Culture media and sterile technique
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In nature, fungi grow upon and within complex materials, from which they derive the organic
and inorganic nutrients for survival. Fortunately, many fungi can grow in the laboratory on
culture media, which are easier to prepare and/or more reproducible than are their natural
substrates.
Culture media often contain complex organic materials like potato water or V8 juice, or
enzymatic digests of proteins (peptone, tryptone, casein or yeast extract). Some are of these
supplemented with sugars (usually glucose, also called dextrose) or pH buffers. Antibiotics can
be added to prevent bacterial growth. Because complex materials come from living matter, they
contain an immense variety of components (in fact it is impossible to know everything that is
present), which together are likely to satisfy all but a few species. There are many formulas for
fungal culture media, partly because there are many kinds of fungi, and also because many
studies require specialized nutritional conditions.
For some studies, particularly regarding nutritional requirements, "minimal" culture media can
be prepared from fully defined components. These must include sources of reduced carbon
(often, glucose), nitrogen, phosphorus, sulfur, and trace elements. Trace elements are prepared
from defined ingredients, and the medium is made with ultrapure water.
Some media are known by a set of initials that indicates their main ingredients, for example,
PDA is Potato Dextrose Agar. The word "agar", which occurs in the names of many media,
indicates a polysaccharide by that name. Agar provides little nutrient value; its purpose is to
convert the medium from a liquid (called a broth) into a jelly. However, unless highly purified
(“agarose” is extremely expensive) agar does contains traces of nutrients. These so called "solid"
media are especially good for growing filamentous fungi, because hyphae are adapted to growing
within a substrate; nevertheless, yeasts and bacteria can grow on agar surfaces. Gelatin, another
common gelling agent, is less useful than agar because manhy species secrete proteases, which
soon turn the jelly into a sloppy mess.
Solid media are usually poured into a low flat dish (called a Petri plate), or into a test tube (called
a slant). The broad surface of a Petri plate provides a working area for manipulating the
organisms. A slant is used for long-term culture. The spores of many species can also be stored
in sterile 15% glycerol at –80°C.
Media must be sterilized before use, or cultures will always have contaminants (in other words,
fungal spores are ubiquitous!). The standard way to sterilize media and other culture materials is
by autoclaving, which involves heating the medium to a temperature of 121°C, under 110 kPa
(~ one atmosphere) of steam pressure above ambient. This also melts the agar. For efficiency,
we will be providing most of the plates and media, but I have included general preparation
instructions and recipes.
Discard old cultures by placing them in the orange autoclave bag with the Biohazard sign. Tape
the plates closed (we provide masking tape) before discarding them! This reduces contamination
hazard and also allows you to retrieve a culture discarded by accident.
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Preparing culture media
Unless noted otherwise, “water” means highly purified water that has essentially no minerals.
This is to make the media more easily reproducible.
Begin with a clean beaker that has a somewhat larger volume than the amount of medium you
will be preparing, a clean magnetic stir bar and stir plate, and about 2/3 the total volume of
water. Measure the dry ingredients into a tared weighing boat, using a clean spatula for each
ingredient. Add the ingredients (liquid and dry) to the beaker while stirring. Experience will
show that dry powders like yeast extract dissolve more easily if sprinkled on the surface of the
solution. These powders are very hygroscopic, meaning that they can absorb moisture from the
air. It is important to keep all these containers tightly sealed when not in use, or the powder will
become a soggy cake.
Adjust the pH, if required, and then make up to the final volume with water. Pour the broth into a
flask or bottle that is ~ 30% larger than the liquid volume. Add agar, if required. Agar is a
carbohydrate that creates a stiff gel at 1.5% w/v. Cap flasks with foil or an inverted beaker. Cap
bottles “finger tight” and then loosen by half to one turn. In this way the lid won’t pop off during
pressure changes, nor will the bottle explode.
