DNA Extraction

Epicentre Kit

Cat# MPY80200, 200 preps (here using half the volume, so yield is 400 preps)

If starting from liquid culture, spin down ~1.5mL and remove supernatant.

1. In an Eppendorf tube, add 150µL Yeast Cell Lysis Solution.

2. Using a sterile toothpick, take a small scraping of yeast and mix with the liquid (skip this step if starting with a pellet from liquid culture). Suspend the cells by vortexing.

3. Incubate for 15 min @ 65ºC

4. Incubate for 5 min on ice.

5. Add 75µL MPC Protein Precipitation Reagent and vortex for 10 sec.

6. Spin down 10 min @ max speed. 

7. Transfer supernatant to a clean tube.

8. Add 500µL isopropanol and mix by inversion.

9. Spin down again.

10. Remove supernatant with pipette or aspiration and discard. Wash the pellet gently with 500µL 70% ethanol, let pellet fall at the bottom and remove ethanol carefully.

11. Spin down briefly and remove any remaining ethanol. Let dry tube open.

12. Resuspend in 25-50µL H2O and store at -20ºC for downstream use.

Smash & Grab

1. In an eppendorf tube, add 30-50µL beads (BioSpec 0.5mm dia Zirconia/silica Cat# 11079105z or Sigma acid washed glass beads PN G8772) and 200µL DNA Breakage Buffer (see Solutions Recipes).ƒ

2. Using a sterile toothpick, take a small scraping of yeast and mix with the liquid. (If starting from liquid culture, spin down and remove supernatant from ~1.5mL culture and start in this tube instead.)

3. In the fume hood, add 200µL phenol:chloroform:PCI (25:24:1)

4. Vortex (or FastPrep or Thermomixer, no heat) for 3 min @ maximum speed

5. Spin down 5min @ maximum rpm

6. In a new tube, add 1mL 100% ethanol. From the original tube, remove 125µL of the top layer of liquid, taking care not to disturb the layers. Add to the ethanol tube and mix gently.

7. Spin down 5 min @ maximum rpm

8. Using a vacuum flask with tubing apparatus and new pipette tip on the end, aspirate ethanol, leaving the pellet behind. Leave the remaining pellet to dry completely (gDNA is relatively stable and can be left dry at ambient temperatures for several days).

9. Resuspend pellet in 50µL water and continue with downstream applications.

 PCR

NOTE: handle enzyme carefully-- keep on ice at all time, and avoid vortexing/shaking/dropping the tube. Primers are typically hydrated to a stock concentration of 20µM in the tube. When running multiple reactions at once, create a master mix (vol per rxn x #rxns x 1.1) and add DNA separately to each tube.

1. Standard PCR with rTaq polymerase:

Component Volume per 25µL rxn

10x Buffer 2.5µL

2.5mM dNTPs * 2µL

rTaq* 0.125µL

Primer 1 (20µM) 1µL

Primer 2 (20µM) 1µL

H2O 17.375µL

DNA template 1µL (If using a plasmid as DNA template: use the 1:25 stock dilution.)

* Notes: Here recipe is for TaKaRa kit PN RR001A. These volumes are correct for 2.5mM dNTPs and separate rTaq, but check concentrations and what is available. GenScript reagents are 10mM dNTPs and 2x MasterMix which contains buffer, dNTPs, and rTaq.

2. High fidelity PCR with Platinum SuperFi polymerase:

Component Volume per 100µL rxn

5x Buffer 20µL

10mM dNTPs 2µL

DNA Pol 1µL

Primer 1 (20µM) 3µL

Primer 2 (20µM) 3µL

H2O 67µL

DNA/Plasmid 4µL (If using a plasmid as DNA template: use the 1:25 stock dilution.)

GC Enhancer (opt) 20µL (if using, reduce H2O to 47µL)

Thermal Cycle: Standard Program for both enzymes

(called WE DOIN SCIENCE) - volume 25uL (to adjust if doing 100uL)

Initial denaturation 95°C - 5min

denaturation 95°C - 30s

Annealing 52°C - 30s 30x cycles

Elongation 72°C - 3min (can be adjusted: 90 s/kb for rTaq, 30 s/kb for SuperFi)

Final elongation 72°C - 9min (=3x elongation time)

10°C - ∞

3. High fidelity PCR with Phusion polymerase:

Component 20 µl rxn 50 µl rxn Final Concentration

5X Phusion HF or GC Buffer* 4 µl 10 µl 1X

10 mM dNTPs 0.4 µl 1 µl 200 µM

20 µM Forward Primer 0.5 µl 1.25 µl 0.5 µM

10 µM Reverse Primer 0.5 µl 1.25 µl 0.5 µM

DMSO (optional) (0.6 µl) (1.5 µl) 3%

Phusion DNA Polymerase 0.2 µl 0.5 µl 1.0 units/50 µl PCR

Nuclease-free water to 20 µl to 50 µl  

Template DNA (50-250ng gDNA) < 250 ng

 (1pg-10ng plasmid)

Buffer: use HF by default, use GC for difficult templates (high GC, or secondary structures).

Thermal cycle :

Temperature (°C) time (min:sec)

Denaturation 98°C 0:30

Denaturation 98°C 0:10

Primer annealing* 55°C (45-72°C) 0:10-30 25/30 times

Elongation 72°C 0:15/kb

Elongation 72°C 5:00-10:00

* annealing temperatures for Phusion tend to be higher than with most other PCR polymerases: use the NEB Tm calculator https://tmcalculator.neb.com/#!/main.

4. Yeast colony PCR with rTaq polymerase:

Use this protocol to genotype yeast colonies without extracting DNA.

In this protocol, DNA template are yeast cells resuspended in 3 µL of NaOH 20 mM. The suspension should look cloudy. Boil the suspension for 10 minutes (use “BOIL” protocol in the PCR machine). 

 Component Stock concentration Volume per 25 µL reaction

Betaine monohydrate 5M 5 µL

TaKaRa buffer 10X 2.5 µL

dNTPs 2.5 mM 2 µL

Primer 1 20 µM 2.5 µL

Primer 2 20 µM 2.5 µL

TaKaRa Taq Pol 5 U/µL 0.25 µL

H2O 7.25 µL

DNA template 3 µL

TIPS: Use controls. Negative controls: a) “empty” well (no yeast sample) to check the behavior of the primers in the PCR conditions; b) “negative” yeast (or gDNA) to check if you get unspecific bands in strains that don’t have the DNA segment of interest. Positive control: gDNA, plasmid, strains.

When using a DNA preparation as control, just add it to the final mix, which is 3µL of NaOH 20mM + PCR mix.

Length of amplicon: this protocol is best for short amplicons, i.e., < 1000 bp.

Number of cycles: to resolve multiple bands in a mulptiplex PCR, reduce the number of cycles to 23 or 25.

Thermal cycle program is called “YEAST COL PCR”:

Temperature (°C) time (minutes)

Denaturation 98 0:10

Denaturation 98 0:10

Primer annealing 55 0:30   30 times

Elongation 72 1:30/kb

Elongation 72 5:00

Running a DNA Gel

Updated 2019/01/15

Standard 1% agarose in 1X TAE, 0.01% EtBr:

1. Determine gel size needed based on number of samples and available combs. Account for at least one ladder per row.

Gel box Comb sizes (x2) Gel volume Buffer volume

B1A 6, 10, 12 wells 50mL 250mL

B1 10, 14 wells 80mL 500mL

B2 12, 16, 20, 24 wells 120mL 700mL

D2 30, 40 wells 120mL 700mL

1. If not available, prepare 1L of 1X TAE (100mL 10X TAE + 900mL DI water).

2. Prepare 1% agarose in 1X TAE in a flask, melt in the microwave in 30s increments (after boiling, hold up to the window to be sure it is completely dissolved), let it cool 5-10min. Under the hood, add EtBr in the flask (1µL per 10mL), gently swirl to prevent air bubbles, and pour in the gel mold, add comb(s) (! too hot agarose can crack the gel box). 

Alternatively: There is a stock bottle of 1% agarose w/ EtBr. Heat in 30s increments in the microwave until completely melted (leave lid slightly unscrewed to prevent pressure buildup). To refill: 400mL 1x TAE + 4.0g agarose, microwave to melt completely, then add 40µL EtBr to the bottle. 

NB: vapors produced by adding EtBr in hot agarose, or by boiling agarose solution containing EtBr, are hazardous.

1. Allow gel to solidify completely, ~30min depending on size/thickness (turns opaque).

2. Place the gel box with the electric leads on the right hand side. Remove the gel tray from the box and rotate so that combs are at the top. Fill the gel box with 1x TAE until gel is covered. Remove combs and rinse.

3. Mix PCR product with 6x loading dye using one of the following methods:

1. On a small piece of parafilm, pipette dot ~1µL of loading dye for each sample. Draw up 5µL sample, adjust pipette volume to 6µL, and dispense the sample into a dot of dye. Pipette mix, draw up into pipette, and load.

2. Add 2µL dye per sample directly in PCR tube. Draw up 6µL of PCR product + dye, and load. NOTE: Do not use this method if downstream processing such as sequencing is required, as the dye will interfere, or you’ll need to purify the sample.

1. Load samples into the wells. Add 5µL ladder, at least one per row.

NOTE: ladder prep: To 10x Thermo 1kb DNA Ladder stock tube (-20ºC), add 167µL 6x loading dye and 733µL H2O. Pipette mix. 

1. Put the cover onto the gel box. Make sure the black lead is at the top and the red lead at the bottom (red is positive, and slightly-negatively-charged DNA will move towards this end). Turn on the power supply and adjust if necessary to 150V and press run.

2. Allow the gel to run for 15-45 min depending on fragment size and wanted resolution. (best if upper and lower dye bands are about 2 cm apart).

3. Image with UV on the imager (1s exposure is plenty enough) to see DNA bands. 

4. Discard gel in hazardous waste (EtBr is toxic/carcinogenic) dedicated to gel. If you soaked your gel in 1X TAE 0.01% EtBr, discard solution in dedicated liquid hazardous waste bottle dedicated for EtBr. Rinse out gel tray, combs, and box with DI water.

Troubleshooting: 

• my gel is over run, the lower bands became invisible as EtBr migrate upward… 

• Or: I added neither sybr nor EtBr

→ In these cases, use a disposable tray, fill with 1X TAE 0.01% EtBr and let sit for ~1hr, and re-image.

Alternatives to standard 1% agarose in 1X TAE to optimize resolution: 

• To separate very long fragment, agarose concentration can be decreased to 0.7-0.8%

• To separate very small fragments, agarose concentration can be increased to 1.2% or more.

• To separate even better fragments up to 1.5kb that are very close in size, use 1X Sodium borate buffer instead of 1X TAE buffer. Recipe for 10X Sodium borate is 500mL boiling water + 9.5g Sodium TetraBorate DecaHydrate. Dilute to 1X when using. This buffer allow to run very fast, at 300V, without heating up too much, thus gives very sharp bands. Usually run for 5-15mn.

Agarose DNA gel staining with Sybr Gold: 

We use Sybr Gold to visualize bands with blue light that does not damage nucleic acids. This is important when cloning.

• A stock of 6X Loading Dye containing 1:1000 Sybr Gold can be stored at room temperature. 

• For ladder+Sybr: To 10x Thermo 1kb DNA Ladder stock tube (-20ºC), add 167µL {6x loading dye + Sybr} and 733µL H2O. Pipette mix. NOTE: if using 1x ladder, add 0.17ul Syb per mL ladder.

• Use this dye+Sybr to make ladder so that the concentration of sybr is the same in ladder and sample (or add 0.17uL per 1mL ladder). Concentration of Sybr in ladder and sample has to be the same otherwise migration will be different.

• Pour agarose gel without adding any nucleic acid dye to it.

• Add Loading Dye containing Sybr Gold to a final 1X concentration to the samples before loading the gel. 

• Visualization: Sybr Gold can be visualized upon excitation with UV light. If you need to cut out a DNA band from the agarose gel, you can use the blue screen in the Symington lab (12th floor).

• Discard Sybr Gold gels in the EtBr waste.

• Running buffer can be reused.

 Sequencing with GeneWiz

→ Sequencing unpurified PCR products:

1. Prepare PCR product for sequencing. For >48 samples a plate can be used, but for less simply use 8-strip PCR tubes. Add 10µL un-purified PCR product per sample per tube. Label tubes from left to right as “LB1”, “LB2”, etc as asked. Prepare primer(s) in microtube(s): 5µL per reaction to 5µM concentration. Label the microtube(s)

2. Create the order on GeneWiz’s website. 

1. Login using leb2210@cumc.columbia.edu and PW “Rim4disorder”.

2. Click “Create Sequencing Order” and select “Standard”, “Create by online form”, and sample type: “Custom” (for PCR purification). Put in the number of samples to submit to create the form.

3. For “DNA Name”, enter something descriptive. Click “Fill DNA name sequentially” to add sequential numbers, if desired. For “DNA Type”, select “Unpurified PCR”. For “DNA Length” enter as appropriate; for “My Primer Name”, enter the primer name such as “LB_31”, for “My Primer Concentration”, enter “5”; for “DNA Conc”, “Genewiz Primer”, “Notes”, leave blank. Below the form, “Description” is optional.

4. Click “Save & Next”, review all entries, and then click “Next”.

5. For payment, select “Credit Card”-- it should be saved into the Genewiz account. Put in quote# DPP1702151 to assure our pricing. Click “Next” and then submit.

6. Print a copy of the order form to send in with the samples.

3. In a ziploc bag, place samples, primer tube(s), and printed order form.

4. Leave bag in drop box: Floor 7, white box labeled “Genewiz sample pickup” by the elevators. Pickup time is usually 7pm. Results are emailed within 1-2 days.

Preparation of Bacterial Competent Cells (Inoue et al., Gene 1990)

Media and Mixes

SOB (from Hanahan, JMolBiol 1983): 2% Bacto Tryptone

0.5% Yeast Extract

10 mM NaCl

2.5 mM KCl

Autoclave

Add 10 mM MgCl2

10 mM MgSO4

filtered, add after autoclaving

pH should be 6.8 to 7.0

TB (store at 4˚C): 10 mM PIPES (store at 4˚C)

15 mM CaCL2

250 mM KCl

adjust pH to 6.7 with KOH

Add 55 mM MnCl2 (store at 4˚C)

filter

Culture cells

1. Streak bacterial strain on LB agar plate (or where appropriate), incubate at 37ºC overnight.

2. Inoculate 10-12 large colonies in 30 ml SOB and grow them overnight at 18ºC with vigorous shaking.

3. Check regularly the culture density to have a rough estimate of doubling time (it should be ~4 hours).

4. Dilute culture in 250 ml SOB and grow till OD=0.6 at 18ºC with vigorous shaking.

Competent cells preparation

Before you start:

1. Cool down centrifuge to 4ºC.

2. Prepare 1.5 ml tubes on racks and cool them down in the -80ºC freezer.

Procedure (all these steps are performed in the cold room)

1. Place flask on ice bath for 10 min to cool down. To ensure homogeneous temperature, rotate the flask from time to time.

2. Aliquot cell culture in 50 ml tubes (250/50=5 tubes) and harvest cells for 10 min at 4ºC at 2500 g.

3. Resuspend pellet in 80 ml of ice cold TB with vortex (that is, in each tube, add 16 ml in 5x50 ml tubes). Vortex at max speed for 1 sec, and put on ice. Repeat until pellets are completely resuspended.

4. Incubate in ice bath for 10 min.

5. Harvest cells for 10 min at 4ºC at 2500 g.

6. Gently resuspend pellet in 20 ml of TB with vortex (that is, in each tube, add 4 ml in 5x50 ml tubes). Vortex with mid-speed for 1 sec, and put on ice. Repeat until pellets are completely resuspended.

7. Combine all aliquots.

8. Add DMSO to final concentration of 7%.

9. Incubate in ice bath for 10 min.

10. Dispense 100 ul cells in 1.5 ml tubes.

11. Immediately chill aliquots in liquid nitrogen.

12. Store at -80ºC.

Test competent cells

1. Transform the competent cells with a concentration series of pUC19 (a commercially available plasmid used for these routine tests; it is in the -80ºC): 100 pg, 10 pg, 1 pg, and 0.1 pg. Use standard protocol for transformation, and plate on LB + Amp.

2. Determine competence: cfu/ug of DNA. A good range of competence is 10^8-10^9 cfu/ug.

Yeast Transformation

Last update: 2019-01-25

Anytime before Day 3: Prepare plasmid or PCR, as needed, using Platinum SuperFi reagents, and check for appropriate bands via gel.

Day 1: Pull out onto YPG yeast strain(s) to transform, fresh from -80°C.

Day 2: Inoculate 25mL YPD and grow overnight @ 30°C. 

Day 3:

1. Dilute culture to OD600 ~0.2 in 50mL fresh YPD (usually 1mL into 49mL YPD).

2. Let grow ~3 hrs @ 30°C on shaker. Harvest cells at ~OD600 0.4-0.7.

3. Transfer culture into 50mL falcon and spin down 1 min @ 3,000g.

4. Wash: Resuspend with ~25mL sterile water and spin down again.

5. Resuspend in 1mL 0.1M lithium acetate and transfer to an eppendorf tube.

6. Spin down 1 min @ 3,000g and discard supernatant. Resuspend in 400µL 0.1M LiAc.

7. Aliquot 50µL cells per transformation reaction. Spin down and remove supernatant.

8. Per reaction, add the following, in this order:

• 240µL 50% PEG-4000

• 36µL 1M LiAc

• 10µL boiled ssDNA (thaw on ice) (tube should be re-boiled every 3-4 use)

• 74µL prepared plasmid/PCR product (if volume is <74µL, add water to 74)

1. Vortex for 1 min.

2. Incubate @ 30°C for 30 min (in warm room).

3. Heat shock at 42°C for 15min (in water bath, better than in thermomixer).

4. Spin down, remove supernatant, and resuspend in 100µL H2O.

5. → If selecting on prototrophy, plate on selective drop-out medium. 

→ If selecting on drug resistance, grow 3h in liquid YPD, then plate on selective medium, or directly plate on YPD, grow overnight and replicate-plate to selective medium containing antibiotic/drug the morning after (do not let the lawn overgrow on YPD).

After 2-3 days onto selective media:

1. Streak out colonies if any to a fresh selective plate to eliminate false positives.

2. Patch individual colonies to YPD to grow up O/N.

1. Collect a scrap of each patch to extract DNA and check that the gene of interest was correctly inserted with PCR, using check primers for the gene of interest (see database).

2. Replica plate to YPD and YPG to check mitochondrial functionality.

Yeast Dissection

→ Usually start with either a diploid strain or two haploid strains mixed on YPD 4% at least ON uiat 30°C. Transfer cells on SPO plate (in a thick patch) or in 7mL liquid SPO in a 50mL flask on shaker. Let sporulate 2 days to 1-2 weeks at 30°C (aka until tetrads are visible under the microscope). Better yield is obtained in liquid SPO. It is also possible to collect cells from a meiotic time course.

→ Note for using the dissection scope: Set the needle at the lowest position with the knob before setting a plate on the scope, and before removing it, to avoid breaking the needle. Set the needle at the lowest position with the joystick to move along the plate in 2D.

1. → If starting from a SPO plate, using a sterile toothpick, resuspend a good amount of cells in 20µL zymolyase in a 1.5mL tube to digest tetrad ascus (1mg/mL if 100T zymolyase, 5mg/mL if 20T zymolyase in 1.2 M Sorbitol Citrate). 

→ If starting from liquid SPO, spin down 500µL of culture and resuspend cell pellet in 20µL zymolyase.

Note: handle zymolyase carefully, do not vortex or drop. 

1. Incubate @ 37°C for 30 mn (can be adjusted).

2. Add 1mL of sterile H2O to dilute and inactivate the zymolyase; invert or flick tube to mix. No pipetting or vortex to not disrupt digested tetrads. 

Note: Digested tetrads can be kept as is at 4°C for 1-2 weeks for later dissection.

1. Lay 15µL of digested tetrad along the center of a YPD dissection plate. You can draw a line on the back of the plate to help, but erase it with ethanol after for easier viewing

Note: do not prepare plates too much in advance, as the spores may start to germinate after ~hours on rich medium.

1. Set the plate on the dissection scope, center the objective on the cell line, and adjust the focus to see cells. Carefully turn the knob to lift the needle until its shadow appears. Use the joystick up and down and left and right to pick up cells. 

2. Look along the center strip of yeast for a tetrad, and lift the needle to pick it up. Real tetrads are small, and the 4 spores are of equal size and similar aspect, and appear disposed in a regular diamond shape.

3. Move the microscope stage over 10 units from the center, and deposit one of the tetrads. Deposit the remaining three spores 5 units over each (usually coordinates 15-20-25-30 and 45-50-55-60 on the x-axis, and coordinates 5 to 45 on the y-axis). Record cell deposition or failure on paper to facilitate interpretation.

4. Move the stage down 5 units and repeat, creating 9 total rows of broken-apart tetrads on each side, to create the pattern seen on the picture. 

