Chapter 3

Collection of Hosts

Except for the few species available in axenic culture, trichomycetes must be studied from material obtained in arthropod hosts. The wide range of host types, size, and habitats requires many different collecting methods. In this section some of the techniques we have found to be useful will be described. Living hosts are more easily dissected than preserved ones and, more importantly, provide superior material for study. Consequently, not only must the arthropods be found and collected, but they need to be kept in a living condition until dissected in the laboratory. Freshly dead specimens may be used. However, as decomposition progresses the thalli degenerate rapidly and sporulating parts may produce artifacts that can lead to misinterpretations. Molts of hosts (shed exuviae) should also be collected, because some stages of the fungi such as zygospores and resistant spores may develop only during ecdysis.

Hosts of trichomycetes can be handled safely without special precautions, with two exceptions. One obvious group consists of those crustaceans with large chelae such as the crabs. The other group is the millipedes, which, when disturbed, may secrete a noxious liquid from glands located laterally along the body. This substance contains cyanide in some species and can stain the skin and has an unpleasant odor, but is not harmful provided it does not contact membranes of the mouth or eyes.

Freshwater hosts are more common in flowing waters than in quiet ponds and pools. They consist mostly of immature insects attached to stones and sticks or to immersed living or dead vegetation, and include the larvae of blackflies and midges and nymphs of mayflies and stoneflies, among others. Their trichomycetes are species of Harpellales and Amoebidiales (Paramoebidium). Clean, small streams are generally preferable to larger rivers. The insects often will be located in riffles and other parts of the stream where water flows rapidly and provides good aeration. Infested larvae of Chironomidae, Dixidae, etc. may be found along waterfalls where they are bathed by running or trickling water. The Dixidae also occur on aquatic vegetation where the water interfaces with air. Winter-emerging stonefly nymphs (Capniidae) are probably in diapause and hyporheic (deep in the substratum) most of the year; they can be collected in the winter and early spring in leaf packs or on sticks and stones in flowing streams when they come up from the substrate to complete their development. Amphipods and insect larvae may live in aquatic vegetation or decomposing plant material near the border of streams, or in pools, and isopods are found on rocks and submerged plants and decomposing vegetation. Some of these crustaceans are hosts to species of Eccrinales.

A pair of fine-pointed (but not sharp) forceps is useful to pick up arthropods from stones and sticks lifted from the water. Mayfly and stonefly nymphs on the underside of rocks often drop into the water when disturbed; consequently, stones should be turned over quickly and, preferably, held over a pan to catch those that do drop off. A net positioned downstream can also be used to catch nymphs that release themselves. A white pan is useful to examine the emptied contents of nets or strainers that have been swept through aquatic vegetation, or to hold plucked vegetation until blackfly and chironomid larvae are transferred to jars. Stones may have chironomid larvae visible on the surface or concealed in silt tubes. Others are found in the algae that coat some rocks, or in decaying wood. Blackfly larvae may occur in profusion on rocks or submerged leaves in streams, but care should be taken not to scrape them off in masses, because under these conditions they become entangled in the silky threads spun from their salivary glands and do not survive long.

Insects from fast-flowing streams should be placed in collecting jars or resealable plastic bags without crowding and with a minimum amount of water. As soon as possible these jars should be placed on ice. Jar lids can be tightened during transport to prevent loss of specimens, but the lids should be opened occasionally to insure adequate aeration. It is generally best to separate the families of insects, and care should be taken that predaceous forms are not included. Aquatic arthropods that contain trichomycetes are not carnivorous, but under crowded conditions in time some may resort to cannibalism. On arrival at the laboratory, the specimens should be further sorted and placed in shallow layers of prechilled stream or distilled water in containers with loose lids such as petri dishes, and kept in a refrigerator. Depending on the species, the care in their handling, and transportation conditions, insects can be maintained alive in a refrigerator without additional food for several days to several weeks.

