Protocols on Two-Hybrid Exhaustive Screens

Micheline Fromont-Racine

Pierre Legrain

Jean-Christophe Rain

A list of protocols is given in this document. These protocols have been designed in the Laboratoire du Metabolisme des ARN and were prepared for the diffusion of the technique (Nature Genetics, 1997, 16, 277-282)

We are very grateful to Rob van Nues who contributed greatly to the writing of many parts of this document.

Laboratoire du Metabolisme des ARN

Departement des Biotechnologies

Institut Pasteur

25-28, rue du Dr Roux

75724 Paris Cedex 15

France

Genetique des Interactions Macromoleculaires (Alain Jacquier)

Table of contents

Protocol A: The Mating experiment

Protocol B: X-Gal-overlay assay

Protocol C: Filter lift X-Gal assay

Protocol D: PCR on yeast colonies

Protocol E: Identification and classification of the candidates

Protocol F: Plasmid rescue from yeast by electroporation

Protocol G: Plasmid minipreps from E. coli

Protocol H: Yeast transformation

Annex: Media

Protocol A : The Mating experiment

The mating procedure allows a direct selection on selective plates because the two fusion proteins are already produced in the parental cells. No replica plating is required. We routinely obtain a 10 to 20% mating efficiency.

Usually, the Y187 strain is transformed either with the FRYL yeast genomic library or human cDNA library. (see annexes A and B for the characteristics of the yeast genomic FRYL library and the human cDNA library, respectively). Independently, bait plasmids are introduced in the CG1945 strain. The Y187 strain contains a sensitive LacZ reporter gene whereas the CG1945 strain has a non-leaky HIS3 reporter gene that enables the selection of positive diploids in the absence of 3-aminotriazol (3-AT).

Alternatively, bait plasmids can be introduced into the L40 strain. This plasmid derived from pBMT116 plasmid. It encodes for LexA fusion protein that can be assayed in a two hybrid system in the L40 strain containing His3 and LacZ reporter genes downstream LexA binding sites. The same Gal4-derived libraries could be used. However, the L40 strain is not deleted for theGAL4 gene and although glucose repression occurs, a residual activation of this gene in the diploid cells promotes the transcription of the parental Y187 strain’s LacZ reporter gene. Thus, in a L40 x Y187 diploid cell, selection is made mostly, not to say only, on the HIS3 reporter gene.

This protocol is written for the use of the FRYL yeast genomic library cloned into the Y187 strain. Any other genomic or cDNA library might be used with minor adaptations to take into account their complexity and the cell density of the frozen cells.

Strains

* CG1945 (MATa Gal4-542 Gal80-538 ade2-101 His3-200 Leu2-3,-112 Trp1-901 Ura3-52 Lys2-801 URA3:: GAL4 17mers (X3)-CyC1TATA-LacZ LYS2::GAL1UAS-GAL1TATA-HIS3 CYHR) tranformed with the pAS2∆∆ bait plasmid (see Annex E for plasmid map)

* Y187 (MATa Gal4∆ Gal80∆ ade2-101 His3 Leu2-3,-112 Trp1-901 Ura3-52 URA3::UASGAL1-LacZ Met-) transformed with the FRYL yeast genomic DNA library (see Annex A).

Day 1 preculture

Materials

* 100 ml flask with 20 ml -W medium

Experiment

Preculture of CG1945 cells carrying the bait plasmid in 20 ml -W medium

Grow at 30°C, vigorous agitation.

Day 2 culture

Materials

* 1 liter flask with 150 ml -W medium

Experiment

at 6 pm

- Estimate OD600 of the -W preculture of CG1945 cells carrying the bait plasmid. Measure OD600 of 4 to 10 fold dilution depending of the preculture. The OD must lie between 0.1 and 0.5 in order to correspond to a linear measurement (1 OD = 107 CG1945 cells using our spectrophotometer).

- Inoculate 150 ml -W at OD600 0.006/ml (may depend on local growth conditions; to be tested with various dilutions).

- Grow o.n. at 30°C, vigorous agitation.

Day 3 mating

Materials

all the material must be sterile

medium and plates (see Annex C)

* 5 YPglu + Tetracyclin (Tet) 6 mg/l plates

* a 50 ml tube with 30 ml -LWH

* 100 ml flask with 20 ml of YPglu

* 75 -LWH + Tet (6 mg/ml) plates

* 2 -L plates

* 2 -W plates

* 2 -LW plates

materials

* multipipettor and 10 ml sterile tips

* glass beads 3 mm

* 50 ml tubes

Experiment

A.M.

- Measure OD600 of the -W culture of CG1945 cells carrying the bait plasmid. It should be around 1; definitely not higher than 1.5.