5. Incubate the plate at 30°C for 2 days until there is sufficient growth. After one day, using a scalpel wiped with ethanol, remove center strip of agar+yeast from the dissection plate, being careful to not disturb or cut off the tetrads

Replica Plating

1. Gather, for each plate to replica plate (= primary plate), 

• Secondary plates: 2 MIN plates (label them as “A” and “α”), 1 YPD plate, 1 YPG plate, and 1 plate for any other media to be tested (label them as a appropriate).

• 2 velvets (kept sterile in aluminum wrap) and a metal block.

1. Prepare Mating test plates:

• Method 1: Add 200µL YPD on each MIN plate. For each tester strain, using a 5mL serological pipette, collect cells from stock plate (stored in under-bench fridge), and evenly spread the YPD+cells mixture onto the designated plates. Let dry plates half-o under the biosafety cabinet a few minutes.

• Method 2: ‘the Copacabana technique’ (you can prepare up to ~8 plates at once.)

• Stack MIN+MATa and MIN+MATα plates in 2 separate stacks. Add ~15 glass beads to each plate. For each tester (stored in 5mL tube in under-bench fridge), dispense 100uL of cell suspension onto appropriate plates. 

• Spread cells over the agar surface by shaking the stack of plates in an horizontal motion (avoid swirling), alternating orientation, until liquid is fully absorbed.

• Discard beads in a beaker containing ~2mL 70% EtOH, aspirate ethanol, and transfer beads to ‘used beads’ collection bottle (contains ~100mL 10% bleach). 

1. Replica-plating per se:

1. Fix a velvet onto the metal block, and press the primary plate onto the velvet to transfer cells (tap it gently but firmly). Make sure that all the yeast colonies/patches are partially (but not fully!) transferred onto the velvet.

2. For each secondary plate, press it over the velvet w/ cells and give it two firm slaps with your palm to stamp yeast cells from the velvet to the plates. 

TIP: Mark with a sharpie any visible large clump of cells transferred to a secondary plate that may be erroneously interpreted as positive growth.

→ Important: You can stamp cells onto several plates successively from the same velvet but it is necessary to switch velvet after a MIN+tester plate, since it contains yeasts at its surface. Thus you need 2 velvets per primary plate. 

NOTE: With SK1 (rough morphology) the second velvet gets usually better transfer from the primary plate than the first velvet. The opposite is true for BY or W303 (smooth morphology). Growth on synthetic media is limited compared to growth on yeast extract based media. 

Thus, optimum plate repartition can be:

First velvet: Second velvet:

- YPD (control of transfer) - YPG (control of transfer)

- brown plates (yeast extract based) - white plates (synthetic media)

- MIN+MATa - MIN+MATα

1. Incubate stack of plates in warm room overnight.

2. Score and record the growth/no growth results the next day. If transfer is poor, or strain is “sick”, it may require an additional day of growth.
   

Basis for mating type test: 

MATa and MATα tester strains (respectively B398 and B399 in our collection) are his1. They can be used to test mating type of any auxotrophic strain (= with nutritional requirements) that is HIS1 (which is the case for virtually all our strains). Your haploid strain will mate with either MATa or MATα tester strain, and the resulting diploid will be prototrophic (= no nutritional requirement) by genetic complementation, and thus will be able to grow on minimum medium (the phenotype is called a-mater or α-mater). Your auxotrophic diploid strain won’t mate with neither MATa or MATα tester strain, thus will remain auxotrophic and won’t grow on MIN+a or Min+α plates (the phenotype is called non-mater). It is wise to add a MATa and a MATα control onto the primary plate when looking for non-maters, to make sure that the test worked!

Adding Strains to Collection

Haploids

1. Start with strains of interest, e.g. successful transformation patches that have been confirmed by PCR or spore from tetrad dissection that have the desired characteristics.

2. Update the Berchowitz Yeast database with the new strain number, description, and characteristics.

3. Plate a full small or half a large YPG plate with the yeast streaked out to create a lawn.

4. Incubate @ 30°C for two days to allow sufficiently dense growth.

5. In a sterile screw-top cryo tube, add 1mL 15% glycerol using a filter tip. Label the tube.

6. Using a sterile stick, scrape up the entire lawn of yeast cells and mix into the glycerol. Vortex and store at -80°C.

Diploids

1. Start with strains of interest, e.g. crossed strains purified in single colony and tested.

2. Update the Berchowitz Yeast database with the new strain number, description, and characteristics.

3. Plate a full small or half a large YPD 4% plate with the yeast streaked out to create a lawn.

4. Incubate @ 30°C O/N. (beware: longer time of growth or growth on a less rich medium such as YPG will cause SK1 diploid to starve and sporulate).

5. In a sterile screw-top cryo tube, add 1mL 15% glycerol using a filter tip. Label the tube.

6. Using a sterile stick, scrape up the entire lawn of yeast cells and mix into the glycerol. Vortex and store at -80°C.

Pulling out Strains from Collection

1. Label YPG* plate with desired strain numbers (*unless specified otherwise in the strain database entry for your strain of interest).

2. Remove the tube from the -80°C freezer (do not thaw!).

3. Wearing clean gloves and using a long sterile stick, take a scraping of the frozen yeast cells, and patch it to the plate at the location of its number.

4. Let grow @ 30°C O/N.

Creating Knockout Primers

Start with gene of interest to be knocked out (eg. BMH1), Berchowitz Yeast database, SnapGene software, and SGD website.

1. Go to www.yeastgenome.org. In the search bar, enter the name of the gene to be knocked out. On the results page, in the section “sequence”, opent the “download” drop-down menu. Select “Genomic DNA +/- 1kb”, and download the file.

NOTE: the sequence of SK1 genome is available in the lab Dropbox. You can use SnapGene software to align the sequence extracted on SGD onto SK1 chromosome to find it and design your primers directly onto SK1.

1. Open the file in the SnapGene software, select “show translation” to find the ORF and switch to the “sequence” tab to see the sequence.

2. (you can also repeat 1 to download the “Genomic DNA” file, which gives only the sequence of the gene of interest and should start with “ATG”. Then align this sequence onto the “Genomic DNA +/- 1kb” to identify the ORF.)

3. Select the sequence of the ORF, select “Feature” from the top of the screen, then “Add Feature”. Enter the name “[gene] CDS” for coding sequence and select OK. This will annotate the actual gene on the genome file.

4. Directly upstream of the gene, select the first 40 basepairs from the coding sequence. At the top, click “Primers” then “Add Primer” and select “Top Strand” for the forward primer (or “Bottom Strand” for the reverse primer).

5. Go the LEB Primers database and create new entries for the primers being created. Make sure all information is filled in.

6. Back in SnapGene, add the name of the primer according to the database such as “LEB_22”. From the popup menu, you can select and copy the primer sequence to paste into the LEB database. For the primers directly up- and down-stream, make sure to add the corresponding 20bp plasmid sequence to the primer. Then the pop-up, select “Add Primer to Template”.

7. Repeat for the reverse primer, except select “Bottom Strand” to make the primer from.

8. Then, make the check primers by moving ~20bp upstream (or downstream for reverse) from the other primers and making a 20bp check primer, repeating steps from above and pasting the sequence into the database.

9. Order primers from IDT (www.idtdna.com select DNA & RNA > DNA oligos > single stranded DNA > order now > enter the number of item to order > fill the name LB_XXX and the sequence, formulation = none, purification = standard desalting. Proceed to order.)

10. When receiving primers, resuspend into sterile H2O to 20uM (v=n/c). The quantity in nmol is indicated on the tube label. If volume is > 2mL it won’t fit in the tube, so make a higher concentration (100uM for example) and from that make a 20uM stock in a screw cap tube with blue cap (store in ‘PRIMERS additional dilutions” box.)

Meiotic Time-Course Experiment 

(Updated: 4/30/18)

Overview:

Plan for 6 days of setup and ~13 hours on collection day for experiment.

1. Any time prior to experiment day (day 5), label all tubes. Per strain (usually 4 strains, 12 time points):

• 1 250mL flasks for day 3, 1 2L flasks for day 4 and 1 2L flasks for day 5 (label with strain#).

• 1 50mL conical tube for cell washing on day 5 (label with strain#).

• 12 15mL conical tube for sample collection (color dot on side)

• If IF: 12 microcentrifuge tube (color dot and label with strain# and hours on top) NOTE: Use Eppendorf brand microtubes for IF as they hold cells slightly better than generic tubes and prevent sample loss.

• 2mL tubes (colored cap labeled with strain# and hours, color mark+hrs on side)

• 12 for Western

• 12 for RNA

• 8 for SDD-AGE

NOTE: SDD-AGE samples are only taken on the hour (8 total); omit for half-hour sample points

• If FACS samples are desired, prepare 12 microtube (regular ones) and label with color dot, strain, and hour on the top.

• If DNA samples are desired to PCR-check genes of interest for the timecourse strains, prepare 1 microtube tube per strain.

⇒ Color-code each set of samples per strain using colored caps for 2mL tubes as well as paint pens for conical tubes and sides of 2mL tubes.

1. Prepare media if not already available:

• YPD: use 1.05 YEP stock bottle, add 50mL 40% glucose, 10mL 100X Adenine, 10mL 100X Uracil, 10mL Tryptophan.

• SPO (0.3% KAc + 0.02% Raffinose): use SPO stock bottle, add 1mL 20% Raffinose.

• (BYTA should be available as stock)

‘Inoculation at specific OD’ cheat sheet:

• Check OD600 of starting culture on spectrophotometer by diluting 1:20 (50µL from each culture into 950µL H2O and blank cuvette: 50µL medium into 950µL H2O). Multiply OD600 reading by 20 to get the actual OD of the culture.

• v1 = c2 x v2 / c1 → v1 = <wanted OD> x <final volume> / <starting OD>

‘Induction ’ cheat sheet:

• If inducing expression of pGAL-NDT80, add β-estradiol to 1µM at t=6h.

• If inducing expression of pCUP1-IME1/4, add CuSO4 to 25µM at t=2h.

• If inducing degradation of AID-tagged protein, with osTIR1 under pCUP1 promoter, add 25µM CuSO4 at t=4h and 1mM Auxin at t=5h. Alternatively, use osTIR1 under pHOP1 promoter, and induce with 1mM Auxin at t=5h or 1mM Auxin at t=5h30.

Day 1

1. Pull out cells from collection onto a YPG plate, grow overnight in warm room.

Day 2

1.  Patch to a YPD 4% plate from the YPG patch, grow overnight in warm room.

Day 3

1.  Around noon: Inoculate each strain into 25mL YPD liquid culture in 250mL flasks, grow overnight on shaker in warm room.

Day 4

1. Add 50µL of formaldehyde in the eppendorf tubes labeled for IF. Move sample tubes to the cold room in racks. (formaldehyde in small volume evaporate easily, so it is better to keep the tubes in cold. If required to keep tubes for days later, keep @ -20)

2. Set up cold room with pipette and tips, aspiration setup, a tube rack for western, IF, and FACS if applicable, and 5% TCA with dispensette adaptor set to 1.8mL.

3. Set up the -80ºC with tubes racks for Northern and SDD-AGE if applicable.

4. Set up warm room with 10mL serological pipettes, mechanical pipettor, prepared 15mL conical tubes, and sharps container. 

5. At noon, dilute into 100mL BYTA to OD600 = 0.3 into a 2L flask and grow overnight on shaker in warm room (usually: ~2mL YPD culture into ~98mL).

Day 5 (experiment day)

1. At ~6:30 am, collect 360 OD600 of cells (usually ~40-55mL of BYTA culture).

2. Set up the cold room with a bucket half-filled with liquid nitrogen.

3. Transfer to a 50mL tube, spin down 1min @ 3,000g, dump supernatant (if >50ml, do it in 2 steps).

4. Wash once in ~25mL H2O, spin down 1min @ 3,000g, dump supernatant. 

5. At 7:00 am, resuspend cells into 200mL SPO at OD600 = 1.8. (Resuspend pellet in 25mL SPO medium, transfer to a new 2L flask, rinse tube with 25mL SPO, transfer to same 2L flask, and add the remaining 150mL SPO). Setup SPO flasks in warm room on shaker.

6. Volume sampling:

• IF: 450µL

• FACS: 1mL

• Northern: 2mL

• Western: 4mL

• SDD-AGE: 2mL

Routinely, standard time points are collected at t=0, 6, 7, 7.5, 8, 8.5, 9, 9.5, 10, 10.5, 11, and 12hrs, with IF, Northern and Western at all time points, and SDD-AGE on the hour only. So <volume>=8.5mL for on-the-hour and <volume>=6.5mL for half-hour samples.

At t=6hrs (right after the t=6h samples have been collected), induce NDT80 expression by adding β-estradiol to a final concentration of 1µM (add 40µL of 5mM stock [stored at -20℃] for an initial 200mL volume).

NOTE: Handle β-estradiol carefully-- be sure to wear gloves and avoid contact with skin. Use a filter tip to add stock to the flask(s). See MSDS for more information.

For each sampling point:

1. In the warm room remove required volume from the flask and dispense into the conical tube. Repeat for all cultures/strains.

NOTE: Do not forget to turn the shaker back on!

1. In the cold room, dispense sample into appropriate tubes for the time point: 450µL into the microtube with formaldehyde for IF, 1mL into the microtube for FACS, and 2mL into each RNA and SDD-AGE screwcap tubes, leaving 4mL in the conical tube for Westerns.

2. Spin down the conical tubes at maximum speed.

3. Meanwhile, spin down the 2mL tubes and the microtubes for FACS 2min @ 3,000g.

4. Dump supernatant from the conical tubes down the sink. Per tube, add 1.8mL 5% TCA (stored in amber bottle w/ dispenser in cold room). Pipette mix and transfer to a 2mL screwcap tube. Transfer to Western rack. Store at 4ºC.

5. Return to 2mL tubes: aspirate liquid and snap freeze by throwing the closed tubes in liquid nitrogen. Store at -80ºC.

6. For the microtubes for FACS, aspirate the liquid and resuspend cells in 1mL of 70% EtOH, transfer to FACS rack and store at 4ºC.

7. As the time course goes, transfer 2mL tubes from liquid nitrogen to pre-chilled tube racks and move to -80ºC (tip: transfer regularly, or at most in between the 11h and 12h time point).

8. NOTE: it may be wise, although not mandatory, to keep the flasks in the warm room overnight or over the week-end to check for presence of tetrads (at least in the control strains) before processing the samples.

Day 6 (or later)

Time course sample processing:

• Samples with Formaldehyde - IF Microscopy; Samples should be prepped the day after taken at least until the Sorbitol citrate step, as longer time spent in formaldehyde makes them more difficult to digest. Once prepped, samples can remain frozen until imaged.

• Samples with TCA - Protein/Western Blot; Samples can be extracted and frozen at any time, but the best is to process samples until the drying step. Alternatively, samples can be completely processed and remain frozen prior to running gel. 

• Spun-down Samples - SDD-AGE Blot can be started at any time but requires 4 consecutive days for all steps.

• Spun-down Samples - RNA/Northern Blot can be started at any time but requires radioactive P-32, which must be ordered as needed, and requires consecutive days. The best is to process samples until freezing in ethanol step to be processed later.

• FACS - Samples are best processed within ~2 weeks. Requires two days of processing with time booked (via iLab) to read samples on the BD LSR II. Once Day 1 is completed, samples should be completed and read the next day as the SYTOX Green dye will deplete over time, reducing efficiency.

IF Microscopy

Solutions:

1M Potassium Phosphate Monobasic: 136.09g/L in 960mL H2O, dissolves easily, autoclave

1M Potassium Phosphate Dibasic: 174.2g/L in 960mL H2O, dissolves easily, autoclave

0.1 M KPi [pH 6.4]: 27.8mL 1 M Potassium Phosphate Dibasic, 72.2mL 1 M Potassium Phosphate Monobasic, 900mL H2O

1.2M Sorbitol Citrate: 17.4 g Potassium phosphate dibasic (K2HPO4), 7.0 g Citric Acid, 218.64g Sorbitol, dissolve completely, add H2O up to 1L, then sterile-filter.

Digestion/Sample Prep:

⇒ Start with microtube containing ~450µL sample + 50µL (37%) formaldehyde (incubated overnight at 4ºC). 

Spin down in cold room 2min @ 3,000g.

1. Aspirate liquid, leaving pellet behind (be careful not to suck any up-- leave behind a small amount of liquid if necessary).

2. Resuspend in 1mL 0.1M KPi [pH 6.4] (Potassium phosphate buffer - Stored @4℃) and vortex.

3. Spin down and aspirate liquid (NB: at this stage, cell pellet is barely visible as cells mostly stick to the tube wall. Pellet will be visible again at later steps).

4. Resuspend in 1mL 1.2M Sorbitol Citrate (samples can be frozen at this point to continue later).

5. Spin down and aspirate liquid.

6. Prepare fresh digestion mixture: per sample, 200µL 1.2M Sorbitol citrate, 20µL Glusulase (4ºC), and 6µL 100T Zymolyase (concentrated 10mg/mL stock is prepared in a microtube @ -20ºC in the enzymes box) For 24 samples, make a stock using 6mL sorbitol, 600µL glusulase, and 180µL Zymolyase and add 226µL/sample tube.

7. Vortex thoroughly to resuspend.

8. Incubate @30℃ in the warm room on the rotisserie rack for 45min, then check on cell wall digestion in the microscope. Leave longer as necessary to ensure adequate digestion. 

NOTE: Check digestion every 30 min. Pipet on a slide 1.5 ul cells twice, add 1.5uL 1% SDS to one spot, cover with round coverslip and check under microscope. You should see ⅓ cells in the spot with SDS compared to the one with no SDS.

1. Spin down and aspirate, leaving behind a brown pellet.

2. Wash with 1mL 1.2M Sorbitol Citrate-- DO NOT VORTEX!-- mix with pipette.

3. Spin down and aspirate.

4. Resuspend in 100µL 1.2M Sorbitol Citrate and freeze until ready to image.

IF Slide Preparation:

1. Either:

1. Prepare slide by placing 5µL of 0.1% polylysine (in 2mL aliquots stored in 4ºC) on each well and letting it sit for 5 minutes. Do not touch well.

2. Or use prepared slide pre-coated with poly-lysine

2. Wash slide under distilled water and air dry completely.

3. Fully resuspend the cells and add 5µL to each well. Do not touch well with pipette tip. Let cells sit in wells for 10 minutes. 

4. Remove sup from each well by placing vacuum tip to the side of the well, without directly touching the well with the aspirator tip.

5. Using forceps, put slide in methanol for 3 minutes followed by 10 seconds in acetone. Let it air dry completely.

6. Add antibody - either conjugated (1 step) or not (2 steps, primary then secondary):

1. Add 5µL of 1:200 anti tubulin FITC (this is conjugated) (or other appropriate antibody) to each well. Prepare mix of 2.5uL Ab in 497.5uL 1% PBS BSA.

Or Add 5µL of 1:200 anti tubulin (this is not conjugated) (or other appropriate antibody) to each well. Prepare mix of 2.5uL Ab in 497.5uL 1% PBS BSA.

1. Incubate in wet chamber for a minimum of 2 hours (can be left overnight). 

1. Remove supernatant and wash each well 5X in 5µL PBS/BSA(contaminates easily, always use aliquot) by pipetting, letting the PBS/BSA sit for ~3 minutes, and aspirating drops 5X. Do not touch wells.

2. If need secondary, repeat steps 6 and 7 with anti rat FITC 1:200.

3. Remove slide from wet chamber, let the wells dry completely, and add 3µL (approximate by eye) of DAPI-MOUNT to each sample. 

4. Carefully put coverslip on top of slide, being sure to completely cover all wells containing cells. Lightly wipe the coverslip with a kimwipe to press out bubbles while holding down the edges of the coverslip so it does not slide.

5. Carefully paint with clear nail polish around the edges of the coverslip to seal it (do not cover wells containing cells with nail polish) and let it dry for 10 minutes before proceeding with imaging. 

6. Turning on scope, be sure to oil the 100X lens and open the if.exp (lowercase) experiment on the computer to photograph and count the cells in each well. Image should include DAPI and FITC channels (blue and green).

 Operating the DeltaVision Microscope:

1. Begin by turning on the computer and microscope by pressing the two power buttons on their respective CPUs (both will light up blue when turned on). The computer CPU is the upper unit and the microscope CPU is the lower unit on the desk. 

2. Turn on the air for the microscope table by turning the knob on the yellow oxygen tank to the left.

3. Log on to the computer with password cn84864 and double click on the desktop icon for “Start SoftwoRks” to open up the microscope software. Go to “File” and click on the “Acquire” option. Confirm that the objective turret is lowered by turning the coarse focus knob on the righthand side of the microscope away from you, then click “Initialize.”

4. When the software is open on the computer, open up the marked points list by clicking on the bullet point list icon, and click “mark point.”

5. To begin an experiment, click on the flask icon at the top of the program. Open a saved experiment by clicking on the “open experiment” icon in the Design/Run experiment window that will pop up. For IF use the if.exp (lowercase) experiment, for the screen use Screen1.exp, and for a movie use meiosis1.exp. 

6. Adjust the settings on the microscope accordingly; to adjust the lens setting of the microscope, use the revolving nosepiece to switch between the 100X or 60X lens and adjust the dial below the oculars to DIC 100 or DIC 60. In the SoftwoRks program, change the lens setting to match whichever lens you are using. For if.exp you should use 100X; for Screen1.exp and meiosis1.exp you should use 60X.

