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High Efficiency Transformation-PDF

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High Efficiency Transformation-PDF

Protocol

Day 0

  • Make sure you have the necessary solutions (instructions for how to make them can be found here):
    1. Single-stranded carrier DNA
    2. PEG 3350 50% w/vol
    3. 1.0 M LiAc
  • Make sure you have enough plasmid DNA for your transformations. Each transformation should use between 0.5 and 1.0 µg of plasmid.

Day 1

Inoculate the yeast strain into 5 ml of liquid medium and incubate overnight.

Day 2

  1. Determine the titer of the yeast culture by pipetting 10 µl of cells into 1.0 ml of water in a spectrophotometer cuvette and measuring the OD at 600 nm. For many yeast strains a suspension containing 1 x 106 cells/ml will give an OD600 of 0.1.
  2. Transfer 50 ml of YPD to the pre-warmed culture flask and add 2.5 x 108 cells to give 5 x 106 cells/ml.
  3. Incubate the flask on a rotary or reciprocating shaker at 30°C and 200 rpm.
  4. Boil a 1.0 ml sample of carrier DNA for 5 min and chill in an ice/water bath while harvesting the cells (the next step).
    • It is not necessary or desirable to boil the carrier DNA every time. Keep a small aliquot in your own freezer box and boil after 3-4 freeze-thaws. But keep on ice when out.
  5. When the cell titer is at least 2 x 107 cells/ml, which should take about 4 hours, harvest the cells by centrifugation at 3000 g for 5 min, wash the cells in 25 ml of sterile water and resuspend in 1 ml of sterile water.
    • If the cell titer is higher/lower than 2×107 cells/ml, you’ll dilute it back down/up in a subsequent step
    • Now would also be a good time to start warming up the 42°C water bath you’ll need later on
  6. Transfer the cell suspension to a 1.5 ml microcentrifuge tube, centrifuge at 3000 g for 30 sec and discard the supernatant.
  7. Add water to a final volume of 1.0 ml and vortex mix vigorously to resuspend the cells.
    • If your cell titer was higher/lower than 2×107 cells/ml, add proportionally more/less water at this step. Also, note that if you split your initial 50ml of culture into multiple tubes, the water should be split appropriately between the tubes.
  8. Pipette 100 µl samples (ca. 108 cells) into 1.5 ml microfuge tubes, one for each transformation, centrifuge at 3000g for 30 sec and remove the supernatant.
  9. Make up sufficient Transformation Mix for the planned number of transformations plus one extra. Keep the Transformation Mix in ice/water.
It’s generally easiest to make a master mix of PEG, LiAc and the SS-carrier DNA and add 326 µl from the master mix to each transformation tube before adding the plasmid DNA + water component.
For the plasmid DNA + water component, you’ll need to calculate, for each transformation, what vol. of your plasmid DNA solution contain the appropriate amount of plasmid DNA (0.5-1.0 µg). Add in the appropriate amount of plasmid DNA solution and then make it up to 34µl with water.
Number of Transformations
Reagents 1 5 (6X) 10 (11X)
PEG 3350 50% w/v 240 µl 1440 µl 2640 µl
LiAc 1.0 M 36 µl 216 µl 396 µl
Boiled SS-carrier DNA 50 µl 300 µl 550 µl
Plasmid DNA plus Water 34 µl 204 µl 374 µl
Total 360 µl 2160 µl 3960 µl
  1. Add 360 µl of Transformation Mix to each transformation tube and resuspend the cells by vortex mixing vigorously.
  2. Incubate the tubes in a 42°C water bath for 40 min.
  3. Microcentrifuge at 3000 g for 30 sec and remove the Transformation Mix with a micropipettor.
  4. Pipette 1.0 ml of sterile water into each tube; stir the pellet by with a micropipette tip and vortex .
    • We like to be a gentle as possible at this step if high efficiency is important. Excessive washing washes away transformants.
  5. Plate appropriate dilutions of the cell suspension onto SC selection medium. For transformation with an integrating plasmid (YIp), linear construct or oligonucleotide, plate 200 µl onto each of 5 plates; for a YEp, YRp or YCp library plasmid dilute 10 µl of the suspension into 1.0 ml of water and plate 10 and 100 µl samples of the dilute onto two plates each. The 10 µl samples should be pipetted directly into 100 µl puddles of sterile water on the SC selection medium.

