for developing protocols de novo. See also Popova et al. (2016, 2019, 2020).
• Engelmann (2011) provided an overview on embryo cryopreservation and on in vitro derived propagules (Wang et al. 2021).
• A vast list of species that have been cryopreserved, using seeds, cell suspensions, calluses, embryonic axes, somatic embryos, and shoot-tips can be found in Gonzalez-Arnao et al. (2014).
• A recent compilation of cryopreservation protocols for different organisms and tissues, including plants, can be found in Wolkers and Oldenhof (2021).
• Roque-Borda et al. (2021) reviewed some cryopreservation methods applied to agronomic plant germplasm.
• A special issue on “ plant cryopreservation ” in the journal Plants (2021:10) includes new protocols, physical and chemical aspects of freezing and drying, cold adaptation and thawing.

The following table offers a list of protocols developed for different plant species:

Bryophyte spores can be dried to low RHs (<50%) and stored dry at low (sub-zero) temperatures. Survival to LN2 exposure after drying to 15%RH has recently been reported, opening the doors to bryophyte spore banking (Tiloca et al. 2022). Bryophyte spores often contain chloroplasts and storage lipids, potentially affecting sample longevity (Ballesteros et al. 2020), thus monitoring is recommended. Bryophytes also produce gemmae, undifferentiated vegetative propagules, that can be collected and placed in small paper envelopes. Envelopes should be placed in a box containing silica gel for ca. 3 h (time may vary depending on the species) for drying. Envelopes containing dried gemmae should be placed in cryovials and directly immersed in LN2 (North et al. 2021).

Fern spores should be dried in environments between 15–75% RH and stored at either -20 °C (Ballesteros et al. 2019), -80 °C or at -196 °C to maintain their long-term (> 10 years) viability (Nebot et al. 2021). However, storage conditions must be carefully defined for each species as some species may age faster at -20 °C when compared to higher (e.g., 5 °C) or lower (-80 °C, -196 °C) temperatures potentially due to lipid crystallisation issues (Ballesteros et al. 2019). For short- (1–3 years) and medium- (up to 10 years) term storage, refrigeration (3–7 °C) can be used after drying at a RH between 15% and 25% (Ballesteros et al. 2019; North et al. 2021). Storage at room temperature or wet storage is not recommended (Ballesteros and Pence 2018). Spores may deteriorate and die, particularly chlorophyllous/green spores, during LN2 storage due to their inherently short lifespan (Ballesteros and Pence 2017; Ballesteros et al. 2019). Longevity will vary depending both on the temperature and the type of spore, with species having chlorophyllous/green spores ageing faster than non-chlorophyllous/non-green spores (Ballesteros and Pence 2018; Ballesteros et al. 2019).

In addition to the spores, bryophyte and fern gametophytes and sporophytes can be cryopreserved for long-term conservation purposes (Ballesteros and Pence 2018). These tissues are generally cryopreserved following the encapsulation dehydration method. This procedure comprises three steps: 1) the encapsulation of tissue in spheres of alginate gel, 2) cryoprotection using a combination of osmotic dehydration with concentrated sucrose and drying, and 3) rapid immersion in LN2. For ferns, whole gametophytes can be used, but if too large, they should be cut into small pieces prior to cryopreservation. The use of abscisic acid during tissue culture likely improves survival during LN2 storage (Ballesteros and Pence 2018). Furthermore, actively sporophyte-growing meristematic tissue, such as shoot tips or green globular bodies survives better than mature non-growing tissues (Ibars and Estrelles 2012). This method has been mainly applied to leptosporangiate ferns, while further research is needed for eusporangiate and lycophyte genera (Ballesteros and Pence 2018). Refer to Mikula et al. (2011) and Nebot et al. (2021) for gametophyte and spore protocols respectively. Further information and protocols can be found in Pence (2008). For bryophytes, techniques similar to those employed for ferns can be used. Rowntree et al. (2005; 2007; 2009; 2011) developed several LN2 cryopreservation protocols for UK bryophyte species. Cold-acclimated bryophyte specimens have also been successfully cryopreserved without the use of cryoprotectants; the protocol can be found in Segreto et al. (2010).

