Media Protocols

Yeast DNA Mini-Plasmid Prep

1. Take a large loopful of cells from a plate (the type of media doesn’t matter). If there are no sufficiently large colonies (ª3 mm in diameter) then scoop up the primary growth streak.

2. Resuspend the cells in 200µl of lysis buffer in a 1.5ml microfuge tube.

Lysis Buffer: 300 mM NaCl

10 mM Tris, pH 8

1 mM EDTA, pH 8

0.1 % SDS

2. Add 300 µl of 0.45 nm diameter glass beads*.

3. Vortex vigorously for one minute. in a VWR multiple tube shaker™(setting 7.)

4. Add 500 µl of Phenol-Chloroform and vortex again for one minute.

5. Centrifuge for five minutes in a mucrofuge.

6. Pipette off the aqueous phase and reextract once more with 500 µl of Phenol-Chloroform and once with 300 µl of Chloroform.

7. Precipitate the DNA by adding two volumes of ice cold Ethanol.

8. Resuspend the pellet in 100 µl of TE.

*The glass beads washed in 1M HCl followed by repeated washes with distilled water. The beads must then be dried completely before use (usually 80°C ON works.)

URA Dropout

Ade 1.6

Arg 1.6

Asp 8.0

His 1.6

Leu 4.8

Lys 2.4

Met 1.6

Phe 4.0

Thr 16.0

Trp 1.6

Tyr 2.4

In vitro ß-Galactosidase Time Assay

1. Grow 3 ml ON cultures in selective media.

2. Dilute the cells, into 6 ml of fresh media, to an OD600; = 0.3 and grow the cultures for 3h to an OD600 between 0.5 and 1.0 (assuming 107 cells per OD600)

3. Measure the OD600; Take 1 ml samples of each culture and pellet the cells in microfuge tubes.

4. Resuspend the cells in 1 ml of Z buffer.

Z Buffer: 16.1 g. Sodium dibasic •7H2O

5.50 g. Sodium monobasic • H2O

0.75 g. KCl

0.25 g. MgSO4•7H2O

Q.S. to 1 l. with H2O.

2.7 µl. ß-mercaptoethanol is added, per ml, just before use

5. Transfer the cells to a glass tube and add 25 µl. of 0.1% SDS and 50µl. Chloroform to each sample.

6. Vortex for 1 min.

7. Preincubate the samples for 5 min. at 28°C.

8. Add 200 µl. of 4 mg/ml ONPG in 100 mM Na phosphate pH 7 to each sample and mix; Record this time as TIME 0.

9. Incubate at 28°C.

Since the amounts of ß-galactosidase will vary between samples the time that it takes for the yellow color to develop will also vary (up to 2h.); therefore, each tube has to be watched carefully.

10. When a sample turns yellow add 0.5 ml. 1M Sodium carbonate to stop the reaction. Record the time you stopped the reaction as TIME t.

11. Pellet the cell debris in a microfuge (2 min.) and carefully remove the supernatent without distubing the loose pellet or removing any Chloroform.

12. Measure the OD420 for each sample and calculate the units as follows.


ß gal units ? 1000 ¥ æææææææ with ?t ??TIME t-TIME 0 (in min.)º

?t v (OD600) and v ??sample volume (usually 1 ml)

Media Recipes

Bacterial Growth Media:

Luria-Bertiani Media (per liter)

10 g Bacto Tryptone

5 g Yeast Extract

10 g Sodium chloride

(for plates add 20 g of Bacto Agar)

Terrific Broth (per liter)

Solution 1 Solution 2

12 g Bacto Tryptone

24 g Yeast Extract 2.31 g Potassium monobasic

4 ml Glycerol 12.54 g Potassium dibasic

Q.S. to 900 ml Q.S. to 100 ml

Combine both solutions after autoclaving

Drug concentrations:

Ampicillin 100 µg/ml

Kanamycin 25 µg/ml

X-gal 20 µg/ml

Yeast Media:

10X YNB: (per liter)

50 g Ammonium sulfate

17 g Difco Yeast Nitrogen Base*

Q.S. to 1L w/Water

Filter sterilize

* w/o Amino Acids and Ammonium sulfate

YPD: (per liter)

10 g Yeast extract

20 g Bacto Peptone

20 g Glucose

(for plates add 20 g Bacto Agar)

Glucose Minimal Media: (per liter)

Combine the following:

20 g Glucose

870 mg Dropout mix

100 µl of 10N NaOH (or 1 NaOH pellet)

900 ml Water

After autoclaving add:

100 ml 10X YNB

5 ml 200X Tryptophan

Galactose Media:

For Galactose media 100 ml 20% Galactose is substituted for the 20g Glucose in each recipe. As a result you should compensate for the volume of Galactose when you add the water.

20% Galactose appears to suffer from

autoclaving so it is usually

filter sterilized.

When making Minimal Media, NaOH serves two functions:

First it aids in the solubilization of some of the amino acids.

Second it protects the agar when autoclaving.

Without the NaOH the agar would be hydrolysed in the acidic solution…This poses a special problem with Galactose Minimal Media plates.

When autoclaving Bacto Agar and the Dropout mix together, the agar will often become opaque and an undesirable shade of gray. Glucose (present when autoclaving Glucose Minimal Media Plates) seems to prevent this color change. As a result I have taken to preparing Galactose Minimal Media Plates the following way:

For 1 liter, 20g agar and 800ml Water

are autoclaved in a 2L flask.

In a 250ml flask, 870 mg of Dropout mix

and 1 NaOH pellet are autoclaved in 100ml

of water.

After autoclaving the contents of the two

flasks are combined, along with 100ml 10X YNB,

the Galactose and 5 ml of 200X Tryptophan.

5-FOA Plates: (per liter)


20 g Glucose

20 g Bacto Agar

870 mg URA Dropout mix

100 µl 10N NaOH (or 1 NaOH pellet)

500 ml Water

Autoclave for 25 min.


(It turns it orange )

Combine the following:

100 ml 10X YNB

1 g 5-FOA

5 ml 200X Tryptophan

6 ml 100X Uracil

Q.S. to 500 ml w/Water

Mix on a magnetic stirrer until the 5-FOA goes into solution

then heat the solution in a 55°C waterbath or in a microwave oven

After autoclaving the agar mixture, combine the agar and FOA solutions and pour