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Trapping Methods FAQs

  1. Can I sample soil cores at regular intervals along a transect to recover a representative collection of fungi colonizing roots in a plant community?
  2. What sampling method(s) can I use if my goal is to recover as many species of a fungal community as possible?
  3. Can roots be sampled, washed and added as inoculum to trap cultures to sample only those fungi forming mycorrhizae?
  4. When using root inoculum, should there be any consideration of the type of roots sampled (such as coarse versus fine, anchor roots, etc.)?
  5. Should inoculum be mixed throughout the pot contents or added as a layer?
  6. What can be done if the amount of inoculum is low (as with roots)?
  7. Why is it so important to use larger pots for trapping?
  8. Why should trap cultures be grown for a minimum of four months?
  9. Why do fungi that don’t sporulate in the field sporulate in pot culture?
  10. Can plants transplanted from the field into pots serve as an inoculum source to trap colonizing fungi?
  11. Can trap cultures still be productive if no sporulation is observed but mycorrhizal colonization has been detected?
  12. Should a plant species native to the sampled site be used instead of a “standardized” host such as sudangrass or bahia grass?
  13. How do I know that most of the fungal species present in a community have sporulated in trap cultures?/
  14. Can trapping help to ascertain changes in fungal community structure in a single plant over time?
  15. Why is so much sand used with the field soil to set up trap cultures?
  16. Some people recommend vermiculite, perlite, or peat moss as a potting medium instead of sand, because it is has less bulk density. Why don’t you ever use these media?
  17. What if I don’t have facilities to produce trap cultures to obtain some idea of the species present in my study site(s)?


  1. Can I sample soil cores at regular intervals along a transect to recover a representative collection of fungi colonizing roots in a plant community?
  2. Yes, if the goal is to examine distribution of fungi already known to be present, and none of the samples are pooled. In this procedure, cores are taken even if plants made not be present. However, in bare areas, inoculum potential any mycorrhizal fungi can vary considerable (and usually is quite low). These samples may or may not lead to sporulation after a single propagation cycle in pot culture. ANY soil core/sample taken outside the root zone will give unpredictable results. Usually only the most aggressive (or persistent) fungi will be recovered because their hyphae and spores are more likely to have spread farther from any “parent” host plant.

  3. What sampling method(s) can I use if my goal is to recover as many species of a fungal community as possible?
  4. Always remember that mycorrhizal fungal propagules (especially the less aggressive colonizing fungi) tend to be concentrated in the root zone. The approach we take is to sample individual plants, with the plants chosen based on the objectives of the study. For example, if the goal is to compare fungal communities colonizing the same host species at different locations, then only that plant species is sampled. If the goal is to examine host preference among neighboring plants, then representative plants are sampled. In any case, we use the following sampling regime:

    Dig around the plant with a trowel or shovel to separate the majority of roots (or root ball) from surrounding soil.

    Use the trowel or other instrument to manually remove soil clumps devoid of roots. It is essential that soil aggregates clinging to roots be retained.

    Manually chop roots into 2-4 cm fragments and mix thorough with soil. We usually use a sharp hatchet on a pre-sterilized cutting board covered with paper back at the lab. This mix is now ready to use as inoculum in a trap culture.

    The final mix should contain at least 50% roots if possible. The roots also are highly infective—often moreso than soil propagules (spores, detached hyphae), and they have a higher probability of containing fungi that haven’t produced enough mycorrhizal biomass to sporulate.

  5. Can roots be sampled, washed and added as inoculum to trap cultures to sample only those fungi forming mycorrhizae?
  6. Definitely yes. But take certain precautions. First, don’t wash the roots (valuable external hyphae will be lost or killed). Take the unwashed roots (usually with some soil attached) and immerse in a large container of water (4 liter volume or more). Let the roots stand for an hour or so, and then reach into the water and gently message the roots to shed soil aggregates. Remove roots and cut into fragments 3-4 cm in length (don’t cut them smaller) and immediately place in moistened growth medium (whatever is going to be used in pots to grow plants).

  7. When using root inoculum, should there be any consideration of the type of roots sampled (such as coarse versus fine, anchor roots, etc.)?
  8. Mycorrhizal fungi are present either internally or externally on most roots growing through the soil. Internal colonization is less abundant in older coarse roots, but fungal hyphae still may be profuse on the external surface. This is especially true of tree species, in our experience. Therefore, the safest coarse is to take a random sampling of ALL roots in the soil. Its as important to obtain a representative sampling of all possible niches as it is to keep inoculum potential as high as possible.

