Preparation of Yeast Media|zoonbio

Preparation of sterile media of consistently high quality is essential for the genetic manipulation of yeast expression. Recipes for media needed in the protocols in this article are provided below.

LIQUID MEDIA

Ingredients for liquid media are dissolved in water to 1 liter, mixed until completely dissolved, and autoclaved in 100- or 500-ml media bottles. Alternatively, liquid media can be filter sterilized, resulting in faster preparation, less carmelization (of dextrose), and faster growth of cells. Recipes for “premixes” are provided to minimize the number of materials that must be weighed each time media is prepared. When preparing premixes, break up any large chunks of dextrose before mixing with other components, then shake the container vigourously until contents are homogenized. It is convenient to make the premix in the empty plastic containers in which 2.5 kg of dextrose is packaged. Throughout this article, YNB −AA/AS refers to yeast nitrogen base without amino acids or ammonium sulfate.

Rich Medium

YPD medium

Per liter:	Premix:	Final concentration:
10 g yeast extract	250 g yeast extract	1% yeast extract
20 g peptone	500 g peptone	2% peptone
20 g dextrose	500 g dextrose	2% dextrose
	Use 50 g/liter
This rich, complex medium—also known as YEPD medium—is widely used for the growth of yeast when special conditions are not required. It is preferable to use a 20% (10×) solution of dextrose that has been filter sterilized or autoclaved separately (and added to the other ingredients after autoclaving) to prevent darkening of the media and to promote optimal growth.

Minimal Media

Minimal medium

Per liter:	Premix:	Final concentration:
1.7 g YNB −AA/AS	68 g YNB −AA/AS	0.17% YNB −AA/AS
5 g (NH₄)₂SO₄	200 g (NH₄)₂SO₄	0.5% (NH₄)₂SO₄
20 g dextrose	800 g dextrose	2% dextrose
	Use 27 g/liter
This minimal medium—also known as synthetic dextrose (SD) medium—can support the growth of yeast which have no nutritional requirements. However, it is used most often as a basal medium to which other supplements are added.

Complete minimal (CM) dropout medium, per liter:
1.3 g dropout powder (table)
1.7 g YNB −AA/AS
5 g (NH₄)₂SO₄
20 g dextrose
(Alternatively, replace last three ingredients with 27 g minimal medium premix)
CM dropout powder, also known as minus or omission powder, lacks a single nutrient but contains the other nutrients listed in table. Complete minimal (CM) dropout medium is used to test for genes involved in biosynthetic pathways and to select for gene function in transformation experiments. To test for a gene involved in histidine biosynthesis one would determine if the yeast strain in question can grow on CM minus histidine (−His) or “histidine dropout” plates. It is convenient to make several dropout powders, each lacking a single nutrient, to avoid weighing each component separately for all the different dropout plates required in the laboratory.

Nutrient Concentrations for Dropout Powders^a

Nutrient^b	Amount in dropout powder(g)^c	Final conc. in prepared media (µg/ml)	Liquid stock conc. (mg/100 ml)^d
Adenine (hemisulfate salt)	2.5	40	500
L-arginine (HCl)	1.2	20	240
L-aspartic acid^e	6.0	100	1200
L-glutamic acid (monosodium salt)	6.0	100	1200
L-histidine	1.2	20	240
L-leucine^f	3.6	60	720
L-lysine (mono-HCl)	1.8	30	360
L-methionine	1.2	20	240
L-phenylalanine	3.0	50	600
L-serine	22.5	375	4500
L-threonine^f	12.0	200	2400
L-tryptophan	2.4	40	480
L-tyrosine	1.8	30	180^f
L-valine	9.0	150	1800
Uracil	1.2	20	240
^aCM dropout powder lacks a single nutrient but contains the other nutrients listed in this table. ^bAmino acids not listed here can be added to a final concentration of 40 µg/ml (40 mg/liter). ^cGrind powders into a homogeneous mixture with a clean, dry mortar and pestle. Store in a clean, dry bottle or a covered flask. ^dUse 8.3 ml/liter of each stock for special nutritional requirements. Store adenine, aspartic acid, glutamic acid, leucine, phenylalanine, tyrosine, and uracil solutions at room temperature. All others should be stored at 4°C. ^eWhile these amino acids can be used reliably when included in autoclaved media, they supplement growth better when added after autoclaving. ^fUse 16.6 ml/liter for L-tyrosine nutritional requirement.

It may be preferable to use a 10× solution of dropout powder (i.e., 13 g of dropout powder in 100 ml water) that has been “sterilized” separately (and added to the other ingredients after autoclaving) to improve the growth rate in this medium.

