Part G : Methods and Materials

Laboratory Methods

A. Sterile Techniques
B. Growth Media
C. Incubation Temperature and Growth Rates
D. Using Toothpicks and Inoculating Loops
E. Spreading Cells
F. Pipets and Pipetting
G. Subculturing Yeast
H. Isolating Single Colonies
I. Replica Plating
J. Estimating the Number of Yeast Cells in a Culture
K. Dilution and Plating Procedures
L. Microscopic Examination of Yeast
M.Yeast Strains
N. Irradiating Yeast With Ultraviolet Radiation

A. Sterile Techniques

Sterile technique is always a relative matter. The precautions required depend on the experimental situation, including the growth media used, the competitive abilities of the experimental organism, the duration of the experiment, and the intended use of the culture. For these experiments the most serious contamination problem is mold. Many common molds grow well on yeast media and compete effectively, over-growing the plates and obscuring those yeast that do manage to grow. Bacteria are less of a problem as most of them do not grow well on these media and, with a little experience, one can easily distinguish them from yeast. Other strains of yeast can be disastrous contaminants if they go undetected. In particular, a red yeast that sometimes shows up can be really confusing, but is distinguished from our red yeast because it doesn't require adenine.

In contrast to short-term experiments, however, sterilizing media must be quite rigorous because storage allows ample time for even slow-growing contaminants to develop. We have concluded from experience that storing media in the cold is actually counter-productive. It does not prevent contamination, but slows its growth. Consequently contamination may go undetected until the cultures are incubated. It is much better to store media at room temperature and detect contamination before the medium is used.

(1) Sterilizing With Moist Heat

Moist heat provided by an autoclave or pressure cooker is an efficient way to sterilize most materials. At a pressure of 15 psi above atmospheric pressure, water reaches a temperature of approximately 121O C before it boils. Most materials are effectively sterilized by 15 minutes exposure to this temperature. If an autoclave is not available, an ordinary household pressure cooker will effectively sterilize media supplies in small batches.

(2) Sterilizing With Dry Heat

 Dry materials such as glass and metal may be sterilized in an oven, but this requires higher temperature (160O C) and longer time (at least 2 hours) than an autoclave. This is most practical for glassware when an oven controlled by an automatic timer is available so that sterilization and cooling can occur overnight. Use of disposable, pre-sterilized, plastic ware makes the oven less important.

(3) Sterilizing by Flaming

The flame from a gas burner effectively sterilizes small glass or metal objects, such as inoculating loops, but one must avoid "frying" the yeast by contact with objects heated in a flame. Dip glass spreaders in alcohol and then use the flame to ignite the alcohol. Place metal tools such as loops in the flame until they are red hot. Cool hot tools by touching them to the agar or inside of a sterile glass tube. However, since flames are dangerous this method can and should be avoided whenever possible. We have found that flaming is really only necessary to sterilize inoculating loops and small instruments. We have not been able to demonstrate any value in flaming the openings of tubes, bottles, or flasks in these experiments.

 (4) Sterilizing With Alcohol

In many experimental procedures, the most effective way to sterilize objects is with ethanol. Either 95% or 70% will work. The latter is actually more effective, but the former is often more convenient. Of course the alcohol must be allowed to evaporate or be burned off before the object is used in contact with the yeast. Use a flame to burn off the alcohol, a candle is generally less expensive and safer than a gas burner. The alcohol, more than the heat, does the sterilizing, so just "light" the alcohol to minimize heating and speed up the process. Also, wipe the bench with alcohol before starting an experiment to remove mold-laden dust, the most common source of contamination. If your skin is not particularly sensitive, wipe your hands with a small amount of alcohol, too.

 (5) Keeping Sterile Things Sterile

The most common sources of contamination during an experiment are dust, from the bench top, from the air and from people. This dictates several obvious principles:
1) Keep things covered as much as possible.
2) Don't touch anything that will come in contact with the culture and if you do touch it sterilize it again before using it.
3) Wipe down the surface around the experiment with alcohol and minimize air turbulence.
4) Avoid talking, singing, whistling, coughing, or sneezing in the direction of things that should be sterile. Long hair, if not tied back, may be a source of contamination.
5) Maintain a suitable area for preparing, storing, and using sterile media. Unfortunately, house plants, animals, and other materials such as Drosophila media, commonly found in biology class rooms, are abundant sources of mold and must be kept far away from the area used for sterile procedures.