7. To adjust the filter settings you are using, first rotate the eyepiece filter wheel to either 1 or 2. 1=channel for D-F-T-C, and 2=channel for D-F-mCH-Cy. Then in the Polychroic drop-down menu in the program, choose the corresponding channel. FInally, clicking the gear icon in the program and going to the “Misc” tab, change the filter wheel sets to the corresponding channel for excitation, emission, and eyepiece, then click Activate. Use channel 1 for IF.exp and channel 2 for Screen1.exp and meiosis1.exp.

8. Going to the “Run” tab of the Design/Run experiment window, Enter “1” for the image file name and adjust the location of the images you will capture in “Settings.” Check the box for “Enable Post-Acquisition Processing,” go to “Processing Tasks,” and click the “+” symbol to add a task. Click on the task and select “Quick Projection” from the drop-down list.

9. Place one drop of oil on the objective and mount the slide coverslip down. Raise the objective turret until the lens is just touching the slide, and focus using the eyepiece. Once a desired area on the slide has been found, rotate the port selector knob located on the front of the microscope base to the camera position. 

10. On the computer, use the black stage window and the camera icon to navigate the slide and acquire images of yeast fields. Focus the image using the Z stage position (A black bar to the right of the black stage window). When you have found a usable location on the black stage window, use the open marked points list to mark a point. Repeat until you have at least 100 cells per well/sample/timepoint. 

11. Click the green arrow icon to run the experiment. Unless you made any important adjustments to the experiment, do not save changes to the experiment before running. After the experiment has completed, clear the marked points, change the location of the images, and repeat step 10. You may need to re-oil the objective occasionally. 

12. To view the images you have taken on the microscope, open the “Home” folder on the computer desktop. Double click on the subfolder “Data 2” to access your personal folder and the subfolders within that containing all experiment files. Open the “PRJ” files to view the images and score them. 

13. After you have completed imaging the slide, close the program. Remove the slide from the slide holder and use a lens wipe to carefully but thoroughly remove the oil from the lens using a blotting technique. Lower the objective turret, turn the knob on the oxygen tank to the right to turn it off, and transfer the image files to a hard drive before shutting down the computer and the microscope. To shut off the microscope, hold the power button on the CPU down until the blue light turns off.

Western Blot

1. In tubes with pellet + 1.8mL 5% TCA, spin down 5min @ 5,000g. 

2. Aspirate TCA, ensuring pellet is dry. (Liquid left behind will affect the pH of the sample)

3. Wash with 1mL acetone, vortex (pellet will NOT resuspend), and spin down again 5min @ max speed. 

4. Aspirate and let dry (~30min-1hr should be sufficient; pellet will appear dry and cracked).

(→ pause point: dry pellet can be kept as is and processed later - if you need to keep your samples for a long time that is the best step to stop at).

1. Make fresh breakage buffer (adjust volumes as necessary) :

For 10 samples:

• 1mL TE Buffer

• 2.75µL 1M DTT (Dithiothreotol, stock is at -20ºC)

• 10µL Halt Protease Inhibitor (4ºC)

• 10µL 1M Tris Buffer, pH 11

1. Into the screw-cap tube with the pellet, add 50µL glass beads and 100µL breakage buffer.

2. Process the tubes in the FastPrep for 45 s @ max speed (6.5M/S) in the cold room.

NOTE: make sure to tightly close tubes, to secure it well in place. Make sure that the disk holding the tubes is well in place, and tighten the screw top. Hold the lid of the machine while running.

1. Add 50µL 3x SDS loading buffer + BME

2. Boil 5min @ 100ºC

3. Spin down 5min @ max rpm

Note: At this stage, if needed, samples can be diluted with 1X SDS loading buffer + BME in breakage buffer to adjust for OD 

(→ pause point: samples may be frozen to run gel later).

1. Load gel: 

• Prepare a BioRad pre-cast gel--there are small/”mini” (10 or 15 well) and large/”midi” (26 well) sizes, and different percentages of gel vs gradient. Be sure to remove the tape strip from the bottom. 

• Rinse the gel and wells with 1x SDS Running Buffer and aspirate out the wells.

• Load gels into the dock-- for mini box, wells should be facing the inside, on the gasket; for the midi box, the sample reservoir should be on the inside. 

• Fill box with 1x SDS Running Buffer. For the mini box, liquid level should cover the tops of the gel wells completely inside and to the appropriate fill line on the outside; for the large box, the outer area should be filled to the fill line and the space for the wells should be filled over the tops of the wells.

• Load 4µL sample per well, with the Biorad ladder in the first well, samples in the remaining wells, and finally a negative/no-tag control (usually: 15055 @ 6hrs, prepared @ -20ºC) on the last well on the right end.

• Run at 200V for approximately 40min, or until the dye reaches the bottom of the gel.

1. Transfer the proteins from the gel to a membrane:

• Gather materials before starting - pre-prepared Transfer Buffer (per 1L: 200mL 5x Transfer Buffer from 4ºC, 200mL Ethanol, 600mL H2O), plastic trays with Transfer Buffer, 2 stacks BioRad blotter paper, Biorad nitrocellulose membrane (1 per gel), piece of parafilm slightly larger than gel, small flat scoopula to open gel case, pencil, pipettor and serological pipette, and mini roller. Label the membrane as appropriate.

• Rinse off the gel cartridges in DI water.

• Use the scoopula to break open the sides of the gel casing and gently remove the gel into a plastic tray with Transfer Buffer (gel should be submerged).

• Soak one stack of blotter paper in the other tray with Transfer Buffer, place the stack inside the BioRad TurboBlot tray. Roll out gently to smooth.

• Wet the membrane and place on top of the blotter paper. Add transfer buffer on top of the membrane, creating a bubble of liquid over the entire surface. 

• Use the parafilm to pick up the gel from the transfer buffer and gently slide it on top of the membrane, taking care not to make it wavy or tear the gel.

NOTE: the ladder should be oriented to the left side of the membrane.

• Wet the second stack of blotter paper and place it over the gel. Smooth out very gently with the roller. Transfer excess Transfer Buffer back into the bottle.

• Repeat for any remaining gels into the TurboBlot trays.

• Close and lock the tray lid and run the pre-set program for Mixed Weight (7min) if using “turbo” blotting paper or Standard SD (30min) if using standard blotting paper. 

Once blotting is complete, 

• Reveal the membrane in Ponceau, rinse w/ DI water. 

NOTE: Ponceau is re-usable - put back into the bottle after use.

• Make fresh Blocking Buffer, 3% milk in TBST (1.5g milk powder + 50mL 1x TBST). 

• Transfer the membrane to a plastic tray w/ Blocking Buffer

• Incubate @ RT for 1hr on the shaker table.

(→ pause point: membrane may be kept in tray w/ 1x TBST @4℃ to stain it later)

1. Dump milk, add appropriate primary antibody, and incubate on the shaker table (see below for typical antibody progression). 

NOTE: primary antibodies are re-usable and pre-prepared at 4℃; dump the entire 50mL tube into the tray. (Final concentrations are 1% milk, 1% BSA, 1:50 sodium azide, and antibody (concentration depends on antibody, see below), in TBST).

1. Rinse twice in 1X TBST and wash twice 10min @ RT on the shaker in 1x TBST.

2. Add secondary antibody (prepare fresh) and incubate. Antibody mixture should be 1% milk and 1% BSA in 1x TBST, and antibody concentration/incubation time is dependent on which primary antibody was used (see typical progression below). 50mL is typically plenty enough, though for smaller mini gels as little as 10mL can be sufficient. Note secondary antibody is the antibody of the primary, and is always animal-based. We use anti-mouse for standard Westerns.

3. Rinse twice in 1x TBST and wash twice 10min @ RT on the shaker in 1x TBST

4. Stain in ECL Prime - 1mL each of the two components A&B, pipette mix, and run over the membranes several times. Move the membrane(s) inside a sheet protector for imaging to prevent drying out and ensure consistency in images. Image via chemiluminescence at multiple exposure times.

NOTE: Ponceau can be used to remove antibodies so the membrane can be re-treated to image a different protein (but due to differences in abundance, this is not always necessary, see below). In this case, the less-abundant protein should be treated for/imaged first.

In some context, you might want to strip the membrane before using another antibody.

Mild stripping method:

• Re-stain membrane in ponceau

• Rinse in DI several times

• Block 1hr in 1x TBST 3% milk

• Proceed to primary and secondary antibodies as usual.

--Typical antibody progression for our standard meiotic time course:

Repeat steps 12-16 for the following antibodies:

(Ponceau is not necessary between these since Clb3-HA is much lower abundance than Pgk1, and Pgk1 is much lower abundance than Rim4-v5.)

Day 1: 12. 1°: ɑ-HA 1:1,000 O/N @4ºC

Day 2: 13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-mouse 1:10,000 2hrs @RT (5µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in 1X TBST

16. Stain with ECL, Image typically 1min, 2min, 5min

→ Rinse in DI

12. 1°: ɑ-pgk1 1:20,000 O/N @4ºC

Day 3: 13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-mouse 1:20,000 2hrs @RT (2.5µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in 1X TBST

16. Stain with ECL, Image typically 30s, 1min

→ Rinse in DI

12. 1°: ɑ-v5 1:2,000 30min @RT

13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-mouse 1:40,000 30min @RT (1.25µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in 1X TBST

16. Stain with ECL, Image typically 15s, 30s, 1min

Typical antibody progression for Ty3 project:

Repeat steps 12-16 for the following antibodies:

Day 1: 12. 1°: ɑ-VLP 1:1,000 O/N @4ºC

Day 2: 13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-rabbit 1:40,000 1hrs @RT (1.25µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in TBST

16. Stain with ECL, Image typically 2mn, 20mn

→ Rinse in DI

NOTE: it can be useful to repeat with stronger secondary:

14. 2°: ɑ-rabbit 1:20,000 1hrs @RT (2.5µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in TBST

16. Stain with ECL, Image typically 2mn, 20mn

→ Rinse in DI

12. 1°: ɑ-pgk1 1:20,000 O/N @4ºC

Day 3: 13. rinse and wash 2x in TBST

14. 2°: ɑ-mouse 1:20,000 2hrs @RT (2.5µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in TBST

16. Stain with ECL, Image typically 30s, 1min

→ Rinse in DI

Typical antibody progression for other tags:

Repeat steps 12-16 for the following antibodies:

ɑ-Flag 

Day 1: 12. 1°: ɑ-Flag 1/10,000 O/N @4ºC

Day 2: 13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-mouse 1:20,000 2hr @RT (2.5µL/50mL in 1X TBST 1% milk 1% BSA)

15. rinse and wash 2x in TBST

16. Stain with ECL, Image typically 1mn, 5mn 

ɑ-Myc

Day 1: 12. 1°: ɑ-Myc 1/1,000 <enter recommended time>

Day 2: 13. rinse and wash 2x in 1X TBST

14. 2°: ɑ-mouse <enter recommended dilution>

15. rinse and wash 2x in TBST

16. Stain with ECL, Image typically <enter recommended time>

House-Made Western Gels

1. Prep casts: add gray gasket(s) to bottom of cast. Fit a back and front piece of glass into the green clamp and shut; slide into cast and clamp on top.

2. Prepare solutions: Main gel and Stacking gel (do not add Ammonium persulfate and TEMED yet) - these volumes are sufficient for 4 gels, refer to chart to make more or less or different percentages.

Main gel is 25mL 10%:

H2O 9.9mL

30% Acrylamide 8.3mL

1.5M Tris pH 8.8 6.3mL

10% SDS 250µL

---------

10% Ammonium Persulfate 250µL

TEMED 10µL

Stacking gel is 6mL:

H2O 4.1mL

30% Acrylamide 1.0mL

1M Tris pH 6.8 750µL

10% SDS 60µL

--------

10% Ammonium Persulfate 60µL

TEMED 6µL

1. Add Ammonium Persulfate and TEMED to Main gel solution only, and use a 10mL serological pipette to mix. Add to gel casts by placing the end of the pipette tip perpendicular to the back of the gel cast and dispensing carefully. Fill to the bottom of the green line.

2. Add 1mL Isopropanyl alcohol to the top of each gel, filling it the rest of the way (this will make sure the top is flat/even)

3. After ~10min when gels have solidified (check the remaining solution in the beaker to see if it is solid), aspirate off the IPA.

4. Add ammonium persulfate and TEMED to the stacking gel beaker, mix, and add ~1mL to the top of each gel to fill it to the top. Place combs with the BioRad logo facing out, being careful to avoid any splashes by holding a KimWipe in front of the gel.

5. Let sit a minimum of 10min prior to running; place into box and proceed as with pre-cast gels.

Staining protein gels

Imperial Blue staining

Sensitivity: up to 3 ng per band.

For SDS-PAGE (acrylamide):

1. After electrophoresis, place gel in a clean tray with 50-100mL ultrapure water.

2. Wash 3x 5min in 50-100mL ultrapure water on shaker.

Tip: wearing gloves, use your hand as a “flat cage” to retain the gel in the container when emptying it.

1. Before adding the Imperial Protein Stain (Thermoscientific), invert bottle a few times to mix.

2. Remove all water and add 30-50mL Imperial Protein Stain. 

3. Place on shaker for a variable amount of time depending on the abundance of your protein (5 min to a few hours to overnight).

4. Discard staining reagent.

5. Wash 3x 5min in 50-100mL ultrapure water on shaker.

6. End by a final wash of a few hours to overnight to destain a maximum and increase sensitivity (increase band intensity in contrast with background).

Tip: add a folded Kimwipe in the container to enhance destaining.

Silver staining

Sensitivity: 10-50 fold more sensitive than imperial blue but more cumbersome.

Tips: 

• Gel should be manipulated preferentially with glass or polyethylene rods, eventually with PVC or latex gloved rinsed with DI water. Never touch gel with metal object of bare skin.

• Volumes and times needs to be adapted to gel size and thickness. Refer to manual if using gels thicker than 1mm or larger than 15x10cm.

• Before and after using any glassware or plastic container, wash them with lab detergent, then cleaned with 50% (8N) nitric acid, and rinsed with ultrapure water. This prevents inconsistency in staining.

For SDS-PAGE (acrylamide):

1. Bring 50mL of pre-made Development Accelerator Solution (less than 3 months old, stored at 4ºC) to room temperature or prepare fresh: 

• The manual suggests to prepare bigger stock: Dissolve 25g Development Accelerator Reagent into 475mL milliQ H20 in a beaker. Stir with a magnetic bar until dissolved. Add milliQ H20 to 500 mL. Maybe aliquot in 50mL falcon?

• For 1 gel: Dissolve 2.5g Development Accelerator Reagent into 45mL milliQ H20 in a beaker. Stir with a magnetic bar until dissolved. Add milliQ H20 to 50 mL.

1. Prepare Fixative Enhancer Solution:

For 1 mini gel (200 mL) For 1 midi gel (400 mL)

Methanol 100 mL 200 mL

Acetic acid 20 mL 40 mL 

Fixative Enhancer Conc. 20 mL 40 mL

DI water 60 mL 120 mL

1. After gel electrophoresis, submerge gel in tray containing Fixative Enhancer Solution.

2. Incubate 20 min on shaker.

3. Decant Fixative Enhancer Solution from tray

4. Rinse gel 2x 10 min with milliQ water (200 mL for mini gel, 400 mL for Midi gel) on shaker.

5. Within 5 min before use, prepare staining solution: Move stirring platform under the hood and in a >200 mL beaker with magnetic stirrer, add:

• 35 mL milliQ H20

• 5 mL Silver Complex Solution

• 5 mL Reduction Moderator Solution

• 5 mL Image Development Reagent

1. Immediately before use: 

• Add 50mL Development Accelerator Solution (RT not cold!)

• Swirl well

• Add solution to the tray with gel.

1. Stain for 20 min or until desired staining intensity is reached. It may take >15mn before the first bands appear.

2. Prepare 5% acid acetic solution.

3. After the desired staining is reached, place the gel in 400 mL 5% acid acetic solution for 15 min.

4. Transfer staining solution in waste container under the hood.

5. Rinse and wash the gel in milliQ water for 5mn: the gel is ready to be dried or imaged.

6. Staining solution disposal: within a couple of days, the silver metal will precipitate from the solution. At this stage the aqueous waste can be decanted from the silver and disposed separately.

SDD-AGE

Updated 5/8/18

This protocol is based on the one outlined here: Halfmann, R., Lindquist, S. Screening for Amyloid Aggregation by Semi-Denaturing Detergent-Agarose Gel Electrophoresis. J. Vis. Exp. (17), e838, doi:10.3791/838 (2008).

Day 1: (NOTE: gel should be started at 5pm and run overnight until 9am the next morning; plan by pouring gel in the early afternoon and starting step 4 by 3:30-4pm)

1. Start with frozen pellet from 2mL liquid culture in a screw-cap vial.

2. Pour a gel: Use Owl EasyCast B2 size gel box fitted with one 20 lane comb (1.5mm side down). In a 500mL bottle, add 50mL 1x TAE, 49mL H2O and 1.7g Agarose (0.5x TAE, 1.7% agarose final gel concentration). Melt in microwave in 15-30s increments until all agarose is completely melted and the solution is perfectly clear. Leave cap screwed on tightly while microwaving to prevent evaporation and use short increments to prevent boiling over-- be extremely careful when opening as pressure will build. Once melted, move liquid around in bottle to re-integrate all of the steam on sides and top of the bottle into the gel. At the bench, add 1mL 10% SDS, being careful to avoid creating bubbles. Swirl gently to mix. Let cool 15-20m, then pour the entire bottle into the gel cast and leave to dry.

3. Once dry, move gels to the cold room and fill boxes with 0.5x TAES. Leave combs in the gel until ready to load to prevent them from becoming misshapen. Set up peristaltic pump to recirculate liquid from bottom to top of the gel box at 20rpm (~3.7mL/min).

4. Prepare tubes: Per sample, 3 Protein Lo-Bind microtubes labeled with sample names, 2 sets in the freezer (-80ºC) and 1 set at RT.

5. Prepare ice tray: in large ice tray w/ice, per sample add 1 15mL conical tube, cap removed, and a 1mL Gislon pipette tip (no filter) inside pointing to the bottom. From freezer, use a frozen metal block to hold samples.

6. Prepare lysis buffer solution in a 5mL conical tube, keep on ice: SDD-AGE Buffer (stored at 4ºC) contains 100mM Tris-HCl pH 8.0, 20mM NaCl, 2mM MgCl2, and 1% Triton-X, then just prior to use add Halt protease inhibitors, 50mM BME (both at 4ºC). Use ratio (per 10 samples):

2mL SDD-AGE Buffer

40µL Halt Protease Inhibitor

7µL BME (β-Mercaptoethanol)

1. Grab samples from -80ºC, plus controls 35408 @t=6 (no amyloid) and 15055 @t=6 (no tag). Put into frozen block.

2. In each screwcap tube, add 1 PCR tube (200µL) of 0.5mm silica/zirconium beads and 200µL lysis buffer.

3. Break in FastPrep, 45 s @ max speed (6.5M/S).

4. Poke a drainage hole in tubes: Use an 18G needle. Turn the screw-cap tube upside down and tamp twice to move contents away. Poke the hole into the angled side of the bottom cone of the tube.

NOTE: Do NOT poke the hole directly at the bottom center of the tube or it will clog. Put the screw-cap tube inside a 15mL centrifuge tube on top of the 1mL pipet tip.

1. Transfer 15mL conical tubes to the centrifuge in the cold room. Turn to max speed and leave 30s-1min.

2. Remove screw-cap tube from 15mL conical tube. There should be only beads remaining; if there is visible liquid, the tubes need to spin down longer. The 15mL conical tube should have ~200µL liquid inside it with the tip (some transfer of beads is normal, but not preferred). Use a pipette and the tip in the conical tube to remove the liquid and transfer into the first set of cold, labeled microtubes.

3. Spin down 10min @ 2,500g in the cold room. Transfer liquid (avoid any solids) into a fresh labeled tube and repeat spin down, reducing time to 5min.

4. Meanwhile, prep the RT set of microtubes with 20µL 4x SDD-AGE loading buffer (RT) [0.5x TAE, 8% SDS, 20% glycerol, bromophenol blue].

5. From sample tube, transfer 60µL supernatant to the microtube with loading buffer and flick to mix. Incubate at RT for 10min.

6. Load the gel:

For 8 samples + 2 controls: Skip the first lane, load 20µL sample/control in every other well, load 10µL ladder in the well between the last two samples (controls).

1. Run the gel at 25V overnight with TAES recirculation via peristaltic pump.

Day 2: Transfer the gel:

Prepare all materials before starting: B2 gel box, 1x TBS, large plastic tray, blotting paper wick (5.5” x 10”), 2 sets of 6x blotting paper (5.5” x 5”), nitrocellulose membrane (5.5” x 5”) labeled with date/strain as appropriate, parafilm (3 ½ squares cut into 4 strips), 4x pieces tape, folded stack of 16x SuperRags into thirds, glass plate, weight, pipettor w/10mL pipette.

