Now that you know what spores, hyphae and mycelia are, and you have gotten used to manipulating microscopes to see fungal structures in detail, it is time to learn more techniques that will help you to better understand fungi.
I. CULTURE MEDIA FOR GROWING FUNGI
Culture media are mixtures of nutrients or substances used in the laboratory for cultivation of many microorganisms. Fungi may be cultured on a variety of media types. These media can be either solid or liquid depending on the experiment to be performed. Most fungi grow in medium containing a high carbohydrate source, nitrogen source, a pH of 5-6, and a temperature range from 15-37 C. Since no medium will sustain growth of every fungal species, the type of medium on which a fungus grows will vary from species to species. Some fungi, such as the downy mildews, cannot be cultured in vitro, and survive only on their plant host. (What do you call these kinds of fungi?)
There are two general types of culture media: natural and synthetic. Natural media are composed of natural substrates such as herbaceous or woody stems, seeds, leaves, corn meal, wheat germ, and oat meal. The exact nature of these substrates vary from time to time and cannot be duplicated exactly. Natural media are usually easy to prepare but they have the disadvantage that their composition is largely unknown. Some examples include corn meal agar, potato dextrose agar, V-8 juice agar, and dung agar.
Synthetic media, on the other hand, contains ingredients of known composition. These types of media can be duplicated with exactness each time they are made and contain defined amounts of carbohydrates, nitrogen, and vitamin sources. Czapek-Dox medium, glucose-asparagine and Neurospora crassa minimal medium would fall in this category.
All media used in culturing fungi must be sterilized before use. Steam sterilization by autoclaving is the customary method of sterilizing most culture media, but it cannot be used with heat labile compounds. Generally, materials are autoclaved for 15-20 min at 15-17 psi, and at temperature of 121 C or 250 F.
Some recipes for making common media are described below. We will not be able to prepare all of them but basically you need to weigh out the specific amounts of ingredients, place them in a specific amount of distilled water, autoclave the media, allow it to cool and dispense into appropriate containers (test tubes, bottles, or petri dishes).
1. Corn Meal Agar (Many fungi grow on this type of medium and often
induces sporulation).**
Corn meal- 20 g
Peptone- 20 g
Dextrose- 20 g
Agar- 15 g
Distilled water- 1000 ml
Add the corn meal to the water and simmer for 30 min to 1 hr. Filter through several layers of cheesecloth. Add the other ingredients to the filtrate, bring up the volume with distilled water, then autoclave for 20 min at 15 psi on slow exhaust.
2. Potato Dextrose Agar (PDA)**
Peeled, diced potatoes- 200 g
Dextrose- 20 g
Agar- 15 g
Distilled water- 1000 ml
Prepare the potatoes as for corn meal. Autoclave as described above.
3. Vegetable Juice Agar (V-8)
V-8 juice- 180 ml
Calcium carbonate- 2 g
Agar- 20 g
Distilled water- 1000 ml
Mix all the ingredients and autoclave as above. Other juices such as tomato or carrot can be substituted for V-8 juice.
4. Saborauds Medium
Peptone- 10 g
Glucose- 20 g
Agar- 20 g
Distilled water- 1000 ml
5. Rabbit Dung Agar
Fresh rabbit dung- 500 g
Place in a dry flask. Autoclave for 30 min. Pour 8-10 pellets into petri dish. Cover with a thick layer of sterilized 2% agar.
**These culture media (plus many others) have been prefabricated by such companies as Difco to make life easier for most microbiologists. Basically, you just need to weigh a certain amount of the prefabricated add a liter of water and autoclave. There is no need to boil potatoes, filter and weigh ingredients separately.
A. Making media slants
Remove caps of sterile tubes and sterilize the mouth of the tubes by passing them through the flame several times. Dispense 2-3 mls of molten PDA into the tube using sterile pipets. Do not lay the cap or pipet down or allow them to touch anything. Replace cap and place tubes at an angle until the medium solidifies. Sterilize mouth of tubes again before replacing cap. During slanting, care should be taken not to permit the molten agar to come in contact with the cap.