- For the mating you must use twice as many bait cells as library cells. A vial of the Y187/FRYL yeast genomic DNA library contains 4.108 viable cells. To get a good mating efficiency you must collected the cells at 4.5.106 cells per cm2.

- Estimate the amount of bait culture (in ml) that makes up 80 OD600 units for the mating with the yeast library (1 OD = 107 CG1945 cells).

- Thaw a vial containing the Y187/library slowly on ice.

- Add the contents of the vial to 20 ml YPglu using a sterile plugged 1 ml pipet; rinse the vial with culture liquid.

- Let those cells recover at 30 oC, under gentle agitation for 10 minutes. Set your timer.

Mating

- Put 80 OD600 units of bait culture into a 250 ml flask. (Do not leave the CG1945 cells without agitation because of the aggregation of the cells).

- Add the Y187/library culture to the bait culture. Mix the library/bait cells by hand. Transfer the mixture of diploids into 50 ml sterile tubes. Centrifuge the cells at 5000 g (4000 rpm) for 3 min. Discard the supernatant

- Resuspend the pellet with 2 ml of YPGlu medium.

- Distribute cells in 400 µl samples on YPGlu plates with glass beads. Spread cells by shaking the plates.

- Incubate plates cells-up at 30 oC for 4 h 30mn.

P.M.

Collection of mated cells

- Mark the -L, -W, -LW and -LWH + Tet plates with date and bait-name. Add 5 to 7 sterile glass beads to every plate.

Wash and flame forceps (before placing them inside a sterile 9 cm plate).

- Place the base of a 9 cm plate against the base of a Bunsen burner. Place a filter, with mated cells up, inside the base. Rinse the cells from the filter with 1 ml of -LWH using a Pipetman. Reapply the cell-slurry a couple of times until all cells are washed off. Pipet the cell-slurry into the collection tube. (The cells are clearly visible as a reddish layer and come off as sheets in the beginning). Wash and re-rinse the filter with another ml -LWH and pipet this into the collection tube. Discard filter with original YPglu+Tet plate.

- Repeat this with all the filters. You will end up with about 30 ml of cell-suspension. Take note of the actual volume.

Plate the controls

- Shake cell-suspension and make a 1:1000 dilution by three 10 fold dilution steps: (50 µl mix to 450 µl of fresh -LWH medium, shake well). Spread 50 µl of the 1000 fold dilution on -L, -W and -LW plates. Incubate cells down at 30°C for two days.

Plate the Screen

- Take multipipettor and sterile 10 ml tips.

- Shake cell suspension and distribute cells in 400 µl samples over the 75 -LWH+Tet plates with glass beads. Spread cells by shaking the plates. Incubate plates cells down at 30°C for three days.

This can be done only if one knows the behaviour of the bait (e.g. it does not require addition of 3-AT in the plates). Otherwise, for an unknown bait, the optimal conditions must first be determined by plating samples onto -LWH plates without and with 3-AT at various concentrations (ranging from 1 to 50 mM).

The cell mixture can be stored at 4°C and plated 3 days after the mating.

Day 5 Estimation of diploid number

- Late in the afternoon: check controls of mating as well as some selective -LWH plates. About a hundred diploid colonies should be visible on -LW plates. Multiplying this number by 20 (50µl sample), 1000 (dilution factor) and 30 (volume in ml of the diploids mixture) gives the amount of diploids (i.e. 60.106 when you counted 100 on the -LW plate: 100 X 20 X 30 X 1000).

- Count colonies on control plates. Estimate the mating efficiency by dividing the number of colonies on -LW plate by the number on the -L plate and multiply by 100.

- Colonies should become visible on -LWH+Tet plates.

Protocol B : X-gal-Overlay assay

Introduction

X-Gal-overlay assay is performed directly on the selective medium plates after scoring the number of His+ colonies. This procedure is less sensitive than the filter assay (Protocol C) but it is less time consuming and has the advantage of a better recovery of cells since it does not require freezing the cells in liquid nitrogen.

Material

* Work under the hood. Dimethylformamide (DMF) is toxic.

* Set up a waterbath. The water temperature should be 50 oC.

* Stock solutions:

- 0.5 M Na2HPO4 pH7.5 (71 g Na2HPO4 per 1 liter + 4 ml orthophosphoric acid). Distributed per 250 ml and stored at 50°C (neither filtrated nor autoclaved).

- 1.2 % Bacto-agar distributed per 210 ml (2.52 g/210 ml). Autoclaved and stored at 50°C.

(Alternatively, phosphate and agar solutions are kept at room temperature and the solutions are mixed just before use after melting the agar in a microwave oven)

- 2 % X-Gal in DMF and stored at -20°C.