Autoclave using the liquid cycle, which means that the pressure will be reduced slowly after the
sterilization time so that the liquid will not boil over (there will be some bubbling, hence the
need for a larger container than the liquid volume). Liquids must always be placed in an
autoclave tray in case of boil over. Generally, 20 min sterilization is sufficient for one litre of
medium. Note that larger volumes usually require longer sterilization – the heat content of steam
is finite! Two litres requires almost 30 min sterilization. If you have not heated your media for
long enough, the agar will not have melted completely, and you will see small granules swirling
in the next step.
When the autoclave cycle is finished, open the door carefully, and (wearing gloves!) remove the
tray and flasks/bottles. Agar-solidified media must be mixed by swirling, which also gives you a
chance to check that the agar is fully melted (shown by tiny particles). Avoid making more
bubbles than absolutely necessary, because you want a smooth surface for your plates and not all
the bubbles will break during cooling. Agar solidifies at about 45°C, so to speed the cooling
process we put the autoclaved flasks in a 47°C water bath. Ring weights keep the bottles/flasks
stable while they are cooling.
We use pre-sterilized plastic Petri dishes (A), which have a rim designed to make stacks more
stable. To pour plates, wash your hands, choose a draft-free area, and wipe it down with 70%
ethanol. Open a sleeve of plates so that the stack is right side up. Pour about 25 ml of medium
per plate (no more than half-full), twisting the flask slightly as you finish each plate to reduce
drips. Avoid slopping over the edges or onto the lid, both of which contribute to contamination.
Leave the plates as a stack to cool, so that the agar can solidify. Cooling takes a little longer this
way, but pouring is more efficient and takes far less space. Plates are labeled as to the medium
they contain by a coloured strip code on the side (efficiently done down a stack). When the agar
has solidified the plates will look cloudy from the side, and have a slight ripple pattern from the
top. Invert the stack of solidified plates so condensation does not form on the agar surface. Leave
plates at room temperature for at least several hours to overnight so that the medium dries out
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slightly. This reduces contamination. Media can also be poured as a test tube slant (B, previous
page) for long term culture storage. Petri plates should stored bagged and inverted. This keeps
any condensate away from the medium surface, and reduces contamination.
To transfer the culture we do the following:
 Take an inoculating needle, usually a thin needle or wire at the end of a long pencil-like
handle, and heat it in an alcohol or gas flame until it glows bright red (Fig A). “When it’s red,
it’s dead”
 Allow the needle to cool for about 15 seconds or touch a piece of sterile agar to cool and
moisten the tip. If the tip is hot, the agar will boil around it. (A hot needle will kill the mould
that is to be transferred).
 Open the Petri dish containing the culture just wide enough to allow entry of the needle.
 With the heat-sterilized needle, cut out a small portion of the colony margin. Hyphal tip
transfers work best as they are usually the most active parts of the culture; in addition,
transfers from the heavily sporulating central portions will result in spores being spread into
the air. Especially in medical work, hyphal tip transfers are essential. The excised colony
margin should be only about 1 mm square (Fig B).
 Transfer the square of colony margin to the sterile plate, making sure that the lid is opened
only wide enough to admit the needle and make the transfer. Place the block at the centre,
withdraw the needle and flame it until it is red hot, to kill all adhering spores and hyphae
(Fig C,D).
 Close the lid; label the bottom of the plate with a marking pen, including name of culture and
date. Depending on the culture and the procedure, you can either wrap a thin strip of Parafilm
around the sides of the plate to cover the opening, or hold the lid in place with couple of
pieces of masking tape. Parafilm sealed plates are more likely to be contaminated by
unwanted (environmentally borne) molds, but they are necessary for species that produce
abundant spores (e. g. Rhizopus and Neurospora). Masking tape sealed plates will tend to dry
out eventually, but the culture will have some access to oxygen.
 Leave the culture to grow in a protected place that has as little air movement as possible.
 Transferring from plates to tubes, tubes to plates, or tubes to tubes is done in a similar
manner. When using tubes, always flame the mouth to kill any spores of airborne moulds.
Never put the cotton plugs or lids of tubes on the table as they will pick up contamination.
Steps in sterile technique. Inoculating needle is
heated in an alcohol flame (A). Small piece of
colony is removed from Petri dish (B) and
transferred to a new dish of agar (C), yielding a
platecontaining a piece of the old culture at its
centre (D).