1. In the plastic tray, add enough 1x TBS to submerge the gel.

2. Prepare the B2 Gel Box with the gel platform turned upside-down on the ledge. Across the platform, place a wick and fill the sides with 1x TBS.

3. Wet the first stack of 6x blotters in 1x TBS and place on top of the wick. Use a glass tube sprayed down with ethanol to roll across the blotter paper to flatten it out.

4. Stop gel running, discard TAES, and rinse with DI water. Remove any extraneous pieces of the gel sticking off and transfer to the tray with 1x TBS. Place on top of the blotters, wells facing down. Gently remove any air bubbles and add 10mL 1x TBS buffer on top of the surface of the gel.

5. Wet nitrocellulose membrane in 1x TBS buffer. Place on top of the gel with the label facing down.

6. Wet and place second stack of 6x more blotters on top of the membrane and repeat rolling the glass tube to flatten, being careful not to crush the gel.

7. Place parafilm around the 4 sides of the blotter/gel stack to prevent anything on top of the gel touching anything below the gel, as this will affect transfer. Tape to the gel box to hold in place.

8. Place folded stack of SuperRags on top of the blotters.

9. Top with glass plate and center a weight (such as a full 1L stock solution bottle)

10. Leave 24hrs to allow the gel to transfer.

Day 3: Primary Antibody

1. Unpack the gel transfer, discarding SuperRags, gel, and blotters. Transfer the membrane to a plastic tray.

2. Prepare Blocking Buffer in a beaker: per membrane, add 50mL TBST and 1.5g powdered milk (3% milk). Pour over membrane and incubate at RT on the shaker table for 1hr. Discard liquid.

3. Incubate in 1° antibody ɑ-v5 overnight on the cold room shaker-- 100mL bottle is prepared in 4ºC and can be reused. Final concentrations are 1% milk, 1% BSA, 1:50 5% sodium azide, and 1:2,000 ɑ-v5 antibody.

Day 4: Secondary Antibody & Image

1. Pour primary antibody back into its container. Rinse the membrane twice in 1x TBST, discarding liquid after use. Then wash by adding enough TBST to the tray to cover the membrane and incubate at RT for 10min on the shaker. Repeat wash step (2x total).

2. Prepare 2° ɑ-mouse antibody: in a 100mL beaker, per membrane add 0.5g BSA and 0.5g milk, then 50mL 1x TBST (1% solution), then 10µL secondary (ECL ɑ-mouse) for a final dilution of 1:5,000. Dump out the TBST rinse from the tray, add secondary solution, and incubate for 2hrs at RT on shaker.

3. Repeat 2x rinses and 2x washes.

4. Do a final stain in the container adding two parts of ECL Prime stain (1mL each solution A & B). Pipette mix, and use the pipettor to pick up stain and rinse over the membrane several times.

5. Transfer the membrane into a sheet protector to prevent drying and ensure images are consistent.

6. Image the membrane using chemiluminescence; start with a manual exposure of 15s and adjust accordingly, taking images at longer and shorter exposures (typically 1min and 2min). Save image file with the tag and exposure time in the comment. Discard membrane.

RNA Extraction & Northern Blotting - last update RL 2021-10-04

 Day 1: Start Extraction

⇒ Start with tubes containing frozen pellets of SPO culture (up to 24x at a time) . 

1. Switch thermomixer to 65ºC.

2. Transfer tubes into two metal tube racks for 2mL tubes inside an ice bucket. 

3. In each tube, add 50µL (small scoop) 0.5mm zirconia/silica beads

4. Add 400µL TES. Vortex to resuspend.

5. In the fume hood, add 400µL Acid Phenol. Vortex to mix.

6. Incubate tubes on thermomixer at 65ºC, 1400rpm for 30min.

7. Meanwhile, label a second set of screwcap tubes with 40µL 3M sodium acetate and 1mL 100% ethanol pre-chilled at -20ºC, and store at -20ºC. 

8. Spin down tubes in the cold room for 5 min @ max speed. 

9. In the cold room, transfer ~300µL of the aqueous (top) layer from the sample tubes into the pre-chilled tubes, avoiding the bottom layer/solids. 

10. Incubate overnight at -20ºC to precipitate. Discard phenol-containing tubes into phenol solid waste container. Move dirty tips to the hood.

(→ pause point: if you don’t need to run your samples soon, that is the best step to stop at).

Day 2: Finish Extraction, Denature, Run Gel, and Transfer

⇒ Finish extraction, Denature, and prepare for loading:

1. Spin down tubes in the cold room for 20min @ max speed. In the fume hood, dump liquid into ethanol-phenol-chloroform liquid waste container.

2. add 1mL 80% ethanol (-20ºC) to wash pellet, spin down tubes 5min @max speed

3. aspirate liquid, and dry out completely in the biosafety cabinet or fume hood (>1hr).

(→ pause point: ON or over weekend OK)

1. Add 25µL DEPC H2O to dry RNA pellet and incubate in thermomixer 15min @37℃, shaking at 1000rpm to help resuspension of RNA, then keep on ice.

2. Measure RNA concentration on the NanoDrop (Export data on USB key)

3. optional : dilute samples to 1ug/uL

(→ pause point: freeze at -80C OK)

1. Make denaturing mix:

per sample, 22µL total 48x Sample Mastermix:

15µL formamide 792µL

5.5µL formaldehyde 290µL

1.5µL 10x MOPS 79.2µL

1. Prepare/label enough 8-tube PCR strips, add 22µL denaturing mix per tube, chill on ice. 

2. Add 8µL of RNA sample at 1µg/µL in each tube:

• If samples were not diluted to 1µg/µ, determine the volumes of RNA and DEPC H2O to add to get 8µg in 8µL. 

1. Save remaining sample at -80ºC. 

2. Incubate 15min @ 55ºC in the thermalcycler (=PCR machine, pre-set cycle is saved). Keep PCR strips at 4ºC in thermal cycler until ready to load.

⇒ Run Gel, and Transfer

1. Prepare gel (1 gel per 24 samples) in B2 gel box fitted with two 12-well combs (thin side for better resolution, thick side for better sensitivity and/or if your sample concentration is low aka larger volume). 

1. For 1 gel, add in 250mL bottle For 2 gels, add in 500mL bottle

• 1.9 g agarose - 3.8 g agarose

• 110 mL H2O - 220 mL H2O

• 15 mL 10x MOPS - 30 mL 10x MOPS

Note: for thinner gels (runs faster, bands are sharper)

For 1 gel, add in 250mL bottle For 2 gels, add in 500mL bottle

• 1.52 g agarose - 3.04 g agarose

• 88 mL H2O - 176 mL H2O

• 12 mL 10x MOPS - 24 mL 10x MOPS

1. Heat in the microwave until boiling (boils quickly!); remove, mix well, repeat until dissolved.

2. In fume hood, add 25mL formaldehyde (50mL for 2 gels). Mix well and pour into gel box. 

Note for thinner gel: add 20 mL formaldehyde (40mL for 2 gels)

1. Once solidified, fill gel box with 700mL 1x MOPS.

1. Add 2µL loading dye to each sample, pipet mix and load 20µL (=5µg RNA) per well

2. run for 2h30mn @ 80V.

3. Set up transfer during electrophoresis. Material needed for each gel:

• an additional B2 gel box, with the gel platform turned upside-down on the ledge.

• ≥500mL 10x SSC (stock of 20x SSC should be available)

• 3x large tupperwares

• 1x 5.5” x 10” blotting paper wick (usually a stack is pre-cut)

• 6x 5.5” x 5” blotting paper (usually a stack is pre-cut)

• 1x 5.5” x 5” piece of GE Hybond N+ membrane, labeled with pencil on top corner.

• 4x strips of parafilm (3 ½ squares cut into 4)

• 1x stack of 16 SuperRag towels folded into thirds 

• 1x glass plate and 1x weight (eg. 0.5-1L stock solution bottle - better not more).

1. Rinse out the gel in DI water and soak it in 10x SSC (submerge it) for a few minutes. 

2. Meanwhile, assemble blotting stack: 

• Across the gel box platform, place the paper wick, wet with 10x SSC, roll across the paper with roller to flatten it out. Add 10x SSC on top.

• Repeat 3x: Add blotting paper on top of the wick, roll across to flatten out. Add 10x SSC on top.

• Remove any extraneous pieces of the gel sticking off and place on top of the blotters, wells facing down. Gently remove any air bubbles under the gel. Add 10x SSC on top.

• Dip the pre-labeled Hybond N+ membrane in DI H2O, then in 10x SSC buffer. Place on top of the gel. (NOTE: if the membrane isn’t wetted in DI first, it will not correctly take up 10x SSC and cannot be used)

• Wet and place 3x more blotters on top of the membrane and gently roll to flatten, being careful not to crush the gel or move the membrane.

• fill the sides of the box with 10x SSC.

• Place parafilm around the 4 sides of the blotter/gel stack to prevent anything at the top of the stack from touching anything at the bottom of the stack. 

• Place the folded stack of SuperRags on top of the stack, top with the glass plate and center the weight.

1. Leave stack overnight to transfer by upward capillarity.

(→ pause point: transfer stacks can stay as is of days or weeks without issue if need be)

 Day 3 - morning: Cross-Link, Stain, and pre-hybridization

1. Pre-heat water bath to 65ºC with 1x 50mL conical tube of Hybridization Buffer (Stored @ -20ºC) inside per membrane to hybridize. Pre-heat thermomixer to 100ºC.

2. Boil salmon sperm DNA 5min, then chill on ice ~1min. 

3. Add 300µL salmon sperm DNA to the Hybridization Buffer, keep buffer @ 65ºC.

4. Discard SuperRags and blotters from gel box. Discard gel into formaldehyde waste container. Move membrane to a glass plate and cover with plastic sheet protector.

5. Cross-link with UV in the Bio-Rad crosslinker (located in room 1620 or 12xx [Lorraine Symington gel room]). Power on, place membrane on glass trays inside, and run a cycle at 1200 (x 100 µJoules/cm2), also called “optimal crossling” on the machine. Remove membrane and power off.

6. Move membranes into trays and cover with Methylene Blue solution (~50mL). Place onto rocking table ~1min, then pour the solution back into the bottle. Rinse with MilliQ H2O (not DI) several times, avoiding to run the water directly on the membrane, then soak on the shaker in water for ~1min to destain.

NOTE: if methylene blue solution has become lighter over time, better to remake one fresh. For 600mL: add 180 mg methylene blue powder into a solution of 80 mL 3M NaAc + 520 mL milliQ H20.

1. Move stained membranes into sheet protector. Scan in black & white at 400dpi, in TIFF format (this is the rRNA loading control).

2. Place the membrane on a wire mesh sheet and roll up from bottom to top, and then place the left end first into a hybridization tube (if the membrane is rolled the wrong direction, it will roll itself into a tight coil on the rotisserie rack in the oven). Make sure that the cap is set with its rubber seal.

3. Add 20mL of Hybridization Buffer per tube, keep the remaining 20mL at 65ºC.

4. Place Hybridization tubes in hybridization oven (room 1517) with cap oriented toward the back of the oven. Incubate with max rotation at 65ºC in the for at least 4 hours-- can be left for several days if necessary, but less time is preferred.

 Day 3 - afternoon: Prepare radioactive probe and hybridization

A. Make PCR product (anytime ahead)

• Prepare ahead PCR reactions in 100uL with rTaq (for CLB3, this is 192·193::1)

• If on gel only one band is visible, purify PCR product with Macherey Nagel PCR Clean-up kit (use 50% NTI buffer) and elute in 2 steps with 25uL 65ºC water each time. Do not use elution buffer: contains EDTA and will interfere with downstream reactions).

• If several bands are visible on gel, do a gel purification. Same elution conditions apply.

• Measure DNA concentration and dilute an aliquot to 5ng/µL. Keep both diluted and undiluted product in freezer.

⇒ see note at the end for radioactive labeling related notes.

B. Label dsDNA probe with [α-32P]dCTP (on Day 3 step 11 or a few days earlier)

1. Switch on heat blocks at 100ºC and at 37ºC heat block.

⇒ 2 kits are available in the lab. Volumes are given for preparation of 1 probe. 

• GE Amersham MegaPrime Labeling Kit

2. Add 5µL purified dsDNA probe at 5ng/uL to a microtube. 

3. Add 5µL Primer Solution from the kit.

4. Boil for 5 min at 100°C (Place tube hat on tube to keep from popping open) to denature DNA, then chill on ice 1 minute.

5. Spin down the tubes briefly, and add to the tube on ice :

• 5µL Reaction Buffer

• 4µL dATP

• 4µL dTTP

• 4µL dGTP

• 16µL H2O

• 2µL enzyme (Klenow)

6. Move to the radiation station, transfer tube to plexi rack and turn on the Geiger counter (0.1x).

7. add 5µL radioactive [α-32P]dCTP (stored at 4℃ room 1517). 

8. Quickly pipette mix. Incubate in water bath @37℃ for 10 min.

9. Add 5µL 0.2M EDTA to stop the reaction.

10. Switch heat block to 100ºC for next step

• Invitrogen RadPrime DNA Labeling System Kit (Cat# 18428-011)

2. For each probe to prepare, add 5µL purified dsDNA probe at 5ng/uL to a microtube. 

3. Boil for 5 min at 100°C (Place tube hat on tube to keep from popping open) to denature DNA, then chill on ice 1 minute.

4. Spin down the tubes briefly, and add to the tube on ice :

• 1µL dATP

• 1µL dTTP

• 1µL dGTP

• 16µL H2O

• 20µL 2.5x RadPrime solution

5. Move to the radiation station, transfer tube to plexi rack and turn on the Geiger counter (0.1x).

6. add 5µL radioactive [α-32P]dCTP (stored at 4℃ room 1517). 

7. Add 1.5 µL Klenow 

8. Quickly pipette mix. Incubate in water bath @37℃ for 30 min.

9. Add 5µL Stop Buffer to stop the reaction

10. Switch heat block to 100ºC for next step

C. Purify labeled probe from unincorporated labeled nucleotides: 

⇒ During the last minutes of the 37ºC incubation step of probe labelling, prep one column per 50uL of probe to purify from the GE Illustra ProbeQuant Kit G50: 

1. Vortex the column to resuspend resin.

2. open the cap ¼ turn to break the seal, twist off the bottom, and place into a collection tube. 

3. Spin down 1min @ 735g. Keep collection tubes on the side, and move columns (which should now have a solid piece of resin inside) to new microtubes.

4. From the labeling reaction tube, slowly transfer up to 50µL to the top-center of the resin (avoid column wall and do not touch resin with pipette tip). Check gloves and pipette for radiation after transfer.

5. Spin down 2min @ 735g.

6. Transfer column back into saved collection tube, close the microtube containing the flow-through.

7. Check effectiveness of radioactive nucleotide incorporation:

1. Set the geiger counter on its side ~ 1 ft / 30 cm away on the right of the radioactive shield.

2. Keep tubes into plexi rack, keeping them on the leftmost part of the radioactive shield (should not be detected by geiger counter).

3. Move one tube at a time to cubic non-plexi tube holder on the rightmost part of the radioactive shield. Read cpm value measured by geiger counter. Compare microtube value and column value (best if ~5x higher or above).

Note: by experience, I have had blots working even if the flowthrough value was equal to or even 5x lower than the column…

1. If using later, the probe can be frozen and stored in a plexi rack @ -20ºC (room 1525).

D. Start hybridization: 

1. Place tube hats on microtubes and boil 3min @ 100ºC. While boiling, remove hybridization tubes with membranes from the oven, discard the hybridization buffer and replace with the remaining 20mL of fresh hybridization buffer kept at 65ºC in water bath.

2. Move microtubes to ice for 2min. Spin down quickly using the pulse button of the microcentrifuge at the radiation station.

3. Add 50µL (entire volume) of the probe to the hybridization tube (make sure to submerge the tip completely in buffer to transfer all volume; use a filter tip; waste must go into radioactive container) and return to the hybridization oven set at 65ºC on the rotisserie overnight.

4. Prepare Wash Buffers and have them warmed at 65ºC ON. It may be convenient to aliquot them into 50mL tubes.

Day 4: Washes + Expose to Phosphor Screen

1. Empty liquid from hybridization tubes into the liquid radioactive waste container, ensuring there are no drips.

2. Add ~30mL pre-warmed Low Stringency Wash to each tube and return to the oven/rotisserie for 5min. Check gloves and bench top for any radiation contamination.

3. Repeat with ~30mL Low Stringency Wash for 10min.

4. Repeat with ~30mL High Stringency Wash for 15min.

5. Repeat with ~30mL Low Stringency Wash for 5min, but empty it into a 50mL tube at the radiation station. Measure cpm value of the wash (should be residual, ≤200).

6. Place a piece of blotter paper on the bench large enough to fit all membranes. Remove the membranes from tube by pulling on wire mesh, not on membrane. Transfer membrane onto blotting paper and let dry for ≥10 minutes (humidity will damage the phosphor screen). You may check radiation level across the membrane with the Geiger counter.

7. Wrap the membrane: spread out a large piece of saran wrap on the bench, drop off the membrane face down onto the saran wrap, cover with a piece of blotting paper slightly larger than the membrane. Wrap up and move to a cassette. Tape down all four sides so the plastic is taut, return phosphor screen on top and close the cassette. Store it in a drawer while it is exposing. Exposition time is usually overnight, but can be as short as 1-2 hrs or as long as than several days depending on signal detected with geiger counter.

Day 5: Scan Phosphor Screen

1. Image on Typhoon (Room 1205), set value to 100µm and clear phosphor screen when done (Eraser, room 1205, long setting). Membrane should be discarded in radioactive solid waste. Phosphor screen should return back in cassette for protective storage.

2. Alternatively:

1. Image on Typhoon (Black Building, 5th floor, Room 536 Eric Greene Lab), select Phosphor, 250V,  100uM, choose file location (Desktop-> Vivek->Berchowitz folder *BRING FLASHDRIVE TO COPY FILES*), select dimensions of scanning area. Clear phosphor screen when done (eraser room HHSC 1205 long setting). Membrane should be discarded in radioactive solid waste. Phosphor screen should return back in the cassette for protective storage.

Radioactive labeling related notes:

⇒ Radioactive labeling can be done a few days ahead, taking into account “as of date” and planned date of hybridization (see 32P decay table), as well as the abundance of your transcript. If so, freeze probe after purification on column.

⇒ Radioactive labeling is commonly done using [α-32P]dCTP, but can also be done using [α-32P]dATP. Adapt the mix of nucleotides to the labelling reaction accordingly. It can also be done with [γ-32P]ATP for 5’end labelling of ssDNA oligos.

⇒ EHS requires us to track our radioactive stocks and wastes the day of use:

• in the yellow binder (“use record and waste log” tab)

• on the waste container labels. Determine the amount (in µCi) that you’ll use (and discard) for your experiment.

• On LATCH labcliq.com/l/columbia/

⇒ We usually order 250µCi. The ‘pig’ officially contains 25µL but in reality contains 30µL, and contains ≥250µCi (see as-of-date to determine actual activity on experiment day). 

⇒ We use 5µL per probe (~50µCi), thus we can make up to 6 probes per pig, which gives 42µCi per probe. Thus we estimate that half and half end up in the solid and liquid waste (21 µCi each). Fill amount in the yellow binder, onto the waste container label and online on LATCH labcliq.com/l/columbia/ .

⇒ EHS requires us to check that the Geiger counter works using the check source every single day that we work with radioactivity using the 137Cesium sealed source. The measure of check source activity must be recorded into the yellow binder, “survey meter” tab. 

⇒ Make sure that the radiation station is set up with the material you need: extra gloves, red and green filter tips (with P20 and P200 pipettes), 50mL tubes in rack to serve as radioactive solid OR liquid waste containers, plexi tube rack, spray bottle of count-off detergent, kimwipes and/or paper towels. When working, always keep a few 50mL tube open, available for disposal of tips, microtubes, columns, kimwipes... During incubation steps, close and dispose of 50mL waste-tubes to the radioactive waste container using a plexi box for transport.

⇒ Check the working area and the pipettes with the geiger counter before and after working. Frequently check your gloves, and change them as needed.

RNA Extraction WILD STRAINS VERSION

Day 1: Start Extraction: (4+hrs)

⇒ Start with tubes containing frozen pellets of SPO culture (up to 24x at a time) . 

1. Switch thermomixer to 65ºC.

2. Transfer tubes into two metal tube racks for 2mL tubes inside an ice bucket. 

3. In each tube, add 50µL (small scoop) 0.5mm zirconia/silica beads

4. Add 400µL TES. Vortex to resuspend.

5. In the fume hood, add 400µL Acid Phenol. Vortex to mix.

6. Incubate tubes on thermomixer at 65ºC, 1400rpm for 60 min.

7. Meanwhile, label a second set of screwcap tubes with 600µL chloroform store at -20℃.

8. Spin down tubes in the cold room for 15min @ max speed. 

9. In the cold room, transfer ~300µL of the aqueous (top) layer from the sample tubes into the pre-chilled tubes, avoiding the bottom layer/solids. 

from tubes phenol, cell pellet and leftover supernatant i.e. ‘THE CLOUD’: 

1. Add 300µL TES. Vortex to resuspend. (total aqueous = 800uL)

2. Incubate tubes on thermomixer at 65℃, 1400rpm for 60min.

3. Spin down tubes in the cold room for 15min @ max speed. 

4. In the cold room, transfer ~300µL of the aqueous (top) layer from the sample tubes into the existing pre-chilled tubes from step 9 (with 600ul chloroform and 300ul super from step 7), avoiding the bottom layer/solids. Vortex to mix. 