* Overlay mixture:

0.25 M Na2HPO4 pH 7.5

0.5 % agar

0.1 % SDS

7 % DMF (SIGMA ref D-4551)

0.04% X-Gal (Euromedex ref EU0012-D)

For every plate you need 10ml overlay mixture:

5 ml 0.5M phosphate buffer pH 7.5

4.2 ml 1.2 % agar

200 µl 2 % (w/v) X-Gal in DMF

100 µl 10 % SDS

500 µl DMF

*-LWH plates

*Sterile toothpicks

Experiment

- Prepare the amount of overlay-mix you need. 500 ml maximum for one batch in order to keep the conditions similar (DMF will evaporate) and to prevent the mix settling too long in the bottle.

- Mix 250 ml 0.5 M phosphate buffer pH 7.5 + 210 ml 1.2 % agar + 5 ml 10 % SDS. Go under the hood and add 25 ml DMF and 10 ml 2 % X-Gal.

- Don’t add X-gal solution unless you will use it immediately. Temperature of the mix should be between 50°C and 45°C. Higher temperatures might affect the cells; lower temperature impairs handling.

- With pipetpump and sterile 25 ml pipet divide the overlay-mix over the plates in portions of 10 ml. Let the mix flow out gradually where there are no colonies growing. Too fast a flow or directly hitting a colony will cause smearing of the yeast cells. Too slow a flow will cause settling of the mix before it has spread equally over the plate.

- Work through the whole batch of plates laid out. Collect them when the top layer is settled (keep the plates horizontal; don’t tip them; the whole overlay layer might move, causing smearing of the colonies). Check whether all colonies are covered by the overlay. If not, apply some mix on the ‘uncovered’ spots.

- Incubate plates cells/overlay-up at 30oC. Note the time.

- Check for blue colonies after 30 min, 1 h, 2 h, 4 h and 6 h incubation time. Mark positives with a felt-tip pen (a different colour/marking-style for each check at the different incubation time).

After 30′ and 1 hour do a quick check for blue colonies. Check very superficially. Do more careful checking after 2, 4 and 6 hours.

- During checking, take the plates to the hood and check the colour against a black background. Don’t inhale the DMF, keep your head out of the hood. At this moment it is not essential whether you are able to pick all blue colonies (some will be pale).

- Number the positives (If you want you can sample them in classes, distinguishing them according to colony-size and intensity of blue-colour).

- Take some fresh -LWH plates (it is important to maintain the selection for the interaction on -LWH plates) and divide into 4 sections. Number them according to the number of positives found. Mark them with date and bait-name.

- Streak the positive colonies after 6h incubation time. Put the rest back at 30oC.

- To streak cells from a single positive colony take a sterile toothpick through the agar-overlay into the colony and collect some cells. Streak the cells into one line. Take a fresh toothpick and restreak cells from the first line into a second one. Take a third toothpick and streak cells from the second into a third line and, after turning the toothpick, into a fourth line.

- Next day, check plates for positives. Streak newly found positives. It is important not to wait too long in doing this as the components in the overlay-mix are not very good for the yeast cells (After 24h incubation time the cells might be dead).

- It takes about two days for colonies to grow.

There are five items to keep in mind:

1) What counts as a ‘real’ positive (= blue colour) can differ from one screen to another. It might be related to the nature of the bait fusion, and also to its expression level.

2) Another factor is how fast the yeast colonies are permeabilized. Therefore you will see blue colonies coming up after an hour whereas for other colonies, it takes six hours. The intensity of the blue colour can also range from deep dark to almost grey-light blue. Sometimes you see a blue halo around the colonies or the colour is only restricted to the colony or its center. Always compare the colour of putative positives with that of colonies on the same plate or on plates from the same screen.

3) Because of these kind of differences keep track of the time at which positives are found, size of colony and intensity of blue-colour.

4) In the overlay method the blue-colour develops with time. Compare the blue colour of the positive colonies at a later time (i.e. after 24h) than when you found/streaked them. Strong positives will turn bluer than weaker ones. Colonies that turned blue at a later time-point than others might have developed a more intense blue colour than the ‘earlier’ ones.

5) Recovery of small colonies can be difficult, especially when streaking after 24h incubation.

Protocol C: Filter lift X-gal Assay

Introduction

The filter lift assay is a fast and sensitive method to check whether the reisolated colonies are positive and homogeneous.

