Isolating fungi from natural substrates
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The same day as the lab presentations on mushrooms, we will be setting up moist chambers from
which we will be isolating mold fungi. See below. If you have a substrate you would like to try,
bring it along! Otherwise (and in addition) we will be using herbivore dung.
1. Set up for mould isolation from natural materials -- Moist chambers
Direct isolation of fungi is often more effective if the natural substrate has been kept moist for
one to several weeks to allow moulds to grow and sporulate. The easiest method involves a
container called a moist chamber (Figure 11). Moist chambers can take any number of forms, but
are basically containers holding a material such as cotton, paper, cloth, sterile sand or soil, or peat
moss that can be kept moist for several weeks. The specimen is placed on top of the moist
material and left until moulds begin to grow on it. You can use glass containers resembling very
deep Petri dishes, and like Petri dishes they have loosely fitting lids or, to save washing, clear
plastic food grade containers with snap lids. For the packing material we nearly always use peat
moss, collected in the field and dried for later use. The moist chamber container is part filled with
moistened peat moss. Peat moss is like a sponge in that it can absorb and hold a tremendous
amount of water. When water is added to the moist chamber it soaks up into the moss. The excess
can be poured off and the moss squashed down until it forms a moist layer no deeper than about
one-quarter of the height of the container. We then place a single or double layer of filter paper or
paper towel over the moss layer so that the specimen does not come in contact with it. Thus
prepared, the moist chamber is ready for the specimen.
In many areas peat moss, in its fresh form, is hard to obtain and it is necessary to find a
substitute. Although many people use cotton for this purpose, I think that it is best avoided as it is
a good substrate for mould growth itself. Instead, try using a layer of sand. This will not hold
water as well as peat, so watch carefully to make sure it does not dry out. But don't make it too
wet or all you will get is mud!
When you are ready to add the specimen, moisten it slightly (unless it is already damp)
and place it on top of the filter paper. Cover the dish and leave it in a place where the temperature
is reasonably constant.
Within a few days moulds will begin to appear on the specimen. Most beginners are
unprepared for the extremely small size of many moulds and tend to overlook them completely.
Often students using moist chambers for the first time complain that nothing is growing on their
specimens, only to have an instructor point out at least half a dozen different moulds! Be sure to
examine the material with a magnification of at least 15-20 times and with good illumination.
Illumination is especially important and should be focused on the area of the specimen that is
under examination. When something interesting is found, it can be removed for microscopic
examination; but more about that part later (lab 2 and appendix 2 on media and sterile technique).
 A moist chamber containing three pieces
of dung
In order to obtain a pure culture of something
in a moist chamber it is only necessary to
follow the direct transfer procedure given
above, making sure that the sterile needle
picks up only the spores that are wanted. If the needle inadvertently touches the surface of the
material in the moist chamber it will pick up all sorts of undesirable contaminants.
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Moist chambers can be used for all kinds of materials. We have used them to incubate
dung, wood, leaves, old stems, corn stalks, bark, seeds, fruits, old fungi, dead insects, and
numerous other things. Natural materials are usually more productive than man-made ones, but
sometimes old pieces of cloth or leather products can be good. You are only limited by your
imagination. Moist chambers are also useful in mycological "detective work", to discover the
cause of a particular decay. Placing the decayed material in a moist chamber often will result in
abundant sporulation of the guilty organism in the area affected. This technique works well with
diseased plant material as well as manufactured products.
With very small specimens, such as insect parts or seeds, it may be easier to use a Petri
dish as a moist chamber. We sometimes simply put a few layers of filter-paper in a Petri dish,
moisten them, and put the specimen on top. Such moist chambers may dry out very easily,
however, and have to be tended closely. A better method is to make up a batch of water agar (a
solution of 20 grams of agar in a litre of water), sterilize it, and pour it into sterile Petri dishes.
