Frequently Asked Questions
There are two possibilities: (i) the culture became contaminated during the culturing process in your greenhouse or (ii) the culture appeared to be pure in our protocol because only one fungus was sporulating in our greenhouse/growth room, but a contaminant fungus differentially grew and began to sporulate with the change from our greenhouse conditions to yours. When such an occurrence arises, contact us immediately! We will immediately check our source cultures and their propagation history (for the presence of the contaminant in previous generations or in the original accession).
If the contaminant species matches a fungus in our records, the problem is ours and we will immediately correct it. If it doesn’t, then check culturing conditions.
The problem is the method by which we ascertain purity of a culture: the presence of spores that can be identified to species. But sporulation occurs only after a fungus achieves some threshold mycorrhizal biomass (which may be isolate or species-specific). Thus, two or more fungi can be colonizing a root system, but only one is a consistent sporulator in one environment. When this environment changes, the other fungi present in low mycorrhizal biomass may be favored, spread more, and then sporulate.
For example, a monospecific culture from Florida yielded spores of four species in three different greenhouses (so it wasn’t a local problem), and worse, the fungus that was supposed to be there barely sporulated! Cultures grown in Florida never produced spores of these contaminant fungi. As another example, a culture of Gi. rosea in its 10th propagation cycle with no sign of contamination produced sporulation by G. intraradices when grown in a greenhouse in Arizona.
There always is the possibility that the contaminant came from an outside source, but this is unlikely if care is taken during the culturing process.
Start from spores, so that other propagules which lead to colonization (hyphae, roots) are excluded. This way, only the sporulating fungus is propagated in the next culture generation. We use this method whenever the possibility of a contaminant exists (and we want to exclude it).
In Lynn Abbott’s lab in Perth, Australia, each culture generation is started from spores, thus minimizing the problem altogether (as long as there is no contamination from an external source). We do not follow this procedure for two reasons: (i) its very time consuming and thus logistically impossible with our limited staffing and high number of active cultures, and (ii) spores of isolates of some species (never predictable) are much less infective (if not uninfective) than vegetative propagules (mycorrhizal roots or hyphae).
Our job is to culture ALL organisms accessed into the collection, so we must always start with whole inoculum and than narrow propagule type to spores once we are certain the spores are highly infective in our culture environment.
No. With only propagules of one fungus, then any constraints by root volume, rate of root growth relative to rate of fungal growth, distribution of inoculum (as a layer versus a homogeneous distribution), etc. affect only the final yield of that culture. Only when two or more fungi coexist in an inoculum is pot size and other variables of major importance because competition for niche space becomes a dominant force.
Either method (below roots or directly onto roots) works. We have used both, but have decided on the root inoculation procedure as a standard procedure because of it is more efficient and also because it gives us positive results for certain fungal isolates when no other method works. We think the reason for this method’s success is that spores are in direct contact with roots at different physiological states, thus optimizing colonization at some point by one or more spores.
Single spore inoculum is useful if: (i) the person preparing the spores doesn’t have enough experience to separate species precisely, (ii) the goal is to try and distinguish genetic segregates (frequency of nuclear segregation, not allelic segregation) or (iii) if the goal is to examine genetic heterogeneity in a single cell.
It is potentially disastrous if the inoculum is meant to maintain all of the genetic heterogeneity of the “population” of that fungal isolate. For our purposes, we use batches of spores (no less than 100, if possible) to minimize loss of genes in low frequency in spores. Of course, we have no idea if or to what extent this is happening in each fungal isolate, so we can’t ignore the potential for its occurrence.
It depends on the genus of the fungus being used as inoculum. Many species (but not all) in Gigaspora and Scutellospora can be started from active cultures without much problem. Isolates of some species, such as Gi. gigantea, can produce 100% cultures from single spores!
Species of Glomus are more heterogeneous, and thus considerably more unpredictable. Species of Acaulospora and Entrophospora generally are difficult to start without a dormancy period. Besides the cumulative genotype due to evolutionary history, much also depends on life history traits at the site where the fungus was found.
Given how little we know about any and all of these fungal genotypes, would you want to take a chance? With the heterogeneity of germplasm in INVAM, we can’t afford that, so we generally follow a standard practice of at least one month storage apart from the living plant host in an attempt to put all fungi on equal footing.
This is a very common pattern that is largely ignored in the literature. Its one for which we are always monitoring because it can occur no matter how careful someone is in preparing spores for inoculation. Invariably, the contaminants are the vesicular-arbuscular fungi (Glomus, Acaulospora, Entrophospora) and rarely (if ever) Gigaspora or Scutellospora.
It was this consistent distinction which led us to believe the source of contamination was hyphal fragments associated with the spores separated and cleaned for use as inoculum. Sometimes these hyphal fragments are not detectable, despite rigorous examination of spores. Even though these fragments are highly infective (much more so than spores for many species), they usually are few in number.
As a result, establishment of an infection unit and subsequent secondary colonization increases slowly and is manifested as sporulation only after several to many culture generations (depending on fungal aggressiveness, compatibility with host phenology, and environment). It is this constant possibility of intrinsic contamination that requires strict examination of all cultures prior to take-down and storage.
Even after sporulation declines or ceases, the fungus/fungi may still be happily colonizing roots and “living the good life”. There is no Viagra for fungi, though. We have been somewhat successful in retriggering sporulation by using one of two methods: (i) reseeding the culture with a genetically distant host to the one being used (e.g., legume instead of grass or vice versa), or (ii) taking a section of plants from a culture, transplanting it to a larger pot, and seeding around it with a different host species. The former is easier and seems to produce more frequent positive results.
The main reason is contamination. We have yet to find ANY cultures that remained monospecific when this was done. Pot contents dry out and when this happens, the dust level increases, and these floating particulates (which may contain fungal propagules) can spread passively between pots. The likelihood of this occuring increases in summer with increased air circulation for cooling. All sorts of things can happen in the glasshouse during this time period which could compromise both purity and health of the pot contents.
The other problem is buildup of populations of microbial saprophytes (bacteria, fungi, actinomycetes) over time, especially in a pot environment where plants have long ceased to grow and root senescence (and release of organic substrates) accelerates. These microbes may not pose a problem for the immediate culture, but they spell trouble for longevity of fungal propagules in storage and infectivity in the culture cycle.
If you have kept up with the literature, then you know some recent studies are being reexamined in light of contamination of spores by non-glomalean fungi. This can happen to the most discriminating reearcher because spores can appear quite clean and healthy yet be either parasitized or dead. The method we use to remove these tiny culprits from spores is to incubate extracted spores at 4oC for various periods and cull out any suspicious looking ones. After extaction (by whatever method), transfer spores to a petri dish containing tap water. Incubate the petri dish in a refrigerator for a minimum of 48 hrs.
Examine spores under a stereomicroscope and remove all those which show signs of contamination or degradation. If the proportion of compromised spores is > 5%, then incubate for another 24 hours and cull again. If more than 15-20% of spores are contaminated, then the whole batch of spores should be discarded and new ones obtained from a different culture. In this case, microbial contamination would be too high to insure its exclusion from a spore sample. If the proportion of compromised spores is < 5% after the first incubation, then pull as many of the healthy spores as needed.