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Recipes
“Complete” Medium (a complicated recipe, but good for Aspergillus nidulans)
10 g D-glucose (a.k.a. dextrose)
2 g peptone
1 g yeast extract
1 g casamino acids
50 ml 20X nitrate salts (see below)
1 ml trace elements (see below)
1 ml vitamin solution (see below)
Adjust pH to 6.5 with 1 M NaOH. Add distilled water to 1 litre. For solid medium add 15 g
agar. Autoclave.
Minimal Medium (a fully defined medium)
50 ml 20X nitrate salts (see below)
10 g D-glucose
1 ml trace elements (see below)
1 ml 1% thiamine
50 ul 0.05% biotin
Adjust pH to 6.5 with 1M NaOH. Add distilled water to 1 litre. For solid medium add 18 g of
agar per litre. Autoclave.
20X Nitrate Salts
120 g NaNO3
10.4 g KCl
10.4 g MgSO4.7H2O (5.2 g if anhydrous)
30.4 g KH2PO4
Add distilled water to 1 litre. Autoclave. Store at 5oC.
1000X Trace Elements
1. Dissolve these salts in 80 ml of distilled water in the order indicated:
FeSO4 ·7H2O (ferrous sulphate)
1.0g
Disodium EDTA (ethylene diamine tetra-acetate) 10.0g
Adjust pH upwards with KOH pellets. A golden yellow solution results above ~ pH 5.5, and
this is sufficient to proceed.
2. Dissolve these salts in 80 ml of distilled water in the order indicated:
ZnSO4·7H2O (zinc sulphate)
4.4g
H3BO3 (boric acid)
2.2g
MnCl2·4H2O (manganous chloride)
1.0g
CoCl2·6H2O (cobaltous chloride)
0.32g
CuSO4·5H2O (cupric sulphate)
0.32g
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(NH4)6Mo7O24·4H2O (ammonium molybdate) 0.22g
Combine Solutions (1) and (2), and readjust the pH to 6.5 using first KOH pellets, then KOH
solutions of decreasing concentration. Bring the final volume to 200 ml with distilled water,
and store at 4-8°C. As with traditionally prepared Hutner's TE, this solution is initially bright
green, turning purple upon storage. Precipitates are never formed.
Vitamin Solution
0.01 g each of biotin, pyridoxin, thiamine, riboflavin, PABA (p-aminobenzoic acid), nicotinic
acid in 100 ml water. Store in a dark glass bottle at 4oC. Riboflavin is light-sensitive.
5YEG (general purpose media)
5 g yeast extract
10 g D-glucose
Add distilled water to 1 litre. For solid media add 18 g of agar per litre.
5GP (general purpose medium)
10 g D-glucose;
5 g peptone
Add distilled water to 1 litre. For solid medium add 18 g agar
V8 agar (a good natural medium – improves sporulation of some strains)
200 ml V8 vegetable juice
3 g CaCO3
20 g agar
1000 ml water
Rose Bengal Agar (good for slowing growth of colonies in a mixed culture when making
isolations, and reducing bacterial and other contaminants)
1000 ml water
10 g D-glucose
5 g peptone
1 g KH2PO4
30 mg Rose Bengal (10 ml of a 3 g/l stock solution)
15 g agar
Autoclave and cool
Add 30 mg streptomycin, swirl to mix, and pour
“Organic” medium for oomycetes
10 g D-glucose
1 g peptone
0.1 g yeast extract
1 g KH2PO4
0.3 g MgSO4.7H2O (or 0.15 g anhydrous MgSO4)
15 g agar (optional)
GY medium for oomycetes
Glucose 1%
Yeast extract 0.5%
Agar 1.5%
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Dilute Salts (for inducing zoosporogenesis in oomycetes) makes 1 litre each
DSA
13.6 g KH2PO4
17.4 g K2HPO4
13.2 g (NH4)2SO4
DSB
7.4 g CaCl2.2H2O
10.2 g MgCl2.6H2O
Store stocks at 4°C.