Because the agar solution contains almost no nutrients it will support little mould growth and
thus serves only as a water reservoir. The specimen can be placed on the agar surface and
examined as in any other moist chamber. Agar media containing nutrients can be used here, but
often they only become overrun by the fastest growing moulds at the expense of everything else.
Whether using a moist chamber or working with freshly collected material, one often
discovers that moulds produce spores inside some kind of structure, such as a pycnidium. Direct
transfer of these structures invariably leads to contamination by other moulds. To avoid the
problem it is necessary to clean off the surface of the structure sufficiently that all foreign spores
and bacteria are removed. I do this either by pushing the structure over, around, and through an
agar medium for 10 minutes or more until it is clean (you’ll be doing this when cleaning
Aspergillus cleistothecia), or by surface sterilizing it for about 60 minutes in a 10 per cent
commercial chlorine bleach solution. The required times and concentrations of bleach in the latter
technique will differ some what from one specimen to another; the technique is most effective if
the treatment can be varied with several samples. The agar technique demands more dexterity but
can be applied to a single specimen with predictable results.
Direct plating
Often it is most convenient to place materials that are of interest directly on a nutrient agar
medium. As already noted, this technique encourages rapidly spreading moulds at the expense of
other fungi but is nevertheless widely used. It is a simple technique, requiring the placing of
small bits of the substance on the surface of the agar or the pouring of melted but cooled agar
over the fragments. After a few days' incubation mould colonies appear on the surface, and can
be transferred into pure culture.
The amount of material placed in the dish varies, depending upon how heavily it is
infected with mould spores. This technique is commonly used in soil studies, requiring only a
pinch of soil, evenly dispersed over the surface of the agar. It is difficult to apply any rules here,
however, because a small amount of material may yield a different set of moulds from a large
amount. If the amount of material is large, the result may be a combination of a plating technique
and a moist chamber.
We find that Martin's Rose Bengal Medium is a good choice for direct plating, as the
Rose Bengal dye and antibiotics in it slow down colony growth and keep the colonies from
growing together at first. We will be using Rose Bengal agar for primary isolations from the dung
cultures, since it tends to keep fast-growing zygomycetes under control.
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Be It Ever So Humble, There's No Place Like Dung
Juliana T. Hauser, Microbiology Department, Carolina Biological Supply Company
Although many people shun dung as a subject fit for study, it is a marvelous resource that is
constantly being produced and deposited in large quantities at convenient locations. Even though
it has passed through an animal's digestive tract, dung retains many nutrients. Thus, it attracts its
own fauna and flora consisting of bacteria, fungi, protozoa, platyhelminths, nematodes, annelids,
and arthropods. Coprophilous (dung-loving) fungi are uniquely adapted to herbivore dung. They
are deposited with dung, and they grow and reproduce there. They disperse their spores from the
heap to a location from which they will be consumed by a herbivore, pass through its gut, and
again be deposited with the dung heap.
Dung as a Home
Dung consists of the macerated and undigested remains of plant food plus vast quantities of
bacteria (mostly dead) as well as animal waste products, such as broken-down red blood cells and
bile pigments. The nature of herbivore dung depends on the efficiency of the digestive tract,
which, in turn, depends on the animal's digestive anatomy and its microflora. Ruminants produce
fine-textured dung of fibrous plant material whereas horses, with a less efficient system, produce
much coarser dung. Dung decomposes rapidly because the macerated material has a high
nitrogen content, available aeration, and a high water content that is protected from fluctuations.
Although there is little available protein in dung, many other undigested food components
are present. Dung is rich in water-soluble vitamins, growth factors, and mineral ions, some of
which are metabolic by-products of the microbes in a herbivore's gut. For example, coprogen, an
organo-iron compound found in dung, is necessary for the growth and reproduction of the fungus
Pilobolus crystallinus. Dung also contains a large amount of readily available carbohydrates.