Arthropods from lakes, ponds, pools, and other quiet waters that are likely to contain Harpellales include the larvae of mosquitoes, midges, caddisflies, and mayflies. In general, ephemeral pools and ditches are less likely to have trichomycetes than more permanent bodies of water, even though the appropriate arthropods may be present. One of the most useful tools we have found to collect insects from aquatic vegetation or muddy bottoms is a round-bottomed, woven-metal food strainer, 15 cm or less in diameter, which we use with the two metal prongs (at the edge opposite the handle) bent backwards against the strainer. Contents of such a strainer, or a net, can be emptied into a pan for examination. Smaller specimens can be removed and transferred to jars using a small pipette fitted with a bulb, larger specimens with forceps. Bloodworms (Chironomus spp.) build tunnels in muddy bottoms of pools, ditches and even slow-running streams, and if the mud is disturbed they can be scooped from the water. They can sometimes be removed effectively from concentrated populations by slowly dragging a small wire loop (~8 cm in diameter) attached to a handle through the mud, which results in the larvae becoming looped around the wire, or by simply dredging with a net. Other midges with trichomycetes are sometimes found in small transparent tunnels, which they build on the surfaces or within the senescent tissues of hydrophytes. Cladocerans are best collected with a plankton net. Collembola (springtails) may be a challenge, but can be collected in a pan and transferred to a jar with water for transport and storage.

As the majority of known marine hosts live in or near the intertidal zone, collecting is easiest at low tide. Mud-burrowing crabs may be caught while they are walking on the surface, or removed from their burrows by inserting a trowel or spade through the mud so as to cut across the burrow and prevent their downward escape. Collecting anomurids such as Callianassa and Upogebia may require deep and considerable digging with a spade, or with a special device in Australia called a "yabbie" pump that consists of a cylinder with a plunger. Nonburrowing isopods and amphipods may seek shelter and moisture under stones, or under wood, seaweed, etc. deposited at the high tide line. Some of these animals that do not live within the water at high tide might more properly be classified as terrestrial, but they are tolerant of water with high salinity. The ubiquitous rock louse, Ligia spp., moves rapidly on rocky shores, seawalls, and docks, and may be difficult to catch. But specimens can often be collected from walls and pilings by placing a pail or plastic bucket under them into which they will fall when disturbed or when prodded from cracks. They are somewhat delicate and must be handled carefully. In rock piles, smaller rocks lifted quickly over a pail may allow catching several Ligias at a time. Most marine arthropods can be kept for days in a cool place in a container with a shallow layer of seawater and some objects onto which they can crawl. Freshwater, rather than seawater, should be used to replenish the water as it evaporates in order to prevent excessive increases in salinity. Aerated seawater tanks may also be used, or fresh seawater can be used to replace the stale seawater once or twice a day.

Terrestrial arthropods with trichomycetes include millipedes, beetles, isopods, and amphipods. They are found predominantly in or around the moist, decomposing vegetation on which they feed. Populations may fluctuate considerably in number in areas where droughts occur. Some millipedes and beetles either normally inhabit logs or seek shelter in logs during drier periods, and can be located by tearing apart the decomposing wood. All terrestrial hosts can be placed in containers with some of their natural substrate for transporting to the laboratory for subsequent maintenance.

The time of year when trichomycetes can be obtained depends on habitat and climatic conditions. Especially important may be the life cycles of some of the insect hosts whose immature forms, but not pupal or adult stages, contain trichomycetes. The timing of collections is generally more critical in temperate climates with severe winters than it is in tropical or subtropical areas of the world where developmental stages of populations of multivoltine species may overlap considerably. Species of Harpellales are essentially restricted to immature insects. At high altitudes and latitudes, the larval stages may be found only during relatively short seasons. Nevertheless, the abundance and variety of larval insects may be so intense in many of these regions that they offer some of the best collecting of Harpellales. When infested hosts consist of adults as well as sexually immature stages of the same species (many crustaceans and millipedes), the fungi may be collected at any season when the hosts can be found. Marine habitats are usually more stable than freshwater and terrestrial ones in temperate climates, so that many marine trichomycetes can be obtained throughout the year. However, even in such cases there may be some variation in abundance and stages of development of the trichomycetes that is tied to environmental factors and molting cycles of the host (Hibbits, 1978).

Living arthropods with trichomycetes may sometimes be obtained from commercial biological supply houses or professional collectors, but such sources are usually unreliable because of unpredictable infestation among different populations. Daphnid and bloodworm cultures (the latter, for instance, available as fish food from merchants in Japan and possibly other countries) can yield excellent Amoebidium parasiticum infestation in some cases. Likewise, it is possible to obtain from commercial sources living hosts like the fiddler crab (Uca spp.) and spiroboloid millipedes containing species of Enterobryus. These may be satisfactory only if shipped soon after they are collected from natural sources.
 

