Protocols for Laboratory of Molecular Pharmacology

Immunofluorescence:

Prepare sterile cover glass in 35mm X 10 mm dish. Remove cover glass soaked in 70% Ethanol and place in 35mm X 10mm dish for appropriate samples, wash with 1x PBS sterile , and air dry in the hood(this can dry while preparing cells). All PBS is –CaCl -MgCl

Take cells from incubator and remove medium and wash with PBS.

Add 2ml Trypsin, cover and remove quickly. Place in incubator for ~10s (incubation time depends on cell types).

Remove and add 2ml of appropriate medium to dish and collect cells.

Take 20 l of cells into cell counter (hemocytometer), counts the number of cells and set at concentration 4x105 cells to seed cells to cover slip, and add appropriate concentration to cover glass slide and add medium to 2 ml, and place in incubator overnight.

Using room temp. drug, add appropriate dose for specific time course. (APH 2 l per 2 ml for 1 g/ml)

After treatment, remove medium and wash with PBS, and expose cells to 500 l of cold 4°C hypotonic lysis solution (10 mM Tris-HCl pH7.4, 2.5 mM MgCl2, 1 mM PMSF and 0.5% Nonidet P-40) for 8 minutes on ice, remove and quick wash with PBS.

Fix cells in 500 l of 4% paraformaldehyde for 10 min (dilute from 20% paraformaldehyde) at room temp., followed by 5 min. PBS wash (2x).

Incubate in 1 ml of cold 4°C 100% Methanol in –20°C for 15 min., followed by 5 min. PBS wash (2x).

Block with 500 l 5% BSA in PBS for 15 min-1hr covered in wet-box in 37°C incubator, followed by quick wash PBS.

Add antibody with 0.5% BSA for 100 l diluted solution per sample to cover glass for 1 hr at 37°C, covered in incubator, followed by quick wash , 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS. (For PCNA, use 1 to 100 dilution)

dd second antibody using same protocol as described above with appropriate concentration. (For -H2AX, use 1 to 1000 dilution)

Add secondary antibody diluted with 0.5 %BSA at appropriate concentration for 100 l per sample and incubate covered for 1hr at 37°c (For PCNA, use 1 to 100 dilution of Alexa 488), followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS.

Repeat at appropriate concentration for additional secondary antibody using same protocol described above. (For -H2AX, use 1 to 400 dilution of Cy-3)

After wash, add 8 l DAPI diluted in Mounting Medium for each sample onto a clean glass slide (up to 3 samples per slide), and place the cover glass inverted, with cells facing into DAPI and store in 4°C. Allow DAPI to set for 3 hours or over night.

All washes unless specified are at Room Temperature (RT).

DNA Fiber Assay:

Prepare cells, seeded, at 4x105 in 10 mm dish for specific sample. Add medium to 2 ml.

For 2 ml of cells add 4l of 10 mM Idu (final conc. 20 m), mix and incubate at 37°C for 10 min.

After 10 min, remove medium and quick wash PBS (3x), then add 2 ml appropriate medium. All PBS is –MgCl –CaCl.

Add appropriate treatment for specific time. (for conc. 1 g/ml APH, add 2 l APH)

Add 4 l of 10 mM CldU (conc. 20 m) and 2 l APH, then incubate at 37°C for 20 min.

After 20 min., remove medium, then quick wash PBS once.

Add 1 ml Trypsin to each dish, quickly remove and incubate for ~10 sec. at 37°C

Add 1 ml PBS to each sample and check concentration with cell counter for and dilute in PBS if necessary for 1x106 cell/ml, then collect cells in 1.5 ml tube and centrifuge for 5 min. at 5,000 RPM.

Check pellet, remove supernatant, and resuspend with appropriate amount PBS for specific conc (1x106 cells/ml). Can be stored at 4°C for a few days.

Add 2.5 l of sample with 7.5 l of SDS lysis buffer(0.5% SDS in 200 mM Tris-HCl, pH 7.4, 50 mM EDTA) (pipette up & down 3-5 times), and place total solution on uncoated glass slide, which should be placed at an angle (we use an old tip box) to allow for solution to run down, for 8 min. covered with foil. Slides are 75x25 mm Daigger Superfrost/Plus Microslides. Cover slip is 24x50 mm.