(6) Study Your Contamination to Avoid It Next Time

 If some of your agar plates become contaminated, you can often tell by examining the plate how contamination took place. If the contaminants are imbedded in the agar, the contaminant was probably poured with the medium. If the contamination is beneath the agar it could have been in the dish before it was poured or entered when the lid was being opened to pour. If the contamination is on top it could have come in before the lid was closed after pouring, while opening the lid to wipe out condensation or while the plate was being inoculated.

 (7) Tailor Your Techniques to Your Needs

 Appropriate sterile technique is essential for successful microbiological experiments, but excessive precautions can be serious distractions. Yeast and bacteria call for different methods from tissue culture. Yeast grow robustly, out-pacing most things, except a few filamentous (fuzzy) fungi and some bacteria. Experiments can become contaminated and still not be lost when students are using yeast because the yeast will grow well enough, in spite of contamination, that the results of an experiment can be read through the offending growth. Therefore, use the most caution while making and pouring media.

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B. Growth Media

 Most of the experiments in this handbook use one or two of the following types of media:
1) a nutritionally complete medium containing a suboptimal amount of adenine (YED),
2) a nutritionally complete medium containing an excess of adenine (YEAD),
3) a chemically defined medium lacking adenine (MV),
4) a chemically defined medium with adenine (MV + Ade),
5) a sporulation medium (YEKAC),
6) a medium used to identify petite mutants (PETITE) and
7) a rich medium used to store yeast strains (YEPAD).

We use media prepared from commercially available ingredients. The small extra expense of these ingredients is more than compensated for by their uniformity and dependability. The savings that can be realized by using "home-made" ingredients are smaller than might be expected because we have not yet found a substitute for the most expensive ingredient, agar. Although common sources, such as various kinds of vegetable juices, can supply most of the other ingredients, we have found the sacrifice of reproducibility to be unwarranted.

 YED medium (Yeast Extract + Dextrose) will be our standard growth medium. It is called a "complex" medium because it is made from natural ingredients (yeast) and its exact chemical composition is not known. It contains everything that is in yeast--including adenine, of course--so it is also a "complete" medium. Red adenine-requiring mutants grow well on this medium and develop the characteristic red pigment.

 YEAD medium (Yeast Extract + Adenine + Dextrose) contains the same ingredients as YED but has excess adenine added. Red adenine-requiring mutants grow well on this medium and will not develop the characteristic red pigment.

 YEPAD medium (Yeast Extract + Peptone + Adenine + Dextrose) contains the same ingredients as YEAD but has peptone added. This is a very rich medium used for storing yeast strains.

 MV medium (Minimal plus Vitamins) is a chemically defined medium, meaning it is made up from the minimum of pure chemical ingredients necessary to support growth of wild type yeast. We will use MV when we need a medium that contains no adenine.

 MV + Ade medium (Minimal plus Vitamins + adenine) contains the same ingredients as MV but has excess adenine added.

 YEKAC (Yeast Extract plus Potassium Acetate) induces diploid yeast to sporulate and undergo meiosis. It is nutritionally unbalanced so very little growth will occur.

 PETITE medium contains glycerol as the carbon source and is used to identify petite colony mutants. Petite cells will not grow on this medium.

 Recipes For Agar Growth Media

 If you are using a prepackaged mix follow the instructions on the package. If you are weighing out dry ingredients use the following recipes. The recipes are give for the preparation of 100 milliliters. If you want to make more, simply multiply the amounts to get the volume you want. Example: to make 500 milliliters multiply each ingredient by 5. Don't put more than 600 milliliters in a 1000 milliliter flask. If you fill containers too full they may boil over during the sterilization process. It takes approximately 25 milliliters of medium to pour one standard 100 x 15 mm plate. (Liquid media may be made from these recipes by omitting the agar.)

 Yeast-Extract Dextrose Medium (YED):

 1 gram Yeast Extract
2 grams anhydrous dextrose (glucose)
2 grams Agar (agar-agar; gum agar)
100 ml water

 Yeast-Extract Adenine Dextrose Medium (YEAD):

 1 gram Yeast Extract
2 grams anhydrous dextrose (glucose)
2 grams Agar (agar-agar; gum agar)
8 ml adenine stock solution
92 ml water

 Yeast-Extract Peptone Adenine Dextrose Medium (YEPAD):

 1 gram Yeast Extract
2 grams Peptone
2 grams anhydrous dextrose (glucose)
2 grams Agar (agar-agar; gum agar)
8 ml adenine stock solution
92 ml water