In general, mature orthodox pollen does not need additional treatments before cryopreservation (Ganeshan et al. 2008; Funnekotter et al. 2021). Protocols for pollen cryopreservation both for desiccation-tolerant pollen and for a desiccation-sensitive pollen species (e.g., corn) can be found in Nebot et al. (2021). Protocols for different commercially relevant species can be found in Ganeshan et al. (2008) and for tropical plant species in Rajasekharan et al. (2013). Further cryopreservation protocols are described in Popova et al. (2015).

Recalcitrant pollen should be quickly and partially desiccated to moisture levels at which no freezable water exists but above levels where desiccation injury is apparent (Towill and Walters 2000; Anushma et al. 2018). Current desiccation techniques can be found in Nebot et al. (2021) and Impe et al. (2022).

Orthodox seeds. In general, orthodox seeds have a low moisture content and can be stored in LN2 without cryoprotectants. Seeds should be dried at 25–32% RH and then, they can be placed into cryovials and immersed in LN2. If seeds have a large size, they can be placed in laminated foil packets (Funnekotter et al. 2021).

Orthodox seeds with short-life spans. Some orthodox seeds have a very short-life span (i.e., Populus deltoides or Salix spp.) and tend to deteriorate and die within a few years (Pammenter and Berjak 1999). Although low temperature storage under low moisture content is possible, cryopreservation may increase longevity of these seeds. However, it has to be noted that drying requirements for LN2 storage will be different from those for conventional storage. Seeds are mostly dried at 32% RH and 18 °C or 5 °C before cryostorage (Ballesteros et al. 2021). The Cincinnati Zoo and Botanical Garden suggests a list of publications and videos on how to evaluate the best storage for short-lived seeds. Also note that seeds may continue ageing and degrading while maintained in LN2 (Walters et al. 2004; Ballesteros and Pence 2017; Davies et al. 2018). Walters et al. (2004) concluded that cryopreservation does not maintain cells in an infinite longevity state, as it has been established that molecules continue moving at temperatures below -130 °C, allowing ageing to advance (Ballesteros and Walters 2019; Ballesteros et al. 2020). To mitigate this, mature seeds must be freshly harvested, should have a high initial quality, and require careful handling (Ballesteros and Pence 2017). Ballesteros et al. (2021) provided cryopreservation methods for orthodox and intermediate seeds, as did Pritchard (2007) and Pritchard and Nadarajan (2008).

Intermediate seeds. To avoid drying damage, intermediate seeds are often dried to higher moisture contents than those for conventional storage. Drying of intermediate seeds can be done at 20-25 °C and 50-75% RH before cryopreservation storage (Ballesteros et al. 2021; Funnekotter et al. 2021).

Recalcitrant seeds. This type of seeds usually has a large size. Hence, the embryonic axes or the embryo can be excised and cryopreserved (Funnekotter et al. 2021). This procedure should take place right after collection in a laminar flow hood under sterile conditions. Embryonic axes need to be processed within two hours after removal. Embryos should be flash-dried using silica gel or saturated salt solutions in a drier for ca. 2-4 hours of rapid drying (Berjak et al. 1993), then to be placed into cryovials and stored in the vapour phase of LN2 (Ballesteros et al. 2021). It is crucial that embryos are at the right developmental stage for a successful cryopreservation (Theilade and Petri 2003). Refer to Ballesteros et al. (2021) for detailed guidance on the excision of embryonic axes method. Recalcitrant seeds can also be germinated in vitro and shoot apical meristems from the seedlings can be used for cryopreservation as an alternative when embryo excision is not possible (FAO 2014). Further protocols for working with recalcitrant seeds can be found in Walters et al. (2008) or Ballesteros et al. (2021). Cryopreservation is thus recommended for long-term storage.

Excision of embryos from orthodox and intermediate seeds is also possible. However, prior seed desiccation should be avoided (Funnekotter et al. 2021).

Marine invertebrates. Lack of knowledge about reproduction biology and physiology of many marine organisms, as well as the short-term availability of gametes are also big challenges for developing cryopreservation protocols (Paredes 2018; Hagedorn et al. 2019). Paredes et al. (2018, see also cited references) provide a summary of protocols applied to different marine invertebrate organisms (molluscs, echinoderms, cnidaria, sandworms, barnacles), whereas Paredes (2015) focused mainly on protocols for sea urchin and bivalve larvae. Moreover, the CRYOMAR protocol toolbox (Paredes 2020a) provides protocols for larvae and sperm of mussels and sea urchins, as well as sperm from sea cucumbers and oysters.