Plating out phage-PDF

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Materials

  • Host culture
  • Phage stock, serially diluted in SM buffer (100 mM NaCl, 25 mM Tris-HCl pH 7.5, 8 mM MgSO4, 0.01% (w/v) gelatin) or in sterile liquid culture medium
  • Petri plates containing solid culture medium
  • Aliquots of molten soft agar for overlays (4 – 5 ml each, enough to cover one plate), held at ~50 ºC in a heat block or water bath
  • Tube racks, pipettors, sterile tips
  • Aseptic working area (near a flame or in an aseptic cabinet)

Procedure

Spot titration of phage

  1. Pour a bacterial lawn
    1. Label the bottom surface of a culture plate with the phage and lawn strain names, and mark the positions where phage dilution spots will be deposited.
    2. Place the plate face up near a flame or in an aseptic cabinet.
    3. Retrieve a top agar aliquot from the water bath.
      The agar will start to solidify when it cools to below about 40 ºC, so you must work quickly for this procedure.
    4. Transfer 100 μl of the host culture into the molten agar.
    5. Vortex the molten agar on medium speed, briefly (about 1 second)
    6. Open the plate and pour the contents of the tube onto the agar surface. Swirl the plate gently to distribute the molten agar across the plate.
    7. Replace the plate lid, and allow the plate to sit undisturbed for a few minutes to let the agar solidify.
  2. Apply phage spots to lawn from serial 10-fold dilutions
    1. Place the plate containing the bacterial lawn you prepared in step 1 face up. Open the lid and set it aside.
    2. Aspirate 10 μl of the lowest phage dilution and deposit it over the corresponding label marked on the bottom of the plate. Keep the pipette tip close to the surface of the plate to prevent the drops from sliding.
    3. Repeat this procedure for each of the phage dilutions, depositing a 10 μl drop of diluted phage over its corresponding dilution marked on the bottom of the plate.
    4. When done, leave the lid off of the plate and allow the plate to dry near a flame or in an aseptic cabinet until the drops are absorbed, about 10-15 min.
    5. Incubate the plates, inverted, at the optimal growth temperature of the host until plaques appear in the lawn. The lower phage dilutions will yield confluent spots of lysis, which should give way to individual plaques in the spots at higher dilutions.
      Individual plaques may be counted in these spots but be aware that the counting error will be high.

Full-plate titration of phage

  1. Determine the approximate titer of your phage stock, by spot titration or previous experience. Determine which serial dilution of your phage stock will yield ~50 to 500 plaques in one 100 μl aliquot. Use this dilution, the dilution above it and the dilution below it for plating.
  2. Prepare 3 plates by labeling them with the bacterial strain name, the phage name and the dilution to be plated. Place them face-up near the working area.
  3. Place 3 empty sterile top agar tubes into a rack, label them with the three dilutions you will be plating.
    1. Aliquot 100 μl of the host culture into each tube.
    2. Aliquot 100 μl of the phage dilutions you will be plating into their corresponding tubes containing host culture. Vortex the tubes briefly to mix.
  4. Retrieve one molten agar aliquot from the water bath.
  5. Pour the aliquot from its tube into one of the tubes containing the host cell-phage mixture, vortex briefly, and pour the contents onto the appropriately labeled plate. Swirl gently, replace the lid and allow to sit undisturbed until the lawn sets. Repeat this step with the other dilutions.
  6. Incubate the plates, inverted, at the optimal growth temperature of the host until plaques appear in the lawn.