 PETITE Medium:

 1 gram Yeast Extract
0.025 gram anhydrous dextrose (glucose)
2.4 ml glycerol
2 grams Agar (agar-agar; gum agar)
98 ml water

 Minimal with Vitamins (MV):

 0.15 grams Yeast Nitrogen Base without amino acids and ammonium sulfate
0.52 grams ammonium sulfate
2 grams anhydrous dextrose (glucose)
2 grams Agar (agar-agar; gum agar)
100 ml water

 Minimal Media plus Adenine (MV + ade):

 0.15 gram Yeast Nitrogen Base without amino acids and ammonium sulfate
0.52 gram ammonium sulfate
2.0 gram anhydrous dextrose (glucose)
2.0 gram Agar (agar-agar; gum agar)
8.0 ml adenine stock solution
92 ml water

 Sporulation Medium (YEKAC):

 1 gram potassium acetate
0.25 gram Yeast Extract
2 grams Agar (agar-agar; gum agar)
100 ml water

 Adenine Stock Solution

 400 mg adenine in 400 ml water (1 mg/ml)
Store at room temperature.
Use 2 ml stock solution for 100 ml of medium; reduce the water added to the medium by 2 ml.

 Adenine Blotter Paper Disks

 1.25 grams of adenine
150 ml distilled water
art blotter paper disks
Add adenine to the distilled water in a large enough beaker so the solution does not boil over. Boil five minutes. Add blotter paper disks and boil five minutes more. Remove the disks with sterile forceps to a sterile covered container such as a Petri dish. Place them in a single layer and allow them to dry for several days.

 Sterile Water

 For many of the short term experiments it is possible to "sterilize" water by boiling.
1) place the water in a loosely covered container,
2) place the container in a pan of boiling water for 15 minutes,
3) after it cools tighten the lid on the water container.

 Probably a better way to prepare sterile water is to place the loosely covered water containers in an autoclave or pressure cooker and then sterilize the water under the same conditions used for growth media.

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C. Incubation Temperature and Growth Rates

 The optimum growth temperature for these strains is approximately 30O C, but they will grow quite satisfactorily over a much wider range. All of these experiments can be done at a comfortable room temperature, but the yeast tolerate almost any temperature that people can. However, the temperature will affect the growth rate. At lower temperatures the cells will grow more slowly, and the rate must be determined by experiment. The incubation times estimated in these experiments are based on incubation at temperatures within a degree or two of 30O C.

 One can, in fact, slow yeast down by chilling them to make experiments fit into class schedules. Simply refrigerate or merely cool plates at different stages to slow down growth. Keep in mind, however, that after a culture has been refrigerated there may be a lag period of several hours before it begins to grow again. Of course, if it has reached the stationary phase it will not grow further at any temperature.

 Yeast strains can actually be stored for prolonged periods in the refrigerator at temperatures just above freezing. Some strains will continue to grow, although very slowly, at these temperatures, but most remain viable for weeks or months, and some for years.

Temperatures above 30O C should be avoided. Although cells will grow at higher temperatures, they accumulate undesirable petite mutations and sporulation is strongly inhibited. The red pigment in adenine mutants is also inhibited at higher temperatures. It is safest to incubate cultures that are being sporulated at a comfortable room temperature.

 Incubate cultures in the dark or reduced light as much as is convenient. Although not critical, prolonged exposure to normal room light or brighter light will reduce the viability of the cells that contain the red pigment. The UV sensitive strain G948-1C/U should not have prolonged exposure to light sources.

For optimal growth, and for the red pigment to develop, the plates must be aerobic. We incubate plates in open plastic food-storage bags. Enough oxygen is available with the bags open to support aerobic respiration. Plastic bags keep the plates from drying out too fast, protect them from contamination and make them easier to carry without spilling.

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D. Using Toothpicks and Inoculating Loops

 For most purposes the easiest way to move yeast from one place to another on agar medium is with a sterile toothpick. The flat style toothpick is easier to use than the round style. The pointed end is useful for picking samples from individual colonies or for making small spots or streaks. The flat end is good for spreading the yeast, for streaking it out to isolate single colonies and for mixing. Toothpicks are sterile directly from the box as long as the box has not been contaminated. Just cut across one corner to make a small hole (about 1/4 to 1/2 inch across) and the box will serve as a dispenser. The toothpicks in a box are pointing in both directions, but you can easily select the end you prefer for any task. Of course, one must take care to handle each toothpick only by the end that will not touch the yeast or the medium.