After 8 min., fix cells in 3:1 methanol/acetic acid (prepare fresh) for 5 minutes in coplin jar at room temperature.

Remove and allow to air dry on tip box covered for 8 min. and store in 70% Ethanol at 4°C overnight.

Incubate with cold 4°C 100% Methanol at room temperature for 5 min., followed by 5 min. PBS wash (2x) in coplin jar at room temperature under yellow light. (At this point everything, i.e washes, addition of antibodies are performed under yellow light and the washes are in the coplin jar).

Add 150 l RT 2.5 N HCL to each slide & cover with slip-cover in wet-box in incubator at 37°C for 1hr, followed by 5 min. PBS wash (2x). (Always remove cover slip for all washes, and for each incubation, use a new cover slip)

Add 150 l 5% BSA for 15 min., covered in wet-box in incubator at 37°C,Followed by quick wash PBS.

Add rat CldU(Anti-BrdU, Accurate Chemical, #OBT0030) to 0.5% BSA (1 to 100 dilution), then add 150 l to each slide for 1hr at 37°C, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS.

Add Alexa 488 anti-rat IgG to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS.

Add mouse Idu to (Anti-BrdU, BD, #347580) 0.5%BSA (1 to 50), then add 150 l to each slide for 1 hr at 37°C, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS.

Add Cy3 aAnti-Idu mouse IgG (Cy-3 mouse IgG) to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS wash (5 min.), then quick wash PBS.

Add 20 l DAPI in mounting medium or use alternative DAPI staining to each slide to counter-stain for DNA and store in 4°C. Allow DAPI to set for 3 hr or over night.

Images of DNA fibers were captured by CCD camera (Roter Scientific; Cool SNAP FX)

with epifluorescence microscopy (Olympus; IX70) using a 100X objective lens.

The length of DNA fiber is measured by Iplab software.

Troubleshooting:

Adjusting the initial concentration of cells per slide may help. I usually use between 6000-10000 cells per 2.5 l.

Alternative wash for antibodies can be 3x 0.1% Triton in PBS for 3 min., then PBS quick wash

Alternative DAPI staining: Take DAPI (.25mg/ml) and dilute 1:1 with PBS and add 100 l to slides and cover with cover glass for 10 min at room temp followed by 2x PBS wash for 3 min. Then add 20 l Mounting Medium to slides, cover with cover slip and store at 4°C.

Western Blotting:

After concentrations of samples are equilibrated (Bradford Protein Assay), boil samples for 5 min., then keep on ice.

Using 10% Tris-glycine gel for PCNA, remove white sticker, label wells and place in clean protein electrophoresis apparatus (Biorad), wells facing in, and pour 1x running buffer (dilute from 10x.) into chamber.

Load protein marker (for See Blue marker, 20 l) and samples (at specific concentrations) into the chamber and set at 120 V for 2 hr.

Cut chromatography paper and cut out a membrane. You need 6 squares of paper and 4 pads for 1 membrane.

After 2 hr at 120 V, fill a tray with transfer buffer (25x to 1x tris-gly transfer 40 ml, 10 % methanol 100 ml to 1 L w/ water) and remove running buffer and wash electrophoresis apparatus filling it with transfer buffer. Also place the 6 papers in a tray with transfer buffer. Also place the 4 pads in a tray and cover with transfer buffer.

Place the membrane in methanol for 5 min and then keep in transfer buffer Remove the gel out of its casing, cutting off the ends of the geland place the gel in the tray with transfer buffer.

In the chamber for the membrane transfer, place two pads, with 3 papers and then the gel with the membrane followed by 3 papers and two pads into the electrophoresis apparatus. Take stack oriented as so, and place in deep box and place in the apparatus lid facing out (Do not contain air bubbles between gel and menbrane).

Set voltage at 40 V for 2 hr or 20 V for overnight.

After electrophoresis, block the membrane in 5% milk (50 ml PBS, 2.5 g dry milk) overnight or 1 hour.