Material

* Plates with healthy, well-grown yeast colonies

* Hybond C-extra filters (Amersham, RPN 82 E)

* incubation plastic boxes

* Whatman 3MM paper

* Millipore forceps

* Saran wrap

* Liquid Nitrogen

* sterile toothpicks

* -LWH plates

* 2 % (w/v) X-gal solution in Dimethylformamide (DMF)

* 1M Na2CO3

* Z-buffer with ß-mercaptoethanol (Z-ßOH):

100 mM Na2HPO4 pH7.5

10 mM KCl

1 mM MgSO4

add 1.8 ml b-mercaptoethanol per 500 ml Z buffer (50mM) just before use (if added in advance keep solution at 4oC).

To 20 ml Z-ßOH buffer add 400 ml 2% X-gal solution.

Experiment:

- Take a sheet of Whatman paper upon which you can work.

- Cut a sheet of Whatman that fits exactly into the bottom of the incubation box.

- Prepare just enough Z buffer-ßOH/Xgal mix for wetting the paper: For one incubation box (about 300 cm2 of paper) you’ll need about 20 ml Z-ßOH/Xgal mix.

- Pour the Z-ßOH/Xgal mix over the paper. Start in the middle and tilt the box so that the paper is uniformly wetted. Get rid of any bubbles. (Try to lift the wet paper with forceps at one of the corners to let the bubbles escape; be careful the paper may tear). Pour off excess Z-ßOH/Xgal mix and keep as back-up. Place the lid on the box.

- Take a Hybond C-Extra filter (or enough to cover those plate-sectors where yeast colonies have formed), remove protecting papers. Take a ball-point and mark the filter with a letter/number and a line.

- Mark the plates with a line and numbers corresponding to the filters.

- Lift the marked filter, with the mark at the underside (i.e. towards the yeast cells), using the forceps (or by hand with gloves) and the lid of the yeast plate.

- Position the filter above the yeast plate so that the lines on the plate and the filter are aligned.

- Gently let the filter descend upon the yeast cells (i.e. not smearing the colonies). Tap the plate in order to enhance the wetting of the filter.

- Close the plate with the lid and, in the case you have to do a lot, apply a filter to the next plate.

- When the filter is uniformly soaked or when all plates are done, lift the (first) filter gently from the plate using the forceps. Directly hang the filter in liquid nitrogen without releasing your grip on the forceps. Count to five, take the filter out of the nitrogen and let thaw cells-up on your paper work-sheet. (Continue with the next filter). Repeat freezing/thawing step a second time.

- When the filters have thawed, transfer them to the incubation box (containing the Z-ßOH/Xgal pre-wetted Whatman filter). Lay filters carefully cells-up and without trapping air bubbles. Close the box and seal the lid with parafilm. Incubate at 30oC.

- Prepare a stop-paper and a wash-paper:

Stop-paper: Take a slightly smaller piece of Whatman paper and place inside an incubation box. Wet the paper with 1M Na2CO3.

Wash-paper: idem, but wet the paper with distilled water.

- After 3 h incubation time, transfer the filters from the incubation box on the stop-paper and let them sit for 1 min. Transfer them to the wash-paper for another min., then dry them on a fresh piece of Whatman paper.

- Store the filters by sticking them to a new sheet of Whatman paper using doublesided tape. Keep the filters that belong to one screen together. Wrap the papers with saran-wrap and seal the edges. Store together the filters belonging to the same screen.

Restreak cells from a positive colony after checking by the filter lift assay (for each clone). Take some fresh -LWH plates and divide into 8 sections. Number them according to the number of positives found. Mark them with date and bait name. Restreak cells from a single positive colony with 3 sterile sticks as described above to get isolated colonies.

Protocol D: PCR ON YEAST COLONIES

Introduction

PCR amplification of fragment of plasmid DNA directly on yeast colonies is an efficient procedure to identify sequences cloned into this plasmid. It is directly derived from a published protocol (Wang.H et al, Analytical Biochemestry, 237, 145-146, 1996). However, it is not a standardized protocol : in our hands it varies from strain to strain, it is dependent on experimental conditions (number of cells, Taq polymerase source, etc.). This protocol should be optimized to specific local conditions.

Materials

* 10x PCR buffer:

500 mM KCl

100 mM Tris HCl pH8.3

15 mM MgCl2

0.1% Gelatine

* 10 mM dNTP (mixture of the four dNTP at 10mM each)

* Taq polymerase (5U/µl) from Pharmacia (ref. 27.0799-02)

* 1 N NaOH

* oligonucleotide upstream 100ng/µl (oligo forword for the W strand of the pACTIIst prey plasmid):

* oligonucleotide downstream 100ng/µl (oligo reverse for the C strand of the pACTIIst prey plasmid):

(see Annex D for the sequence of pACTIIst plasmid).

Experiment

- Take one colony with a toothpick. Resuspend the cells at room temperature in 10 µl 0.02N NaOH in an Eppendorf tube by turning the toothpick for several seconds in the NaOH solution. Prepare 0.02N NaOH just before use (5 µl 1N NaOH + 245 µl water).

