Habitats rich in arthropod-dispersed fungi include dung bark beetle galleries, mushrooms, and decaying vegetation. One approach to observing the presence of sticky (phoretic) spores from a particular substrate is to examine the surfaces of arthropods collected in the substrate by examining them with very high-powered dissecting microscope or a compound microscope.  Insects, especially beetles and flies, can be caught in traps as they are attracted to or leave the substrates. The life cycles of the fungi usually are synchronized with those of the dispersers, so the diaspores are mature as the arthropods themselves are ready to disperse. Phoretic spores also can be found by looking through insects and mites in arthropod collections, although this procedure may be very tedious, because low numbers of spores are present in the general arthropod population from unspecified substrates. We can also study fungi with phoretic spores by manipulating the close contact of potential dispersers with the fungus, as is described in the next few paragraphs.

Basidiobolus ranarum is well known to mycologists as a fungus that can almost always be isolated from the dung of toads, frogs, and lizards as well as many other animals.  Some isolates of the fungus are mammalian (including human) pathogens, so care must be used in handling these fungi. For this study a known non-pathogenetic culture should be acquired from a culture collection.

An interesting feature of the life cycle of Basidiobolus is that it may produce several types of spores; usually, depending upon its microhabitat. The ballistospore is forcibly discharged to stick to substrate.  On the other hand, the capilliconidium is not forcibly discharged, but it has a weak spot in the conidiophore so that upon contact the spore is released, and capilliconidium with a sticky blob on glue on the tube-like haptor at its distal (outward) end can attach to anything it contacts, often a passing insect or even a growing fungus hypha.  Capilliconidia may develop directly from a hypha or from a mature ballistospore. Over time the attached capilliconidium becomes cleaved into internal segments and the attachment glue darkens due to oxidation. The dark material makes it easy to spot a tiny colorless capilliconidium on the surface of a light-colored arthropod.

Many mycologists had focused their attention on the intriguing, forcibly discharged spores of Basidiobolus, and it was only in 1956 that the capilliconidial stage was discovered adhering to culture mites. Even more recently (1989) it was suggested that the attached capilliconidia provides the explanation of a small mycological mystery: Is Amphoromorpha truly a fungus as it was described to be in 1918 and 1920? Amphoromorpha was described as a fungus that is associated with insects.  The thallus of Amphoromorpha was described as a sac-like, internally cleaved structure that becomes attached to insects and mites, and long lists of hosts have been drawn up.  The “synthesis” of Amphoromorpha from Basidiobolus capilliconidia attached to mites was first suggested by the accurate observations and drawings of mycologist Charles Drechsler. 

• Petri dishes containing a layer of half strength agar (8g corn meal agar, 12g Bacto agar/L H₂O
• Basidiobolus culture (obtained from culture collection to insure that it is non-pathogenic to mammals
• Cow dung moist chamber cultures; use dung that is fairly fresh, but not so fresh that insects have not had a chance to visit it.  Also, this experiment needs to be done at a time when the weather is warm enough for insect activity.
• Living mites; the long-legged, fast-moving predaceous mites found in about 3-6 day-old dung chamber cultures are easy to locate using a dissecting microscope to observe the dung surface. Light-colored insects also may be used; termites collected in logs or dead branches are usually easy to collect by tapping the wood so that the termites rain down into a plastic bag.  The light color makes it easier to spot the attachment point of the fungus that is dark in color.
• Disposable gloves to protect hands from contact with dung
• Microscope slides and cover glasses
• Glycerol-acid fuchsin (Benjamin, 1971) (optional)
• Dissecting microscope
• Compound microscope

• When the cow dung moist chambers are about 6 days old a clean cover glass can be grasped at one corner with forceps.
• Take a cover glass held parallel to the dung surface, and lower it until it almost touches the dung. Ideally it should touch only the tips of the diverse aerial fungal growth extending from its surface.
• Place the cover glass on a slide with the side that touched the moist chamber facing down on the slide.
• Water or a water soluble mounting medium with a slight tint of stain such as glycerol-acid fuchsin should be introduced with a pipette into the space between the cover glass and the slide.
• Use a compound microscope at low magnification to determine if sticky spores that might attach to an insect are present on the slide.
• If spores are present (usually there will be several so that there will be no doubt that they have this mechanism for dispersal), you might want to try touching the cover glass to the dung culture again, but this time actually play the disperser and transfer the spores to a new agar plate.
• If you are careful not to damage the microscope lens, use the low power lens of the compound microscope to look through the cover glass to monitor the development of the spores you transferred to the surface of the agar on the cover glass. Observations may continue over a period of a week or even longer if successful.

• Inoculate the agar in the Petri dishes with Basidiobolus by streaking from the original culture about four days before the experiment is to be performed. The presence of some capilliconidia on the Petri dish lid is an indicator that the cultures are ready for the rest of the experiment.
• With a light touch use forceps to pick them out of the cow dung moist chamber or the container in the case of termites. The insect and mite dispersers may need to be cooled so that you can catch them without harming them.
• Place the dispersers in the Basidiobolus cultures. You will need to observe the disperser behavior for some time after they are introduced into the plates, because some of the mites and termites may burrow into the agar or go to the top of the Petri dish. Walking on the top of the Petri dish is not a real problem because many spores (both ballistospores and capilliconidia) are found on the lids where they were shot from the agar or on which they develop.  A desire to burrow into the agar is a problem because the dispersers need to contact the capilliconidia if dispersal is to be effective. Prod the wayward burrowing dispersers gently to get them back onto the agar for a period of time.
• Harvest several of the potential dispersers from Petri dishes each day to determine if any capilliconidia are attached to them.  This is necessarily destructive sampling because the best observation comes from mounting the potential disperser between two cover glasses and carefully examining it on both sides to look for the spores.  Observations should be continued for 3-4 days.
• The following conditions may be scored over time: 1) number and position of attached capilliconidia per disperser, 2) color of attachment site (colorless of tan or brown), 3) cleavage within a capilliconidium present or absent. Determine and describe the development  process beginning with Time 0 as the time of introduction of the potential dispersers into the cultures.

• What is the cost of carrying fungus spores to the arthropod?
• Why can Basidiobolus usually be found in the dung of amphibians and reptiles?
• If the object of spore dispersal is to hit a target (for which a fungus has the enzymes to decompose), is air or phoretic dispersal a better method?  Or there any qualifications to your answer?

• Blackwell, M., and D. Malloch. 1989. Similarity of Amphoromorpha and secondary capilliconidia of Basidiobolus.  Mycologia 81:735-741.
• Drechsler, C.  1956.  Supplementary developmental stages of Basidiobolus ranarum and Basidiobolus haptosporus.  Mycologia 48: 655-676.
• Thaxter, R.  1914.  On Certain Peculiar Fungus-Parasites of Living Insects.  Bot. Gaz. 58:235-253.
• Thaxter, R.  1920.  Second Note on Certain Peculiar Fungus-Parasites of Living Insects.  Bot. Gaz. 69:1-27.