 

 

Layered plates-PDF

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Layered plates-PDF

Overview

The cells being plated are sandwiched between layers of cell-free agar. The submerged cells grow into smaller colonies and many more colonies can be counted per plate (I’ve been able to distinguish over 1000 yeast colonies per plate using this method–Jgritton 14:46, 22 Aug 2005 (EDT)). In addition by avoiding the use of a spreader, you can eliminate the error due to cells sticking to your spreader. The result is more accurate estimates of cell numbers in a sample.

Procedure

  1. Prepare petri plates with 0.4 cm of base agar medium.
    • LB (or other nutrient) with 1.2% agar for bacteria
    • YPD (or other nutrient) with 2% agar for yeast
  2. Prepare 0.7% agar medium (soft agar medium)
  3. Warm small test tubes to 45 C and add 3 mL of soft agar medium.
  4. Pipette sample to be plated onto inside rim of test tube.
  5. Immediately pour soft agar onto the base agar, pouring over the pipetted sample.
  6. Swirl plate to spread sample/agar mix and allow to set
  7. Pipette or pour 2 mL of soft agar onto plate and swirl to spread. Allow to set.
  8. Incubate plates

Materials

YPD Soft Agar

  1. In a 2L flask add:
    • 10g Yeast Extract
    • 20g Peptone
    • 7g Agar
  2. Add water to 950 mL
  3. Autoclave
  4. Add 50 mL 40% glucose
  5. Dispense into bottles (~330 mL into each of three 500 mL bottles) and allow to cool
  6. Microwave to melt agar before use

Reference

Koch, A.L. Growth Measurement in Methods for General and Molecular Biology 1994 pg 249-277

  • MIT Library Call #: QR65.M26 1994 (Hayden Library)

Lysate for Western-PDF

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Protocol

  1. Grow cells
  2. Harvest
    • Spin down
    • Wash with PBS
    • Spin down enough cells to collect a 50-100 µL cell pellet in a FastPrep tube
  3. Prepare lysis buffer (beforehand) and chill on ice
    • PBSMT:
      • 2.5 mM MgCl2
      • 3 mM KCl
      • 0.5% Triton X-100
      • in PBS
    • Add protease inhibitors
      • PMSF: 1 mL per 100 mL buffer
        • Use fresh PMSF: dissolve 0.035g per 1mL 100%EtOH
      • PLAAC: 100 µL per 100 mL buffer
      • 0.5 M Benzamidine: 260 µL per 100 mL buffer
  4. Add 50-100 µL of cold lysis buffer to cell pellet and vortex
  5. Add 1-2 scoops glass beads (bead level should be just below liquid level)
  6. Mix everything together by vortex
  7. Lyse in FastPrep bead beater
    • 6.5 m/s
    • 40 second lysis
  8. Add 500 µL lysis buffer (keep everything on ice!)
  9. Spin down at maximum speed in tabletop centrifuge for 10 min. (in cold room)
  10. Remove supernatant (lysate) to another tube.

Colony PCR-PDF

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Zymolyase Solution:

  • Regular stock of zymolyase is 10 mg/ml diluted in water, mix by inverting, not all will go into solution (filter it) store aliquots at -20°C (minimize freeze-thawing if possible)
  • Make 2.5 mg/ml zymolyase solution in 0.1 M sodium phosphate buffer pH 7.5 immediately before the PCR experiment

Lysis of Yeast:

  • Aliquot 15 µl 2.5 mg/ml enzyme solution (in 0.1 M sodium phosphate buffer pH 7.5) into thin-walled PCR tubes
  • Scrape a small amount of yeast colony and resuspend in a PCR tube (one lysis reaction when dilute will yield ~15 PCR reactions, only 5 µl is used for a template in each PCR reaction)
  • Incubate on the bench (RT) for 20 min
  • Place in PCR block and heat to 37°C for 5 min and then 95°C for 5 min

PCR Step:

  • Dilute the lysis solution 1:5 by adding 60 ul ddH2O to each PCR tube
  • Place 5 ul of this dilution into a fresh PCR tube
  • Add PCR mix (usually 45 ul total per reaction):
    • 2.5 µl 5’ Primer (10 µM)
    • 2.5 µl 3’ Primer (10 µM)
    • 5 µl Thermopol Buffer
    • 1 µl dNTPs (10 uM)
    • 33.5 µl ddH2O
    • 0.5 µl AmpliTaq or Vent DNA Polymerase
  • Run the Lyse Program (LysAn50), which takes about 5 hours
  • Check product by running 10 µl PCR sample + 2 µl 6x DNA dye on 1% agarose gel

Assaying mating-PDF

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Assaying mating-PDF

Setup

You have yeast strains that are deficient in mating (eg Ste12 knockouts) and would like to test whether transforming them with a plasmid that contains genes that are supposed to complement the mating deficiency restores the ability to mate.