The toothpick is an alternative to the traditional inoculating loop, which is available commercially, even in platinum. We make our own inoculating loops from a 6 to 8 inch piece of brass rod (as used for brazing) and nichrome wire. Drill a small hole near the end of the rod to anchor the wire, insert the end of the wire into the hole, wrap several turns around the rod and leave an inch or two of wire straight. In the end of the straight portion form a loop with a pair of needle-nose pliers. Flame the wire in a bunsen burner until it glows red to sterilize it before each use, and of course, allow it to cool before it touches (or fries) the yeast. Touching it to the agar cools it immediately.

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E. Spreading Cells

 There are several ways to spread cells. They use various types of pipetting equipment and various methods of moving the cells on the agar surface.
Quantitative pour plating method
This method requires the preparation of a separate final dilution tube for each plate and then the entire contents of the tube are poured onto the agar surface. This method has the advantage of not requiring the use of a flame. It has the disadvantage of not being as reproducible.

1. Make a 1 to 10 dilution series of yeast cell suspensions.
2. Pipet 0.1 ml of the appropriate suspension into a sterile tube containing 0.9 ml of sterile water. Swirl the tube to suspend the cells and then pour the entire contents of the tube onto the surface of the agar medium. Tilt and rotate the plate to evenly distribute the suspension over the surface of the agar.
3. If there are spots that don't get covered use a sterile toothpick to spread the suspension over the uncovered areas.

The suspension must be allowed to soak into the agar before the plates are exposed to the experimental treatment. To speed up the process make the agar plates several days in advance and let they dry out a bit before you pour on the yeast suspension.

 Video section Serial Dilution & Viable Cell Counts illustrates the pour plating method.

 Quantitative spreading method
When you want precise data, it is better to use an alternative plating method. The cell suspension is accurately pipeted onto the surface of the agar and then spread with a sterile spreader. There are two types of spreaders:

 No flame paper clip spreader
You can easily make an inexpensive reusable spreader by straightening out two bends in a large paper clip, leaving a hairpin-shaped end for spreading and a straight handle at right angles to it. You can sterilize several spreaders together in covered beakers or wrapped in foil or paper.
Flamed glass spreader
If you have the equipment and skills to work with glass rod you can make a simple glass spreader by heating and bending a one inch section at the end of a short piece of glass rod. These spreaders can also be purchased.

1. Make a 1 to 10 dilution series of yeast cell suspensions.
2. Use a fresh pipet tip to pipet 0.1 ml of the appropriate suspension onto the agar surface. Thump the tube before taking the sample. If you are doing multiple plates you can pipet cells into each plate using the same pipet tip and then spread them without flaming the spreader between each plate.
3. Sterilize the spreader in 95% ethanol and then pass the spreader through a flame to ignite the alcohol. Hold the spreader carefully until all the alcohol on the spreader completely burns. The use of alcohol flames is a dangerous step and requires care and close supervision by the teacher.
4. Cool the spreader by touching it to the agar . Use the flat side to spread the suspension over the surface; then use the curved part of the spreader to make overlapping strokes. Turn the plate 90 degrees and repeat the process of making overlapping strokes. The lid can be held over the plate to minimize contamination. Avoid spreading the suspension to the edge where it can run down between the agar and the side of the Petri dish.

Video section Using Yeast to Measure Solar UV, illustrates a flamed glass spreader plating method.

Qualitative pour plating method to produce lawns of cells
This method produces a thick even growth of cells (lawn) over the surface of the agar plate. This is a quick method and is useful for a variety of qualitative experiments.

 1. Make a turbid suspension of yeast cells.
2. Pipet 1.0 ml of the suspension onto the surface of the agar medium. Tilt and rotate the plate to evenly distribute the suspension over the surface of the agar.
3. If there are spots that don't get covered use a sterile toothpick to spread the suspension over the uncovered areas.

As in the quantitative pour plating method the suspension must be allowed to soak into the agar before the plates are exposed to the experimental treatment. If you make the agar plates several days in advance and let they dry out a bit before you pour on the yeast suspension you will speed up the process.

 Video section Effect of Solar UV on Cells illustrates the pouring method of plating for a lawn.