Fungal Dung Inhabitants
Considering the great variation in the feeding habits, habitats, and digestive systems of
herbivores, it is surprising how universal coprophilous fungi are. All classes of the Kingdom
Fungi are found on dung, with the Zygomycetes usually appearing first, followed by the
Ascomycetes, and finally the Basidiomycetes. Their distribution is influenced locally by the
number of herbivores in an area. Some species are restricted to a particular herbivore; for
example, Lasiobolus cainii is found only on porcupine dung. However, many coprophilous fungi
grow indiscriminately on any herbivore dung. The greatest variety of fungi have been reported on
cow, rabbit, and horse dung, but this could be because the majority of research has focused on
these animals.
Adaptations of Fungi to Dung
Coprophilous fungi are highly specialized for growth on dung, and some never occur elsewhere.
While some dung fungi show few modifications peculiar to their habitat, most do have some
unique features. Many exhibit some very specialized structures to ensure survival in their unique
habitat.
Herbivores do not graze near their own dung; therefore, the spores must be propelled
beyond this "zone of repugnance." Thus, the spores or spore masses are relatively large and
heavy. In the Zygomycete Pilobolus, for instance, the entire sporangium is discharged as a unit.
In the bird's nest fungus, Cyathus stercoreus, the peridioles (the "eggs") containing many spores
are violently discharged when a raindrop hits the peridium (the "nest"). The spores/masses,
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because of their weight, do not remain in the air long, but follow a parabolic trajectory landing on
nearby grass without the aid of air currents. The sporangium and sporangiophore of Pilobolus
measure about 0.5-1.0 cm, yet the sporangium has been propelled as much as 1.8 m vertically and
2.1 m horizontally.
Some coprophilous fungi exhibit a phototropic response that determines the direction the
spore mass will be projected and ensures that the spores clear the substrate. Spore discharge is
always during the day. The entire sporangium of Pilobolus, for example, grows toward the light
source. To demonstrate this phototropic response, place the mature culture of Pilobolus inside a
container with a hole punched into one side of the container top. Wrap the container inside
aluminum foil and punch another hole in the foil aligned over the container hole. Place
transparent tape over the hole, and set the container in a window. The following day, remove the
foil and container. The majority of the ejected sporangia will be found stuck to the tape or around
the light source in the container top.
The spores are dark to protect them from ultraviolet light until they are consumed by a
herbivore. In some fungi, melanin is present in the spore walls; in others, a dark membrane
covers the spore mass. Coprinus comatus, a mushroom that fruits on dung, exhibits these dark
spores.
The spores/mass are often mucilagenous so that they stick to vegetation upon impact, and
the mucilage, when dry, cements firmly. In Cyathus, the peridiole has a sticky piece of hypha, the
funiculus, which attaches to vegetation upon impact and wraps the peridiole firmly around it.
Other coprophilous fungi, such as Mucor hiemalis, form a sticky droplet around their spores.
When an insect visits the dung, the spores stick to the insect's body. If the insect rests again on
other vegetation or another dung heap, the spores rub off and adhere to the new environment.
Many of the coprophilous fungal spores will not germinate until after passing through an
herbivore's digestive tract: They must be heated inside the gut, digested by the gut enzymes
and/or bacteria, or stimulated by the higher pH of dung.
Fungal Dung Succession
Some observers have noted a true ecological succession on dung in that first the Zygomycetes,
then the Ascomycetes, and finally the Basidiomycetes appear. Early researchers suggested that
this succession was a nutritional one. They postulated that the Zygomycetes, or sugar fungi,
appear first, because their spores germinate quickly and their mycelium grows rapidly, exploiting
the fresh substrate; that is, they utilize the simple sugars and hemicelluloses present in dung. The
Zygomycetes are not capable of utilizing the cellulose and lignin found in dung. When the
simpler carbon sources are metabolized, the Zygomycetes disappear and are replaced by the
Ascomycetes, which can utilize cellulose. These are then replaced by the Basidiomycetes, which
can utilize both the cellulose and the lignin.