B. Pouring media plates
Open a bottle of molten Saborauds medium and flame mouth. With your left hand (or other free hand) raise one side of the petri dish cover just enough to insert the mouth of the bottle. Pour 20-25 ml of medium into the plate. Replace cover immediately and gently rotate the plate to spread the agar evenly over the bottom of the dish. Allow the plates to cool sufficiently for the medium to harden before using them. (NOTE: Observe aseptic conditions all the time when working with culture media).
II. CULTURE TRANSFERS
Many fungi can be kept indefinitely depending on how their "owners" care for them. Fungal isolates can be kept for 6 months to a year before they have to be transferred to another agar plate in order to replenish their nutrient source. Some fungi in their dormant state can survive up to ten years without a food source before becoming inviable. Normally fungi are stored on agar slants at 5 C with periodic transfers occurring around six month intervals. Naturally, not all fungi cooperate with this time table and must be checked for viability more frequently.
Other methods of long term storage include maintaining spores or mycelium in 20% glycerol at -80 C to -130 C. Cultures maintained in such a state are viable for several years.
A. Culture tube preparation
1. Sterilize the transfer loop or needle by holding the wire in the flame until it is red hot. Allow loop to cool.
2. While holding the sterile loop and the old culture, remove the cap.
3. Briefly heat the mouth of the tube in the flame before inserting needle or loop.
4. Get spores, mycelia or agar block containing fungal structures from old culture and transfer onto your fresh agar slant. Heat the mouth of tubes and replace the cap.
5. Incubate at appropriate temperature until growth has appeared. Store at 5 C for future use.
B. Plate culture
Same techniques as in tube culture preparation, only this time, spores or mycelium or agar block containing these fungal structures are aseptically transferred onto plates with solidified agar media.
III. ISOLATION OF PURE CULTURES
A. Hyphal Tipping
To separate and isolate one species of fungus from an undesired contaminant(s), a technique called hyphal tipping is usually performed. This technique is also used when isolating a fungal species from infected plant tissues. Materials required:
Dissecting scope
Scalpel knife
Contaminated plates
Lots of patience
Using the dissecting microscope, view the species of interest at high magnification. Locate individual strands of hyphae of fungus. After flaming the scalpel, cut the end of the hyphal strand (about 1 mm from end). Now take the small piece of agar where the hyphal end is located and transfer to another agar plate. Several hyphal tips may be required in order to ensure that you have isolated the fungus of interest.
B. Single Spore Isolation
Another way to isolate a pure culture of a particular species is to remove individual spores from the species of interest. This procedure can only be used if the fungus sporulates in culture or in planta, otherwise, hyphal tipping is your best option. Depending on what you are doing, there are two ways to isolate single spores. When working with plant material, it is sometimes possible to observe individual spores from an infection lesion on the plant using a dissecting microscope. Dip the needle in sterile distilled water to wet the end and try to remove single spore from the infected tissue. Place the spore onto an agar medium.
If the spores are small, are difficult to manipulate, or there are other contaminating fungi, it may be necessary to remove several spores in bulk and dilute them in sterile distilled water. Even though there may be other contaminating fungi along with your fungus of interest, the idea is to dilute them away from each other. It will then be possible to isolate the fungi from each other either by hyphal tipping or by single spore isolation without worry of nearby contaminants.
Plant materials and contaminated plates will be used so that you can practice these techniques.
IV. DILUTION SERIES AND DILUTION PLATING
Another important technique used by mycologists, bacteriologists and microbiologists is the use of dilution series and dilution plating. These two techniques are essential for many of the experiments we will carry out this semester so it is crucial that you become comfortable with these concepts. An example of a dilution series is as follows: You have just found a fungal isolate that sporulates profusely and wish to plate about five spores of the fungus per agar plate. One way to achieve this is to add water to the isolate in order to get a spore suspension. The spore suspension may contain millions of spores per ml. By using a haemacytometer, you can determine the number of spores per ml and then dilute them whereby approximately 5 spores can be placed onto an agar medium.
Example: You have a test tube that contains 1 x 106 spores per ml. You wish to plate approximately 100 spores per plate. If you dilute the mix 100 fold (10-2), the concentration will now be 1 x 10-4/ml. If you dilute this mix again by 100 fold (now at a dilution of 10-4 or 10,000 fold dilution from the original concentration), the final concentration of the spores is 102 spores per ml or 100 per ml. So, if you plate 1 ml of this dilution, your goal of 100 spores per plate is achieved.