A review on cryopreservation protocols in crustaceans and other marine invertebrates is provided in Guo and Weng (2020, see also supplementary material) and Aquino et al. (2022). Sperm protocols are described in Morales-Ueno and Paniagua-Chávez (2020) for crustaceans, in Demoy-Schneider et al. (2020) for bivalves, and in Hagedorn et al. (2019) for corals. Further research on the cryopreservation of gametes and larvae of echinoids, ascidians and polychaetes is currently being carried out by Paredes (2020b) and Paredes et al. (2021b). Toh et al. (2022) provide a somatic cell cryopreservation protocol for corals.

Campos et al. (2021) reviewed oocyte cryopreservation challenges in aquatic invertebrates. Coral larvae, as well as zebra fish larvae, have been successfully cryopreserved using vitrification and laser warming (Khosla et al. 2017; Daly et al. 2018).

Helminths. Many helminth species can only be cryopreserved using vitrification methods. Eggs, larvae and microfilariae are frequently cryopreserved applying a protocol that includes the addition of ethylene glycol in two steps, followed by rapid cooling to -196 °C (James 2004). Cryopreservation protocols for parasitic worms can be found in James (1985, 1990, 2004) and Eckert (1988). Other protocols have been developed for specific genera such as Heterorhabditis (Nugent et al. 1996), Schistosomula (James 1982), Dirofilaria (Shirozu et al. 2020; Zinser et al. 2021), and Pristionchus plus other free-living nematodes (Matthias Herrmann, pers. comm.). Elsen et al. (2007) focused on tropical and some other plant-parasitic nematodes, whereas Beraldo and Pascotto (2014) focused on animal-parasitic nematodes.

Insects. Cryopreservation techniques have been applied to the house fly, ladybird beetles, spined soldier bugs, fireflies, fruit flies, silkworms, honeybees, and eventually to mosquito species known to act as vectors for human pathogens (Gallichotte et al. 2021; Viert et al. 2021). Note that a developmental stage that is permeable to water and cryoprotectants has to be used for cryopreservation. Hence, embryos and eggs with chorion, wax layers or vitelline membranes are not recommended (Gallichotte et al. 2021). Ethylene glycol is generally used as a cryoprotectant in insects, and often trehalose or polyethylene glycol is added to avoid damage during the vitrification process (Gallichotte et al. 2021).

A robust and easy to implement protocol for fruit fly embryos can be found in Zhan et al. (2021). A protocol for honeybee semen can be found in Auth and Hopkins (2021), whereas Whitman et al. (2019) described a freezing preservation (-80 °C) protocol useful for molecular studies of terrestrial arthropods.

Fish. Many fish sperm cryopreservation protocols have been developed, mostly for marine species (Rawson et al. 2011; García et al. 2016; Martínez-Páramo et al. 2016). Sometimes it may be necessary to pool samples from different individuals to obtain the required volume (Martínez-Páramo et al. 2016). Generally, 10–15% DMSO is effective for fish, but glycerol, trehalose, and propylene glycol have proven more effective in some species (Browne et al. 2019).

Cryopreservation protocols for sperm and germ stem cells, as well as the current state of cryopreservation of oocyte and embryonic cells can be found in Betsy and Kumar (2020), Routray (2020), and Cabrita et al. (2022). Additional protocols are provided in Kopeika et al. (2007) and Rawson et al. (2011). Dhanasekar et al. (2022) developed a protocol for cobia sperm, and Yang et al. (2022) for brown-marbled grouper sperm. Protocols specifically designed for shark and ray sperm can be found in García-Salinas et al. (2021). Refer to Šćekić et al. (2019; 2020) for cryopreservation protocols of eel gonadal tissue and ovarian stem cells, and to Tiersch (2001) for sperm cryopreservation of aquarium fishes. A summary of germ cell cryopreservation in different teleost species is provided in Robles et al. (2017). See references cited in Martínez-Páramo et al. (2016) for further protocols.

The AQUAEXCEL project developed a protocol booklet for different fish species (Labbé 2018), as well as a validation procedure for the isolation, cryopreservation and transplantation of trout and carp germ stem cells (AQUAEXCEL 2020). In addition, the AQUAGAMETE project included the development and standardisation of techniques for cryopreservation of fish sperm.