Principle

You’ll need two strains to use as mating tester strains, one MATa strain, and one MATalpha strain; these tester strains should have mutations in an unusual auxotrophic marker, like lys1.

The principle of the assay, which is simply done on agar plates, is that if the experimental strain, which should have at least one of the more common auxotrophic markers (e.g., trp1 or ura3), is mated to the mating tester strains, any resulting diploids will be able to grow on synthetic medium with no added nutrients but neither haploid will be able to grow.

Alternatively, or in addition, using the same principle, the strains can be mated to each other eg by transforming the MATa strain with, for example, any URA3 plasmid (like the pRS316 vector) and transforming the MATalpha strain with, for example, any TRP1 plasmid (like the pRS314 plasmid) and then mating the two strains together–only the diploid will grow on the minimal plates.

Materials

  1. MATa and MATalpha tester strains with unusual auxotrophic markers (eg lys1)
  2. YPD plates
  3. Synthetic dropout plates with added nutrients to grow your tester strains
  4. Minimal synthetic plates with no added nutrients (just YNB, glucose, agar, water)
  5. Velvets and a replica block for replica plating
  6. Strains to be tested, with and without the plasmid(s) transformed into them

Protocol

  1. Patch your strains and appropriate positive and negative controls on the same plate E.g., you can make 1 cm x 1 cm patches on a single -aura plate of the following strains: your MATa strain transformed with an empty URA3 vector, your MATa strain transformed with a URA3 plasmid containing the genes that are supposed to restore mating, a control MATa strain that is URA3, a control MATalpha strain that is URA3
  2. Grow for 1-2 days at 30 degrees
  3. Take a glob of tester strain (a sphere with a diameter of about 1-1.5 mm) with a toothpick from a colony or patch on a plate, and resuspend in 1 ml YPD liquid in a 1.5 ml tube. Vortex vigorously to mix.
  4. Pipet 200 ul of tester cell suspension on a YPD plate and spread evenly. Allow to dry for 5-60 min.
  5. Replica plate cells from your patch plate (made in 1 above) onto the plate that has evenly spread tester cells (made in 4 above).
  6. Incubate at 30 degrees for 6-18 hrs to allow mating to occur
  7. Replica plate cells from your mating plate (made in 5 above) onto a minimal synthetic plate with no added nutrients
  8. Incubate at 30 degrees for 2-3 days
  9. Diploids should grow and haploids should not grow, allowing you to detect mating as growth

Dropout plates for yeast-PDF

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Dropout plates for yeast-PDF

Materials

(Solutions are all available from the media room)

  • 200ml bottle of 2x SD
  • 200ml bottle of 4% agar — make sure to sign it out
  • 40% glucose
  • CSM minus the relevant marker(s), as powder
  • Stack of plates (above the -20 freezers)

Yeast growth medium generally consists of three parts: a nitrogen base (supplied by SD), a sugar source (supplied by glucose), and necessary amino acids (supplied by CSM). These three components are what you’re going to mix and then add to agar to make plates.

Making amino acid dropout (CSM) solution

Generally easiest to make a 10x master solution that you can dilute back when you’re pouring plates. Let’s assume you’re going to make 200ml of a 10x master solution:

  1. Grab a 500 ml conical flask
  2. Figure out how much of your dropout powder you’re going to need and weigh it out
    • For example, the CSM – TRP mix we have is 0.74g/L, so for a 200ml of a 10x solution, you’ll need 10*0.74/5 = 1.48g.
  3. Dump CSM into conical flask and fill it up to 200ml mark with deionized water (from white tap above the sink)
  4. The CSM will need a bit of help dissolving, so put in a stir bar (rinse it off first with dI water) and put it on a stirrer for 1-2 hours (speed around 6-7, heat 3-4).