In this portion of the lab, you will be given a known amount of material which you will have to dilute in order to plate 50, 100 or 200 spores/plate.
V. USING THE HAEMACYTOMETER
The hemacytometer is a single piece of glass with an H-shaped trough forming 2 counting areas with an engraved grid upon each. It has supports that hold a special cover glass the proper distance (0.1 mm) above the grid areas. The counting chamber and coverslip should be cleaned with water and alcohol and wiped dry with Kimwipes. A reproduction of a grid area showing the dimensions of the various squares and rectangles are shown in the accompanying handout.
A. The counting chamber
1. There are two grids per slide. Each grid is divided into a 3 x 3 pattern of large squares, each 1 mm wide. Where there are 3 bordering lines, the center is the boundary. Some of the large squares are subdivided into smaller squares or into rectangles; some of these are further subdivided. The basic dimensions on the above diagram provide the data needed to calculate the area of each size of square or rectangle (there are 400 small squares in the central square millimeter).
2. Center the special coverslip so that the long sides are parallel with and border onto the outer troughs of the slide and the short sides are equidistant from the edges of the slide. With the coverslip in place, the distance between the bottom of the coverslip and the grid is 0.1 mm. With this information you can now calculate the volume of each size of the small areas on the grids.
3. At this point, do a dry run with the coverslip in place over the grid. See if you can locate the grid using the low power objective. Watch from the side for the initial downward adjustment of focus. After locating with the low power, switch to high power.
B. Filling the chamber
1. Thoroughly agitate a given spore suspension, let the liquid motion stop; suck the liquid into and expel several times from a Pasteur pipet, then "load" the pipet.
2. Holding the pipet at a 45 angle, dropwise fill one depression (the pier on the slide containing the grids is actually depressed at the outer edges) until the suspension starts to cover the grid area; stop the addition; capillarity will cause the grid to be sufficiently covered. If you fill the chamber properly no liquid will go into the troughs and there will be no bubbles under the coverslip. If the troughs overflow the coverslip will float; you must then disassemble, rinse, and dry before starting again. Do not move the coverslip after the chamber is filled.
C. Counting
1. Use the appropriate objective to locate conidia on the grid.
2. Decide: (a) what size grid area you will use for making the counts; (b) how to count spores on border lines; (c) how many, and in what consistent pattern, areas will be chosen for counts; and (d) how many repetitions of counts are needed from each dilution of suspension.
3. Let the prepared haemacytometer stand for 1-3 min to allow all units to settle at the bottom of the chamber.
4. Make a quick evaluation of the number of cells occurring on the grids. There may be too many or too few. If so, start over with a different dilution. You should be able to count units in at least two dilutions to check accuracy and dilution procedures.
5. Count units in both grid areas. It is recommended that an average of 200-250 units should be counted on each grid since with that average the standard deviation will be approximately 15 and the total count for stock will be accurate within 10-15%.
NOTE: Units can refer to conidia, zoospores or sexual spores like oospores, ascospores, and basidiospores.
D. Calculations
1. Note the size of square suitable for counting and the number of these squares per square millimeter (mm2). Make note of this number (16, 20, 25, 80, 400, etc).
2. Count the number of spores in several squares and obtain an average number of spores per square.
3. Multiply this value (average) by the number of squares (of the size counted) per square millimeter. (This value represents the number of spores per 0.1 mm3).
4. Multiply by 10 to obtain number of spores per mm3.
5. Multiply by 1000 to convert mm3 to ml. NOTE: 1 cm3 = 1 ml = 1000 mm3
6. Multiply by dilution factor to get spores per ml in stock.
VI. QUESTIONS
1. If you were to develop a synthetic medium for a particular species of fungus, how would you go about doing this?
2. What are some 'criteria' you can use to distinguish the hyphae of your fungus of interest from hyphae of other fungal contaminants growing in the same media?
3. How would you go about separating your favorite fungus from contaminating bacteria in the culture medium?
4. How can you tell if the media provided for this exercise were sterile? How can heat-labile ingredients of culture media be sterilized?
5. You have a spore suspension with 2.75 x 108 spores/ml. If you diluted this down 1000 fold, and plated 100 _l of the resulting suspension onto an agar medium, and assumed that all the spores are viable, how many colonies would you expect growing? Show your calculations.