- Incubate 5 min. at 100°C. For a large series, vortex the tubes before the incubation at 100°C to counteract the sedimentation of the cells.

- Put on ice immediatly. Centrifuge the tube briefly to spin down the drops of condensation on the cap.

- Vortex to resuspend the pellet and transfer 2 µl of the cell extract in a 0.5 ml PCR tube preincubated in an ice/water bath.

- Prepare the PCR mix for X reactions (at least one more -or 10 % more for large series-than the actual number of reactions).

for one reaction:

24.6 µl H2O (qsp 30 µl)

3.2 µl 10x PCR buffer

0.7 µl 10 µl dXTP

0.5 µl oligo for

0.5 µl oligo rev

(0.5 µl Taq polymerase)

- Mix all the components except the Taq enzyme.

- Go to the thermal cycler. The block of the thermal cycler must be at 94°C when you put your samples inside.

- Add Taq enzyme to the PCR mix just before you distribute 30 µl of PCR mix per reaction.

PCR program: PTC-200 (MJ Research) or UnoII (Biometra)

- step 1 3 min 94°C

- step 2 94°C 30 sec.

- step 3 50°C 45 sec.

- step 4 72°C 3 min.

(31 cycles step 2 to 4)

- step 5 72°C 5 min.

- step 6 15°C for ever.

- Check the quality, the quantity and the length of the PCR fragment on 1% agarose gel. Take 3 µl from the PCR reaction and mix with 10 µl loading buffer (1.3 µl 10x blue loading buffer + 8.7 µl H2O; 10x blue loading buffer is: 10 mM Tris pH8, 1 mM EDTA, 30% glycerol, 0.2% bromophenol blue, 0.2% Xylene cyanol blue).

- The length of the cloned fragment is the estimated length of the PCR fragment minus 300 bases that correspond to the amplified flanking plasmid sequences.

Protocol E: IDENTIFICATION AND CLASSIFICATION OF THE CANDIDATES

1) SEARCH IN DATABASES

Introduction

This protocol varies from one lab to another depending on the local conditions for databases searches.

Materials

* DNA Strider software (Marck. C; 16, 1829-1836, 1988)

* SGD blast program (http://genome-www.stanford.edu ; SGDB, Stanford, California).

* YPD (http://www.proteome.com ; YPD, Proteome Inc., Beverley, Massachussetts).

* NCBI blast program (http://www2.ncbi.nlm.nih.gov/BLAST).

Experiment

An example of a table of results is given in Annex F

Sequence N-terminal fusion of the preys.

- Each clone is sequenced. Create one DNA Strider file per clone.

- Read and enter the sequence from GGG ATC C. It is the BamHI site used to clone the yeast FRYL library. Starting with the 3Gs gives the coding frame of the Gal4 domain.

The 20 bases after the BamHI site are from the linker added to the genomic DNA fragments for the making of the FRYL library.

It is important to check if the linker sequence contains a mutation (especially frameshifts).

- Read at least 50 bases of the insert.

- Search chromosomal coordinates (chromosome number, strand w or c, position) with the SGD blast program.

- Enter the data into in a column of an Excel Microsoft® table.

- Search the ORF corresponding to chromosomal coordinates (or the names of the two flanking ORFs). Extract and copy the sequence of the ORF into a DNA Strider file (+ 1000nt flanking regions).

- Biological information on the ORF can be extracted from the YPD database.

- Compare the insert sequence with the ORF sequence to get the exact location of the beginning of the insert related to the initiation codon.

- Enter the data in a column of the table.

- Compare the translated sequence of the clones with the protein sequence to check the coding frame.

- Insert in a column of the table the length of the cloned fragment as estimated by agarose gel electrophoresis of the PCR fragments.

2) CLASSIFICATION OF THE CANDIDATES

Considering the size of the library - a fusion point once every four bases - one can expect to find a given ORF several times as independent clones in a complete screen of such a library. However, the probability of the selection of a given fusion depends on the length of the interacting domain and on the position of the interacting domain along the coding sequence.

These parameters predict that all candidates fall into one of the following categories.

The categories A1, A2, A3 and A4 correspond to potentially encoded yeast ORFs. Their inserts start either inside the ORF coding sequence or upstream the initiation codon *.

The B category corresponds to fused polypeptides located in an intergenic region, in the reverse orientation of an ORF, in a non-polypeptide encoding region (rDNA, telomeric DNA, mitochondrial DNA) or in a Ty retrotransposon element.

The A1 category consists of candidates found several times as distinct independent clones.

