Exercise 3 (EX3) Enzymes produced by mold isolates
Lab 5
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Goals for this lab exercise and during the upcoming weeks. Consult appendices on sterile
technique, slide culture, dichotomous keys, media and staining, and glossaries.
1) Isolate fungi that are growing on decaying materials in moist chambers, using sterile
technique. You will have to check your cultures and your moist chambers periodically to
monitor for growth, sporulation, and whether you have isolated pure or mixed cultures. Lilia
and I will be available on request (email or during class time) to let you into 307 Biology, but
only in the mornings. Other courses use the lab in the afternoons.
1. Ultimately, you should generate a collection of at least 8 different asexual fungi (no more
than one zygomycete).
2. Determine the growth rate for each of your pure isolates, using the plate culture method,
reviewed later in this handout. You will be using your isolates in a degradation lab in a few
weeks' time.
2) Learn basic spore forms of asexual fungal spores by working with known and unknown
isolates that are provided in pure culture.
 Prepare adhesive tape mounts of conidia and conidiogenous cells for examination and
identification.
 During the first lab period, make sketches of conidia and conidiogenous cells for at least
three "knowns" and three "unknowns". We will provide pictorial and written keys, as well as
a glossary of standard terms to aid you in identification. Learn to use the key to identify these
isolates.
 Ultimately, you will be identifying your moist chamber isolates as Zygomycetes or
Deuteromycetes, and then determining the genus of as many isolates as possible. Where
unambiguous identification to genus is not possible, suggest plausible possibilities.
Background
Fungi grow on or in substrates, which can be solid, semi solid, or liquid. In natural settings,
substrates often have more than one physical characteristic, or the predominant characteristic
changes over time. For example, imagine a piece of fruit that has a small spot of decay. Initially
most of the fruit is firm, but as the decay organism(s) releases degradative enzymes the fruit
softens or even liquifies.
In natural settings (unlike a lab culture plate) mold fungi can occur as communities, at a very
small scale. Even a tiny area can support more than one species. However, most studies use pure
isolates of specific strains, and so it is essential to isolate individuals from the community. The
only practical way to get down to the small scale of individual moulds is to use a binocular
dissecting microscope with high enough magnification to allow spore-bearing structures to be
seen. Then, a sterile needle can be used to remove the spores of a given mould to a Petri dish
containing culture medium. This is called direct transfer (described below) which you will use to
isolate colonies for pure culture. Today we will be using media on which most species will grow
(potato dextrose agar, V8 juice agar) and a medium containing the dye, Rose Bengal, and an
antibiotic to slow down the growth of Zygomycetes and bacteria to assist in isolating slower
growing deuteromycete species.
Exercise 3 (EX3) Enzymes produced by mold isolates
Lab 5
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As we have discussed in lecture, hyphae from different fungal species can have distinctive sizes,
shapes (e. g., smooth vs undulating), and growth rates. However, the greatest structural variety is
seen in how different species reproduce, that is, in their sporulation. Our challenge for the second
part of this lab exercise, then, is to begin to learn the characteristic structures of deuteromycete
sporulation. You will be making adhesive tape mounts (described below). Using these mounts
and a compound microscope, you will begin identifying isolates.
Draw pictures of the conidia and the cells that form them – this will assist you in examining your
slides and retaining and accurate image. Remember that you will have to scan to find good
examples of various parts – seldom do these look like textbooks, at least not on the first attempt.
Dichotomous keys are an efficient way to guide you through the identification process. I have
devised a glossary of terms (all-important in this process) which will help you to interpret the
descriptions in the key.
Steps in making a direct transfer.
1. Wipe down your benchtop with 70% ethanol and allow it to dry. Wash your hands and tie
back long hair.
2. Label the underside of Petri plates containing appropriate medium with your name, the date,
the substrate from which you are isolating and, if possible, a brief description. An alternate
method is to give each of your isolates a number (for example, I might call mine SKj1, SKj2
and so on) and then record the your descriptions in your lab book. The purpose of the
description (and sketch, if appropriate) is to remind you what you were hoping to isolate. For
example, if you were hoping to isolate an orange mould, and your plate eventually grew
more than one colour, the description would indicate which was your intended isolate, which
you could purify from your "first-round" plate.