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F. Automatic Micropipettors, Serological pipets, and Disposable Bulbed Pipets

 Automatic pipettors are the workhorses of modern molecular biology and microbiology. They are fast and accurate, and save a great amount of time and frustration. The experiments in this manual often call for the use of a 0.1 ml pipettor. Although they are relatively expensive, they make the job so fast that one can easily be shared by several students or lab groups. You will use it for making serial dilutions and for plating cells on agar. The sterile tips are packaged in reusable plastic dispensers. Practice using the pipettor with water until you feel comfortable with it. (If automatic micropipettors are not available it is possible to substitute disposable bulbed pipets, serological pipets or graduated capillary tubes)

1. Press the small end of the pipettor firmly into the open end of a tip while the tip is still in the dispenser.
2. To fill it, slowly depress the plunger until you feel resistance, but not as far as possible. Immerse the tip in the solution and then slowly release the plunger. If any drops adhere to the tip, wipe them on the edge of the container.
3. Place the tip into the liquid in the tube you are pipetting into and slowly depress the plunger. This time press it as far as possible to expel all of the sample. Remove the tip of the pipettor from the solution, release the plunger, and discard the tip.

Serological pipets and pipet pumps may be used instead of automatic pipettors. These pipets will give very accurate volume measurements. It is possible to purchase presterilized serological pipets in several sizes. You must have a set of pipet pumps available for students to use with these pipets. Remember it's never good practice to use your mouth to draw liquid into a pipet. Not even sterile water!

 Disposable bulbed pipets are also acceptable for most of the experiments in this handbook. Three drops from a one milliliter bulbed pipet is approximately equal to 0.1 ml. The volume measurements are not as accurate but these pipets are cheap and do not require the use of pipet pumps. These pipets may also be purchased presterilized and individually wrapped. The wrappers of presterilized pipets provide a convenient sterile holder for the pipet during the experimental procedure.

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G. Making a Subculture of Yeast

 Yeast nutrients can be supplied by a solid or a liquid medium. When the yeast strains come from a supplier they are usually growing on agar slants. If you keep these slants tightly closed and refrigerated you can store them for up to one year. The smaller slants allow you to use toothpicks to move the yeast. It's best to have fresh cultures for your experiments. So the day before you intend to use the yeast transfer a small number of cells from the stock culture to fresh medium (usually YED). You can use a flamed loop to transfer the cells but a sterile toothpick is usually easier. Touch the toothpick to the cells in the slant; you don't need to transfer many cells. Make a streak of cells on the surface of the agar; try not to gouge the surface of the agar. This procedure of growing fresh yeast is called subculturing. You should subculture at least 12 hours before you intend to use the yeast.

 Sometimes cultures will be sent from yeast stock centers on milk papers. To make a fresh subculture: Open the foil sterilely and using sterile forceps place the small square of milk paper on a sterile agar YED plate. If you wipe the paper across the agar several times you should get single colonies to use and a large spot of cells will grow where the paper is placed. It will take from 3 to 5 days for many strains cultured in this way to grow to a suitable size. Cultures sent this way are less vulnerable to inclement weather conditions and long travel times.

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H. Isolating Single Colonies

 Many experiments require growing colonies from isolated single cells. This is easily accomplished by streaking cultures out on an agar plate with the flat end of a toothpick. The purpose of the technique is to dilute the cells on the surface of the agar in successive, overlapping streaks, until individual cells are separated or distributed to form isolated single colonies. Figurebelow illustrates the technique, which you should practice until you are proficient.

Figure 11: Isolation of Single Colonies

 The first step is to make a single, short streak of cells (about 2 cm long) near the edge of the agar in a petri dish. With a fresh toothpick, make a second, overlapping streak, at an angle of about 15 degrees to the first one, dragging cells from the first streak onto fresh agar. Make several more parallel streaks with the same toothpick. Then take a new toothpick and repeat the process starting within the end of the last streak. Repeat this sequence until the whole plate has been covered. Since the population of cells that is produced from a single parent cell is a clone, this is a process for cloning yeast. If a culture is not pure, for whatever reason, this provides a method for isolating pure clones. The same result can be achieved using a flamed and cooled loop at each step where a fresh toothpick is used. This approach is slower, and risks frying the cells.

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I. Replica Plating

 In some experiments we transfer many cultures from one kind of medium to one or several other kinds, to determine the cultures' growth requirements. This can be done using toothpicks or loops, but if there are very many strains to be transferred it becomes laborious. A simple, rapid method -- replica plating -- played a major role in the development of microbial genetics and is one of the great labor-savers of all time. The experiments described below can be done without this technique, but it is fascinating to do and worth the trouble to set it up. Even with modest experiments, the effort is a good investment in the long run.