The three other A categories correspond to candidates found only as a single fusion, even if the same clone is found several times:

The A2 category consists of fusions starting close to an initiation codon of a yeast ORF and at a distance smaller than 150 bases from the in-frame stop codon located upstream of this ORF. These candidates correspond to amino-terminal interacting domains. For such interacting domains, fewer candidates are expected since in-frame nonsense codons upstream of the yeast ORF interrupt its translation.

The A3 category candidates contain large coding inserts (over 1000 bases). This category may correspond to preys with a large interacting domain. Since the average size of inserts is 700 nt, candidates with large interacting domains are underrepresented.

The A4 category contains the other candidates. We cannot predict why these clones are found only once, although several hypotheses can be proposed, such as incorrect folding or toxicity of fusion proteins.

*Out of frame fusions are found. We do not discard these candidates but we label them with a asterisk to keep in mind that they may encode bonafide yeast polypeptides through a frameshifting event (see Annex G).

Protocol F: Plasmid rescue from yeast by electroporation

introduction

This experiment allows the recovery of a plasmid from yeast cells by transformation of E. coli with a yeast cellular extract. In the two-hybrid screening experiment, the diploid cells contain two different plasmids carrying the TRP1 (bait) and the LEU2 (prey) markers, respectively. We use a bacterial strain (MC1066) that carries the trp and leu auxotrophies that can be complemented by TRP1 and LEU2 yeast genes. Usually we select for the prey plasmid carrying LEU2 gene.

materials

Plasmid rescue

* glass beads 425-600 µm (Sigma ref:G-8772)

* Phenol/chloroform (1/1) premixed with isoamyl alcohol (Amresco ref: 0883)

* Extraction buffer:

2% Triton X100

1% SDS

100 mM NaCl

10 mM TrisHCl pH8.0

1 mM EDTA pH8.0

* Mix Ethanol/NH4Ac: 6 volumes Ethanol with 1 volume 7.5 M NH4 Acetate

* 70% Ethanol

* yeast cells in patches on plates.

note: This protocol can be performed with frozen cells prepared from colonies or patches on plate, mixed in water and frozen directly at -20°C. ( in 50 µl H2O).

electroporation

* SOC medium

* M9 medium

* Selective plates: M9-Leu + Ampicillin

* 2mm electroporation cuvettes (Eurogentec ref: CE0002-25).

Experiment

Plasmid rescue

- First, add 400 µl of glass beads in each 1.5 ml eppendorf tube; second, add 200 µl extraction buffer (use multipipettor in case you do a large number of extracts); third, put the cells of each patch in the tube using blue tips (the cells must coat 2 mm of the tip).

- resuspend the cells in extraction buffer with the blue tip. Continue under the hood.

- Add 200 µl Phenol/chloroform and vortex cells vigorously for 7 min.

- Spin tubes 10 min, 15000 rpm.

- Transfer 140 µl supernatant to a sterile eppendorf tube and add to each 500µl Ethanol/NH4Ac. Vortex.

- Spin tubes 15 min 15000 rpm at 4oC.

- Wash pellets with 250 µl 70% Ethanol. Dry pellets (5′ in Speed Vac).

- Resuspend pellets in 10 µl water. Store extracts at -20oC.

electroporation

For a large number of electroporations:

- Take a large ice-bucket and line up the electrocuvettes according to their number.

- Alongside the cuvettes line up sterile eppendorf tubes numbered as the cuvettes but also carrying the clone-name. To every tube add 1 µl of yeast plasmid DNA-extract.

- Mark the selective plates M9-Leu with the date, the name of the clone as well as the number of the cuvette.

- Fill sterile eppendorf tubes marked with cuvette number and clone name with 1 ml of SOC medium.

- Thaw a vial with electrocompetent E.coli (Strain MC1066 for selection on Leu and Trp). Keep vial on ice.

- To every 1 µl yeast DNA sample add 20 µl electrocompetent cells; mix and transfer the mix to the cold electroporation cuvette.

- Do the electroporation directly. Set the Biorad electroporator on 200 ohms resistance; 25 µF capacity; 2.5 Kvolts. Wipe cuvettes dry with a tissue paper and tap them to remove any trapped air-bubbles. Check if suspension makes contact with both electrodes. Place cuvette in the cuvette holder. Do the electroporation; time constant should be similar for every electroporation (around 4.7).

- Directly add 1 ml SOC into the cuvette and transfer the cell-mix into sterile Eppendorf tube.

- Let cells recover for 30 min at 37oC, spin the cells down 1 min, 4000 g and pour off supernatant. Keep about 100 µl medium and use it to resuspend the cells and spread them on selective plates (e. g. M9-Leu plates). Pipet the suspension over the glass beads and directly shake the plate to prevent a clustering of cells on the place where they have been pipetted (especially when plates are a bit dry).