Maintenance and Preservation of Hosts

Successful collecting expeditions invariably require that the living arthropods be maintained for at least a few days in the laboratory while they are being dissected and their fungi studied. Longer periods of maintenance may be desirable or necessary where collecting sites are distant from a laboratory, or when hosts must be available during experimental investigations involving them. With some species the percentage of infested individuals in laboratory populations can actually increase over time, whereas in others the infestation may decrease as the hosts molt and shed their fungi and then do not become reinfested, or if they are fed an unsuitable diet. Many arthropods molt in the laboratory, and thus provide a stage of fungal development that may not always be recoverable in nature.

Insect larvae and nymphs from streams can be kept alive in a refrigerator in shallow layers of water, as described previously. Those that remain alive may molt, pupate, or metamorphose into adults. The larvae of some dipterans cannot be identified to species easily, if at all; consequently, it may be helpful to the specialist to be given preserved specimens of pupae or adults that are known to have developed from immature forms identical to those that have been dissected.

Hosts from quiet waters include dipteran larvae and cladocerans. Most can be kept at room temperature in jars half filled with water, or in a refrigerator if arrestment of the rate of host development is desired. Fungal transmission can be fairly good among crowded mosquito larvae (with Smittium spp.) and cladocerans such as daphnids (with Amoebidium parasiticum). If sample specimens in such collections are examined and found to be uninfested, the remainder can be left a few days to a week or more for subsequent examination on the possibility that infestation undetected in a few individuals has spread to others. It is best to remove the pupae of mosquitoes as they appear in order to avoid problems later with escaping adults; with one exception (Sweeney, 1981a), only larval stages of mosquitoes are known to harbor trichomycetes. Bloodworm larvae that live in muddy bottoms will survive better in shallow layers of water, either with or without mud. Infested cultures of cladocerans, in contrast to insect larvae, can be kept indefinitely under appropriate culture conditions.

Many marine hosts, such as hermit crabs, true crabs, and isopods, can be maintained satisfactorily in the laboratory provided there are facilities with running seawater or aquaria with aeration and filtration systems. Intertidal species should be provided with stones or other objects onto which they can climb. The authors have kept fiddler crabs and isopods (Sphaeroma quadridentatum and Ligia exotica) for several months to more than a year in small aquaria with slowly recirculating filtered seawater, feeding them small amounts of rolled oats. The Sphaeroma isopods, however, lost their infestation by Palavascia sphaeromae under these artificial conditions (Lichtwardt, 1961b).

Most terrestrial arthropods can be kept in terraria for prolonged periods of time. Many species of millipedes and pill bugs (Armadillidium, etc.) will go through one generation after another if provided with ample decomposing leaves and wood, and will maintain their trichomycete infestation. The substrate must be kept moist, but not wet. Passalid beetles can survive for more than a year on decomposing wood. When collecting these kinds of arthropods, extra supplies of their natural substrates can be gathered and kept in plastic bags, and added to the terraria as needed. Substrates obtained from gardens or urban sites are sometimes contaminated with pesticides and should be selected with caution.

All hosts of trichomycetes can be satisfactorily preserved in 70% ethanol for use as vouchers or for subsequent identification. Specimens that have been dissected may be used for identification provided essential anatomical parts are not missing. Generally, however, it is better to preserve undissected specimens from the same collection for this purpose.

We know of no satisfactory method to kill and preserve hosts that are to be dissected and studied critically at a later date. Fixatives such as alcohol or formalin-alcohol harden the host tissues making dissection considerably more difficult, as well as causing the thalli of trichomycetes to become brittle and subject to damage, especially if they are intermixed with undigested contents of the gut lumen. Later complete dissections are done more easily if the guts are removed from the host and fixed in 10% lactophenol (Amman's medium). Reasonably satisfactory slides can be prepared from such material stored for a few weeks.
 

Dissection Techniques

Several methods can be employed to dissect the guts of arthropods, depending on the investigator's individual preference and objective. The methods described in this section have been found to be satisfactory for general trichomycete studies.