 The principle of replica plating is that of a rubber stamp . 

Figure 13: Replica Plating

 A cotton velveteen stamp is used to stamp a replica of the pattern of cells growing on one plate to one or several other plates. The apparatus consists of a cylindrical holder for the velveteen that is just the right size to fit inside the bottom of a petri plate, and a ring of some sort to hold the velveteen in place. For standard 10 cm plastic plates the appropriate diameter is approximately 3.3 inches. The holder can be a cylinder of wood, metal, plastic, or any other material you can fabricate. It can even be a one pound food can or other heavy cylindrical container if the bottom is flat. (A circle cut from a foam meat tray makes a good flat surface to put on a food can) The ring to hold the velveteen can be a large hose clamp, some other found object, or even a large rubber band.

The velveteen, of course, must be sterilized, and newly cut cotton velveteen usually needs to be de-linted with a piece of masking tape before it is sterilized. Autoclave and dry it thoroughly. We use six-inch squares, which we stack and wrap in paper, autoclave, and then dry on the vacuum setting. If an autoclave is not available, dry them in a warm oven or just let them sit on the counter until they dry. To reuse the velveteen squares, rinse them out in plain water and resterilize. Other fabrics such as several layers of cotton gauze may be used instead of cotton velveteen.

To replica plate, invert the master plate, the one with the yeast on it, over the velveteen on the holder and press the plate firmly against the sterile fabric surface. Then slowly peel the plate off, leaving a replica or print of the cell pattern on the velveteen. Transfer the replica on the velveteen to one or a series of sterile plates by pressing each plate gently onto the velveteen and popping it off. It is possible to make as many as 20 replicas from a single master plate if necessary. The procedure is illustrated in Figure 13.

The manner in which you remove the plate from the velveteen controls the amount of yeast transferred from velveteen to plate or from plate to velveteen. If you peel the master plate off the velveteen with a slow, rolling motion, more yeast will remain on the velvet than if you pop it off with a quick vertical motion. It is desirable to transfer as few cells as possible and still be able to see the print of the cells on the replica plate.

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J. Estimating the Number of Yeast Cells in a Culture

 There are many ways of counting cells directly, the most common making use of counting chambers for the microscope or electronic particle counters. These have little instructive value in genetics, and are not suitable for general classroom use. Therefore, we have designed experiments that don't depend on accurate particle counts. When cell counts are needed, we determine them from the numbers of colonies on the plates (see Dilution and Plating Procedures).

Many experiments, however, do require making approximate estimates of the number of cells in a liquid suspension in order to get a reasonable number of colonies plated. For this you will use one of the most sophisticated and sensitive optical instruments in existence: the human eye. With surprisingly little practice, you can learn to estimate the number of cells in a suspension by just looking at it.

 Visual estimation of cell density is based on the eye's fairly sharp threshold for observing turbidity. When viewed in a standard 13 x 100 mm tube, yeast suspensions of less than about 1 million cells per ml are not visibly turbid. Above this threshold density they are visibly cloudy. By adjusting the number of cells in a suspension until just barely visible, you can obtain a suspension of known density (approximately 1 x 106 cells/ml) and then use serial dilutions to obtain other concentrations. (Also see the experiment Serial dilutions and viable cell counts)


a. Place 1 to 2 ml of sterile water into a tube.
b. Use a cooled, flamed inoculating loop or sterile toothpick to suspend a small amount of yeast, about the size of the head of a pin, in the water.
c. Mix the suspension thoroughly by holding the tube loosely near its top between thumb and forefinger, and thumping it near the bottom with the palm side of one or more fingers of the other hand, imparting a swirling motion. (The thumping motion approximates the gesture one would make to beckon someone to come hither.)
This suspension should contain approximately 1 x 107 cells/ml, but this density is difficult to judge, so we will dilute it before judging its density.
d. Pipet 0.9 ml of sterile water into a tube with a sterile 1.0 ml pipet.
e. Dilute 0.1 ml of your cell suspension into this tube and thump it. Compare it with a tube containing sterile water. If the tube with cells is barely turbid, then it contains very nearly 1 x 106 cells/ml.
f. If the tube is not visibly turbid, then add additional 0.1 ml volumes of the original suspension until it is. If you think it is more turbid than the limit of visibility, then add more sterile water. Adjust the volumes of water and cells until you are convinced. Keep track of the volumes you add so you will know the final dilution you have made.