- Incubate plates for 36 h at 37oC

Note: Wash the electrocuvettes and the caps with tap-water (apply high pressure to force out any remaining cells); rinse them with double distilled water and then with ethanol and let them dry. Close the cuvettes with the caps and store them in the supplier case according to their number.

Preparation of electrocompetent cells

- Inoculate 1 liter LB with a 10 ml o.n. pre-culture of the E.coli strain of interest (MC1066 for Two Hybrid-plasmid rescue)

- Grow to an OD600 of around 0.6. (It should not be higher than 0.8).

- Chill culture on ice and in cold-room for 15 min. From now on keep cells cool and handle with care.

- Pellet cells 15 min 3000 g at 4oC

- Resuspend cells in 1 liter cold sterile water

- Pellet cells 15 min 3000 g at 4oC

- Resuspend cells in 500 ml cold sterile water

- Pellet cells 15 min 3000 g at 4oC

- Resuspend cells in 20 ml sterile 10% glycerol

- Pellet cells 15 min 3000 g at 4oC

- Resuspend cells in 2 ml sterile 10% glycerol and complete up to a final volume of 3 ml (about 3.1010 cells /ml).

- Distribute in aliquots of 50 µl or multiples of that. (You’ll take 50 µl per normal electroporation and 20 µl when you have to rescue a large number of plasmids)

- Freeze the aliquots in dry Ice/EtOH but take care to prevent traces of EtOH entering into the tubes) and store at -80oC.

Protocol G: PLASMID MINIPREPS FROM E.COLI

Materials

* STET buffer :

8% Sucrose

50 mM EDTA pH8

10 mM TrisHCl pH8

0.5% Triton X100

* 20 mg/ml freshly made Lysozyme (dissolved in water)

* EtOH/NH4Ac: mix 6 volumes of ethanol with 1 volume of 7.5 M NH4 Acetate.

* 70% Ethanol

* TE pH8

STET/Lysozyme: for 10 minipreps: 3 ml STET + 250 µl 20 mg/ml Lysozyme

Experiment

- Start with 2ml bacterial cultures grown overnight.

- Transfer 1.5 ml to non-sterile Eppendorf tubes, pellet cells by centrifugation 20 sec, discard the supernatant by overturning the tubes. Keep about 100 µl medium and use it to resuspend the cells with vortex.Be careful! all the cells must be well resuspended.

- Add 300 µl STET/Lysozyme buffer

- Keep on ice for 5 min.

- Incubate 2 min at 100oC (use hot-block rack for transfer).

- Put on ice.

- Spin 15 min at 4oC

- Take out the pellet with a toothpick

- Add 750 µl EtOH/NH4Ac to supernatant, vortex

- Spin 10 min, wash with 500 µl 70% EtOH, dry pellet (5 min in speed-vac).

- Dissolve pellet in 50 µl TE by incubation at 65°C for 5 min and vortex.

Protocol H: YEAST TRANSFORMATION

Adapted from: Gietz et al, Yeast 11, 355-360, 1995

Material

* 2 ml eppendorf tubes

* 50 ml tubes

Media

* liquid medium for preculture: YPglu or selective medium when the yeast strain harbors a plasmid.

* plates for plasmid selection.

Solutions

* Yeast carrier DNA (Clontech ref: K1606-A)

* 1M Li Acetate (filter sterilized) (Fluka ref: 62395)

* 10x TE (100 mM Tris pH 7.5 ; 10 mM EDTA) autoclaved

* Sterile distilled water

* Prepare 0.1 M LiAc/TE solution by mixing:

100 ml 1M LiAc

100 ml 10x TE

800 ml sterile distillated water

* 40% PEG 4000 (Merck ref:807 490 ) filtrated on Nalgene (0.2µ).

40 g of PEG 4000

10 ml 1M LiAc

10 ml 10x TE

add H20 to a final volume of 100 ml.

Experiment

* For one transformation you need 10 ml of culture (OD = 0.6; with our spectrophotometer, 1 OD corresponds to 107 cells per ml) which will be finally resuspended in 50 µl 0.1M LiAc/TE.

* all steps are performed at room temperature

- Set up a preculture of the strain in 20 ml of appropriate medium the day before

- Grow o.n at 30°C until saturation

- Inoculate 50 ml of medium (for 5 transformations) at 0.15 OD with the o.n preculture

- Grow until OD= O.6

- Centrifuge the cells in 50ml sterile tubes at 5000 g (4000 rpm)for 3 min

- Discard the supernatant by overturning the tube

- Resuspend the pellet with 2 ml of sterile water

- Transfer into an 2 ml eppendorf tube.