3. Avoid opening your Petri plate further than, faster than, or more often than necessary. Lifting
the lid creates currents that can pull spores inside, or can disturb spores growing on your
colonies. Both lead to contamination.
4. Flame a dissecting needle or inoculating needle (this is slightly finer) until it is red hot. Then
dip it into 70% ethanol to cool. Flame briefly to dry. Even though the needle will not be
glowing, it will still be very hot, and deadly to touch to a tiny cell. Lift one side of your
labelled Petri dish and cool the needle by touching it to the sterile agar surface of the medium
until the hissing noise stops. This has the additional function of wetting the needle so spores
will adhere more easily.
5. Select a colony that is producing spores and make sure that your sterile needle picks up only
those ones. Transfer the spores to the agar in your Petri plate. Remember that spores are very
tiny and you might not be able to see that you have transferred anything, only the area that
you touched during the collection. Close the plate with masking tape (not sealed around the
edges, just so that it cannot open accidently – you can accumulate a stack of plates and tape
as a group) and wait a few days for the mould to grow into a new colony. Because we
incubated our moist chambers at room temperature, you can leave your plates at room
temperature as well
Exercise 3 (EX3) Enzymes produced by mold isolates
Lab 5
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6. At this stage, you might have to do a second round of isolation to get pure colonies. We will
be available for you to check your plates and re-isolate if necessary.
Growth rates of your isolates (as per Ex2)
When you have isolated pure strains, you will need to determine their growth rate for a future
lab. To do this, inoculate a fresh Petri plate, noting the place on the agar and the time of
inoculation. Examine the plate once or twice a day, and mark the edge of the colony with a fine,
permanent felt-tipped pen; note the time for each growth mark. Use this data to calculate a
growth rate. When we do the degradation lab, you'll be able to use more than one isolate per
plate as long as they have similar growth rates.
Adhesive Tape Mounts
1.
Use two slides. To one, add a small drop of lactofuschin dye, and place a drop of 95%
ethanol on the other. The ethanol drop will need replenishing if you're slow with the next
two steps, and should be replaced between making mounts of different isolates.
2.
With forceps and scissors, cut a small (< 1 cm) piece of transparent cellophane tape that
doesn't have any fingerprints. Touch the sticky side gently to the surface of a fairly young
region of a fungal colony, and then remove it.
3.
Wet the adhering cells briefly in the drop of 95% ethanol, then hold the tape sticky side
down in the drop of dye. Turn the tape over, so that the cells are facing up, and use a
dissecting needle to dislodge the tape from the forceps. The tape should be floating in the
lactofuschin, with the sticky side up.
The purpose of the ethanol step is to help the lactofuschin "wet" the fungal structures. The
aerial portions of fungal hyphae are often coated by hydrophobic molecules, which the
ethanol rinse washes away.
4.
Add another drop of stain to the surface of the tape, and examine it under a coverslip.
5.
Slides made with lactofuschin will resist drying out for several days. They are made
permanent by sealing the edges with nail polish.
a. Be sure the glass is clean and dry. All oils and stains and mounting fluids and water
should be wiped away, and the surface wiped lightly with alcohol.
b. Paint a smooth thin band of nail polish around the edges, lapping at least a couple of
millimeters onto both the slide and the coverslip. Use colored polish, to see clearly
where you have put it.
c. Let the polish dry thoroughly. Apply a second coat, and dry again. Label your slide.
d. Be gentle if removing immersion oil from the coverslip – rubbing can dislodge it.
6.
If it is important for you to know the colour of the spores (although this will be obvious
from inspecting the culture on the plate) or other structures. We will also supply mounting
solution without lactofuschin, which will preserve your sample without staining it.
IMPORTANT! Adhesive tape is not sterile – any culture examined by this method can become
Exercise 3 (EX3) Enzymes produced by mold isolates
Lab 5
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contaminated. If you intend to examine an important culture this way, subculture it first, to have
an uncontaminated culture as backup.