The digestive tract of arthropods is divided into three principal parts, which differ considerably in their anatomy, relative length, and functions among various groups of arthropods. The hindgut and foregut arise embryologically from the ectodermal layer, whereas the midgut has an endodermal origin. The hindgut and foregut are thus an invaginated extension of the exoskeleton, and are lined with a noncellular substance composed largely of chitin. It is to this cuticular layer, produced by the underlying epithelial cells, that the thalli of trichomycetes are normally attached. As arthropods grow in size or metamorphose and shed their outer integument, they also shed these gut linings including any attached thalli.

The midgut is the primary region of digestion and assimilation in many arthropods. In some groups the midgut is lined with a chitinous peritrophic membrane. Two major types are recognized. One of these is produced more or less continuously from a ring of cells at the anterior end of the midgut, and it consists of a transparent tube that is unattached to the epithelial cells of the midgut except at its origin. This type of membrane is found in several families of dipteran larvae and is the site of growth of most of the Harpellaceae. Young thalli attach and begin to grow at the anterior end of the peritrophic membrane and are slowly shifted posteriorly as the membrane grows, with the result that mature sporulating thalli occur near the posterior end of the membrane. Peritrophic membranes in some Diptera may extend some distance into the hindgut, where the membrane disintegrates. Dissection methods to recover the peritrophic membrane will be described later.

Fungal growth in the foregut region can be found in species of two genera of Eccrinaceae, Arundinula and Enteromyces. Their hosts are crabs, anomurans (shrimplike crabs), and crayfish. Species of Arundinula occur in the hindgut as well. The anomurans and most crabs live in marine or brackish habitats, consequently, they should be dissected in seawater or seawater diluted up to 50% with freshwater. Care should be taken that evaporation does not result in excessive buildup of salinity during dissection procedures. The carapace can be cut dorsally with scissors along two parallel lines, one on either side of the center, beginning near the eyes and extending down the abdomen to the telson. Removal of the skeletal strip between the cuts will expose the large stomach and the hindgut. The abdomen of crabs is folded ventrally under the body. It can be lifted backward and torn off; this piece will include the entire hindgut. Stomachs and hindguts can be excised and cut open using fine iris scissors with the aid of a dissecting microscope and flushed with seawater to remove intestinal debris. Crayfish and freshwater crabs should, of course, be dissected in freshwater. Preparation of the fungal material for microscopic examination will be described in the next section of this chapter.

The remaining genera of trichomycetes (except for the external species, Amoebidium parasiticum) live in hindguts. Those of millipedes are easily and quickly exposed by cutting off the anterior portion of the animal about one-quarter of the distance down from the head with a sharp razor blade or scalpel, then cutting off the last couple of segments near the anus. The posterior part of the hindgut is grasped with forceps and pulled out of the body. Alternatively, the gut can be removed by making lateral incisions on both sides of the body (Reichle, 1978). The junction of the hindgut with the midgut can be recognized by the presence of a sphincter muscle and by the zone of attachment of the Malpighian tubes. Starting with the anterior end, the hindgut is cut open carefully with fine scissors so as not to penetrate too deeply, and is flushed with water. In addition to Eccrinaceae, there may be numerous filamentous bacteria, protozoans, or nematodes within the hindgut. Thalli of Enterobryus spp. may attach to the cuticle of nematodes in some species of millipedes. Many Eccrinaceae select particular regions of the hindgut for attachment and growth, such as the very anterior part, whereas others may be found throughout its length.

The hindgut of beetles can be cut out of the abdomen after removal of the elytra and wings. However, the authors have found it possible in most instances to remove their hindguts by firmly grasping with forceps and pulling at the anal plates of the intact beetle. Coleoptera often have looped hindguts that are structurally complex and long. Most isopod hindguts can also be removed by placing the host on its back and grasping the anal region and pulling. It may be desirable first to tear off some of the pleopods in those isopods in which they are large. With amphipods and nymphs of mayflies and stoneflies, the hindgut can be obtained by grasping the posterior segment and associated appendages and pulling; the unwanted parts can then be torn away from the hindgut with fine forceps. Some of these guts are too small to be cut open with scissors, and one may wish to use a minuten insect needle mounted in a handle, or some similar instrument of small size.