Once the CSM has gone into solution, you’ll need to filter-sterilize it:

  1. Get a 200ml orange screwtop bottle and a vacuum filter
  2. Flame the bottle & top, remove the filter from its packaging and screw it onto the bottle. The filter is sterile, so try not to touch it to anything else
  3. Attach the adapter to the filter and the vacuum nozzle to the adapter (vacuum should be off)
  4. Pour the CSM solution into the top of the filter
  5. Turn on vacuum and allow the solution to get sucked through the filter and then turn off the vacuum
  6. Unscrew the filter, flame the bottle and top again, recap it and label it.
  • Alternatively, see here to make DO stocks from individual amino acids.

Mixing growth medium

  1. Add 20ml of 40% glucose and 40ml of 10x CSM solution to 200 ml of 2x SD
  2. Heat up mixture to 65 degrees in water bath

Preparation of LB Agar

  1. Melt in microwave:
    • loosen the cap
    • use 50% power (enter time, press Power, 5, Start)
    • monitor as you melt
    • takes approx. 3-5 minutes for a 200ml bottle.

Making agar-growth medium mix

  1. Mix agar and growith medium pouring the growth medium into the agar bottle — no need to cool down the agar first, it gets cooled by mixing with the growth medium
    • Since the total volume is now ~400 ml, you now have a 1x everything, 2% glucose solution, as desired.
  2. Swirl gently to mix, put in water bath for a bit until bubbles disappear

Actual Pouring

  1. Using sterile technique (flame the top of the bottle) pour the agar mixture into the plates.
    • Cover the base of the plate, and then just a bit more after that.
  2. Recap each plate upon pouring. If there are lots of bubbles in your plates (i.e., more than one or two on the edge), you can flame the plate using the small bunsen burner to eliminate bubbles. (See a demo on this). Another way to remove the fine bubbles that may be in your flask before puring is to mist the inside of the flask with a 75% ethanol spray bottle.
  3. Leave plates to dry and cool for a while (overnight even).
  4. Store the plates in their original bags – upside down, so that the gel is hanging downwards (this keeps condensation off the gel).
  5. Label the bags, using correct color tape for the dropout medium
  6. Put bags in a refrigerator

Spheroplast Transformation-PDF

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Materials

  • YPD plates
  • YPD
  • 1 M sorbitol 182 g/l (Sigma S7547)
  • 2 M sorbitol 36.4 g/100 ml
  • SCE (per liter)
    • 1 M sorbitol (182 g)
    • 100 mM citric acid trisodium salt dihydrate (29.4 g)
    • 10 mM EDTA 20 ml of 500 mM EDTA solution
    • final pH to 5.8 with HCl, autoclave, store at RT
  • 1 M DTT solution Sigma 43816
  • Calf-thymus DNA (phenol chloroform purified and precipitated)
  • Lyticase
    • Sigma L4025
    • final concentration 10,000 units/ml in
      • 20 mM phosphate pH 7.5
      • 50% glycerol
      • store in single use aliquots at -80C
  • Sorb-Trp-Ura plates (per liter)
    • 1 M sorbitol (182 g)
    • 0.17% w/v yeast nitrogen base w/o amino acids w/o ammonium sulphate (Difco 0335-15) (1.7 g)
    • 0.5% ammonium sulfate (5 g)
    • 0.6 g/l SC-Trp-Ura dropout mix (0.6 g)
    • 2% agar (20 g)
    • Adjust pH to 5.8 with 1 M NaOH
    • water to 900 ml
    • autoclave, cool to 55C
    • Add 2% w/v glucose (100 ml of a 20% solution) to final 1 liter volume
  • Sorb-Trp-Ura top agar (per 100 ml)
    • 1 M sorbitol (18.2 g)
    • 0.17% w/v yeast nitrogen base w/o amino acids w/o ammonium sulphate (Difco 0335-15) (.17 g)
    • 0.5% ammonium sulfate (0.5 g)
    • 0.6 g/l SC -Trp-Ura dropout mix (60 mg)
    • 2.5% agar (2.5 g)
    • Adjust pH to 5.8 with 1 M NaOH
    • water to 90 ml
    • autoclave, cool to 55C
    • Add 2% w/v glucose (10 ml of a 20% solution) to final 100 ml volume