Amphibians. Spermatozoa from anurans (as from fishes) can be kept at 4 °C for days to weeks without losing viability (Browne et al. 2019). When cryopreserved, 5–10% DMSO or DMFA (dimethyl formamide) should be used as cryoprotectants, but sucrose and trehalose have also proven to be successful (Browne et al. 2019). Rawson et al. (2011), Poo and Hinkson (2019), and Burger et al. (2022) have developed sperm cryopreservation protocols that can be applied to different threatened anurans. Further details regarding cryopreservation of maternal-haploid and embryonic-diploid genomes can be found in Clulow et al. (2022). Strand et al. (2020) also reviewed the topic and produced a summary of protocols and references.

Reliable methods for urodeles have not been established yet (Hagedorn et al. 2019). However, McGinnity et al. (2022) have provided the first steps for the storage and cryopreservation of semen from the North American giant salamander and Guy et al. (2020) for newt species. No storage or recovery protocols exist for caecilians (Clulow et al. 2022a), but some reproduction technologies are under development (Browne et al. 2022). Cryopreservation protocols of somatic cells can be found in Bui-Marinos et al. (2022).

Reptiles. Successful protocols for cryopreserving reptile spermatozoa are scarce (Campbell et al. 2020; Strand et al. 2020), and so far, no reports of offspring production using cryopreserved sperm have been published, most likely due to sperm degradation (i.e., low motility recovery) following cryopreservation (Clulow and Clulow 2016).

A cryopreservation protocol for spermatozoa of squamate reptiles (snakes, lizards and amphisbaenians) has been optimised by adding caffeine to increase motility of sperm after thawing (Campbell et al. 2020; Sandfoss et al. 2022). Young et al. (2017) developed one of the first sperm cryopreservation protocols in lizards. Young et al. (2022a) also compared cryoprotectants and possible combinations to optimise sperm cryopreservation protocols in lizards. Hobbs et al. (2022) assessed different extenders and freezing rates to enhance a sperm cryopreservation protocol for the Australian skink. Young et al. (2021, 2022b) and Blank et al. (2022) developed different protocols for snakes. Furthermore, Clulow at al. (2022) provided a review regarding the latest cryopreservation studies both for reptiles and amphibians.

So far, the only cryopreservation protocol among crocodilians is for spermatozoa of the saltwater crocodile (Johnston et al. 2017). Lamar et al. (2021) published the first collection, characterisation, and storage protocol of tuatara semen. Sirinarumitr et al. (2010) determined the efficiency of certain extenders in olive ridley turtles and hawksbill turtles. Ravida et al. (2017) developed a protocol for desert tortoise sperm, which may also be applicable to other members of the Testudinidae family.

Birds. Sperm cryopreservation is more challenging in birds than in any other vertebrate, since the filiform shape of the spermatozoa makes them more vulnerable to injury from manipulation procedures (Gee et al. 2004; Çiftci and Aygün 2018; Cardoso et al. 2020a). Furthermore, avian sperm is very susceptible to the osmolarity changes that occur during the freezing-thawing process (Cardoso et al. 2020a; Castillo et al. 2021). Some strategies, such as the addition of zinc oxide, lipids, or sialic acid have been included to maintain sperm viability after cryopreservation (Zhandi et al. 2019; Castillo et al. 2021). Further details can be found in Wishart (2007) and Woelders (2021).

Most protocols have focused on poultry semen cryopreservation (Iaffaldano et al. 2016; Thélie et al. (2019; Tkachev et al. 2020; Lin et al. 2022). However, sperm cryopreservation protocols have also been developed and optimised, e.g., for the common parakeet (Dogliero et al. 2017), the peregrine falcon (Cardoso et al. 2020a), and the common pheasant (Castillo et al. 2021).

Gonadal tissues have also been cryopreserved when semen collection is not possible and as a viable alternative to preserving the female genome, with the aim of obtaining poultry offspring, using the allotransplantation technique (Liptoi et al. 2020; Fujihara et al. 2022). Hu et al. (2022) have also developed a cryopreservation protocol for poultry embryonic gonads. Somatic feather follicle cell cultures can be cryopreserved following Cardoso et al. (2020b).