After blocking, remove milk and wash PBS to membrane in tray

Add antibodies diluted to 1% milk to specific pieces of the membrane, (for 1 ml PCNA, add 200 l 5% milk to 800 l PBS and 5 l PCNA mouse antibody at 1 to 200 dilution) then add 500 l to wax and place membrane on top, mixing it. Add other half of antibody to wax where the bands should be and place the other wax piece on top so that there is no bubbles between them and incubate for 1 hr, followed by 0.1% PBS-Triton for 5 min. (3x) on shaker. Place in fresh PBS after wash.

Add secondary antibody specific (for 1 to 20,000 dilution at 20 ml, add 4 ml 5% milk to 16 ml PBS and 1 l secondary antibody, for mouse PCNA, donkey IgG-HRP) to primaries and incubate on shaker in tray for 1 hr, followed by 0.1% PBS-Triton for 5 min. (3x) on shaker. Place in fresh PBS after wash.

Prepare chemiluminescent substrate by adding 1 ml of luminol enhancer and 1 ml stable peroxide solution and mix.

Place membrane, (if membrane cut, all add pieces back together) after removing excess PBS, in small tray and add substrate, mixing and allow to incubate for 5 min.

Take cassette and prepare for membrane by placing saran wrap over the cassette where they overlap in the middle, taping the bottom piece.

After 5 min., place the membrane in the cassette and fold over top piece of saran wrap making sure it is flat against the membrane and tape up the edges.

Take film and develop in the dark room.

Protein Extraction and Measurement of Protein Concentration:

Prepare cells at 2 x10 cm dishes per sample.

Remove medium from plate and add fresh medium and treat accordingly.

After treatment, remove medium and quick wash with PBS.

Use sterile cell scraper and collect cells adding two plates per tube, on ice

Centrifuge tubes at 5,000 RPM for 5 min., then remove supernatant.

Resuspend pellet in 200 l lysis solution(10 mM Tris-HCl pH7.4, 2.5 mM MgCl2, 1 mM PMSF and 0.5% Nonidet P-40) on ice for 8 min.

Centrifuge tubes at 735g for 5 min. and remove supernatant (soluble protein) to tube.

Lyse pellet with 200 μl of buffer II (25 mM sodium phosphate buffer pH 7.4, 0.5 M NaCl, 1 mM EDTA, 0.5% Triton X-100, 10% glycerol, 5 mM MgCl2 and 1 mM PMSF). for 20 min. rotate in cold room.

Centrifuge at max for 15 min.

Collect the supernatant. This is insoluble protein.

Measurement of protein concentration

Create standard for protein assay by adding BSA at .2, .4, .8, 1.2, 1.5 mg/ml in separate tubes.

Add 25 l of each BSA conc. with 125 l reagent A, then vortex.

Add 1 ml of reagent B at room temp. for 15 min.

Create samples by taking 5 l of protein to 20 l water.

Add 125l reagent A and 2.5 l reagent S, then vortex.

Add 1 ml reagent B at room temp for 15 min.

Create blank by adding 25 l water to 125 l reagent A, then vortex.

Add 1 ml reagent B at room temp. for 15 min.

Run in spectrophotometer by setting standard curve (use methods TS), then running samples. Usually concentration set for 100 g/ml.

Comet Assay:(From Trevigen CometAssay Kit, # 4250-050-K)

Prepare cells in dish at 4x105 cells and incubate overnight.

Treat cells with specific drug for time-points and incubate.

Melt agarose gel in boiling water for 5 min. and incubate in 37°C until needed.

Cool Lysis solution in 4°C until needed

After treatment, remove medium, wash with 1x PBS, and add 1 ml Trypsin and incubate ~10s in 37°C, then add 1 ml PBS. Set concentration for 1x105 cells.

Place 80 l of warmed agarose into 1.5 ml tube for each sample and keep in incubator at 37°C.

Remove 8 l of sample and place in agarose tube, pipetting twice. Place 75 l of mixed solution onto Comet slide and spread around circular area. Repeat for all samples.

Place samples in covered box for 30 min. at 4°C.