5. Discard ‘cloud’ tubes into appropriate hazardous waste container. 

6. Spin down in the cold room for 15min @ max speed. 

7. Meanwhile, label a second set of screwcap tubes with 500µL chloroform. 

8. In the cold room, transfer ~500µL of the aqueous (top) layer from the sample tubes into the chloroform tubes. Vortex thoroughly and spin down in the cold room for 15min @ max speed. 

9. Discard phenol tubes into appropriate hazardous waste container. 

10. Meanwhile, label a second set of screwcap tubes with 40µL 3M sodium acetate and 1mL 100% ethanol pre-chilled at -20℃, and store at -20℃. 

11. In the cold room, transfer ~400µL of the aqueous (top) layer from the sample tubes into the pre-chilled tubes, avoiding the bottom layer of chloroform. Vortex.

12. Incubate overnight at -20℃ to precipitate. Discard chloroform tubes into appropriate hazardous waste container.

(→ pause point: if you need to keep your samples for a long time that is the best step to stop at).

Day 2: Finish Extraction (1hr)

1. Spin down tubes in the cold room for 20min @ max speed. In the fume hood, dump liquid into ethanol-phenol-chloroform liquid waste container.

2. add 1mL 80% ethanol (-20ºC) to wash pellet and vortex

3. spin down tubes 15min @max speed

4. In the fume hood, dump liquid into ethanol-phenol-chloroform liquid waste container.

5. add 1mL 80% ethanol (-20ºC) to wash pellet and vortex

6. spin down tubes 15min @max speed

7. In the fume hood, dump liquid into ethanol-phenol-chloroform liquid waste container.

8. aspirate liquid, and dry out completely in the fume hood overnight.

Day 3: Quantify RNA

1. Add 25µL DEPC H2O to dry RNA pellet and incubate in thermomixer 15min @37℃, shaking at 1000rpm to help resuspension of RNA, then keep on ice.

Measure RNA concentration on the NanoDrop (Export data on USB key)

DNA analysis by FACS

1. Harvest cells during time course by aliquoting 1mL into a 1.5mL tube. Spin down 1min @ 3,000g, aspirate, and resuspend in 1mL 70% ethanol. Leave overnight to fix (up to two days is okay).

2. Centrifuge, aspirate, and resuspend in 800 µL 50mM sodium citrate pH 7. Repeat wash step.

3. Sonicate cells for 10 seconds at 2 amplitude.

4. Add 200µL sodium citrate pH 7 with 0.25mg/mL RNAse A and incubate 1hr at 50°C or overnight at 37°C 

5. Add 5 ul of proteinase K and digest 1 hour at 37°C

6. Centrifuge, aspirate, and resuspend in 500 µL 50mM sodium citrate pH 7 with 1:5000 dilution of SYTOX Green (For 15mL of sodium citrate add 1uL of 5mM stock SYTOX Green)

7. Transfer to 5 mL polystyrene Falcon tubes.

8. Let stain for 1 hour or overnight at 4°C, keeping protected from light.

9. Sonicate cells for 10 seconds at 2 amplitude.

10. Analyze tubes on BD LSR II using 488nm excitation and 525nm emission. Voltage should be around 250. Program is saved under Berchowitz account-- right-click on experiment and click “Duplicate Without Data”, then delete sample names and start fresh. Name the first tube the strain name (will be hour 0), save data by clicking “record”, then when done reading click “Next tube” and it will be named “[strain number]_001”.

11. When finished reading all tubes, with the Experiment highlighted select “File” and “Export”, “Export FCS Files” and save to the desktop. Once exported, move to the server on the desktop.

12. Analyze data as you choose but Flowjo is the easiest

Sonication (for which experiment??)

1. Start with pelleted sample in a 50mL conical tube; thaw pellet(s) on ice.

2. Per 50mL conical tube of pellet, add Tris Lysis Buffer to 30mL, 300µL 0.1M PMSF, 300µL Halt, and 5µL Turbo DNAse. Ethanol-sterilize a spatula and use to break up/mix the pellet. Try to avoid shaking the tube as it will create a lot of foam.

3. Prepare sonication assembly: Use a 4L beaker with an upside-down tri pour trimmed to fit in the bottom, pack with ice, and set a conical tube rack on top of the tri-pour. Ice should come up to the top of the tube but be careful it doesn’t get inside.

4. Take assembly, samples, ethanol, and paper towels to to sonicator (Equipment room with ice bin). Power on from the switch located at the back of the panel. A program can be set up (saved as Setting 3) with the desired cycles. Wipe off the needle before inserting into sample.

5. Sonicate at 50A for 12x cycles of 30 s ON, 60 s OFF. Keep tube is submerged completely in ice (sonication will heat up the sample significantly). Inside the sonicator, the beaker will need to be lifted with tube racks so the sonicator needle is completely submerged in liquid (if it is too high up, it will create a lot of foam while running). Repeat for all conical tubes.

Plasmid Transformation/Amplification/Prep in E. coli

Start with: Enough DH5α tubes for the desired number of reactions, ice, water bath or heat block heated to 42℃, plasmid DNA, LB + Amp plates, 2x LB liquid media, 1x LB liquid media, 1000x ampicillin, autoclaved disposable culture tubes w/foil tops, Plasmid Prep Kit

1. Per plasmid to be amplified, thaw 1 tube of DH5α ON ICE.

2. Add 1µL of plasmid DNA to the tube and flick gently to mix.

3. Incubate on ice 10min.

4. Heat shock at 42ºC for 30 s.

5. Add 200µL 2x LB to each tube. Incubate 1hr inside shaker incubator set to 37ºC and 180rpm.

6. Prepare dilutions: typically, 200µL 1:50 dilution is sufficient (prepare 4µL DH5α into 196µL LB). Inoculate a lawn with glass beads onto an LB + Amp plate and incubate overnight at 37ºC.

7. Prepare LB + Amp stock (50mL LB + 50µL 1000x ampicillin) and aliquot 5mL into an autoclaved disposable culture tube, 2x per plasmid. Pick 2x separated colonies from each plate and inoculate one per culture tube.

8. Incubate all tubes in 37ºC shaker overnight.

9. Aliquot 1.5mL to an Eppendorf tube and spin down 1min @ 6,000g. Aspirate solution.

10. Purify plasmid using appropriate kit.

11. Check concentration of prepared DNA on the NanoDrop and write the concentration on the side of the tube. Prepare a 1:25 working dilution for use in yeast transformations.

NOTES: For NovaBlue Competent Cells (Millipore PN 70181), thaw 1 tube of competent cells plus SOC medium. Reduce ice incubation time to 5min, add SOC medium instead of LB (in biosafety cabinet), and plate directly for Ampicillin resistance or incubate 30min @ 37ºC for Kanamycin resistance.

NucleoSpin Plasmid EasyPure Protocol - Macherey Nagel Kit

1. Start with pellet prepared from 1.5mL E. coli culture containing plasmid of interest.

2. Add 150µL Buffer A1 and vortex to resuspend cells (Stored at 4ºC).

3. Add 250µL Buffer A2 and invert 5x to mix (DO NOT VORTEX!)

4. Incubate 2min @ RT

5. Add 350µL Buffer A3 and invert until lysate turns colorless

6. Centrifuge 3min @ max speed

7. Load clear supernatant into a spin column/collection tube

8. Centrifuge 30s @ 2,000g and discard flow-through

9. Add 450µL Buffer AQ to spin column and centrifuge 2min @ max speed

10. Transfer spin column to a new microtube

11. Add 50µL Buffer AE warmed to 65ºC directly to the membrane and incubate 1min

12. Centrifuge 1min @ max speed

13. Measure concentration on NanoDrop and create at 1:25 dilution Working Stock. Store @ -20ºC.

Preparation of DH5ɑ Stocks

1. Inoculate 5mL LB with 5µL Invitrogen stock and incubate O/N @ 37ºC, 180rpm.

2. In a 2L flask, add 396mL LB + 4mL O/N culture. Grow @ 37ºC, 180rpm to OD600 = 0.5 (check at t=2hrs)

3. Prepare ice bucket, centrifuge @ 4ºC, and chill 8x 50mL conical tubes + 80x labeled microtubes, and buffer (Per 100mL, 10mL 0.6M CaCl2, 5mL 200mM PIPES, 15mL 100% glycerol; sterile-filter).

4. Chill 2L flask in ice water bath for 10min.

5. Aliquot culture to 8x 50mL conical tubes and spin down 8min @ 6,000g.

6. In cold room, dump effluent and place tubes on ice. Add 40mL buffer to the first tube and resuspend cells with a sterile stick (note: cells are very flocculated). Dump into the next tube and repeat, resuspending all cells.

7. Incubate on ice 10min.

8. Spin down 5min @ 2500g and dump effluent.

9. Gently resuspend in 8mL buffer.

10. Incubate 2hrs on ice.

11. Dispense 100µL aliquots to pre-chilled tubes and freeze @ -80ºC.

Gibson Assembly

Start with: Prepared PCR reactions of interest, gel-purified; cut insertion plasmid; Gibson Enzyme-Reagent mastermix (Enzymes & Buffers box, -20ºC; check expiration); NovaBlue competent cells kit (Millipore PN 70181); Heat block at 42ºC; LB + Amp plates

Day1: 

1. In a PCR tube, combine for a total volume of 5µL each PCR of interest and cut plasmid in equimolar ratios. Add 15µL Gibson Enzyme-Reagent mastermix and pipette mix. Spin down briefly and incubate at 50ºC for 1hr.

2. Meanwhile, on ice thaw one tube of NovaBlue competent cells and SOC medium.

3. Add 1µL of product from Step 1 and flick gently to mix.

4. Incubate on ice for 5min.

5. Heat shock at 42ºC for 30s.

6. Add 200µL SOC Medium. If plating to LB+Kan, incubate for 30min @37ºC; if plating to LB+Amp, skip incubation.

7. Prepare dilutions as necessary: 200µL 1:10 (20µL + 180µL SOC) and 1:50 (4µL + 196µL SOC).

8. Incubate as a lawn onto a plate using glass beads and incubate O/N @ 37ºC.

Day2: 

1. Prepare LB + Amp/Kan stock and incubate successful colonies in 3-5mL liquid @ 37ºC @180rpm O/N.

Day3 

1. Aliquot 1.5mL to an Eppendorf tube and spin down 1min @ 6,000g. Aspirate solution.

2. Purify plasmid using appropriate kit.

3. Check concentration of prepared DNA on the NanoDrop and write the concentration on the side of the tube. Prepare a 1:25 working dilution.

4. Check for success with appropriate PCR reactions, using 1:25 DNA as the template. Submit successful PCRs for sequencing.

One-step SLIC 

How it works: An exonuclease (T4 DNA polymerase w/o dNTPs) generates single-stranded DNA overhangs to linearized vector and inserts mixed together. Homology introduced by design allows annealing. The annealed complex (now circular containing single-strand nicks) is directly transformed in bacteria, which will produce recombinant DNA by gap-repair. By modulating the junctions, one can also delete or insert restriction sites or tag sequences.

Preliminarily:

1. Linearize vector by restriction enzyme digestion or PCR.

2. Prepare insert(s) by PCR with primers with 15-40 bp tail homologous to each end of linearized vector (15 bp homology tail between inserts).

3. Purify all fragments with commercial PCR purification kit. 

NOTE: do not elute in TE or H20, elute in Elution Buffer from kit or 10mM TrisCl pH8.0-8.5. Measure concentration of purified fragments.

Cloning per se (see table below for help)

1. Mix vector + insert(s) in molar ratios into ≤8.8uL (up to 5 inserts!).

2. Add 1uL NEB buffer 2.1, and add H20 to 9.8uL if needed.

3. add 0.6U of T4 DNA polymerase (0.2uL as stock is 3U/uL) 

4. incubate at RT for 2mn30s (generates 5’-overhangs) 

5. incubate on ice for 10 mn (stops exonuclease and allows single strand annealing).

NOTE: longer incubation on ice is fine, as well as storage at -20C at this stage.

1. Transform competent DH5α bacteria with 1uL of the mixture (proceed with dedicated protocol).

4’. 30s 50ºC

5’. 10 min ice

4’’. 1mn 37

5’’. 1mn 60ºC

Notes:

• Homology design: 

• It is possible to design the homologies of the insert primer tails not directly to the ends of the vector, but a few bp away, to remove a restriction site for example. Non-homologous stretches of at the ends of annealed sequences are trimmed away during the gap repair process.

• For very short insertions, one can design 60 nt oligos with 20 bp overlap and 40 nt 5’ overhangs, to add after the exonuclease treatment, before the annealing.

• Fragments purification: 

• Gel purification is preferable if multiple bands are visible on gel or if vector is linearized by digestion to exclude undigested vector.

• Treatment with DpnI enzyme or Gel purification is preferable if vector is linearized by PCR, to eliminate plasmid template.

• Determination of volumes for molar ratios:

• http://www.insilico.uni-duesseldorf.de/Lig_Input.html calculate for you the amount of insert based on vector/insert sizes and wanted molar ratios.

• Routinely 100 ng of vector is good, but as low as 5ng has been shown to work.

• You can also select several vector:insert ratios from 1:1 to 1:7, but people suggest 1:2, 1:2.5, 1:5 as the best ratios. If several inserts then vector:insert:insert ratios would be 1:1:1 to 1:7:7. It is suggested to try a couple of ratios in parallel to maximize success.

IP/iCLIP

Use samples collected as 5mL, UV-treated with 1500µJ, pelleted, and flash-frozen.

Solutions/Reagents Needed:

• Anti-v5 agarose affinity beads (Sigma PN A7345-1mL, 4ºC)

• iCLIP Lysis Buffer: 50mM Tris-HCl pH 7.4, 100mM NaCl, 1% NP-40, 0.1% SDS, and 0.5% sodium deoxycholate; sterile-filter and store at 4ºC

• Halt Protease Inhibitor (Thermo Scientific PN1861279)

• RNAse I, 1:500 dilution in Lysis Buffer (prepare fresh) (Invitrogen PN AM2294)

• Turbo DNAse (Invitrogen PN AM2238)

• PK Buffer: 100mM Tris-HCl pH 7.4, 50mM NaCl, 10mM EDTA

• Proteinase K (Roche PN 03115887001)

• PK Urea Buffer: PK Buffer + 7M Urea, prepare fresh (see below)

• High Salt Wash Buffer: 50mM Tris-HCl pH 7.4, 1M NaCl, 1mM EDTA, 1% NP-40, 0.1% SDS, and 0.5% sodium deoxycholate; sterile-filter and store at 4ºC

• Wash Buffer: 20mM Tris-HCl pH 7.4, 10mM MgCl2, 0.2% Tween-20; sterile-filter and store at 4ºC

• Acid phenol chloroform

• 3M Sodium Acetate, pH 5.5

• Glycoblue (Invitrogen PN AM9515)

• 100% Ethanol

Pre-prep the following tubes (per sample) and solutions:

• 3x Eppendorf tubes for:

• Beads (pre-chill @ 4ºC)

• Lysate (pre-chill @ -20ºC)

• RNA (pre-chill @ -20ºC)

• 2x 2mL screwcap tubes with caps, for:

• Lysis prep (add 200µL 0.5mm zirconia/silica beads and pre-chill @ -20ºC)

• RNA (add 400µL acid phenol chloroform before use, store in fume hood)

• iCLIP Lysis Buffer + Halt: 3mL for washes and 1mL x #samples + 1:50 Halt Protease Inhibitor; for 12x samples, 15mL Lysis Buffer + 300µL Halt (keep working solution on ice)

• Enzyme Mastermix: Prepare 1:500 dilution of RNAse I (499µL Lysis Buffer + 1µL RNAse I, -20ºC). Per sample use 10µL RNAse I dilution and 2µL Turbo DNAse; for 12x samples, 132µL RNAse I and 26.4µL Turbo DNAse.

• PKurea Buffer: For 12x samples, add 1.26g Urea to a 5mL tube + 2mL PK Buffer to reach 3mL; sterile-filter

Protocol:

Prep Beads

1. On ice, to a new Eppendorf tube add (20µL x #samples, 240µL for 12x samples) Anti-v5 agarose affinity beads using a pipette tip with the end cut off slightly so beads do not get stuck. Mark the volume level.

2. Wash beads: add 1mL Lysis Buffer + Halt, spin down 30s @ 2,000rpm, and aspirate with a fresh tip (err on the side of leaving liquid behind). Repeat for 3 times total.

3. Resuspend with Lysis Buffer + Halt to ~270µL.

4. Distribute 20µL to Eppendorf tube and keep at 4ºC until use.

Lyse Samples (all steps on ice/in cold room unless specified otherwise)

1. Remove 5mL tubes with UV samples from -80ºC and put on ice. Move 2mL screwcap tubes with zirconia/silica beads to a frozen block and put on ice. Set up 15mL conical tubes (1 per sample) cap removed with a 1mL pipette tip tip-down inside and put on ice.

2. Add 200µL Lysis Buffer + Halt to 5mL tube and vortex to mix. Transfer to screwcap tube.

3. Process in FastPrep for 45 s at 6.5M/s.

4. Use an 18G needle and heat tip on a Bunsen burner until red hot to poke a hole in the bottom of the screwcap tube to drain; do NOT poke the hole directly in the center, but off to one side. Place screwcap tube into conical tube.

5. Spin down conical tubes at max speed 30s-1min to drain lysate. Discard screwcaps with beads.

6. Use the pipette tip in the conical tube to transfer the lysate to an Eppendorf tube (pre-chilled @ -20ºC), avoiding transfer of beads as possible.

7. Add 800µL Lysis Buffer + Halt to each tube to bring total volume to ~1mL.

Deplete RNA

1. Add 12µL Enzyme Mastermix to each tube (1:500 RNAse I + Turbo DNAse) using filter tips.

2. Incubate 3min @ 37ºC 1100rpm.

3. Spin down 5min @ 5,000g.

4. Transfer lysate (avoid pellet) to prepared anti-v5 bead tube.

5. Incubate on rotisserie rack in cold room for 2hours.

Proteinase Prep

1. Spin down sample tubes 30s @ 2,000rpm.

2. Aspirate with a new tip.

3. Add 1mL High Salt Wash, spin down, and aspirate. Repeat High Salt Wash.

4. Add 1mL Wash Buffer, spin down, and aspirate. Repeat Low Salt Wash

5. Add 200µL PK Buffer and 10µL Proteinase K, and incubate 20min @ 37ºC 1100rpm.

6. Add 200µL PK Urea Buffer, and incubate 20min @ 37ºC 1100rpm.

RNA Extraction

1. Set ThermoMixer to 65ºC and move 3M NaAc and 100% EtOH to -20ºC. Put GlycoBlue on ice to thaw. Add 400µL acid phenol chloroform to second set of screwcap tubes.

2. Spin down samples 30s @ 2,000rpm.

3. Add 400µL lysate from sample tubes to screwcap tubes with acid phenol in the fume hood. Vortex.

4. Incubate 30min @ 65ºC 1400rpm.

5. Spin down 5min @ max speed in the cold room.

6. Transfer 300µL of the aqueous (top) layer to the third set of chilled Eppendorf tubes.

7. Add 40µL chilled NaAc.

8. Add 1µL Glycoblue, mix.

9. Add 1mL chilled 100% ethanol.

10. Incubate overnight at -20ºC.

11. Spin down 20min @ max speed in the cold room.

12. Dump liquid and wash with 1mL 80% ethanol.

13. Spin down 5min @ maximum speed in the cold room.

14. Aspirate and let dry.

15. Resuspend in 5µL DEPC-treated H2O and incubate 15min @ 37ºC 1000rpm to resuspend.

NEB Depleted RNA Protocol

Prep: 4x sets of PCR tubes, 3x sets of Eppendorf tubes, 6 Eppendorf tubes for mastermixes (Frag & Prime, First Strand Synth, Second Strand Synth, End Prep Rxn, Ligation Rxn, and PCR Enrichment). All PCR tubes and mastermix tubes should be on ice.

All thermomixer programs are set up on LifeTech Thermalcycler under “NEB Depletion Protocols” - Queue up programs ahead of time, as many of the lower-temperature protocols require a long “pre-heating” to cool down the cover.

Go through Manufacturer’s protocol.

IP/MS (incomplete)

For each sample, you will need the equivalent of 8 Western sample (aka 4mL SPO culture at OD 1.8 spun down and resuspended in 1.8mL 5% TCA, processed as per Western blot protocol, aka broken cell pellet in 100uL breakage buffer+50uL 3x SDS loading dye+BME boiled). 4 will be pooled and used with primary Ab, 4 will be pooled and used as a control without primary Ab.