  9. Should inoculum be mixed throughout the pot contents or added as a layer?
  10. We consider layering of any inoculum for trapping that consists of a species mixture to be a big NO-NO for two reasons. First, propagules are concentrated in a narrow band. As roots pass through, these propagules establish entry points. Initially, all secondary growth of the fungi must come from mycorrhiza formed in this narrow zone of colonization and only the most aggressive fungi are likely to spread widely into roots growing rapidly outside this zone. Second, niche space for initial mycorrhizal development is at a premium in a narrow band, so that less aggressive fungi (or those with few propagules) may not even become established. They certainly are less likely to increase in biomass in the presence of aggressive fungi because they can’t keep up with other fungi or even with root elongation.

    In contrast, inoculum dispersed throughout a medium provides spacing so that isolated entry points are formed by whatever propagules are near the roots (whether in large numbers or not). More important, each established fungus likely has more room to spread in different directions and increase in biomass, a requirement for sporulation to be triggered.

  11. What can be done if the amount of inoculum is low (as with roots)?
  12. A homogenous mixing of inoculum in the pot growth medium still is important, but one of two approaches can be taken to insure adequate colonization to stimulate sporulation: (i) use a smaller pot (preferably deeper rather than wider), but don’t use pots with a volume less than 500 cm3, or (ii) use a larger pot, but grow plants for a longer period of time. The former is likely to increase the rate of initial colonization and subsequent spread and the latter is likely to compensate for a low rate of initial colonization. In our experience, the latter seems to be more effective in maximizing number of species recovered (although sporulation may be low). For example, sporulation is abundant on sorghum in 15-cm diameter pots within 3 months for most fungal species, but it may not even be detected until 4-5 months in 30-cm diameter pots.

  13. Why is it so important to use larger pots for trapping?
  14. Its all a matter of space for roots and space for the fungi to colonize those roots. The larger the pot, the more abundant the roots and the more space for initial and secondary colonization by fungal propagules (assuming they are widely dispersed in the medium).

  15. Why should trap cultures be grown for a minimum of four months?
  16. This recommendation was made to accommodate a wide range of host species. Four months is not any sort of magical time period. What is important here is that the host plants should have all the time they need for roots to fill the pot and then to cease topgrowth and then to maintain the culture for another month during which plants are static in their growth. A lot of anecotal observations indicate to us that a high proportion of sporulation occurs during that last 30 days. One possibility for this behavior is partitioning of resources between host and fungus during different phases of growth. We observe repeatedly that arbuscular fungi DO NOT sporulate in response to stress or other adverse stimuli (like a lot of culturable fungi). We see that a fungus sporulates spontaneously to some extent while the plant is actively growing, but it is low compared to when the plant has matured or no longer is growing in pot culture. Maximum investment in vegetative growth appears to take place when roots (and shoots) are most actively growing. Minimum investment occurs when root growth has slowed or ceased and carbon now may be reinvested in sporulation. Possibly, one reason why sporulation in the field rarely compares to that in pot cultures is that constraints on root/shoot growth by pot boundaries does not occur in the field.

  17. Why do fungi that don’t sporulate in the field sporulate in pot culture?
  18. In part, it goes back to the dynamics of root growth in pots versus field soil (see question 7 above). In the field, roots are unconstrained and indeterminate in growth patterns Mycorrhizal development follows a parallel pattern (unconstrained fungal vegetative growth and indeterminate spread). Sporulation, if it occurs at all, likely increases only in roots that have ceased growth and host carbon and nutrients are reallocated from mycorrhizal structures to spores. In pots, root growth becomes constrained relatively quickly and all fungi that have established sufficient biomass will then have host resources to sporulate. In most cases, I think its a juggling act between rates of growth by each symbiont and source-sink relationships at each point in time.

  19. Can plants transplanted from the field into pots serve as an inoculum source to trap colonizing fungi?
  20. Yes. However, the results obtained often do not match those involving whole soil or root inocula discussed in previous questions. In most of the cases with which we are familiar, fewer species are recovered and sporulation is lower. More complicated interactions probably are occurring in this situation here. First, the plant from the field may not be adapted to greenhouse conditions (temperature, light, watering regime, pot environment) and grow poorly, in which case the colonizing fungi also will grow poorly. Second, results are a function of “predisposition”: the distribution and abundance of colonization by different fungal species at the time of transplanting. There does not exist the open “playing field” of new roots passing through widely dispersed propagules found in pots with mixed soil or root inocula. Age of transplants could greatly affect trap culture outcomes, but too little work has been done to offer clues as to which age might optimize species richness and spore abundance in pots.