- Centrifuge the cells at 4000g (6500 rpm) for 1 min.

- Repeat the washing twice with 2 ml of water and twice with 2 ml of LiAc/TE solution.

- Centrifuge at 4000g 1 min. after each washing and resuspend each time the cells by vortexing.

- Resuspend finally the pellet with 200 µl of LiAc/TE solution. Adjust to 250 µl.

- In the meantime, mark your tubes. Do not forget one tube without plasmid DNA.

- Distribute in sterile 1.5ml Eppendorf tube:

- 1µl plasmid DNA (from a mini-prep or at 0.1 mg/ml)

- 5 µl Yeast carrier DNA

- 50 µl of cells

- 350 µl of 40% PEG

- Mix by inversion

- Incubate 30 min at 30°C

- Heat shock cells at 42°C for 20min.

- Add 700 µl of water. Mix by inversion.

- Spin cells for 1 min in a microfuge at 4000 g. Discard supernatant.

- Resuspend the cells with 100 µl of water

- Spread the cells on appropriate medium plates and incubate at 30°C for two-three days.

Annex: MEDIA

YEAST MEDIA

YPGLU

1% yeast extract

2% bactopeptone

2% glucose

10 g Bactopeptone (Difco ref: 0118-17-0)

10 g Yeast Extract (Difco ref: 0127-17-9)

20 g Glucose (Merck ref:1.08342)

Fill to 1 liter with distillated water

Shake until all ingredients are dissolved

For plates add 20 g Bacto-agar (Difco ref:0140-01)

Autoclave 20 min, 110 oC (at higher temperatures sugars and amino acids might be degraded), let cool to 60oC, (when required add antibiotics) mix gracefully so that no bubbles are formed

Pour sterile plates: 25 ml per plate

Drop-Out yeast medium

6.7 g Yeast Nitrogene Base w/o amino acids (Difco ref:0919-15-3)

20 g D-glucose (Merck ref:1.08342)

2 g drop-out powder mix (see below)

Fill to 1 liter with distilled water

Shake/stir until all ingredients are dissolved

For plates: add 20 g bacto-agar (Difco ref:0140-01), mix

Autoclave 20 min, 110 oC (at higher temperatures sugars and amino acids might be degraded), let cool to 60oC, (when required add antibiotics) mix gracefully so that no bubbles are formed

Pour sterile plates: 25 ml per plate

The four Two-hybrid drop-out mixes are:

-W (selection on bait plasmid);

-L (selection on library plasmid);

-LW (selection on both plasmids);

-LWH (selection for both plasmids and two-hybrid interaction inducing the HIS3 reporter).

-LWH + AT (AT may be added at a concentration between 1 mM to 50 mM depending on the bait)

* Tetracycline (12 mg/ml stock solution) is added after cooling the agar to 60°C

* 3-Amino 1,2,4 Triazole (3-AT Sigma Ref: A-8056) is added after cooling the agar to 60°C using a 1M stock solution.

Drop-out powder mix:

Mix 2g of each component:

Adenine; Alanine; Arginine; Asparagine; Aspartic acid; Cystein; Glutamic acid; Glutamine Glycine; Histidine; Isoleucine; Lysine; Methionine; Phenylalanine; Proline; Serine; Threonine; Tyrosine; Tryptophan; Uracil; Valine

plus 4 g of Leucine.

In total 22 essential components from which the underlined ones have to be omitted in the case of the two-hybrid drop-out mixes.

BACTERIAL MEDIA

SOC buffer

2% Tryptone (20 g/l)

0.5% Yeast Extract (5 g/l)

10 mM NaCl (0.58 g/l)

2.5 mM KCl (0.19 g/l)

10 mM MgCl2 (2.03 g/l)

10 mM MgSO4 (2.46 g/l for MgSO4.7H2O)

20 mM glucose (3.6 g/l)

M9 drop-out plates

1g drop-out mix (the same as for yeast medium) (for library plasmids M9-leu: -L; for bait plasmids M9-Trp: -W)

20 g bacto agar in 878 ml double distilled water,

Autoclave for 20 min at 110 oC

Let cool to 60 oC;

Add asceptically:

100 ml 10 x M9 (autoclaved)

10 ml 20% glucose (autoclaved)

2 ml 1M MgSO4 (autoclaved)

10 ml 20 mM CaCl2 (autoclaved)

1 ml 1000 x Ampicilline (100 mg/ml stock)

Pour plates (20 ml/plate)

for 1 liter of 10X M9:

60 g Na2HPO4;

30 g KH2PO4;

5 g NaCl;

10 g NH4Cl

Adjust to pH 7.4

* You’ll need one plate for every rescue