Material prepared just before use

  • STC (per 60 ml)
    • 0.98 M sorbitol 58.8 ml of a 1 M solution
    • 10 mM Tris pH 7.5 600 μl of a 1 M solution
    • 10 mM CaCl2 600 μl of a 1 M solution
    • prepare from sterile solutions just before use
  • STC + Calf thymus DNA
    • Add 50 μg/ml of calf-thymus DNA (phenol chloroform purified) to STC
    • prepare from sterile solutions just before use
  • PEG solution (for 100 ml)
    • 19.6% PEG 8000 w/v (98 ml of a 20% solution Sigma P2139)
    • 10 mM Tris pH 7.5 (1 ml of a 1 M solution)
    • 10 mM CaCl2 (1 ml of a 1 M solution)
    • prepare from sterile solutions just before use
  • SOS (for 30 ml)
    • 1 M sorbitol (15 ml of a 2 M solution)
    • 7 mM CaCl2 (210 μl of 1 M solution)
    • 25% v/v YPD medium (7.5 ml)
    • .0025% w/v uracil (75 μl of a 1% solution)
    • water 7.2 ml
    • prepare from sterile solutions just before use
  • Microwave Sorb-Trp-Ura top agar, hold at 50 C

Protocol

  1. Pick a single colony from a YPD plate containing the correct strain into 5 ml of YPD medium and grow overnight at 30C with shaking. Do not inoculate from a plate stored at 4C.
  2. Add 200-800 μl of the overnight culture to 350 ml of YPD medium in a 1 liter flask and grow 14-18 hours to a final concentration of 5 x 107 cells/ml (mid log phase).
    1. inoculating several different amounts of overnight culture into different flasks allows one to choose the flask which is ready at the time of measurement.
  3. Centrifuge 2x 150 ml at 22C 1000-1200g for 5 minutes in flat bottom centrifuge tubes.
  4. Wash the cells in 20 ml of sterile water
  5. Wash a second time in 20 ml of 1 M sorbitol
  6. Resuspend in 20 ml of SCE and transfer to two 50 ml centrifuge tubes
  7. Add 100 μl of 2 M DTT and the optimal amount of lyticase
    1. See below on determining the amount of lyticase
  8. Incubate samples at 37C for exactly 15 minutes
  9. Centrifuge at 200-300g at 22C for 5 minutes
  10. Remove supernatant by careful aspiration
    1. The pellet will be soft and fluffy; not all cells will be pelleted
  11. Gently add 20 ml of 1 M sorbitol down the side of the tube and swirl to resuspend (do not vortex)
  12. Centrifuge and remove supernatant, as above
  13. Wash again with 20 ml of STC
  14. Resuspend in 6 ml of STC + 50 μg/ml of calf-thymus DNA and combine into a single tube
  15. With cut-tip add 300 ul of spheroplast suspension to sterile 15 ml tubes.
  16. With cut-tip add up to 50 ul of ligated YACs or other DNA for each 300 ul of spheroplast suspension
  17. Incubate samples for 10 min at room temp. (If DNA is added to cells every 30 sec. then 20 transformations can be done in 10 min.).
  18. Add 3 ml of PEG solution and stir carefully resuspend the spheroplasts.
  19. Incubate at room temp for 5-10 min.
  20. Centrifuge at 22°C and 200-300g for 5 minutes.
  21. Remove and discard s/n without loosing the spheroplasts (better with pipette).
  22. With cut-tip add 1 ml of SOS and gently resuspend by slow pipetting.
  23. Incubate at 30°C without shaking for 30 minutes.
  24. Plate spheroplasts as follows:

A) Standard platting: 1. Invert the tube once and add 5 ml (12 ml if 15 cm plates are used) of molten top agar. 2. Close the tube and mix twice by inversion. 3. Pour onto a 37°C pre-warmed sorb-containing plates 4. Tilt the plate to distribute evenly the agar. 5. Incubate upright for at least 15 min. at room temp. 6. Grow at 30°C for 5-6 days (preventing drying after the 2nd day).