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K. Dilution and Plating Procedures

 In many experiments one must spread an approximately known number of cells on a plate to grow into colonies. We use serial dilutions, making several small dilutions, one after another, instead of making one big dilution. The method saves material and avoids the use of large volumes that are difficult to measure and sterilize.


Making Serial Dilutions:
The following procedure yields a series of dilutions that vary by factors of ten. You will be able to use this procedure for most of the experiments that require serial dilutions (Figure 12).

Figure 12: Serial Dilution-Plating Strategy
a. Prepare a suspension containing approximately 1 x 106 cells/ml in sterile water using the procedure described in Estimating the Number of Yeast Cells in a Culture.
b. Pipet 0.9 ml of sterile water into each of three tubes and label them. We recommend making a diagram in your notebook showing the procedure and the meaning of your labels.
c. Resuspend the cells in the original suspension. Remove 0.1 ml with the micropipettor and a fresh tip and transfer it to one of the tubes of water. Then thump the tube to suspend the cells.
d. Repeat the procedure serially, using a fresh pipet or pipet tip for each of the remaining tubes, so that each tube contains 1/10th the number of cells as the previous one. The third dilution should contain approximately 1 x 103 cells/ml.

Plating the Dilutions:
If you expect all the cells to grow, then you will only plate the final dilution, as the other tubes would be too concentrated and the plates made from them would contain too many colonies to count. However, in the radiation experiments you will use the dilutions containing more cells to compensate for the reduced number of cells that survive irradiation. This way you will obtain an optimal number of colonies to count on each plate, even when most of the cells are dead.

 e. You will have better data if you do all platings in duplicate or triplicate. First label the plates using a marking pen (a Pilot SC-UF or a Sharpie). (You will also use this same kind of pen for marking the colonies when you count them.) We recommend that you identify each plate with the strain number, dose (for radiation experiments), dilution plated, and your initials, but you may prefer to code this information in your notes and put the code letters or numbers on the plate. Write only on the lid or in small letters on the edge of the bottom, leaving the bottom clear for marking the colonies when you count them.

f. Resuspend the cells in each tube by thumping it before taking a sample, then pipet 0.1 ml onto the center of each of the duplicate plates. If you are using paper clip spreaders take a sterile spreader from the envelope. If you are using a glass spreader take it from the alcohol, light it in a flame, and after the flame burns out, touch it to the agar of one of the plates away from the cells. Spread the cells over the surface with either spreader, taking care to keep the cells at least 0.5 cm from the edge of the agar. (If you are using the pour plating method pipet 0.1 ml of the suspension into 0.9 ml of sterile water. Mix the cells into the water. Pour all of the mixture onto the agar surface then tilt and rotate the plate until the mixture is evenly spread over the surface of the agar.)

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L. Microscopic Examination of Yeast

 The easiest way to examine yeast under the microscope is to look at them directly on the plate. This only works with low-powered objectives (10x) because higher power ones get too close to the agar and fog up. To see more detail, make a wet mount slide. Put a drop of water on a microscope slide. Transfer a small amount of yeast to the water with a sterile toothpick and stir a little. Then put a coverslip over the yeast suspension on the slide. If preparations are to be viewed by a number of students, seal the cover slip to the slide with finger nail polish and the slide will be viewable for several hours. Oil immersion should not be necessary and for merely observing the shapes of cells it is not necessary to stain them. With suitable staining methods the nucleus can be seen, but the yeast chromosomes are too small to see with most light microscopes.

 For classroom demonstrations you may wish to use a video microscope. There are a number of very good video microscopes available on the market. If you don't have one it's possible to get many of the same results with a classroom microscope and a home video camera. When we look through a microscope our eyes are focused at infinity. If we place a video camera close to the ocular of the microscope and set the camera focus at infinity it will "see" the same thing as our eyes. It helps to cut out light entering from the side by wrapping the camera lens and the microscope ocular with a piece of dark cloth or aluminum foil.