Although this nutritional hypothesis of dung succession was an attractive one that seemed
to fit the observations, it ignored some important ecological and physiological facts. For example,
some spores had already germinated or had their dormancy broken in the herbivore gut. Also, no
scientist had yet considered the interference/competition/enhancement effects of the various dung
inhabitants on each other.
The nutritional hypothesis was based solely on the order of appearance of fruiting
structures. Mycelial development and growth may not have been proceeding at the same rate, so
a second hypothesis, the reproduction hypothesis of dung succession, was developed. This
hypothesis was based on the time it takes each kind of fungus to fruit. The simple sporangia of
the Zygomycetes could develop much more quickly and require much less energy than the more
Isolating fungi from natural substrates
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complex Ascomycete fruiting body. The even larger Basidiomycete fruitification would require
the largest amount of energy expenditure and would, therefore, take the longest amount of time to
appear. This hypothesis still neglects the interrelationships between fungi themselves and
between fungi and other dung inhabitants.
The dung heap is not inhabited exclusively by fungi. As mentioned previously, vast
populations of bacteria, protozoa, platyhelminths, nematodes, annelids, and arthropods coexist
with the fungi. These compete for resources, and some parasitize or consume the fungi while
others provide substrates for them. Also, these organisms can deplete the dung of nutritionally
necessary compounds, such as nitrogenous ones, and can produce waste products that enhance or
retard the growth and reproduction of some fungi. For instance, ammonia, a waste product of the
bacterial degradation of proteins, stimulates sporangial production of Pilobolus, which grows
better in the presence of other microbes.
Coprophilous bacteria enhance the growth of some of the coprophilous fungi, but retard
the growth of others. Also, the number of fly larvae, Lycoriella mali (Diptera: Sciaridae) that
survive to pupate increases as fungal competition increases. It could be that as fungal growth is
inhibited, more resources are available to the fly larvae, or that larvae may be favored by the
enzymes produced by fungi to inhibit other fungi!
There is evidence that some fungi excrete products that inhibit fungal competitors. A few
weeks after dung is deposited, the Zygomycetes and Ascomycetes disappear, but the
Basidiomycetes continue fruiting for months. Obviously, the substrate has not been depleted.
Some Basidiomycetes suppress some of the Zygomycetes and Ascomycetes by hyphal
interference. Within minutes of contact, the sensitive species' hyphae undergo vacuolization and
lose turgor. This is followed by a drastic alteration of cell membrane permeability and subsequent
death of the hyphae.
In the classroom, dung succession is easy to observe and is sure to promote the interest of
students. Fresh or weathered dung can be placed in empty containers. If the dung is dry, add a
small amount of distilled water. Plastic wrap secured with a rubber band will allow observation
and prevent evaporation of the necessary moisture. The dung culture should be observed over a
period of three to four weeks to study fungal succession.
The success of coprophilous fungi, as measured by their ability to produce and maintain a
fruiting body, is not simply a matter of competing against other fungi. Instead, it is a complex
web of interactions, some combative, some inhibitive, some mutually or exclusively beneficial
between and among the bacteria, fungi, protozoans, platyhelminths, nematodes, annelids, and
arthropods. It's a jungle in there!
Francis, S. M. 1988. Fungus guns 2. The Mycologist 2:78-79.
Ing, B. 1989. Why not look at the dung fungi? The Mycologist 3:33.
Kendrick, B. K. 1985. The Fifth Kingdom. Mycologue Publications, Waterloo, Ontario.
Safar, H. M., and R. E. Cooke. 1988. Interactions between bacteria and coprophilous
ascomycotina and a Coprinus species on agar and in copromes. Transactions of the British
Mycological Society 91:73-80.
Stanley. A. 1992. Fun with fungi. Address to the North Carolina Science Teachers' Meeting,
Charlotte, NC. Ms. Stanley is curator of the Fungal Herbarium Collection, UNC-Charlotte.