Week 2 and 3
You should have at least six distinct deuteromycetes and/or zygomycetes isolated. For each
isolate you should have labeled drawings of representative structures with indications of their
sizes. You will have discovered that it is not always possible to come to an unambiguous
identification. In these cases, explain how you arrived at your diagnosis, the reason(s) for the
ambiguity, and which of the alternatives make the most sense.
Enzyme assays
Degradative enzymes can be assayed by growing up mycelium, grinding in buffer, and
testing extracts for enzymatic activities. Problems for survey level assay – enzymes might not
survive extraction, might be in low concentration; buffer characteristics must be optimized for
individual enzymes; cells will be homogenized which may lead to antagonistic interactions;
some enzymes are induced by their substrate and mightn’t be present
Alternative – grow fungal isolates on solid media containing specific complex substrates;
compare colony growth and assay substrate degradation. Note that the media contain a minimal
level of nutrients to support initial growth in case the enzyme is substrate-induced.
These are two part assays: grow the isolates on specialized media, and then (for some assays)
develop the plate to reveal substrate degradation. For all these tests, wait about 10 min after
applying the developing reagent for full colour to show. Note that lack of substrate utilization
will also reduce growth rate and/or hyphal density due to relative nutrient lack.
Pectolytic activity
Medium: 50 ml 20x nitrate salts, 1 ml trace elements, 1 g yeast extract, 15 g agar, 5 g pectin,
1000 ml ddHOH. Buffer to pH 7 to detect pectate lyase (pectate transeliminase) activity.
Buffer to pH 5 to detect pectinase (pectin depolymerase) activity.
Assay: flood with 1% aqueous hexadecyl trimethyl ammonium bromide (may need to warm
slightly to dissolve). HTAB is also called CTAB, and is a common part of plant DNA
isolation protocols. It precipitates intact pectin, leaving a clear zone around a colony with an
otherwise opaque medium.
Amylolytic activity
Medium: Difco Nutrient agar plus 0.2% soluble starch, pH 6.
Assay: Flood the plate with dilute iodine, about the colour of light maple syrup. Amylolytic
activity indicated by a yellow, starch-depleted zone
Cellulolytic activity
Medium: two-layered plates.
Bottom layer (16 ml): 50 ml 20x nitrate salts, 1 ml trace elements, 1 g yeast extract, 15 g agar,
1000 ml ddHOH.
Top layer (8 ml): 12.5 g carboxy methyl cellulose, 15 g agar, 500 ml ddHOH
Assay: flood plates with 10% copper acetate, which will reveal clear zones where soluble
cellulose has been degraded. Fibrous cellulose gives blue precipitate where degraded
Exercise 3 (EX3) Enzymes produced by mold isolates
Lab 5
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Lipolytic acticity
Medium: Autoclave the two solutions separately, and then combine:
3) 10 g Difco peptone, 5 g NaCl, 0.1 g CaCl2.2H2O, 15 g agar, 900 ml ddHOH, pH 6.
4) 10 ml Tween 20 + 90 ml ddHOH. After autoclaving, add the detergent solution (Tween 20 is
polyoxyethylene sorbitan monolaurate) to the sterilized and cooled medium. Mix very gently,
avoiding froth. Pour.
Assay: formation of lipases will cause a visible precipitate of Ca-laurate crystals, followed by a
complete clearing of the crystals due to continued degradation.
Write up:
This is lab writeup has two parts – isolation and identification of moulds from a variety of
substrates in moist chambers, which we have been doing for the past few weeks, and assay of
enzyme activities for those moulds.
Your write up should include the following information for each mould isolated:
a pure culture,
a sketch of conidia and conidiogenous cells, including spore dimensions in µm,
a slide mount.
Identify each isolate to genus using the keys in Barnett and Hunter (Deuteromycetes) and
O’Donnell (Zygomyctes).
Determine the enzyme(s) secreted by each species (there may be more than one) and relate them
to the substrate from which you isolated them. How do the enzymes relate to possible
ecological activities of these fungi?
This lab report will be due February 16st. Your report should be typed, single-sided,
double-spaced, 12-point font, one inch margins. No page limit, but brevity is encouraged
Reference: Hankin, L. and S. L. Anagnostakis 1975 The use of solid media for detection of
enzyme production by fungi. Mycologia 67: 597-607.