All in the cold room:

1. For each 2mL tube, add NP-40 buffer up to 1.5mL

→ Note: could be also NP-40+BSA but not always suitable

1. Spin down 10 mn max speed

2. Combine all 4 tubes in 1 15mL conical tube for each set - be careful to not transfer any glass bead to the conical tube

3. Add primary Ab to 1/50 or 1/100 to the +Ab tube, add same amount of Ab buffer to the -Ab tube (here: 1/100=60uL, Ab buffer=100mM glycine pH 2.5)

4. Let sit 1h (with rotation?)

5. Add 20uL protein A slurry using cut tip

6. Incubate with rotation for 2h

7. Spin down on grey centrifuge at setting 2 for 2mn.

8. Remove supernatant (be careful to not aspirate beads - leave 100uL for safety)

9. Add 1mL NP-40 buffer

10. Using cut tip, transfer to 1.5mL tube

11. Wash 3 times:

1. Spin 30 sec 2000 rpm / 400g

2. Aspirate carefully with vac (leave 100uL)

3. Add 1mL NP-40 buffer

12. Aspirate with pipette, not vac, to remove all liquid, up to the level of the beads - not inside the pellet of beads.

At RT:

1. Add 20uL {breakage buffer + 3x SDS loading dye w/ BME}

2. Boil 5mn

3. Load onto house made acrylamide gel

4. Stain gel in Imperial blue

5. Cut band of interest

6. Send to MS.

Plate Screen Experimental Overview

On day 1, pin cells from collection onto YPG overnight (onto 96-well plate). Separately, plate control strains (37725, 37728, 37731, and 37734).

On day 2, pin to YPD 4% overnight. Move control strains on four points of no growth (double check screen spreadsheet to confirm) on same YPD 4% plate and be sure to note where they are located.

On day 3, inoculate by pinning into 1.5mL YPD liquid culture in a 96-deep-well plate, leaving on warm room shaker overnight. 

On day 4, prepare 5mL BYTA in 50mL Erlenmeyer flasks. Measure OD of a few representative wells (dilute 1:3 first), calculate volume needed to reach OD600 = 0.3 in BYTA (typically ~200µL), inoculate, and grow overnight.

On day 5 (experiment day): 

Prepare cultures: 

Measure OD of a few representative vials and calculate volume needed to reach OD600 = 1.8 in SPO. Pipette out this volume from each flask into a 15mL conical tube and spin down (1min @ 3,000g). Dump supernatant from the conical into the sink, and wash (vortex to resuspend) in 2.5mL H2O. Repeat spindown and dump, then resuspend into 5mL SPO by vortexing. Pour liquid from conical tube into a fresh 50mL Erlenmeyer flask and move to shaker incubator (30℃).

Initiate Meiosis: 

At t=6hrs after starting flasks in SPO, add 5µL 1mM stock of β-estradiol (Stored at -20ºC), making sure the drop is incorporated into the liquid.

Take samples: 

At t=8hrs after starting flasks/2hrs after adding β-estradiol, take samples by removing 450µL culture to a prepared tube with 50µL formaldehyde. Store samples at 4ºC. 

Plate Screen Microscopy Sample Prep

1. Add 450µL cells from SPO culture to 1.5mL Eppendorf tube containing 50µL 37% formaldehyde (total volume 500µL). Fix at 4℃ O/N.

2. Spin down 30s @ 10,000g.

3. Aspirate liquid, being careful to avoid the pellet.

4. Resuspend in 100µL KPO4/sorbitol + 1% Triton-X and vortex. 

5. Incubate 5min @ RT.

6. Spin down and aspirate.

7. Make a fresh stock of KPO4/sorbitol + DAPI: DAPI stock is @ -20ºC and 100x. Dilute to 1x in KPO4/sorbitol to a final volume of (#samples * 100µL * 1.1).

8. Add 100µL KPO4/sorbitol + DAPI to each tube and incubate 5min @ RT.

9. Spin down and aspirate.

10. Add 20µL KPO4/sorbitol and vortex to resuspend.

11. Store @ 4ºC for imaging up to 1 month, or freeze @ -20ºC for long-term storage.

DAPI stock: 0.1 mg/ml in methanol

KPO4/sorbitol stock (100mL): 

60mL 2M Sorbitol

10mL 1M Potassium phosphate [pH 7.5]

30mL H20

Protein Purification in E. coli

Prepare E. coli using LOBSTR:

1. Thaw 1 tube of LOBSTR ON ICE.

2. Add 1µL of p105 plasmid DNA to the tube and flick gently to mix. (p105 is RIM4-3C-HIS, p60 was amplified to include 3C cut site sequences using inverse PCR. Plasmid recovered by PNK, blunt end ligation and transformation back into E. coli)

3. Incubate on ice 10min.

4. Heat shock at 42ºC for 30 s.

5. Add 200µL 2x LB to each tube. Incubate 1hr inside shaker incubator set to 37ºC and 180rpm.

6. Prepare dilutions: here, 200µL of 1:2 dilution was sufficient. Inoculate a lawn with glass beads onto an LB + Kan plate and incubate overnight at 37ºC.

7. Inoculate a single colony into 25mL LB + Chloramphenicol + Kanamycin and incubate in 37ºC shaker overnight. Save LB + Kan plate @4℃ and use a fresh colony to grow up an overnight culture for every set of E. coli flasks to be grown.

E. coli Growth and Induction:

1. Prepare 2x 2L baffled flasks with 500mL LB, 5mL 40% glucose, and 500µL each chloramphenicol and kanamycin. Inoculate with 2mL liquid from the overnight culture.

2. Grow @ 37ºC to OD600 = 0.6-0.8 (~3hrs). Meanwhile, set up water bath shaker in the cold room set to 18ºC.

3. Take Uninduced sample (1mL culture in a screwcap tube, add 50µL TCA to reach a final concentration of 5%).

4. Transfer the flasks to the cold room water bath shaker and let incubate 1hour to stabilize. (Note: E. coli + Rim4-14HIS, without 3C, can successfully grow for three hours at 37°C instead of overnight. Skip step 4 if using this.)

5. Induce by adding IPTG to a final concentration of 1mM (500µL of 1M stock), and let grow overnight.

6. Pre-chill the centrifuge to 4ºC. Take Induced sample (as before).

7. Pour the E. coli culture into 3x 500mL centrifuge containers and spin down in chilled centrifuge 5min @ 6,000rpm.

8. Dump supernatant. Resuspend in 20mL Tris Lysis Buffer per centrifuge container (pellet will NOT dissolve completely) and transfer all into to one 50mL conical tube. Spin down, dump supernatant, and freeze pellets at -80ºC.

9. Repeat the process if more protein is desired, starting with a fresh E. coli culture each time, for a total of 10L of spun down culture (On subsequent days, use new centrifuge containers and conical tube). Proceed with sample preparation.

Sample Preparation for FPLC on Emulsiflex:

1. Thaw pellets on ice.

2. Resuspend pellet in 20mL Tris Lysis Buffer + Urea.

3. Take resuspended pellets on ice (bring extra conical tube(s) as needed to collect 

4. contact: Meagan)

5. Prepare Emulsiflex for running: Open main valve and outlet valve on air tank and ensure the line is pressurized. The metal funnel apparatus should have the lid on. On the outlet line, move the metal coil completely underneath the ice, and the end of the tubing to a tripour as waste. Attach the outbound end of the gas line to the top of the funnel for a few seconds to blow air through and clean out the Emulsiflex. Remove the gas tube and lid from the funnel apparatus. Turn the “Air Regulator” knob up until the small pressure gauge reaches ~110psi (completely opened up).

6. Pre-rinse the funnel apparatus: Add DI H2O to the funnel up to the line, making sure the sides are rinsed. Turn the red “Stop” knob, press the green “Start” button, and watch water level in the funnel. It will start flowing through the outlet—if there is any movement difficulty, use the poker tool to clear any blockages. Press the red “Stop” knob when water is almost not visible.

Note: Do NOT let the funnel get completely dry, it can break the machine!

1.  Pour all resuspended E. coli into the funnel (vortex again just before adding). Press start again and watch the outflow. As soon as it changes color to indicate E. coli instead of water and the machine begins to make a repeated popping noise accompanied by a pulse in the large front pressure gauge, move the outlet line to the extra conical tube for collection. Keep an eye on the funnel volume level and stop running before it empties completely by pressing the “Stop” button.

2. Dump the lysate back into the funnel to run through again. Repeat for a total of three run-throughs.

3. Return outflow line to tripour/waste, and rinse funnel with water, ensuring it is rinsed well to the level of E. coli used. Watch level and once almost completely empty, repeat with ethanol.

4. Return lysate back to the lab and take Total Lysate sample (50µL into 2.5µL TCA).

5. Pre-chill centrifuge to 4℃. Transfer sonicated culture to high g centrifuge tubes, balance by weight, and spin down 20m @ 25,000g (14,500rpm). Take Clarified Lysate (50µL into 2.5µL TCA) and Inclusion Bodies (toothpick of solids into 2mL 5% TCA) samples. Transfer lysate to a fresh tube, avoiding pellet.

Running FPLC with SuperLoop and HisTrap Column:

1. Set up FPLC to run by making sure lines A & B have Tris Lysis Buffer + Urea and Water to start. Turn on machine, load HisTrap HP method unchecking anything to save the run info, and start flowing 100% water through and select “column down flow” on the software, and hook up HisTrap HP column(s) to the red lines. Use “drop to drop” before attaching columns by allowing a few drops to pool into the inlet before screwing in the ferrule. Confirm water is coming out the bottom before attaching tubing. Check for any leaks or pressure issues, and use syringe to remove any air bubbles from the bottom four outlet channels.

2. Load the SuperLoop and connect to the FPLC: Start by making sure the inside stopper is at the 50mL measurement end of the SuperLoop inner glass column, using a serological pipette to move it. Attach the end stopper to the top (50mL marker) of the SuperLoop, and attach the longer green tube + ferrules to the top. Pour lysate in; if volume is <50mL, add Lysis Buffer to reach just over 50mL (zero marker). Place bottom stopper in—careful! To make sure there are no air bubbles, the column should be just slightly over-filled so a small amount of liquid pushes out from the tubing when the bottom stopper is placed. The shorter green tubing + ferrules should be attached to the bottom (0mL end). To attach the SuperLoop to the FPLC, the top line is screwed into the port labeled “LoopE” and the bottom line into the port “LoopF”.

3. Run lysate through the SuperLoop on the FPLC onto a HisTrap HP column; if there is >50mL lysate, run “His trap HP superloop load only” program first with the option checked not to save the file, re-load fresh lysate into superloop, and then run “His trap FF” program; if <50mL lysate, just run “His trap HP” program. While lysate is moving through the SuperLoop, the lines must remain with Lysis Buffer (A) and Water (B), but after the SuperLoop is empty, line A must be switched to Wash Buffer and line B switched to Elution Buffer for the remainder of the program. Load empty 2mL tubes for elutent to empty into in the carousel and line up the white flag with tube position.

4. As the program runs over the next few hours, the UV readings on the computer screen will show protein output. Where there is a spike, the highest amount of protein is to be expected.

5. Measure concentration of the two tubes with the highest protein on NanoDrop using the Protein A280 program and 2µL elution buffer as the blank.

Check Success of HisTrap Elution with a Gel:

1. Prepare Uninduced, Induced, Total Lysate, Clarified Lysate, and Inclusion Bodies samples with standard Western sample prep.

2. Prepare the first 9 FPLC elution samples in a PCR tube with 10µL elution and 5µL 3x SDS Loading Buffer. Heat on thermalcycler 5min at 100C.

3. Load samples onto a Western gel (15-well mini size, 4-15%), 6µL for Uninduced, 3µL for Induced, Total Lysate, Clarified Lysate, and Inclusion Bodies, and 15µL (all) for FPLC elution samples.

4. Run gel, stain 30min-1hr in Imperial Blue, and then rinse/soak in DI water until bands are visible.

5. Scan gel for record-keeping.

Removal of 3C Tag: (Use step 1 only if a plasmid without 3C is initially used)

1.  Combine all FPLC elutent into a single tube.

2. From NanoDrop measurements, calculate the approximate total amount of protein in all elution tubes.

3. Calculate the appropriate amount of 3C protease to add: 1U 3C protease cleaves 100µg protein, and stock is 2U/µL. For calculating volume, double the approximate total protein as a margin of error. i.e. if there is ~2,000µg protein, calculate enough for 4,000µg protein, or 40U which is 20µL of 3C protease.

4. Create controls in PCR tubes: For positive control, 10µL cleavage control protein, 1µL 3C protease, and 1.2µL buffer. For negative control, 10µL cleavage control protein + 1.2µL buffer.

5.  Incubate overnight at 4°C.

Condensing Elutent in Millipore Filter Tubes:

1. Prep a filter column Millipore tube: Important numbers are “Amicon Ultra – 4” where 4 is the mL capacity of the tube, and “Ultracel – 50K” where 50K is the kiloDalton size of the protein that the filter catches. (REF UFC805008)

2. Prechill the centrifuge to 4°C and get an ice bucket.

3. Pour 4mL FPLC elution buffer into the Millipore tube and centrifuge 1min @ 3,000rpm to prime/hydrate the filter. Discard liquid from top and bottom compartments.

4. Load up to 4mL of the combined FPLC fraction elutent to the top compartment.

5. Spin down 15min @ 3,000rpm.

6. Dump flowthrough into a fresh conical tube.

7. If there is space, add more elutent to the top compartment. Mix tube twice by inversion.

8. Repeat spindown, pouring off flowthrough, adding more elutent if there is space, and mixing after each spin down. Repeat until all of the elutent has been transferred, and then continue spinning down and mixing between spins until the volume left in the top compartment is 250µL.

9. Remaining 250µL is ready for injection into the FPLC to run through Superose6 column.

Clarifying Lysate on the FPLC Using Superose6 Column:

1. Open the method for the Superose6 column. Check options to not save data for initial setup.

2. Attach the Superose6 column to the FPLC. Run water through, the switch to 50mM KPi buffer.

3. Attach sample loop where the SuperLoop was (coil of green tubing).

4. Using a 1mL syringe fitted with 22g blunt-end needle, inject 1mL KPi buffer into the inlet, making sure no air bubbles are introduced. Repeat for 3x total and leave the needle in place to prevent leaking.

5. Set up 2mL tubes for catching elution.

6. Press play to start the FPLC program for the Superose6 column, and name file to save results.

7. Draw up sample (should be ~1/100th of column volume, 250µL) being sure to avoid air bubbles. Push out any air from the syringe. Inject and leave the needle in place.

8. Switch inlet B to 20% ethanol for final column rinse.

Processing of Elutent from FPLC:

1.  Cap elutent tubes to prevent evaporation.

2. Prepare samples for a gel with 10µL from each fraction tube and 5µL 3x SDS Loading Buffer + BME into 8-strip PCR tubes. Heat in thermalcycler 5min @ 100°C. Spin down briefly.

3. Load entire volume (15µL) onto a Mini TGX gel and run @200V ~40min or until sample runs to the bottom.

4. Remove gel from casing into a tray and cover with Imperial Blue protein stain.

5. Put on the shaker table @ RT for 1hr.

6. Rinse several time in DI water, then leave to soak in DI water ~30min or until individual bands are visible.

7. Confirm band of interest is present in final samples.

8. Condense fraction tubes that contain the protein of interest and submit.

Solutions Used:

Tris Lysis Buffer is from McKnight Protocol, final concentrations are 50mM Tris-HCl pH 7.5, 500mM NaCl, 1% Triton X, 20mM BME, and 40mM Imidazole. If using Urea, final concentration is 2M.

Wash Buffer final concentrations are 20mM Tris-HCl pH 7.5, 500mM NaCl, 20mM BME, and 40mM Imidazole.

Elution Buffer final concentrations are 20mM Tris-HCl pH 7.5, 500mM NaCl, 20mM BME, and 300mM Imidazole.

KPi Buffer final concentrations are 50mM potassium phosphate pH 7.2, 150mM NaCl.

All solutions for FPLC must be filter-sterilized.FPLC – Rim4 Native Purification

Start with:

Fitz Buffer without EDTA

BME (ß-Mercaptoethanol)

Halt Protease Inhibitor

20% Sarcosal

BioSpec 0.5mm zirconia/silica beads

10mL pellet of desired yeast sample (such as 48), flash frozen

Tubes (per sample): 3x 2mL tubes w/ 2mL 5% TCA; 4x empty Eppendorf tubes; 1x empty 2mL tube; 2x 15mL conical tube with 1mL pipette tip (no filter) inside, tip down

Bunsen burner

18G needle

Prepared polyacrylamide gel, 10%, w/ 15-well 1.5mm comb

Protocol:

1. Start FPLC program before sample prep, allowing ~45mins to equilibriate (if not finished with sample prep at this time, pause program). Program is “Superose6 GF”; save data to “Rim4 Native Purification” folder. During downtime of sample prep, wash the loop twice with 1mL Fitz Buffer (no BME, Halt), then 1x with remaining Fitz Buffer + BME + Halt. Make sure the fraction collector is prepared with 2mL tubes.

2. Prepare Fitz Buffer: Per 1mL Fitz Buffer, add 20µL Halt and 3.5 µL BME. To the 10mL pellet tube on ice, add 200µL buffer and 200µL zirconia/silica beads (one PCR tube filled).

3. Process sample(s) in the FastPrep, 45 seconds at maximum power (6.5M/s).

4. Turn on Bunsen burner. Flame-heat the needle until red-hot, and use to poke a hole in the bottom of the sample tube. NOTE: Hole should be on the outside edge of the bottom; if it is directly in the center, it will clog during spin down. Put the 2mL tube into the 15mL conical, on top of the pipette tip. Spin down tube at max speed in the cold room, for ~30s, or until liquid has moved from the 2mL tube into the 15mL conical. Discard the 2mL tube + beads. Use the 1mL pipette tip into the conical tube to transfer the supernatant (avoid beads) into a new Eppendorf tube.

5. From the supernatant, take the first sample, “Crude”, as 5µL into one of the tubes with 2mL TCA.

6. Spin down the remaining supernatant 1m @ 3,000g in the cold room. Transfer the supernatant to a new 2mL tube.

7. Add 20µL 20% sarcosal and mix. Spin down 20m @ 20,000g in the cold room.

8. Remove supernatant to a new Eppendorf tube. From this, take the second sample, “Supernatant”, as 5 µL sample into the second tube of 2mL TCA. Remainder of supernatant can be discarded.

9. To the 2mL tube with the pellet, add 190µL Fitz Buffer + BME + Halt and 10µL 20% sarcosal. Vortex to mix.

10.  To the 2mL tube, add 200µL zirconium/silica pellets, and vortex until the pellet is completely resuspended.

11.  Repeat step 4.

12.  Spin down the Eppendorf tube with supernatant 2min @ 3,000g in the cold room.

13.  Transfer the supernatant to a new Eppendorf tube. From this, take s sample labeled “Pellet”, as 5µL sample into the third 2mL TCA tube.

14.  Inject the remaining ~190µL supernatant into the FPLC, which should now be equilibrated and ready to fractionate the sample.

15.  While FPLC is running, prep the 3x samples as per standard Western Prep protocol.

16.  As soon as the fractions are completed, prepare them by selecting the sample tube that corresponds to the right output peak (usually tube #3), and as many after that as will fit on the gel (15 wells – 1 ladder – 3 samples taken during prep = 11 fractions). In PCR tubes, add 20µL fraction + 10µL 3x SDS buffer + BME. Boil 5min @ 100°C in the thermalcycler.

17.  Meanwhile, in the remaining volume of the fraction tubes, add 75µL 100% TCA, invert mix, and move to 4°C O/N. If the gel is successful, these will be used to submit for mass spec analysis.

18.  On the gel, load ladder (4µL), 3x samples taken during prep (vol?µL), and then the 11 prepared fractions (entire 30µL volume).

Ime2 Protein Purification

Part 1: Growing and Collecting Yeast

Day 1: Pull LEB collection strain 114 to a YPG plate and grow O/N

Day 2: Inoculate into 25mL SC –Leu medium (be sure glucose is added)

Day 3:

• First thing in the morning, in a 2L flask, add 75mL fresh SC –Leu medium and dump entire overnight culture of 114 in. Grow throughout the day.

• At 16:00, inoculate the cells into YEP + 2% glycerol + adenine, using 8x baffled flasks with 500mL medium each. They should be grown such that, with ~3.5hr doubling time, the OD is ~1.2 the following morning. Note: In first trial, this was calculated as ~5 doubling times (between 16:00 – 9:00), so started at OD = 0.0375. The following morning the culture was only at OD ~ 0.60, so starting with OD = 0.075 for second trial.

Day 4:

• First thing in the morning, measure OD of cultures; they should be ~1.2. Add galactose to 2% to induce Ime2 expression (50mL per flask of 20% galactose stock), and let grow another 4-6 hours.

• Pre-chill centrifuge with 500mL collection flask rotor to 4°C, plus centrifuge with 50mL conical tube rotor to 4°C. To harvest cells, fill 6x collection flasks and spin down 5min @ 3,000g. Dump liquid and repeat with remaining culture until the entire volume has been pelleted. Add water to wash, and combine all cells down to 2 flasks (it should resuspend fairly easily). Repeat spin down and pour off liquid.