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M. Yeast Strains

 The experiments make use of the following strains:

 Wild type strains: will grow on MV or YED media
                HA0: Mating type a
                HB0:  Mating type a

 Red, adenine-requiring strains; wild type for ADE2; will grow on YED or MV + adenine media
                HA1: Mating type a ade1
                HB1: Mating type a ade1

Red, adenine-requiring strains; wild type for ADE1; will grow on YED or MV + adenine media
                HA2: Mating type a ade2
                HBR: identical to HA2
                HB2: Mating type a ade2
                HBR: identical to HB2

Red, adenine-requiring double mutants; will grow on YED or MV + adenine media
                HA12: Mating type a ade1 ade2
                HB12: Mating type a ade1 ade2

White, tryptophan requiring mutants: will grow on YED or MV + tryptophan
                HAT: Mating type a trp5
                HBT: Mating type a trp5

Red, adenine-requiring, tryptophan-requiring double mutants; will grow on YED medium
                HART: Mating type a ade2 trp5
                HBRT: Mating type a ade2 trp5

Red, adenine-requiring, leucine-requiring; grows on YED medium
                HA1L: Mating type a ade1 leu2

Ultraviolet sensitive strain; deficient in excision repair, error-prone repair and photoreactivation;
      grows on MV or YED media
                G948-1C/U: Mating type a rad1 rad18 phr1

The strains are also known by the following alternate designations
                      BSCS Green            Yeast Genetics
                        Version Text              Stock Center
HA0                a1                         XP837-S10
HB0               a1                         XP837-S11

HA1                                             XP300-38B
HB1                                             XP823-S34

HA2              a2                            XP832-S25
HAR              a2                            XP832-S25

HB2             a2                              XP820-S4
HBR             a2                              XP820-S4

HA12                                               XP300-29C
HB12                                               XP300-40C

HAT              a3                               XP837-S5
HBT              a3                               XP837-S6

HART            a4                                XP837-S8
HBRT           a4                                XP837-S9

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N. Irradiating Yeast With Ultraviolet Radiation

Standard germicidal UV-C lamps, which are low-pressure mercury vapor discharge tubes, make an ideal UV source
for irradiating wild type yeast with normal DNA repair enzymes. The sun or quartz halogen yardlights are good
sources of UV for irradiating mutant strains of yeast that are deficient in normal DNA repair enzymes.

Standard germicidal UV-C lamps are common fluorescent tubes without a fluorescent coating and with an envelope
that is transparent to ultraviolet. The low-pressure mercury spectrum is a line-spectrum with the most prominent
line having a wavelength of 253.7 nanometers. This is fortuitously close to the 260 nanometers that is optimal for
absorption by DNA.

The design of the U.V. Radiation Chamber is dictated by several considerations, the most important being safety.
(See Instructions for Building a UV Radiation Chamber) The radiation emitted by germicidal lamps will cause
painful and dangerous burns to eyes and skin so they must be effectively shielded. Exposure to the cells cannot be
controlled by turning the lamp on and off because the output intensity of the lamp is strongly
temperature-dependent. Therefore, it must be enclosed in a shielded box with a shutter mechanism so it can warm
up to a constant temperature before being used. The door and the shutter must be interlocked so they cannot both
be open at the same time.

Operating procedure for UV radiation chamber:

1. Plug in the lamp and turn it on. You can safely tell when the lamp is on by seeing the indicator light.
2. Allow the lamp to warm up for 30 minutes, if possible.
3. Center the petri dish to be exposed on the mark in the bottom of the box, remove the plate lid, gently lower the
door, and open the shutter. Measure the exposure time from when the shutter opened. When the time has elapsed,
close the shutter, gently raise the door, put the lid on the petri dish and remove it. Moving the door too quickly
causes air turbulence which is likely to cause contamination of the agar plates. If the box is fitted with a UV-B bulb
the same procedure may be followed.

Exposure procedure using the sun or quartz halogen yardlights:

1. Cover the dish with dark paper and while it remains covered position the dish at a 90 degree angle to the light
source. If you are using a quartz halogen light remove the glass cover from the light. While the sun and the
yardlight produce less UV radiation than the germicidal tube one should still take reasonable precautions. Don't
look directly at the source and avoid any long term direct exposure to skin.

CAUTION: A quartz halogen bulb gets very hot. The use of this lamp should be under the direct supervision of the

2. Remove the dark cover from the plate and start measuring the exposure time. When the time has elapsed cover
the dish and move it to the incubator. It is usually not necessary to remove the lid from plastic Petri dishes when
you use the sun or quartz halogen yardlights as the UV source. Most plastic Petri dishes are transparent to the
wavelengths of UV produced by these sources. You may want to run a trial experiment on each new batch of Petri
dishes. If you are getting survival rates higher than you expect you may wish to remove the lids during UV
exposure. If contamination becomes a problem you may wish to cover the dishes with thin plastic wrap or a
sandwich bag. These thin plastic films will not absorb significant amounts of UV.

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Last updated Wednesday, 24-Feb-99 02:04:12