  21. Can trap cultures still be productive if no sporulation is observed but mycorrhizal colonization has been detected?
  22. This is a frequently outcome of many trap cultures from almost any environment, and the reasons are complex because they can involve a combination of host, fungus, and/or environmental variables. As long as there is colonization, then the fungi present usually can be induced to sporulate. We use two successive approaches. The first approach is to reseed the culture and obtain a new flush of root growth to increase mycorrhizal biomass. Usually, this alone will produce a positive result. When it doesn’t, though, the second approach is to reseed with a different host species. If the original host is a grass, try a legume, or vice versa.

  23. Should a plant species native to the sampled site be used instead of a “standardized” host such as sudangrass or bahia grass?
  24. Possibly. However, the ideal host species is not necessarily one best adapted to the sampled site, but one which is best adapted to local greenhouse conditions. Native plants can become “foreign” plants to native fungi when thrust into pots in a greenhouse. It seems as though the greenhouse environment (especially light and temperature) have the strongest influence, perhaps because these variables act on both the host and the fungi when in the extraradical phase. Always remember, the trap culture host is analagous to a living “petri dish”. It will serve as a “selective medium” when (i) root development is slow and low in volume (so native plants which have coarse roots with little branching or are slow growing are unsuitable) or (ii) plants are less mycotrophic (lower plateau of colonization, less niche space for ingress by all the different fungi that might be present). Both of these conditions should be avoided at all costs!

  25. How do I know that most of the fungal species present in a community have sporulated in trap cultures?
  26. There is no way current technology provides a definitive answer. All that we can do is conduct exhaustive sampling until the number of species reaches a plateau. Usually, this means several successive cycles of trap culturing, trying several trapping regimes (whole soil, roots, transplants, or different host) and then returning to the field site with some knowledge of the species present and sample directly over several seasons. This is a lot of work, and should only be undertaken when the research questions require this level of characterization.

  27. Can trapping help to ascertain changes in fungal community structure in a single plant over time?
  28. At the present time, this is a goal being tackled by molecular biologists to develop probes that can directly measure the amount of colonization by species present in a fungal community. Of course, this means developing quite a few probes, because a lot of species can colonize the same root system. Another approach, using trapping methodology, can address this goal more indirectly. Roots are sampled at intervals during key periods of plant development (e.g., period of rapid vegetative growth, flowering, seed set, senescence, etc.). The main constraint here is the amount of roots produced by the plant. If the more roots available, the greater the number of potential sampling times. A large sample size is not needed. Just make sure to disperse roots throughout pot contents and grow plants until good sporulation is detected (taking cores from pots to check periodically). Data thus collected should always be interpreted with caution and a healthy dose of skepticism because there always is the possiblity that one or more fungi in trap cultures may not be sporulating.

  29. Why is so much sand used with the field soil to set up trap cultures?
  30. The biggest problem with the pot environment is compaction of soil due to the confined space and repeated watering. Sandy soils might only require a 1:1 v/v dilution, but clay soils often must be diluted 1:4 or 1:5 v/v with sand to create an optimum environment for root infiltration and aeration (let alone fungal expansion into the soil matrix). When in doubt, try several dilutions in small “cone-tainers” or other small pot, seed and watch to see if there is any standing water (a no-no), poor seedling emergence, or slowed growth (relative to the highest dilution). Signs of any problems generally become apparent within days or 1-2 weeks.

  31. Some people recommend vermiculite, perlite, or peat moss as a potting medium instead of sand, because it is has less bulk density. Why don’t you ever use these media?
  32. There is nothing wrong with any of these media to grow plants. And as long as the plants are growing fine, arbuscular fungi in the roots also are likely to be growing well. These components are problematic when it comes time to extract fungal spores/hyphae from a pot. Their light density leaves many particles suspended in sucrose following centrifugation, so they are present with spores after extraction. Not only are spores sometimes hard to see, they also are difficult to manually remove and clean up for mounting, inoculating, or other purposes. If the extract sits for more than 15-20 minutes at room temperature, then aggregation of hyphae, spores and particulates occurs. Spores become almost impossible to collect under this circumstance. The presence of organics is especially distracting. Just try to count spores in almost any of the commercial inoculants on the market (assuming any healthy spores are even present)! A difficult challenge, indeed!

  33. What if I don’t have facilities to produce trap cultures to obtain some idea of the species present in my study site(s)?
  34. In the absence of a greenhouse, it might be possible to set up a modified culture system in situ at the study site. We can’t comment on potential for success, since we have no direct experience with the method. However, as long as the plants are growing fine, mycorrhizal fungi in the roots also are likely to be growing well. To try this method, we would suggest using an annual host with high dependency (e.g., C4 grass, legume) that produces prolific fibrous roots. One benefit of this approach is that many nondestructive samples can be take to monitor frequency of sporulation (which will need to be watched, since spatial and temporal constraints imposed by size and shape of pots is absent).