Aquatic larvae of blackflies, mosquitoes, chironomids, and other Diptera are among the smallest hosts of trichomycetes. Some measure only a few millimeters in length. The Collembola can be a special challenge for the dissector. Despite the small size of such insects, with experience it is often possible to prepare slides for microscopy in less time than it takes with some of the larger hosts. Dissections should be made under the lenses of a dissecting microscope with variable magnification. Either incident light oriented onto a dark, opaque background or light coming from beneath a glass stage can be used. The most valuable tools are high-quality fine pointed jeweler's forceps and micro dissecting needles. The head and anal segment are cut off with a sharp razor blade, and the gut is removed from the posterior end of the body. The dissection should be made in a drop or two of water in a petri dish or on a slide.

With Diptera, careful removal of the hindgut often withdraws the peritrophic membrane as well, or the membrane can be withdrawn by careful removal with the head region. The peritrophic membrane may contain masses of algae and other ingested materials. These are conveniently removed by grasping one end of the peritrophic membrane with forceps and lifting it several times from the water until cleared of all loose materials. The membrane is sufficiently transparent that it need not be cut open, and it can be mounted directly on a slide for microscopic examination. It is necessary to remove the epithelium of the hindgut, however. A simple way to accomplish this with most small dipteran guts is to grasp the hindgut at one end with forceps and pull it gently through the partially spread tips of another pair of forceps so as to strip off part of the epithelium. The remaining epithelium can be stripped off by grasping the other end and repeating the procedure. If desired, the unopened gut can be examined microscopically for the presence of thalli before final preparation. The guts can be either torn open carefully with two pairs of forceps, or a micro needle can be used while one end of the gut is held in place.

Most small thalli of trichomycetes can be seen in opened guts at magnifications of  x30 or lower, but because of their lack of pigmentation they do not always stand out against the background of the gut lining. With larger guts, where the chitinous lining is not easily separated from the epithelial tissue for quick examination at higher magnifications, it is possible to overlook small thalli. Before discarding apparently uninfested material, a drop or two of lactophenol cotton blue can be placed on the inner surface of the gut and then flushed off with water after a few seconds. Trichomycete thalli, if present, will stain sufficiently to be seen against a white background.
 

Preparation of Specimens for Microscopy

Standard methods for fixing and staining trichomycetes can be used for light microscopic studies, of course, but we prefer to study living material with phase-contrast microscopy for most purposes. This produces fewer artifacts and has the additional advantage of permitting limited stages of development to be observed over time in some of the unculturable species. Water mounts on microscope slides may show maturation, release, or germination of spores in some species of Harpellales, Asellariales, and Eccrinales; or amoeba release, locomotion, and encystment in the Amoebidiales. Water mounts can be kept in moist chambers one to several days before all development ceases or the thalli begin to deteriorate. Cysts of the Amoebidiales can be kept for longer periods of time so as to obtain cystospores and cystospore release.

Arthropod molts and small intact gut linings with the epithelial layer removed can be mounted on slides in distilled water. Trichomycete material from some marine hosts can also be mounted in distilled water often without undesirable osmotic effects, although 50% seawater may be preferable if developmental stages are sought. With some larger guts, it may be difficult during the intermolt stage to separate the entire lining from epithelial tissue. If the lining adheres tenaciously, small pieces of gut lining can be peeled off for mounting, or individual intact thalli can be removed by careful manipulation with a fine needle.

It is sometimes desirable to be able to prepare slides containing all stages of the fungus from one gut. The following technique works well with foreguts and hindguts whose linings cannot otherwise be removed easily. The gut is placed to one side in a 100-mm petri dish with sufficient water to cover it. About 20 drops of full-strength lactophenol (~1 drop/ml of water) are added to the inside of the dish opposite the gut and allowed to diffuse through the water. After a few hours, or overnight, the lining usually can be peeled from the tissues with ease. It should be rinsed in dilute (10%) lactophenol, and the excess liquid drained off by holding the lining momentarily against the side of the dish, before spreading the lining on a slide in full-strength lactophenol with cotton blue. As much proteinaceous material as possible should be removed before final mounting, because its coagulation in lactophenol can produce a slide of inferior quality.