Sporulation medium,per liter:
10 g potassium acetate (1% final)
1 g yeast extract (0.1% final)
0.5 g dextrose (0.05% final)
This nitrogen-deficient “starvation” medium contains acetate as a carbon source to promote high levels of respiration, which induces diploid yeast strains to sporulate. Sporulation can be carried out in liquid media or on plates. If nutrients are required, add them at the concentrations listed in table.

Alternative Carbon Sources

Wild-type yeast can use a variety of carbon sources other than glucose to support growth. These include galactose, maltose, fructose, and raffinose. In particular, galactose is often used to induce transcription of sequences fused to the GAL10 promoter. All are used at a concentration of 2% w/v (20 g/liter) and should be made by replacing dextrose in the recipe for minimal or complete minimal (CM) dropout media. For a nonfermentable carbon source—which will not support the growth of petites (cells lacking functional mitochondria; see glossary)—2% potassium acetate (w/v), 3% glycerol, 3% ethanol, or 2% glycerol and 2% ethanol (v/v each) can be used. YPA medium (2% potassium acetate,2% peptone, and 1% yeast extract) is excellent for inducing high levels of respiration in cells prior to sporulation.

SOLID MEDIA

Making solid media for yeast is—for the most part—no different from preparing plates for bacteriological work. For all plates, agar is added at a concentration of 2% (20 g/liter). A pellet of sodium hydroxide (∼0.1 g) should be added per liter to raise the pH enough to prevent agar breakdown during autoclaving. In addition, add a stir bar to facilitate mixing after autoclaving.

After autoclaving, flasks are left for 45 to 60 min at room temperature until cooled to 50° to 60°C. (Drugs and other nutrients are added after 30 min at room temperature.) Just prior to pouring, put the flask on a stir plate at medium to high speed and mix until contents are homogeneous (∼5 min). After pouring, a few bubbles can be removed from the agar surface by passing the flame of a Bunsen burner lightly over the surface of the molten agar (“flaming” the plates). One liter of media will yield 30 to 35 plates.

While a specific brand of petri plate is not required, we recommend Fisher plates (100 × 15 mm, #8-757-12), which have ridges around the tops of the covers to allow easy stacking, making plate pouring less cumbersome.

When preparing the plate recipes below, follow the general guidelines for mixing and autoclaving in the introduction to liquid media. Most plates can be stored at room temperature for ≤4 months. Plates containing drugs (cycloheximide, 5-fluoroorotic, canavanine, and L-α-aminoadipic acid) or Xgal are stable for 2 to 3 months when stored at 4°C.

Minimal Plates and Rich Plates

YPD, minimal, and CM dropout plates

Per liter: Follow recipes for liquid media above, adding 20 g agar and a pellet of NaOH.
Premixes: Follow recipes for liquid media premixes above, adding 500 g agar for YPD premix and 800 g agar for minimal premix. To prepare plates, add one NaOH pellet and the following amounts of premix (per liter):
YPD plates—70 g YPD plate premix
Minimal plates—47 g minimal plate premix
CM dropout plates—47 g minimal plate premix + 1.3 g dropout powder

Specialty Plates

The recipes for α-aminoadipate, canavanine, and cycloheximide plates are included even though no specific use for them is described in this chapter. They are commonly used in negative selection experiments in the same way that 5-FOA plates are used. One can select against the wild-type LYS2, CAN1, and CYH2 genes by growth on plates that select for cycloheximide, α-aminoadipate, or canavanine resistance, respectively.

5-fluoroorotic acid (5-FOA) plates

To a 2-liter flask (containing a stir bar), add:
1 g 5-FOA powder
500 ml H₂O
5 ml 2.4 mg/ml uracil solution
Stir with low heat ∼1 hr until completely dissolved; filter sterilize
To a separate 2-liter flask, add:
1.7 g YNB −AA/AS
5 g (NH₄)₂SO₄
20 g dextrose
20 g agar
1.3 g uracil dropout premix
H₂O to 500 ml and autoclave
(Alternatively, replace first four ingredients with 47 g minimal plate premix)
When the molten agar cools to 65°C, gently add the sterile 5-FOA/uracil solution to the uracil dropout medium by pouring it down the inside wall of the flask containing the latter. Swirl gently to mix and pour the plates.
URA3⁺ strains are unable to grow on media containing the pyrimidine analog 5-fluoroorotic acid (Boeke et al., 1984). This observation has led to methods that use 5-FOA to select against the functional URA3 gene. This type of selection—termed “negative selection” (Brown and Szostak, 1983)—can also be used to select against the wild-type LYS2, CAN1, and CYH2 genes.