B) Platting facilitating “colony breakthrough”: 1. Invert the tube once and add 10.5 ml (with 10 ml plastic pipette) of molten top agar. 2. Close the tube and mix twice by inversion. 3. With the same pipette used in step 1 distribute the spheroplasts-agar onto a big (15 cm) sorb-containing plates pre-warmed to 50°C. 4. Tilt the plate to distribute evenly the agar (this is critical). 5. Incubate upright for at least 15 min. at room temp. 6. Grow at 30°C for 5-6 days (preventing drying after the 2nd day).

  1. Yeast Artificial Chromosomes, Green E.D., Hieter, P., and Spencer F. A., chapter 5 in Genome Analysis: A Laboratory Manual, Vol. 3, Cloning Systems, Birren et al. (eds.), Cold Spring Harbor Press, New York, 1999[Green99]

Sporulation-PDF

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Overview

To sporulate yeast cells need to be starved for nitrogen in the presence of a nonfermentable carbon source. This protocol uses acetate as the carbon source.

Materials

  • 1% Potassium Acetate (autoclaved)

SPORULATION Mark Hickman uses this protocol and regularly has sporulation efficiencies of >50%. 1. Grow a 5 mL overnight culture; dilute 1:50 in the morning and grow for 4 hours at 30°C (to log phase). 2. Pellet cells, wash in 1 mL of 1% potassium acetate, and resuspend in 3-4 mL of 1% potassium acetate. 3. Incubate at room temperature on a roller wheel for 3 days (can incubate longer, if desired). NOTE: When sporulating, if diploid is homozygous for an auxotrophic mutation, add that nutrient to the potassium acetate (about ¼ the amount listed for addition to SD media… any more and it could be used as a nitrogen source).

Stock Solutions

1% Potassium Acetate

Protocol

  1. Grow a 5 mL overnight culture; dilute 1:50 in 5 mL in the morning and grow for 4 hours at 30°C (to log

phase).

  1. Pellet cells, wash in 1 mL of 1% potassium acetate, and resuspend in 3-4 mL of 1%

potassium acetate.

  1. Incubate at room temperature on a roller wheel for 1 day and then transfer to 30°C for 2 more days (can incubate longer, if desired).
  2. Check culture for spore formation on day 2 and continue checking until you see tetrads.

References

 

 

Electroporation of E. coli-PDF

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Procedure

1. Chill the electroporation cuvettes by floating them in an ice bath.
2. Remove vials containing 100μl electro-competent cells from the -80°C freezer and thaw them with the iced cuvettes.
3. Prepare micro-centrifuge tubes containing 900μl SOB media.
4. Turn on electroporator and set voltage to 1.5 kV (1mm cuvettes).
5. Add 5μL of ligated DNA sample to 100μl thawed electrocompetent cells on ice. Swirl the tip around gently in cells to mix DNA and cells.
6. Place cells back on ice to ensure they remain cold.
7. Pippette 100μL of cell-DNA mixture to the cuvette.
8. Wipe off excess moisture from outside of the cuvette.
9. Place cuvette in chamber of electroporator.
10. Pulse the cells by pressing the button on the electroporator twice.
11. Quickly use a pipette to remove the electroporated cell suspension from the cuvette and add it to a tube containing 1ml SOB.
12. Let cells recover at room temperature for 30-60 minutes.
13. Plate 100μl of electroporated cells onto a prewarmed LB-agar plate supplemented with appropriate antibiotics. Incubate plate overnight at 37°C.

 

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