Linings or thalli can be mounted directly from water into lactophenol cotton blue, using normal procedures for making slides of fungi. The coverslips can be sealed with clear fingernail polish or other preferred sealant. Small guts or thalli mounted in water on a slide can be permanently preserved without disturbing the delicate thalli by allowing excess water to evaporate from the edges of the coverslip, then placing a drop of lactophenol cotton blue on one edge and allowing it to infiltrate passively under the coverslip. The three clean edges can be sealed, and after the sealant has hardened, the edge with excess lactophenol can be washed with running water, dried, and sealed. This technique is especially useful when specimens have been studied with phase-contrast microscopy and photomicrographed, because the structural components can be relocated after a permanent slide has been prepared.

The use of fixatives and stains other than lactophenol cotton blue may be necessary for special cytological studies with the light microscope. When these are employed there is a tendency for the nonsporulating parts to plasmolyze or to become twisted during fixation or dehydration, or long thalli (those of some Eccrinales measure more than 1 cm) may break during the embedding process. For these reasons, the use of lactophenol cotton blue is recommended for routine permanent slide preparation.

In recent years, electron microscopic studies have provided valuable information on the biology and phylogeny of the trichomycetes. Preparation procedures for transmission electron microscopy are similar to those generally used with other fungi, except for the dissection of hosts and selection and orientation of suitable material to be sectioned. The quality of fixation may vary depending on the trichomycete species, morphological structure, and the fungal condition at the time of fixation. However, good results have been obtained by several investigators whose studies are cited elsewhere in this book.

Morphological structures to be processed for electron microscopy should be dissected with special care and with a minimum of handling. Larger guts must be cut open to expose the fungi, and either the entire gut or pieces of the gut or its lining can be fixed. Peritrophic membranes and small guts that have had the epithelial layer removed from the lining for visibility need not be cut open, but should be cleansed of all loose detritus in the lumen. Aquatic hosts often contain minute grains of sand and diatoms in their guts that may interfere with sectioning. Removal of unwanted substances can be accomplished by lifting the lining through the surface of clean water several times. Where damage to delicate thalli may be a factor, it is preferable to handle the material as little as possible, even at the expense of not cleaning it.

Good fixation is possible with a 2% glutaraldehyde:3% acrolein mixture in 0.1 M cacodylate buffer at pH 6.8 for 2-3 hr at room temperature, postfixing in 2% osmium tetroxide in the same buffer for 2 hr at 0–4'C. Some trichomycete material will fix well in freshly prepared 1% aqueous potassium permanganate for 1 hr in the dark at room temperature. The chitinous lining of arthropods is often hydrophobic, or may tend to float, owing to air bubbles that become entrapped in folded portions of the lining. Consequently, one should insure that the material in fixative does not float. Sometimes it may be necessary to place it under vacuum for a brief period. Dehydration should be gradual, such as 15 min in each of 10% steps in a graduated ethanol series.

Shallow polypropylene dishes or special flat-embedment plates are useful for the final embedment in plastic. This permits microscopic examination and better selection and orientation of material to be sectioned. Selected portions can be cut out and cemented to short plastic stubs of suitable diameter to fit the microtome chuck.

Scanning electron micrography can be very informative, as with other biological subjects, and its use with trichomycetes probably will increase. Once the proper material to be studied has been located, it can be fixed in glutaraldehyde, permanganate, formalin-aceto-alcohol, etc., then dehydrated, critical-point dried, and coated in the conventional manner. The technique of freeze-fracturing has not been employed in published trichomycete studies to date.
 

Axenic Culturing

Axenic cultures of trichomycetes have led to a variety of studies involving their physiology, morphogenesis, phylogeny, and host relationships. Unfortunately, relatively few of the total known trichomycetes—all in the Harpellales and Amoebidiales—have been available for this kind of experimental work. The majority are species of the largest genus of Harpellales: Smittium. Currently available isolates maintained in the University of Kansas Culture Collection are listed on this web page, together with host and isolation data. The section here deals primarily with the methods that have been employed to obtain primary axenic isolates of trichomycetes, whereas the results of experimental studies using such isolates are covered in Chapter 9.

It is perhaps not surprising that the first species to be cultured was Amoebidium parasiticum, an ectocommensal trichomycete with a wide host range. This was accomplished by Whisler (1960), who retrieved sporangiospores released from thalli attached to Cladocera sp., washed them several times in sterile pond water, and streaked them on 0.1% tryptone agar. In 1962 Whisler published a study of the nutritional requirements of Amoebidium in axenic culture, and obtained maximum growth in a thiamine-enriched tryptone-glucose-salts medium. He was also able to grow the organism in a defined medium by substituting methionine for tryptone.