Xgal plates,per liter

1.7 g YNB −AA/AS

5 g (NH₄)₂SO₄

20 g dextrose

20 g agar

0.8 g dropout powder (omitting appropriate amino acids; see table)

NaOH pellet

(Alternatively, replace first four ingredients with 47 g minimal plate premix)

Dissection plates,

Follow the recipe for YPD plates, keeping in mind that dissection plates used for tetrad analysis should be of uniform thickness and free of imperfections. After the plates have been poured they should be flamed. Stack the plates on a level surface in piles of six and move gently in a circular motion to “even out” the agar. Certain batches of agar produce plates that have microscopic precipitates embedded in the agar, often looking much like yeast spores. If this occurs, an agar of higher purity can be used, such as Noble agar (Difco) or agarose.

α-aminoadipate plates

1.7 g YNB −AA/AS
20 g dextrose
20 g agar
H₂O to 1 liter
Add ingredients to a 2-liter flask and autoclave. When the molten agar cools to ∼65°C, add 34 ml of a solution of 6% L-α-aminoadipic acid (prepared by dissolving α-aminoadipate in 100 ml water and adjusting the pH to ∼6.0 with 1 M KOH). The final concentration of α-aminoadipic acid in the plates should be 0.2%. Swirl gently to mix and pour the plates.

Lys2− yeast use α-aminoadipic acid as an alternate nitrogen source. Since yeast can use certain amino acids as nitrogen sources, only those amino acids which are required by the particular strain being used should be added to these plates. These can be added from liquid stocks prior to autoclaving. When using this media for isolating lys2− yeast, lysine must be added at 30 µg/ml.

Canavanine plates

Follow recipe for complete minimal (CM) dropout plates, omitting the nutrient arginine. When the 1-liter autoclaved solution has cooled to ∼65°C, add 10 ml of 6 mg/ml filter-sterilized canavanine sulfate solution. The final concentration of canavanine should be 60 µg/ml.

The sterile canavanine sulfate solution can be stored frozen.

Cycloheximide plates

Follow the recipe for YPD plates. When the agar cools to ∼65°C, add 1 ml of 10 mg/ml filter-sterilized cycloheximide solution. The final concentration of cycloheximide in the plates should be 10 µg/ml.

The sterile cycloheximide stock solution can be stored frozen.

STRAIN STORAGE AND REVIVAL

Yeast strains can be stored at −70°C in 15% glycerol (viable for >5 years), or at 4°C on slants consisting of rich medium supplemented with potato starch (viable for 1 to 2 years). Both methods are described below.

Preparation and Inoculation of Frozen Stocks

Make a solution of 30% (w/v) glycerol. Pipet 1 ml into 15 × 45-mm, 4-ml screwcap vials. Loosely cap the vials and autoclave 15 min.

To inoculate vials for storage, add 1 ml of a late-log or early-stationary phase culture, mix, and set on dry ice. Store at −70°C. Revive by scraping some of the cells off the frozen surface and streak onto plates. Do not thaw the entire vial. Cells can also be stored in the same way by adding 80 µl dimethyl sulfoxide (DMSO) to 1 ml cells (8% v/v) and storing at −70°C.

Preparation and Inoculation of Slants

1. For 250 slants, add the following ingredients to a 1-liter flask:
70 g YPD plate premix
20 mg adenine (hemisulfate salt)
20 g potato starch
H₂O to 500 ml

Excess adenine prevents ade− mutations from being lost.

2. Set the flask in a 4-liter beaker filled with 1 liter of water. Place the beaker on a heat-controlled, magnetic stirring apparatus and stir with the heat setting on high.

With smooth and continuous stirring, the contents should not burn and should be molten after ~1 hr.

3. With the entire setup in place, begin pipetting 2-ml aliquots into 15 × 45-mm, 4-ml screwcap vials. When all vials have been filled, put the caps on loosely, pack in the original boxes, and autoclave 15 min.

4. Lean the boxes against a support at an angle of ~70°. Allow the slants to dry 2 days and screw the caps on tightly.

Slants can be stored at room temperature for at least 6 months.

5. To inoculate a slant, smear cells from the flat end of a sterile toothpick onto the agar surface of the slant. Cap loosely and incubate 1 or 2 days at 30°C. After growth, screw the cap on as tightly as possible and store at 4°C.

Slants are a convenient way to store and mail strains. They can be mailed immediately after inoculating since sufficient growth will occur in transit. See another method for mailing strains below.

Mailing and Reviving Strains

Yeast strains can be conveniently mailed as slants. Alternatively, transfer cells to a piece of sterile Whatman 3MM paper by pressing the paper onto the desired yeast colony using forceps that have been dipped in ethanol and flamed. Wrap the paper in sterile aluminum foil and mail to recipient.

Revive the strain by placing the paper (yeast side down) on the surface of an agar plate. Incubate the plate at 30°C. A thick patch of yeast should be visible after lifting the paper.