After time add Trevigen Lysis solution to cover each sample (~200 l) for 30 min at 4°C.

Prepare Alkaline solution as follows:

NaOH pellets: 0.6 g200 mM EDTA: 250 lddH20: 49.75 l

Shake to dissolve and set in room temperature before use.

After 30 min. lysis, add Alkaline solution to cover each sample (~200 l) for 30 min., covered in dark at room temperature.

Prepare 1xTBE buffer.

After Alkaline, remove excess solution and wash samples with TBE buffer for 5 min at RT (x2)

Setup electrophoresis apparatus and pour TBE buffer. Set voltage to 19 V for 20 min. Place slides from negative to positive.

Add 70% Ethanol to slides, enough to cover sample for 5 min. at room temp.

Air dry samples to bring cells into single plane for observation at RT.

Once agarose is completely flat on slide, add 50 l of diluted Sybr Green onto each sample and cover with cover glass.

Store at 4°C.

Allow Sybr Green to set for 3 hr or over night.

Fluorescence In Situ Hybridization:

(From BioPrime Labeling kit #18094-011 )To make probe dissolve 100 ng/ml DNA in 5 l TE buffer.

On ice, add 20 l 2.5x Random Primers.

Denature by heating for 5 min. in a water bath. Immediately cool on ice.

On ice, add 5 l 10x DNTP Mixture, then add water to total volume 49 l.

Mix briefly, and quickly add 1 l Klenow Fragment. Mix gently, but thoroughly, and centrifuge for 15-30 sec.

Incubate at 37°C for 1 hr.

Add 5 l stop buffer.

Store probe in -20°C.

Prepare and fix cells on slide using DNA fiber protocol. Slides are 75x25 mm Daigger Superfrost/Plus Microslides. Cover slip is 24x50 mm.

Store slides in 4°C.

Prepare probe solution for 1 slide as follows. Multiply as necessary. Items should be on nice.

Probe (45ng): 2.2 lSsDNA (Salmon Sperm): 0.8 l Cot-1: 2 lWater: (Add to 22 l) 17 l

Add 2.2 l 3M Sodium Acetate per slide at RT. Multiply as necessary

Add 51 l 100% Ethanol per slide at RT. Multiply as necessary

Place in -80°C for 30 min., then centrifuge at 14,000 rpm for 15 min., remove supernatant

Add 200 l 80% Ethanol, wash but do not resuspend.

Centrifuge at 14,000 rpm for 5 min., and remove supernatant.

Add 6l water for each slide/probe into the tube with pellet.

Create Hybrid Mix as follows (enough for 2 slides) :

Formamide: 37.5 l50% Dextram : 15 l20x SSC: 6.25 lWater: 3.75 l

Add 25 l per slide into tube. If tube contains 3 slides worth of probe add 75 l of hybrid mix.

Vortex, and place in 80°C for 10 min.

Immediately place on ice for 5 min. followed by incubation for at least 30 min.

Prepare slides for denaturation as follows:

Remove slides from 4°C, and remove Ethanol.

In 50 ml tube add as follows:

4N NaOH: 700 l100% Ethanol: 12 mlWater: to 40 ml

Place slides (up to 2, face out) in the solution for 3 min.

Then using cold Ethanol place slides in tubes as follows:

70% Ethanol for 3 min.

80% Ethanol for 3 min. 90% Ethanol for 3 min.

100% Ethanol for 3 min.

Allow to dry for 8 min. at angle on tip box.

Once dry, add probe on slide and seal with rubber cement. Usually probe amount 25-32 l, depending on amount of probe solution available and number of slides.

Seal slides with rubber cement and place in wet-box in 37°C incubator over night.

After overnight, remove cement, cover slip, and place slides in 2x SSC while preparing the following in 50 ml tubes (3-50%Formamide/2xSSC(cold 4°C) & 3-2xSSC and place in water bath set to 45°C:50% Formamide/2x SSC 3 min. x 3 times

2x SSC 3 min. x 3 times

5% BSA, 200 l covered by cover glass for 30 min. in 37°C. Use wet-box. (All washes are without cover slip and in coplin jar under yellow light. PBS is –CaCl -MgCl) Quick wash PBS

Alexa 488-Strepavidin for one hour in 37°C created as followed for one slide:Alexa 488-Strep 1 l5% BSA 13.3 lPBS 188.7 lUse 200 l per slide, and cover with cover slip

Wash with 0.1% Triton-PBS 3x 5 min.