Whisler's success at axenic culturing was followed by that of Clark et al. (1963), who obtained from mosquito larvae the first endocommensal pure cultures. Clark et al. surface-sterilized larvae prior to removal of the hindguts, and used antibiotics in the dissection water as well as in a series of washes before plating on a blood agar medium (SNB-9) used for culturing hemoflagellates. By this method they were able to isolate Smittium culicis and a Smittium species that we have identified as S. culisetae. Subcultures grew also on Difco brain-heart infusion agar with a 2% neopeptone overlayer and in NIH thioglycolate broth. However, they reported that trichospore production occurred only after 1 month of growth in the initial isolates, a remarkably slow development. Subsequent studies (El-Buni and Lichtwardt, 1976a) have shown that the use of brain-heart infusion at the concentration recommended by the manufacturer, and certain other cultural conditions, are inhibitory to spore production in Smittium spp.; see Chapter 9.

Lichtwardt (1964a) reported the isolation of two new species, S. culisetae and S. simulii, from a mosquito and a blackfly larva, respectively, using dilute brain-heart infusion and potato dextrose-yeast extract agar media. Since that time he and other researchers have obtained a large number of isolates of Smittium and other genera of Harpellales from several families of dipteran hosts. The simplest methods that have been successful in the authors’ experience are described below.

When these media are used with agar in petri dishes, they should be flooded with a shallow layer of sterile distilled water before inoculation. In agar slants, about 1 ml of distilled water is added to the test tube, and the tube is tipped daily so as to flood the agar surface during initial stages of growth. We also add thiamine and biotin to BHI/10 medium in most instances, but it has not been determined that these vitamins are actually required. However, supplemental thiamine has been shown to be stimulatory to growth (see Chapter 9).

The antibiotics most commonly used have been prepared as a single stock solution consisting of 40,000 units of Penicillin G and 90,000 units of Streptomycin sulfate per milliliter of distilled water. It can be filter sterilized and kept in serum bottles in a refrigerator for 6 months or longer, and is dispensed with a sterile syringe.

For isolating Smittium species, the hindgut is dissected from a dipteran larva and the epithelium is removed. Microscopic examination of the unopened gut lining mounted in water with transmitted light will usually reveal whether or not Smittium is present. The fungus can then be processed in this condition, or the lining can be torn open carefully to release debris from the lumen. Individual thalli can also be removed and used, but greater care is necessary in handling and the results usually have not been as good as when the entire hindgut lining is used.

Amoebidium parasiticum is located on external parts of various aquatic crustaceans and insects, and hosts with mature thalli should be selected. Animals like small cladocerans or first instar mosquito larvae can be used intact. Parts with Amoebidium on larger arthropods (the antennae, anal papillae, etc.) should be excised and used for culturing.

Washing should be done in small containers with antibiotics. The authors prefer 35 x 10 mm sterile plastic petri dishes partially filled with water to which is added 0.05 ml of Penicillin-Streptomycin stock solution per dish. When bacteria are abundant, larger amounts of stock antibiotics, as high as 0.2 ml per dish, can be used without apparent harm to the fungi. The material to be isolated is placed in this wash water for 15-60 sec and transferred serially and aseptically through two or more washings, then to the agar medium with a water overlayer. Antibiotics may be avoided at this stage, or concentrations up to 0.2 ml per dish may be added to the water. It is generally best to vary the concentration in replicate cultures. The use of 60 x 15 mm plastic petri dishes for the initial isolations allows one to examine the inoculum under a x5 microscope objective (x50 magnification) to check on growth and contamination (Fig. 3.1). If growth occurs it is usually evident within 2-4 days. Part of the growing clump of fungus should be transferred to new medium without antibiotics. Transfer of Smittium can be done with a small metal loop, but a Pasteur pipette or micropipette may be desirable for spores and young thalli of Amoebidium that have been released onto the agar surface. Careful monitoring for contamination during the first few days may permit contaminated material to be saved, before it is overcome by bacterial growth, by rewashing in antibiotics and replating on medium with antibiotics. Contamination by yeasts and filamentous fungi is generally more difficult to control, but is less common. Such contaminants can sometimes be diluted out by serial washings of the trichomycete material.