To use the Dilute Salts stocks, put 500 ml of distilled water in a 1000 ml flask, add 0.5 ml DSA
and 1.0 ml DSB, swirl to mix well, and add distilled water to one litre. Swirl again. Mixing
stocks directly causes precipitation.
To induce zoosporogenesis: grow oomycete cultures of in liquid OM, overnight, starting with
very small inocula, ~1mm3. Remove OM by aspiration, and replace with dilute salts. Remove
replace two or three times in the first hour, and then once an hour. Should have good zoospore
release by 4-5 h.
Cotton blue in lactophenol
20 g phenol crystals
20 g lactic acid
40 g glycerine
20 g water
0.1 g cotton blue
NB: lactophenol polymerizes/esterifies with time. This changes the osmolality of the solution so
that it causes plasmolysis. Other blue dyes are probably similar to cotton blue -- trypan blue or
toluidine blue; Dave Malloch suggests ink blue
Lactofuschin
100 ml 85% lactic acid
0.1 g acid fuschin
Good general cytoplasmic stain, deep pink
IKI solution (for carbohydrates)
100 ml water
5 g potassium iodide (KI)
1.5 g iodine metal
Stains starch deep blue (amyloid reaction). Stains some spores brown (dextrinoid reaction).
Chlorazole Black E stain (CBE) for mycorrhizae
 Fix roots for an hour or longer in 3.7% formalin buffered to pH 7 with 50mN Na-K
phosphate buffer
 Place fine clean roots in an autoclavable container for clearing and staining.
 Clear roots in 10% KOH  this must be optimized for each type of root. Keep careful
notes, and start with the most gentle treatment.
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o Delicate roots may disintegrate during autoclaving. Clear at 60°C – 1 h at 60°C is
approximately equivalent to 5 min at 15psi 121°C. Tough roots may need
autoclaving for up to 15 min.
o Wash out KOH using70% ethanol
o Heavily pigmented roots may need post-KOH treatment in chlorine bleach. Note
that a fine, heavily pigmented root was not decolourized by KOH but was
completely cleared by subsequent treatment 1 h in full strength bleach at room
temperature. Bleaching can also use 1% H2O2 in 10% NH4OH
Stain roots  in the autoclave for up to 5 min in stain solution, or at 68C fir 3h, or at
room temperature overnight, or for up to a week at 4°C
o In freshly made chlorazole black E (CBE), 0.03% in 1:1:1 distilled water:lactic
acid:glycerol (DLAG). Make CBE at 0.09% in water, then add equal volumes full
strength (85%) lactic acid and glycerol.
 CBE can be reused if strained to remove fine roots, until it gets too dilute
to give satisfactory results. Anne Ashford makes hers fresh each time.
 Range of possible CBE concentrations 0.003% – 0.1%
o In lactofuschin (described above)
Destain roots before observation by incubating in 47C DLAG. Mount in 50% glycerol
Staining callose and beta-1,3-glucans with aniline blue
(see Smith and McCully 1978a Stain Technology 53:59-85 and 1978b Protoplasma 95:229-254)
 Prepare 0.05% aniline blue (Polysciences gives good fluorescent yield) in 0.67M
phosphate buffer pH 8.5.
 Should be prepared fresh or stored in a dark bottle at 4°C. Replace when starts to turn
greenish
 Staining takes 8-10 minutes, and there is no need to destain. Can use aldehyde fixed or
fresh material, or dis-embedded sections, or Epon?
 Control with buffer alone if staining specificity is important, since some plant
components autofluoresce under alkaline conditions.
Czapek's Solution Agar
Sucrose
30 g
NaNO3
3.0 g
K2HPO4
1.0 g
MgSO4.7H2O
0.5 g
KCl
0.5 g
FeSO4.7H2O
0.01 g
Agar
15 g
Distilled water
1000 ml
Czapek's Solution Agar is a synthetic medium widely used in mycological laboratories. Many
moulds produce very characteristic colonies on it and may also exude pigmented substances.
Aerial growth is often suppressed and sporulation may be enhanced. Some moulds, however,
grow poorly on this medium and may even fail to sporulate altogether, often because of their
inability to synthesize vitamins.