Mammals. No standard procedure exists for the cryopreservation procedure of domestic mammal gametes, except for bulls (Walters et al. 2009; Yánez-Ortiz et al. 2021; Layek et al. 2022). Still, these protocols have been adapted to cryopreserve sperm from wild animals, such as elephants and capuchin monkeys, with some success. Wild species usually require de novo methodologies, since the quality of sperm is lower and less uniform, compared to the sperm from lab and livestock animals (Durrant et al. 2019). Cryopreservation of oocytes and embryos –especially blastocysts– may be possible, but it is still limited due to the low availability of wild animals as gamete/gonad donors (FAO 2012; Bhat and Sofi, (2021). Ovarian tissue can be a good source of oocytes, as it can overcome oocyte cryodamage and super-ovulation issues (Bhat and Sofi 2021). Testicular tissue has been widely cryopreserved, and its perspectives and current advances are described in Silva et al. (2020). Naitana et al. (2015) reviewed the main advantages and disadvantages caused by cryopreservation of sperm, oocytes, and embryos from domestic animals.

Note that in mammals, cryopreservation protocols can vary among species and breeds, and inter-individual differences can also occur, probably due to age or inbreeding (Holt 2000; Toledano-Díaz et al. 2020). Yánez-Ortiz et al. (2021) provide a summary about sperm cryopreservation in farm animals, as well as a detailed explanation regarding structural and molecular sperm alterations caused by the procedure. Some sperm cryopreservation protocols for domestic animals can be found in Hiemstra (2003), Walters et al. (2009), Sieme and Oldenhof (2015) and Galarza (2019). Livestock cryopreservation protocols have also been developed under the support of different projects such as EuReCa (2010), Globaldiv (Ajmone-Marsan and Globaldiv consortium 2010), and IMAGE. The Jackson Laboratory provides a step-by-step cryopreservation protocol in mice. O´Brien et al. (2019) investigated the effect of two freezing protocols on sperm from several endangered wild species. A special issue on “New Challenges in Cryopreservation” in the journal Animals (2022:12) includes recent developments in the use of cryoprotectants, protocols and strategies for cryopreserving mainly mammal germplasm. A summary of cryopreservation methods for ovarian follicles from different species can be found in Campos et al. (2019).

Embryos to be cryopreserved should follow the standard methods approved by the International Embryo Technology Society (Stringfellow et al. 2010). Saragusty and Arav (2010) reviewed the progress in oocyte and embryo cryopreservation by slow freezing and vitrification and summarised the protocols applied in different mammalian species. Furthermore, Bhat and Sofi (2021; and references therein) reviewed the current state of ovarian tissue vitrification in several wild animal species, whereas Tharasanit and Thuwanut (2021) did so for domestic animals. Keros and Fuller (2015) developed a cryopreservation protocol for mammalian oocytes, whereas Fuller and Paynter (2007) developed one for mammalian embryos.

Somatic cells (e.g., from ear tissue) can also be cryopreserved for future use (Woelders et al. 2012). Workflows describing the set-up of somatic-cell banks are found in Groeneveld et al. (2008) and Sekolar et al. (2018). Protocols are usually more generic across mammalian species, and some are referred to in de Lira et al. (2021). A well-detailed protocol can be found in Siengdee et al. (2018).

A list of mammal cryopreservation methods by taxa from 1970 to 2016 can be found in Charlton et al. (2018). The following list mentions some cryopreservation protocols published from 2017 onwards:

Vials containing spores should be warmed at room temperature for 20–40 min. Spores are considered viable when the outer wall of the spore has ruptured and the rhizoid or the first chlorophyllic cell has emerged. A second assessment to calculate the laminar development percentage can take place during the gametophyte transition from one-dimensional filamentous growth to two-dimensional laminar growth.

Spore germination and germination speed can be initially checked daily and then every three days after the 10^th day, for a total of 30 days (Ballesteros et al. 2006). The easiest way to assess viability of fern spores is by germinating 100 spores, randomly selected, for a determined number of days (e.g., 10 days for green spores and 25 days for non-green spores) to obtain the germination percentage (Ballesteros et al. 2006; Ballesteros and Pence 2018). Note that the germination media may need to be optimised for each species and can affect the viability assessment results (Pence 2008). Other indirect biochemical assessments are also available and can be revised in Galán and Prada (2011).

For production and acclimatisation of gametophytes and sporophytes in soil, follow Ballesteros and Pence (2018) and Ibars and Estrelles (2015).