• Meanwhile, prepare 50mL Lysis Buffer (ATP stock must be added fresh) and use ~35mL Lysis Buffer to completely resuspend yeast cells. Divide evenly into 2x 50mL conical tubes. Use ~10mL remaining Lysis Buffer to completely rinse out centrifuge containers (make sure 5mL Lysis Buffer remains).

• Repeat spin down and dump liquid. To the remaining 5mL Lysis Buffer, add protease/phosphatase inhibitors, and then add 2.5mL to each conical tube and resuspend the cells into a slurry.

• Prepare a Styrofoam box with a cold-safe Tupperware container and add liquid nitrogen. Use a 1mL pipette to dropper the yeast slurry into the liquid nitrogen to make “Dippin’ dots”. Once completed, move the Tupperware to -80°C. Once the liquid nitrogen has evaporated, move pellets to a pre-labeled 50mL conical tube.

Cleaning and Sterilizing Square/RoToR Plates

1. Scoop out the agar using one of the glassware cleaning brushes in the tissue culture room, being careful to minimize scratching the plates.

2. Soak the empty plates and lids in a large tub of water and bleach (using the dirty glassware tub is fine) overnight.

3. Wash the plates and lids either in the Rothstein Lab Fisher brand washing machine in room 1611. Add a small (approximately 3 grams) amount of Contrex dish washing soap in the washing machine and run one wash cycle. Alternatively, hand-wash with dish soap. Note: Do NOT use the glassware washing machine on the 15th floor, the temperature is too hot and it will melt the plates.

4. Rinse the washed plates and lids in DI water and allow to air dry.

5. Once fully dry, close plates and move to the biosafety cabinet. Expose to UV source for one hour.

6. After sterilization, use cling wrap to enclose the plates in stacks of 8. Store in the tissue culture room with other plates.

Yeast Transformation Using Sorbitol Plates

Solutions:

• SED: 1M sorbitol, 25mM EDTA pH 8, 50mM DTT (5mL per strain being transformed into) – prepare fresh before using

• SCE: 1M sorbitol, 0.1M sodium citrate pH 5.8, 10mM EDTA (5mL per strain being transformed into)

• 1M Sorbitol (10mL per strain being transformed into)

• CaS: 1M sorbitol, 10mM calcium chloride, 10mM Tris pH 7.5 (store @ 4ºC) (11mL per strain being transformed into)

• PEG Solution: 20% PEG-4000, 10mM calcium chloride, 10mM Tris pH 7.5 (1mL per transformation reaction)

• SOS: 10mL 2M sorbitol, 6.7mL YPD, 0.13mL calcium chloride, 3.17mL water (150µL per transformation reaction

• Sorbitol Plates: Per 500mL (1 sleeve of plates), 100g sorbitol, 4g Difco Yeast nitrogen base, 2g -Trp -Ura powder, 11g agar, autoclave; post-autoclave add 25mL 40% glucose and 5mL 100x tryptophan. Pour into plates ahead of time and store at 4ºC.

• Top Agar: Per 50mL, 10g sorbitol, 0.4g Yeast nitrogen base, 0.2g -trp -ura powder, 0.5g agar, autoclave; post-autoclave add 2.5mL 40% glucose and 0.5mL 100x tryptophan. Keep heated at 65ºC until use. (6mL per transformation reaction)

Additional Materials and Reagents:

• Autoclaved culture tubes

• Heat block at 45ºC to fit culture tubes

• YPD Medium

• Glusulase (50µL per transformation) (Perkin Elmer PN NEE154001EA)

• ssssDNA - Prepare 10µL of 1:10 dilution (1mg/mL) (5µL per transformation reaction)

• Plasmid to be transformed (2µL per transformation reaction)

Protocol:

1. Inoculate 25mL YPD with yeast strain to be transformed into – either direct from glycerol stock, grown up on a YPG plate, or 50µL from liquid culture – and grow O/N @ 30ºC.

2. Inoculate 20mL YPD with 1mL O/N yeast (OD600 ~9-12) and incubate 4-5 hours, until OD600 = 3.0.

3. Meanwhile, prepare Top Agar and keep at 65ºC. Pre-heat a block with one culture tube per transformation reaction to 45ºC. Pre-heat a sorbitol plate per transformation to 30ºC. Pre-label tubes and plates for transformations.

4. Transfer entire volume of liquid YPD to a 50mL conical tube and spin down 3min @ 1500rpm. Dump liquid out into sink.

5. Resuspend in 5mL SED and incubate 10min @ 30ºC.

6. Repeat spin down and aspirate.

7. Resuspend in 5mL SCE. Add 50µL Glusulase and incubate at 30ºC.

8. Every 5-15min, check the condition of the cells under the microscope for spheroplasting: add 3µL to a slide plus 1µL 1% SDS on a slide and look for “burst” cells. Continue incubating until ~85% of the cells have spheroplasted.

**11/29/17 this took ~40min, and 6µL 10mg/mL Zymolyase was added after 15min.

1. Resuspend the pellet in 10mL 1M sorbitol gently and pipette mix-- DO NOT VORTEX.

2. Repeat centrifuge and aspiration.

3. Wash the pellet in 10mL CaS, repeat spin down and aspiration, and resuspend in 0.5mL CaS (DO NOT VORTEX). Move tubes to ice.

4. On ice in an Eppendorf tube, add 5µL diluted ssssDNA (1mg/mL) and 2µL of plasmid DNA to be transformed into the yeast.

5. Add 100µL yeast + CaS to each transformation reaction and pipette mix.

6. Incubate 10min @ RT.

7. Add 1mL PEG solution and pipette mix.

8. Incubate 10min @ RT.

9. Centrifuge 3min @ 1200rpm (do not use a higher rpm than this!) and remove liquid with a pipette.

10. Resuspend the pellet in 150µL SOS solution.

11. Transfer 6mL of Top Agar to a pre-heated culture tube @ 45ºC and let incubate 5min.

12. Transfer the entire yeast resuspension in SOS to a tube of Top Agar. Pour onto the top of a sorbitol plate and rotate to spread evenly across the top. Once dry, incubate @ 30ºC until colonies grow.

Image analysis

Transfer of PRJ files by command line

Aim: transfer all (and only) the PRJ files for a specific set of experiments for analysis on your computer.

1. Open a text editor on the microscope computer (for example gedit).

2. Open the template called “command_line_to_transfer_PRJ_files.txt” saved on the DeltaVision computer’s desktop. 

Note#1: This template is designed to transfer the data onto the black hard drive called “BERCHOWITZ” ; change name (highlighted) if using another device.

Note#2: This template is designed to transfer the PRJ files that are stored on the DeltaVision computer, in the following folder path: 

data2 > lab_member > exp_date > sample_id > time_point > your_PRJ_files. 

You don’t need to change any of the text that is not highlighted, unless your folder organization is different.

( for LAB_MEMBER in Julius; do (for EXP_DATE in 112517 113017; do (for SAMPLE_ID in 5 48 33024 38075; do (for TIME_POINT in 0 6 7 7.5 8 8.5 9 9.5 10 10.5 11 12; do (mkdir -p /media/BERCHOWITZ/${LAB_MEMBER}/${EXP_DATE}/${SAMPLE_ID}/${TIME_POINT}/; cp /data2/${LAB_MEMBER}/${EXP_DATE}/${SAMPLE_ID}/${TIME_POINT}/*PRJ* /media/BERCHOWITZ/${LAB_MEMBER}/${EXP_DATE}/${SAMPLE_ID}/${TIME_POINT}/); done); done); done); done; echo “---task completed---”)&

1. Adapt the command line: change the highlighted items to fit your needs: for each of them, you can put one or more element(s), separated by space.

2. When your modifications are done, open the terminal, copy and paste into the terminal (using the mouse or using CTRL+SHIFT+C/CTRL+SHIFT+V), and press enter. The task may take a few second to complete: wait until “---task completed---” appears on the terminal.

3. You may if you wish save your modified version of the command line for later usage.

4. Then safely remove the hard drive.

Beware: 

1. Be consistent with upper and lower case usage. 

2. If in any of the element you change contains a space, you need to add a ‘\’ character before the space, otherwise the computer will interpret the space as an element separator and not as a character. Let say for example your name is ‘Maria Teresa’: you would replace “Julius” by “Maria\ Teresa” and not by “Maria Teresa” otherwise the computer will understand that you want to take the images from 2 users, “Maria” and “Teresa”, and you’ll get an error message saying “No such file or directory”. 

3. The same goes for any special character, as they have a meaning for the computer, such as ( ) $ # & ; (not exhaustive). 

4. To avoid such problems it is safer to use only ‘A-Z’ ‘a-z’ ‘0-9’ and ‘_’ to name your folders and documents. 

5. Last but not least, when you create ‘variables’ (here LAB_MEMBER, EXP_DATE, SAMPLE_ID, TIME_POINT are variables), do not use simple names such as DATE, USER, HOME, or PATH: those are already existing and used by the computer, and you would overwrite them and likely mess up the computer.

Southern blot from Pulsed-field gel electrophoresis or from gDNA digestion

Protocol adapted from Murakami et al., 2009, from L. Argueso (Lorraine Symington lab) protocol, and from Alain Nicolas lab protocol.

Southern blot from Pulsed-field gel electrophoresis

STEP1: Preparation of agarose-embedded DNA plugs for non-meiotic cells

Specific material:

• Plug molds 

• Plug cutter (home-made: attach 2 razor blades together with anything in between, like small strips of blotting paper, so that they are from 3.5mm to 7mm apart.)

1. For each sample, grow cells ON in 5mL YPD at 30°C on shaker. Alternatively, for pooled plugs, grow cells in 1mL YPD in deep 96 well culture plates.

2. Measure OD600 of the cultures. Collect the appropriate volume for about 40.10^7 cells if haploid and 20.10^7 if diploid (goal: 6ug DNA, 2ug per plug), transfer in a 5mL tube.

3. Spin down 3 min 3000g, discard supernatant. (→ pause point: cell pellets may be frozen @ -20°C)Wash 1x cell pellets in 4mL EDTA 50mM pH 7.5 (pipet mix), spin down 3 min 3000g, discard supernatant. 

4. Weigh one empty 5mL tube for each sample, with a precision of 0.1mg. Write on the tube its weight.

5. Resuspend pellet in 4mL EDTA 50mM pH 7.5 (pipet mix), transfer in the pre-weighted 5mL tubes, spin down 3 min 5000g, discard supernatant (higher speed to obtain a hard pellet).

6. Remove any excess of liquid, and weigh the pellets. Resuspend the smallest pellet (normally around 25-30mg is the minimum expected) in 100uL of 50mM EDTA, and adjust the volume for all the other tubes normalized on the smallest pellet (200uL for 60mg etc...).

1. Label the plug molds, 3 wells per samples. Seal up the flat side of the mold with tape.

2. (under the hood) Prepare fresh spheroplasting solution (5% BME, 1mg/mL zymolyase 100T, in SCE) into a 5mL tube.

Need ~40ul per sample. It is difficult to prepare less than 1mL

(weigh zymo first and adapt volumes to it)

Zymolyase 100T 1mg 

SCE solution 950uL 

BME 50uL

1. Prepare fresh 1% LMP agarose mix, (1% low melting point agarose, 0.125M EDTA pH 7.5), adjusting volume to number of samples.

NOTE for next time: it is possible to do 2% agarose to obtain less fragile pellets if no restriction digests are planned on the plugs.

Need ~200uL per sample, but plan ~1.5x more

LMP agarose 100mg

0.5M EDTA pH 7.5 2.5mL

H2O 7.5mL

Heat gently in a closed 15mL conical tube in a water bath, and keep @ 42°C.

1. Transfer the same volume of each cell suspension (usually around 110uL) in 1.5mL tubes, equilibrate 1 min @ 42°C.

2. (under the hood) Set up the labelled plug molds, a 200mL beaker filled with hot water, with inside the conical tube with melted agarose and an empty 40mL beaker. Aliquot spheroplasting solution by 200uL into 1.5mL tubes. Then process samples by 5 at a time: 

1. Take 5 samples out of the 42C water bath.

2. add 1mL LMP agarose to a 1.5mL tube with 200uL spheroplasting solution. Pipet mix well and place the tube into the tiny beaker (for no more than 5 min, to avoid zymolyase inactivation).

3. process each sample one at a time: add 200uL of spheroplasting+agarose to the 1.5mL tube with 100uL cells in EDTA, pipet mix well, and distribute immediately into 3 wells of the plug mold strip (about 90uL each). 

4. Discard the remaining sphero+agarose.

5. Repeat a and b until all samples are processed.

6. Let the plugs solidify 10-40min @ 4°C.

3. (under the hood) Prepare RNAse solution (0.45M EDTA pH 7.5, 0.01M tris HCl pH 7.5, 7.5% BME, 10ug/ml RNAseA) into 50mL tube(s): 

Need ~3mL per sample. For 80mL (enough for 26 samples)

0.5M EDTA pH 7.5 2x36mL

1M tris HCl pH 7.5 2x40000uL

BME 3x3mL

H2O 2x560uL

RNAseA 10mg/ml 2x40uL (add at the last minute - filter tip!)

1. (under the hood) Aliquot RNAse solution into 5ml tubes (3mL per tube - filter tip).

2. (under the hood) Using a flat toothpick or a spatula, extrude each plug into the corresponding tube with RNAse solution (3 plugs per tube), and incubate @ 37°C for 1h30 or overnight on a rotating wheel.

3. Prepare fresh proteinase K solution:

Need ~3mL per sample. For 80mL (enough for 20 samples)

Sarkosyl buffer 2x40mL

Proteinase K 8mg or 2x205uL of 19.5mg/mL stock (filter tip!)

1. (under the hood) Carefully pour off the liquid over a 50mL tube covered with gauze fabric (use a scoopula to transfer any fallen plug back in the tube) or use p1000 to remove the liquid and replace with 3mL proteinase K solution, invert gently several times.

2. Incubate overnight @ 50°C on a rotating wheel.

Day 4

1. Carefully pour off the liquid (the plugs should have become more transparent?).

2. Wash 2x in 3mL 50mM EDTA for 15 min on a rotating wheel @ RT.

3. Wash 1x in 3mL storage buffer for 15 min on a rotating wheel @ RT (the plugs will start sinking?).

4. Replace with 3ml fresh storage buffer, store plugs @ -20°C.

STEP2: Pulsed-field gel electrophoresis

NOTE: gel runs for 38h or 25h depending on desired setting. Start electrophoresis in the afternoon if 38h, in the morning if 25h.

1. Take out of the freezer the 5mL tubes containing plugs in glycerol, select one plug for each sample, wash 2x to 4x in 2mL 500mM EDTA 10mM Tris pH 7.5 for 1h on rotating wheel.

2. Prepare fresh 4 L of 0.5X TBE (200mL commercial 10X TBE + 3800mL milliQ H2O).

3. Save 150mL 0.5X TBE to make the gel, Fill the tank of the Bio-Rad CHEF Mapper apparatus with the rest of 0.5X TBE, switch on the circulating pump and set temperature to 14°C. Leave for 1 h.

4. Prepare 1% agarose (1.5g biorad PFGE certified agarose + 150mL 0.5X TBE), dissolve in microwave and keep in 50°C incubator.

5. for each sample: 

1. Using a scoopula, harvest the plug, and deposit it onto a clean parafilm.

2. Using a the cutting tool, cut the plug and transfer the central slice onto the tooth of the comb. Discard the remaining plug fragment or save it for later reuse.

6. From the bottle of melted agarose, take out 1-2mL with p1000, and place a drop above the plug on each tooth of the comb: the drop will spread and cover the top of the plug, sealing it to the comb. It should be solidified in 30 s or 1min.

7. Prepare the gel casting stand and comb in the cold room: screw the left and right parts tightly to the central piece, while making sure that the black insert is securely placed in between.

8. Add the comb in the dedicated slot. The slots are larger than the comb: either put the comb against the back or the front of the slot, but make sure it is not angled.

9. Gently pour the 150mL 1% agarose from the very bottom of the gel cast, slowly, until it reaches the comb and until the plugs are covered. If plug slices are thin, no need to use the whole 150mL.let solidify ~15min.

10. Open the gel cast and gently remove the comb. Hold the gel by the black insert, and deposit it in the dedicated slot in the CHEF apparatus (make sure that it is well held at the bottom by the rims so that it doesn’t move during electrophoresis). Optional: Leave in place for 30 min to let the gel equilibrate to the temperature.

11. Set Initial Switch Time to 45 s, Final Switch Time to 95 s, Voltage Gradient to 6 V/cm2 (=120V total), Time to 38h, and Switch Angle to 120°. (alternatively: Set Initial Switch Time to 60 s, Final Switch Time to 120 s, Voltage Gradient to 6 V/cm2 (=120V total), Time to 25 h, and Switch Angle to 120°. The result will be very different).

12. Stop electrophoresis after planned time.

13. Transfer the gel into a large tray and incubate it with fresh EtBr+TBE solution at 0.5 ug/mL for 30 min or with sybr gold (concentration?).

14. In the meantime, if required, empty the tank in a bucket by opening the pipe and switching on the pump. Rinse the tank with milliQ H2O and empty it again. Switch off pump, temperature, pump device, amd electrophoresis device. In Lorraine’s lab, leave it filled.

15. (Rinse gel 5 min with milliQ H2O, and) take a picture with multiple exposure times. (Be careful when choosing the exposure times, the signal fades very fast.) (with sybr gold, was not an issue)

16. Either directly proceed to transfer, or incubate gel in a tray with milliQ water, close and store at 4°C overnight.

17. Irradiate gel in UV crosslinker (Stratagene stratalinker) to nick chromosomes and facilitate transfer. For PFGE UV is preferred to depurination. Place gel in the stratalinker device, onto a sheet protector.

→ Choose ENERGY mode, type in 1800, press start. (UV irradiation = 180mJ/cm2)

1. Incubate the gel in 250mL denaturation solution for 2x 15 min on shaker.

2. Cut a ~15x20 cm Hybond-N+ (Amersham Biosciences) membrane for transfer, dip in in H2O, then dip it in transfer buffer for 1min, and place it on the vacuum blotter. Cover with the plastic mask (gasket).

3. Dispose the gel onto the membrane, making sure the gel seals the plastic mask. A tiny air leak can be repaired by placing a parafilm piece in between the mask and the gel, and adding a little bit of depuration buffer. A larger one (broken gel, opened wells...) can be repaired by pouring 0.8% agarose in 1X TBE to seal it up.

4. Open vacuum to 60-80 mbar.

5. Cover the gel with depurination buffer, leave for 30 min.

6. Pipet out depurination buffer, and cover with denaturation buffer, leave for 30 min.

7. Pipet out denaturation buffer, and cover with transfer buffer, leave for 1h30.

→ regularly check the constant pressure, and add more transfer buffer after 45 min.

1. Wash membrane in a tray with 0.4N NaOH for 15 min.

2. Wash membrane in a tray with 0.5M Na2HPO4 pH7.3 for a bit, up to a few hours.

3. Wash in DI water, and let dry inside absorbent paper cover (Whatman 3MM) for at least 1hr. Can be stored as is in a drawer.

STEP4: Transfer DNA onto membrane

→ proceed as for Southern blot from gDNA digestion protocol.

STEP5: Hybridize with radioactive probe

→ proceed as for Northern, but with Church Buffer instead of Northern Hybridization buffer.

Southern blot from gDNA digestion

STEP1: Extract DNA using epicentre kit (but more carefully than our regular DNA extraction with the same kit)

1. Harvest cells: 7.5 to 15 OD of cells is largely enough (4 to 8mL of OD 1.8 meiotic culture or adjust accordingly), spin down and aspirate supernatant. Process right away or snap freeze and store pellet in -80ºC.

2.  Add 300uL Yeast Cell Lysis solution, vortex or pipet to mix.

3. Add 1uL of 5ug/uL RNase A, vortex or pipet to mix.

4. Incubate @ 65ºC for 15mn.

5. Incubate on ice for 5mn.

6. Add 150uL MPC Protein Precipitation Reagent and vortex to mix for 10s.

7. Centrifugate max speed for 10mn.

8. Prepare new set of tube with 500uL isopropanol.

→ From now on, be gentle: no harsh pipetting, no vortexing, to keep DNA molecules the largest possible. Some even cut their tips.

1. Transfer supernatant to tube with isopropanol and mix by inversion to precipitate DNA.

2. Centrifugate max speed for 10mn to pellet DNA.

3. Aspirate supernatant and add 500uL 70% ethanol, mix by inversion. 

4. Let DNA pellet settle down and aspirate ethanol. Quick spin and reaspirate to remove any remaining ethanol. Let dry.

5. Resuspend DNA pellet in 35uL H2O. store DNA @ 4ºC or -20ºC depending on storage duration.

STEP2: Digest DNA with restriction enzyme

1. Choose restriction enzyme according to your needs, and check on the provider’s website if the enzyme is efficient for long digestions.

(→ Here to probe Ty3 and discriminate Ty3 endogenous locus from Ty3 cDNA: HindIII enzyme give 11kb vs 5kb respectively. BamHI and EcoRV would have worked too but HindIII more efficient ON)

1. Measure DNA concentration on Nanodrop.

2. Prepare digestions in PCR strip, final volume 30uL:

• 10ug gDNA

• 40U or 2uL enzyme at 20’000 U/mL (here HindIII-HF).