The authors have attempted to culture many other genera of trichomycetes from all four orders without success, and many other investigators have made attempts with some of the genera (Lichtwardt, 1954, 1964; Manier, 1954, 1955; Whisler, 1963; Chapman, 1966; Williams, 1971; El-Buni, 1975). Of course, most unsuccessful results are not reported in the literature. Many nutritional and physical parameters that might be significant to these gut fungi have been taken into consideration. It is obvious that the axenic culture conditions that will support growth of Smittium and Amoebidium are less demanding than for other genera. This is emphasized by the fact that the authors on many occasions have attempted to isolate species of genera such as Genistellospora, Pennella, and Paramoebidium growing with Smittium spp. in the same blackfly (Simuliidae) host, and only Smittium has grown.

In 1971 Lichtwardt was able to make about a dozen isolations of Trichozygospora chironomidarum from chironomid larvae collected in Abisko, Sweden. Two of these survived for as long as 18 months, during which time they were transferred repeatedly and grown under a variety of conditions without ultimate success. Likewise, he obtained very limited axenic growth of Enterobryus sp. from the fiddler crab, Uca crenulata, but only over an 8-week period (Lichtwardt, 1964). In addition to the probability that most trichomycetes have unusual and as yet undiscovered requirements for vegetative growth, there is another factor that must be considered. Extrapolating from data obtained with Smittium cultures and from other genera removed from hosts, trichospores and other kinds of spores do not ordinarily germinate except within the host gut (see Chapter 9). Therefore, thalli that cease to grow vegetatively when sporulation commences, such as the unbranched Eccrinales and Harpellaceae, will cease to develop further when their spores mature if those spores are incapable of germination under the culture conditions used. The factor(s) that induces spore germination may therefore have to be determined and incorporated into the axenic culture conditions in order to permit such trichomycetes to complete their cycles.

Only a few of the many isolates of Harpellales produce spores that germinate in vitro. Consequently, when transferring these fungi to dishes or test tubes of new medium the colonies should be broken up sufficiently to provide new clones. This can be done with a small loop when working on agar media. For experiments using liquid medium, young cultures can be chopped in a Waring-type blender to obtain a homogeneous inoculum as is done with other filamentous fungi.

Cultures can be stored on slants of medium in a refrigerator after growing at room temperature for 7-10 days. It is recommended that stock cultures be transferred to new slants every few months, but some refrigerated isolates may survive for up to 1 year or more. Freeze-drying Smittium and Amoebidium cultures has not been successful. The preferred method for long term storage is in liquid nitrogen.

The value of using axenic isolates to help elucidate the biology of the trichomycetes has been considerable, despite the few taxa that have been available. Many fundamental questions raised in the succeeding chapters remain unanswered, however. Their solutions will be greatly facilitated by, or even require, the axenic cultivation of species that are at present unculturable.
 

Preparation of DNA for molecular systematics

Modern phylogenetic studies in most organisms have come to rely on various techniques for analyzing DNA. In trichomycetes, genomic DNA can be isolated either from cultured material (obtainable only from certain species of Harpellales and Amoebidium parasiticum) or from thalli dissected from the host gut. The latter requires PCR amplification and usually cloning to select appropriate regions of DNA. The techniques used in purifying, processing, and sequencing have been basically those used with other fungi. The co-authors of this monograph have found that commonly used fungal primers used for the 18S and 28S rDNA genes usually are not sufficiently specific for amplifying the DNA of trichomycetes taken directly from arthropod guts, because often host tissue or other gut microorganisms are amplified. Consequently, they have developed several specific primers for different taxa of these unculturable fungi (currently unpublished).

The most commonly sequenced gene has been the nuclear small subunit (SSU) 18S rDNA (O’Donnell et al., 1998; Benny and O’Donnell, 2000; Benny, 2001; Gottlieb and Lichtwardt, 2001), using equally weighted maximum parsimony analysis of the dataset with PAUP* software for constructing trees. Sequencing is being done currently in the authors’ laboratory for as many species of unculturable Harpellales, Asellariales, and Eccrinales as can be obtained in order to construct cladograms to determine the affinity of different orders of trichomycetes (Chapter 12).