Prior to testing, pollen should be slowly rehydrated to avoid damage from excessively rapid water intake (Towill and Walters 2000). Pollen viability highly differs within and among crop species, hence tests should be standardised for each species/variety (Sidhu 2019). Note that pollen can be viable by being alive and metabolising, but this does not necessarily mean that it can still germinate or fertilise an egg (Towill and Walters 2000). The best choice is to run a combination of viability tests for indications of the pollen´s ability to germinate and grow (Impe et al. 2020).

Pollen stainability includes the use of Triphenyl Tetrazolium Chloride (TTC or TZ) and FDA. Pollen viability percentage is determined by dividing the number of viable pollen grains by the total number of pollen grains. Usually, the quantification of stained/non-stained pollen is done manually, but this task is laborious and time-consuming (Tello et al. 2018). Novel automated approaches have been conceived to assist in this task and they can be applied to different taxa. Ascari et al. (2020) established a procedure based on fluorescence microscopy to assess pollen grain automated images, whereas Tello et al. (2018) developed PollenCounter, an open-source tool for the high-throughput phenotyping of pollen viability. The open-source software ImageJ (US National Institutes of Health) is also used for calculating pollen germination frequency (Smith and Wallace 2020).

In vitro germination comprises pollen tube growth measurements and it requires taxon-specific liquid or solid media (Impe et al. 2020). Pollen grains are considered viable when tube lengths are longer (>1.5×) than the grain diameter (Costa et al. 2020; Anushma et al. 2018). In vitro germination assessment can be performed using the hanging drop culture, the sitting drop culture, the suspension culture, or the surface culture method, as described in Gowthami et al. (2019). Refer to Costa et al. (2020), and Impe et al. (2020) for detailed protocols, and to Tushabe and Rosbakh (2021) for a compilation of pollen germination media.

In vivo pollen germination and seed/fruit set percentage methods are other alternatives to assessing pollen viability, although they may be more time-consuming (Gowthami et al. 2019).

Viability tends to decline with time, regardless of the storage conditions, and therefore viability assessments should be carried out periodically (FAO 2014). Furthermore, seeds of different species and even seed lots of the same species may vary in their longevities under standard conditions (Probert et al. 2009; Colville and Pritchard 2019). Note that seeds can be considered viable by germinating but they will not necessarily develop into a viable seedling (Filip Vandelook, pers. comm.).

Ideally, a germination viability test should be performed before storage, and no later than 12 months for orthodox seeds or 3–4 months for recalcitrant seeds after collection. Preferably, 100–200 seeds are used for testing, but the viability sample size will depend on the accession size (FAO 2014). Reliable results can already be obtained by using 3 × 25 seeds for one testing condition (Filip Vandelook, pers. comm.). Monitoring should take place at least every ten years for base collections and every five years for active collections (MSBP 2015). If the viability rate lies below 85%, regeneration or recollection should be implemented (Engels and Visser 2003; MSBP 2015). A lower threshold (70%) can be set for some landraces and wild/forest species that cannot achieve viability rate of 85% or higher. While international standards for seed banking (FAO 2014) are mainly tailored to crop species in genebanks, it should be remembered that the diversity encompassed by all seed plants is dramatically greater; and processes and standards will probably need to be adapted and relaxed to accommodate wild species preservation in seed banks (Hay and Probert 2013).

Germination tests of excised embryos and embryonic axes should be assessed by the root production rather than shoot development, as the latter will not occur (FAO 2014).

While the germination test is usually the method of choice for seed viability assessment, vital stains (e.g., TTC) can be informative in species where dormancy is difficult to break (Gosling 2003). TTC is also used to assess the viability of embryos and embryo axes (Normah and Makeen 2008).

All data regarding germination abnormalities, and viable:inviable seed rates should be recorded to determine whether deterioration is occuring (FAO 2014). Ballesteros et al. (2020) provide a list of gene-banking strategies for maintaining germplasm viability and extending the longevity of seeds.

Different methods can be applied to cultured plant cells:

• FDA. The solution fluoresces green under UV in viable cells, whereas dead or damaged cells remain unstained.
• Evan´s blue staining. In this case, inviable cells are stained. This test usually complements the FDA test.
• Phase contrast microscopy is used to assess the presence of a healthy nucleus (Bhojwani and Dantu 2013).
• Micrografting of cryopreserved material.

Plant material that has been maintained for more than ten years in culture is susceptible to somaclonal variations. Hence, these accessions can be grown in the field or a greenhouse for an integrity check, including morphological, cytological, and molecular characterisations (Panis et al. 2020).