• 3uL appropriate 10X buffer (here cutsmart buffer).

• H2O to 30uL

Note: you can make a master mix of enzyme + buffer but some enzymes do not like to be in 10X buffer so include H2O to the master mix. (here used 10uL H2O + 3uL buffer + 2uL enzyme for master mix, and diluted all gDNA to 667ng/uL in 15uL)

1. Incubate on thermal cycler: 37ºC for 16h, 80ºC for 20mn to inactivate enzyme, 4ºC ∞.

STEP3: Separate fragments by electrophoresis

1. Use a B2 gel box, and prepare 120mL of 0.8% agarose in 1X TAE. 

2. Add 5 uL loading dye + sybr gold to sample (vf = 35uL), and load on gel.(Here used 1 24-well comb, which fit 17uL max each. The whole 35uL would have fitted in 12-well comb.)

3. Use 10uL generuler 1kb DNA ladder with sybr gold.

4. Run at 80V for ± 3h or until migrated enough (shorter if using 2 combs).

STEP4: Transfer DNA onto membrane

NOTE: for efficient transfer of fragments ≥10 kb, it is recommended to break the DNA either by UV irradiation of the gel or by depurination (used here). Depurination step induces breakage between a fraction of purine bases and their sugar, followed by decomposition of the sugar and thus creating a single strand breaks. Denaturation step induces separation of DNA strands, which in combination with depurination will produce smaller, single stranded, fragments.

1. Transfer gel into pyrex dish.

2. Cover with Southern Depurination Buffer (0.25M HCl) and incubate for 10mn on shaker. Rinse 3x in milliQ H2O.

3. Cover with Southern Denaturation Buffer (0.5M NaOH 1.5M NaCl) and incubate for 10mn on shaker. Rinse 2x in milliQ H2O.

4. Set up transfer in Alkaline Transfer Buffer (1.5M NaCl 0.25M NaOH) overnight using the same method as for Northern blot protocol.

5. After ON transfer, incubate membrane in 0.4M NaOH for 15mn on shaker.

6. Rinse membrane in 0.5M Na2HPO4 pH7.3 for a few minutes.

7. Rinse in milliQ water and TEST: stain in methylene blue, destain in milliQ water, and image, as for Northern.

8. Prewarm 1 falcon of Prehybridization Buffer, and add 300uL boiled ssDNA.

9. Incubate in rotating oven at 65ºC for few hours to ON.

STEP5: Hybridize with radioactive probe

→ proceed as for Northern, but with Church Buffer instead of Northern Hybridization buffer.

Radioactivity Use Protocols

General Notes about Usage, Storage, and Disposal:

1. Material can only be ordered (via PO) by authorized users. Coordinate ordering with Luke.

2. Radioactive material comes with a CUMC usage sheet that must be placed into the yellow binder. During monthly audits, incoming material will be compared with usage on file.

3. On days of use, the Geiger counter must be checked with a checksource to ensure it is working correctly. Use the Uranium ore and log check into the yellow binder under “Survey Meter Calibration” tab.

4. On days of use/disposal, radioactive material must be logged in two places: in the yellow binder under “Use Record & Waste Log”, and on the waste tag on the disposal container. Check previous entries for details on how to log; be sure the waste container number listed on the usage sheet matches the tag on the actual waste container.

5. Once a pig is completely used up, it can be discarded in solid waste.

6. If radioactive material isn’t used up and expires, note on the use form “Decayed - Discarded pig to solid waste on [date]” and discard.

7. When waste containers are full, go to https://research.columbia.edu/content/radioactive-waste-guidelines and click the link for “Radioactive Waste Pickup Request” to request pickup/get a new container. Make sure all labeling in the binder moving forward reflects the new tag once it arrives.

8. Monthly wipe reports must be done every single month, unless there is no use (in which case the No Use Log in the yellow binder must be filled out). See full protocol below. If a month is missed, a Note To File must be placed in the yellow binder indicating why.

Monthly Wipe Report – Radiation Safety

Materials needed:

Monthly wipe report sheet (extras in binder, or print more avail online)

12x liquid scintillation vials, 20mL (stored in room 1517)

6x Q-tips (stored in room 1525 drawer with toothpicks)

Liquid scintillation solution (stored in room 1517)

Liquid Scintillation Counter (LSC) (room 1205)

Vial racks and flags (stored in room 1204)

Steps:

1. Prepare Monthly Wipe Report sheet with your name (“Authorized User”), Date, and Location. Check the Liquid Scintillation Counter box and write in the machine serial number. Fill in the locations to be surveyed—typically the same ones each month, see previous reports and the lab map for details.

2. Prepare vials: label the tops 1 (Blank), 2-12. Cut q-tips in half and use the cotton end to wipe down the indicated areas (about 10x10cm). Place the q-tip, cotton-side down, into the vial. Add one pump (3mL) of solution to each vial and cap. Bring to Floor 12, room 1205. Edit: lazslo suggest that we add way more than that. ⅔ of vial (12mL).

3. Check out racks and flags for LSC run—they are stored in Lab 1204, far right aisle when facing the windows, left side, middle of the bench, second drawer from the bottom on the left. Be sure to sign out in the log the racks/flags you are using—typically Flag 12. The log book is on the bench across from the drawer. Each rack fits 12 samples.

Usually: control= rack 127 flag SNC

samples= rack 8 flag 12

1. Set up run on LSC software: Right-click on the flag being used (i.e 12) and select “Associate Assay”. A menu will come up; scroll to the bottom and select “ROUTINE WIPES”. Fill in the necessary dialog boxes (user, report name, output location, etc.) and click ok.

2. Load sample vials into rack and place appropriate flag on the end into the slot. Make sure the slider bar is pushed to the left or the machine will not be able to read the flags and just send them around in circles. Set the racks into the machine with the flag end of the rack to the left, facing the user.

3. Press the play button (green flag) on the software to complete the run (takes ~20min for 12 samples). At the end of the run, the report will automatically print out. Hole punch and place in the binder. The column labeled “CPMC” is the counts for P-32 and should be transposed onto the “DPM” column of the wipe report sheet. CPMA and CPMB are counts for 3H and 14C; they should be low (3H <100, 14C <25).

4. Return racks and flags to their home and sign out on the log.

5. Discard liquid scintillation vials in the bin for them in room 1517. Fill out waste logs accordingly.

NOTE: On the report, CMPA, B, and C are the channels set at specific intensities that correspond to H-3, C-14, and P-32 respectively. Since we don’t use these, high numbers in these categories mean the LSC is contaminated and should be reported.

NOTE: Per Angela 1/11/2018 training, the controls/standards inside the machine shouldn’t be used to calibrate because they expired in 2013 and are thus useless. Per Lazslo: they should be used anyway, because of the decay time they are not bad…

Reminder: efficiency=cpm/dpm

dpm=cpm/efficiency

Background = <200dpm

Geiger check when working: check count in a non-working area. Check working area: should be less than 2x non-working area..

NB: in reports, don’t write 0, rather write <0.1.

Polysome Profiling with Sucrose Gradient 

Preparation of solutions

1. 10x Polysome Lysis Buffer:

97 mL (for ~360 samples)

1M TRIS-HCL, pH 7.5 20 mL

1M MgCl2 10 mL

3M KCl 16.66 mL

Autoclaved MiliQ H2O 53.33 mL

1. 1x Polysome Lysis Buffer with DTT and Cycloheximide:

10 mL (for 6 samples)

10x Polysome Lysis Buffer 1 mL

10 mg/mL Cycloheximide 100 uL

1M DTT 30 uL

Autoclaved MiliQ H2O 8.96 mL

→ Add DTT, Cycloheximide, and Triton X (to 1%) just prior to use. Mix by vortexing.       

1. Light Sucrose Solution:
   10% Sucrose + 1x PLB 40 mL (for 6 samples)
   
   

1. Heavy Sucrose Solution:
   50% Sucrose + 1x PLB 40 mL (for 6 samples)
   
   

Preparation of Lysate

1. Transfer pellet with 1 mL of 1xPLB+DTT+Cycloheximide to 2 mL screw-cap tube and spin down at 800 g for 1 min (3’000rpm).

2. Aspirate supernatant and resuspend in 330 uL PLB+DTT+Cycloheximide and add 1 PCR tube of small glass beads. NOTE: from this point, keep all samples on ice.

3. Process the tubes twice in the FastPrep for 45 s @ max speed (6.5M/S) each in the cold room.

4. Heat a 20G needle to red hot with a bunsen burner. Turn the screw-cap tube upside down and tap twice to move contents away. Poke a drainage hole in the side of the bottom of the 2 mL screw-cap tube. Place the screw-cap tube in a 15mL conical tube containing a 1mL pipette tip inside.
   NOTE: Do NOT poke the hole directly at the bottom center of the tube or it will clog. Put the screw-cap tube inside a 15mL centrifuge tube on top of the 1 mL pipette tip.

5. Transfer 15mL conical tubes to the centrifuge in the cold room. Turn to max speed and leave 30s-1min.

6. Remove screw-cap tube from 15mL conical tube. There should be only beads remaining; if there is visible liquid, the tubes need to spin down longer. The 15mL conical tube should have ~300 uL liquid inside it and might have a pellet with the tip (some transfer of beads is normal, but not preferred). Use a pipette and the tip in the conical tube to remove the liquid, resuspend the pellet and transfer into the first set of cold, labeled microtubes.Make sure the beads do not get transferred to the microtube by placing the pipette tip against the wall of the tube.

7. Spin down microtubes for 5 minutes at 800 g (3’000 rpm).

8. Remove supernatant and transfer to a new eppendorf.

9. Spin down for 20 minutes at max speed.

Preparation of Sucrose Gradient

1. Add Light Sucrose Solution to ultracentrifuge tubes to the mark.

2. Add Heavy Sucrose Solution at the bottom of the tube (i.e under the Light Sucrose Solution) and fill up to the mark.

3. Place caps on tubes and set gradient maker to 10-50 gradient. Make sure to level the pedestal.

Preparation for Ultracentrifugation

1. Using the Figurski lab rotor and swinging buckets (Beckman SW41Ti rotor) (NOTE: be very careful to not drop the rotor!), place the ultracentrifuge tubes in the buckets and balance their weights by adding buffer such that 1 and 4, 2 and 5, and 3 and 6 are balanced. Make sure to measure their mass with the caps on.
   HINT: Tare balance to 0 with the heavier swinging bucket, tube and cap of the pair and then balance to 0 with the second by adding PLB. Tare to 0 again for the next pair.

2. Add 300 uL of sample to tubes, carefully not to disturb the gradient. If there are blanks, add PLB in same volume.

3. Set buckets on rotor and place rotor on (Beckman LE-80K) ultracentrifuge. Make sure it is well hooked and balanced before starting the ultracentrifuge!! Set ultracentrifuge to 38’000 rpm for 2 hrs. Hit ENTER+START. Make sure it gets to 3’000 rpm (~1 min) and then check on Ultracentrifuge every 30 mins.

†

ɑ-factor induction time course and mating time course for Ty3 induction

• on Zymo website, it is said to use 5uM for bar1∆ cells and 100uM for BAR1 cells. Our strains are BAR1 so I used 100uM. I have not tried 5uM. NOTE actually used 10uM

• Bilanchone 2015: they use 6uM ɑ-factor, their strain is BAR1

• Menees 1994: they use 0.34uM ɑ-factor their strain is bar1∆, they use 3.5uM for BAR1 cells

Day1: pull MATa (or MATa and MATɑ) strains of interest on YPG

Day2: inoculate strains in YPD ON (10 mL culture in 50 mL flask is enough).

Day3:

Adapt culture volumes to sample needs, including some or all of the following:

• 3.6OD for Northern blot

• 7.2OD for Western blot

• 270 OD for ribotrap

• 45 OD for polysome / fractionation

• ɑ-factor induction time course:

1. At 8:00 am, dilute ON culture to OD ~0.2 in 50 mL fresh YPD in 250 mL flask. 

2. Let cells grow and reach log phase (~4h) at 30ºC on shaker.

3. At 12:00 pm, measure OD and collect 36 OD of cells (OD=0.36, 36 OD = 45mL).

4. Spin down cells and resuspend in 100 mL at OD=0.36 in 1L flask.(it should be about 12:30 pm now)

5. At 12:30 pm

1. collect t0h time point (OD=0.36, 10.8 OD for NB+WB = 30mL)

2. induce with ɑ-factor at 16ug/mL / 100uM (stock is 5mg/mL / ) : here 224uL in 70mL.

6. At 2:00 pm, measure OD and collect t1h30 time point.(usually OD~0.6, 10.8 OD for NB+WB = 18mL)

7. At 3:30 pm, measure OD and collect t3h time point. (usually OD~1.2, 10.8 OD for NB+WB = 9mL)

8. At 8:30 pm, measure OD and collect t8h time point.

• ɑ-factor induction time course with pulse chase:

1. At 8:00 am, dilute ON culture to OD ~0.2 in 50 mL fresh YPD in 250 mL flask. 

2. Let cells grow and reach log phase (~4h) at 30ºC on shaker.

3. At 12:00 pm, measure OD and collect 36 OD of cells (OD=0.36, 36 OD = 45mL).

4. Spin down cells and resuspend in 100 mL at OD=0.36 in 1L flask.(it should be about 12:30 pm now)

5. At 12:30 pm

1. collect t0h time point (OD=0.36, 10.8 OD for NB+WB = 30mL)

2. induce with ɑ-factor at 16ug/mL (stock is 5mg/mL) : here 224uL in 70mL.

6. At 2:00 pm, measure OD and collect t1h30 time point.(usually OD~0.6, 10.8 OD for NB+WB = 18mL)

7. Harvest culture and wash in YPD

8. Resuspend in same volume of YPD and let grow 4 more hours (it should be about 2:30 pm now)

9. At 6:30 pm, Measure OD and collect t5h30 time point. 

• Mating time course:

1. In the morning, dilute ON culture to OD ~0.2 in 50 mL fresh YPD in 250 mL flask. 

2. Let cells grow and reach log phase (~6h) at 30C on shaker

3. Measure OD, determine volume to collect t0h samples and determine volume to collect 36 OD of cells.

4. Collect t0h sample

5. Collect and spin down cells and resuspend each in 5mL YPD

6. Combine MATa and MATɑ in 50 mL flask (makes 10 mL at OD=3.6).

7. Collect t1h30 time point.

Sum1-AID pGAL-NDT80 induction in YPG - MATa haploid cells

Day1

- Pull strains on YPG.

Day2

- Inoculate strains in YPG ON in 100mL in 1L flask.

Day3

- Dilute cultures in YPG to OD 0.3 in 150mL YPG in 2L flask.

- Leave 2-3h for cells to adjust to new medium and start growing again. Measure OD, adapt collection volume to OD.

- Prepare tubes for sample collection:

- Collect 3.6OD for Northern blot

- Collect 3.6OD for Western blot

- Collect T0h time point. Volume after T0h= V0

- Induce with Auxin 1mM (stock 500mM)

- Collect T1h time point. Volume after T1h= V1

- Induce with ß-estradiol 1uM (stock 5mM) 

- Collect time points up to T7h.

Sum1-AID pGAL-NDT80 induction in KAc - MATa haploid cells

TO FINISH TO EDIT

Day1

- Pull strains on YPG.

Day2

- Inoculate strains in YPG ON in 50mL in 1L flask.

Day3

- Dilute cultures in YPG to OD 0.3 in 150mL YPG in 2L flask.

- Leave 2-3h for cells to adjust to new medium and start growing again. Measure OD, adapt collection volume to OD.

- Prepare tubes for sample collection:

- Collect 3.6OD for Northern blot

- Collect 3.6OD for Western blot

- Collect T0h time point. Volume after T0h= V0

- Induce with Auxin 1mM (stock 500mM)

- Collect T1h time point. Volume after T1h= V1

- Induce with ß-estradiol 1uM (stock 5mM) 

- Collect time points up to T7h.

Transfection Protocol:

Written by Tom Wood 06/08/2021 - last update 2022/08/03

Equipment/reagents needed:

• Media: DMEM, DMEM SUPP, OPTI-MEM

• 24 Well Plate(s)

• Trypsin EDTA for passaging in day 1 or day 3 step. 

• Eppendorf tubes in TC hood

• Desired construct + corresponding GFP-expressing vector + corresponding empty vector, all at 0.5-1 µg/ µL

Day 1:

1. Plan out the transfection map: number of conditions, controls, number of wells per conditions.

2. From a maintenance dish, count and seed cells into x number of wells, with 500ul DMEM SUPP in each well, for a 24 well plate at the desired cell density (250k per well for Hek293T is best). 

3. Place these cells back in the incubator to rest until transfection  (do up-down-left-right shaking in the incubator to distribute cells)

Day 2:

1. Pre-warm DMEM SUPP and OPTI-MEM media at 37C. 

2. Prepare medium mix: 250ul DMEM SUPP and 200ul OPTI-MEM to scale up to n+1 number of wells.

3. Aspirate the medium from wells and replace by 450ul of the medium mix to each well. 

4. do up-down-left-right shaking to distribute the liquid all over the well surface.

5. Place these cells back in the incubator for ~24 hours (also do the up-down-left-right shaking in the incubator)

6. Prepare DNA-Lipofectamine 3000 complexes in 2 steps:

• Step 1: make master mixes (see table)

For n+1 reactions

n= 4 reactions

Tube 1

(make 1 per Lipofectamine concentration*)

Tube 2A

(No DNA control)

→ For 1 well

Tube 2B

(Empty Vector control)

→ For 1 well

Tube 2C

Desired DNA (make one per experimental condition

→ For 1 well

Tube 2D

Next condition…

→ For 1 well

- x ul Lipofectamine 3000 (1 µL per reaction/well)

- 1ul P3000 Enhancer reagent

- 1ul P3000 Enhancer reagent

- 1ul P3000 Enhancer reagent

- 1ul P3000 Enhancer reagent

- x ul OPTI-MEM (25 µL per reaction/well)

- 25ul OPTI-MEM

- 25ul OPTI-MEM

- 25ul OPTI-MEM

- 25ul OPTI-MEM

• Step 2: Combine diluted Lipofectamin with diluted DNA (add tube 1 to tube 2) at a 1:1 ratio, by adding the diluted Lipofectamine to the diluted DNA.
  

NOTE:

• Scale up Tube 1 to n+1 number of reactions

• *Prepare 2 sets of Tube 2s if testing 2 Lipofectamine concentrations

• Scale everything up to the number of wells 

Procedure:

• Dilute Lipofectamine 3000 Reagent in Opti-MEM Medium (“Tube 1”), mix well.

* for new transfections, usually test 2 concentrations: 0.75 µL and 1.5 µL Lipofectamin.

• Dilute DNA in OptiMEM Medium, then add P3000 Reagent (“Tube 2”), mix well.

→ minimum 4 reactions:

• transfection of your desired DNA(s) (“Tube 2D”, 1 per desired condition):

• transfection of a GFP-expressing DNA control to assess transfection success (“Tube 2G”), 

• transfection of an empty vector DNA control (“Tube 2E”) which will be analyzed in the same way as the desired experiment DNA transfection

• no DNA control (“Tube 2N”) to assess cell health (NOTE: see with practice if necessary)

1. incubate DNA-Lipofectamine 3000 mixes for 10-15 mins at RT in the TC hood. 

2. Add 50ul of DNA-Lipofectamine 3000 mix to its corresponding well, slowly in a dropwise fashion directly above the middle of the well. 

3. Do up-down-left-right shaking to ensure even distribution of the lipofectamine mix

4. Place these cells back in the incubator for ~24 hours (also do the up-down-left-right shaking in the incubator)

Day 3:

1. Visualize samples and take images of each well (currently using Hachung lab microscope in their TC room), making sure to save GFP and Trans images. Image J/FIJI can be used for analysis. 

2. To collect cells for samples (e.g. western or northern)

1. prewarm trypsin EDTA

2. aspirate the OPTI-MEM, DMEM SUPP, lipofectamine mix from the well

3. add 200-250ul of prewarmed trypsin EDTA to each well (do up-down-left-right shaking to distribute the liquid all over the well surface). 

3. Place the plate back into the incubator for 3 mins. 

→ Label 1.5 mL tubes for sample collection (1 tube per well, pool if needed post counting and viability measure).

1. After incubation, add 4x the volume of DMEM (800 µL - 1mL) to each well and gently spray the well with the media to remove as many cells as possible from the plate. 

2. Move the ~1ml of DMEM/trypsin EDTA mix to the pre-labelled eppendorf tube. 

3. Spin the tubes at 300 RCF for 5 mins. 

4. Return to the TC hood and aspirate the supernatant, without disturbing the pellet.

5. Resuspend the pellets in 1ml of DMEM SUPP. 

6. Count cell: Mix 10ul of trypan blue with 10ul of resuspended cells (give tubes a shake/pipette to make sure cells have not settled back down) and load 10ul of this mix into a cell counting slide. Count cells on the countess and take as many cells as needed for your samples, pooling tubes from wells containing the same conditions if needed. 

For now: 250-500K cells per western sample (adjust volumes of other reagents as needed), use all other cells for RNA extraction for the Northern sample.