Biotin-Strepavidin (Vector, BA-0500) for one hour in 37°C created as followed for one slide:

Biotin-Strep 2.4 l5% BSA 13.3 lPBS 184.3 lUse 200 l per slide and cover with cover slip

Wash with 0.1% Triton-PBS 3x 5 min.

Alexa 488-Strepavidin for one hour in 37°C created as followed for one slide: Alexa 488-Strep 1 l

5% BSA 13.3 lPBS 188.7 lUse 200 l per slide and cover with cover slip

Wash with 0.1% Triton-PBS 3x 5 min.

Alternative DAPI staining. Take DAPI (.25mg/ml) and dilute 1:1 with PBS and add 100 l to slides and cover with cover glass for 10 min at room temp followed by 2x PBS wash for 3 min. Then add 20 l Mounting Medium to slides, cover with cover slip and store at 4°C. Allow Mounting Medium to set for 3 hr or over night.

For IdU and CldU staining of 400 ml RL4-B18:

Take 175 ml of cells into large centrifuge tube (x2) and incubate with 35 l (to each tube) of 100 mM IdU (final conc. 20 m) for 10 min. at 37°C.

Remove tubes and spin down at 1,000 RPM for 5 min.

Remove medium and wash 2x with 10 ml PBS (-MgCl –CaCl) (resuspend pellet) at 1,000 RPM for 5 min.

Add 175 ml of fresh medium to each tube and add 35 l (to each tube) of 100 mM CldU (final conc. 20 m) for 20 min. at 37°C.

Remove and collect 50 l of cells into 15 ml tube for control and then spin down at 1,000 RPM for 5 min. and remove medium.

Add 1 ml PBS to each tube and resuspend pellet. Collect cells in 50 ml tube. Repeat.

Add PBS to 10 ml and resuspend cells thoroughly.

Prepare fractionation by turning on machine and setting as follows:

Rotor: JE-5Speed: 2,000 RPMTime: Hold

Press start to activate centrifuge.

Turn on flow rate machine and set to 30 ml/min and change from water to PBS in tube and run for 5 min.

Label 50 ml and 15 ml tubes according to fraction rate (14, 16, 18, 20, 22, 24, 26, 30).

Check centrifuge for air, and if block has air, quick spin centrifuge to remove it.

Close the line from PBS to centrifuge and cells into syringe and add into collecting line, open line and allow cells to go in.

Close line and then open line to PBS and centrifuge.

Set flow rate to 12 ml/min and allow for collecting tube to become clear, then set flow rate to 14 ml/min and collect to 50 ml in labeled tube by removing collecting tube from waste into appropriate tube.

Once tube approached 50 ml, set flow rate to next fraction (16 ml/min) and place into next tube, while taking 10 ml into 15 ml tubes for FACS. Repeat for all fractions and place on ice.

Once complete, turn off centrifuge, remove tube from PBS and replace with water, flowing and removing cells from centrifuge block.

Set flow to 60 ml/min to completely clean and then shut-off.

Cover 50 ml tubes on ice until needed.

Prepare for FACS:

Centrifuge 15 ml tubes for 5 min. at 5,000 RPM.

Remove PBS and add 2 ml 70% ethanol and keep in 4°C for at least 30 minutes.

Prepare fresh PI solution as follows in 15 ml tube:

Add 50 l of stock PIAdd 4 l RNase AAdd PBS to 2 ml

After 30 minutes, remove ethanol and wash with PBS.

Add 200 400 l to each tube, plus control, mix and transfer to 5 ml round-bottomed tube and take to FACscan.

Restart computer and open Cell Quest.

Turn on FACS and run with water for 5 min. followed by isotone for 5 min.

In Cell Quest, go to acquire and connect to cytometer.

Open in cytometer, settings 1-4.

Go to instrument settings and open settings. Find appropriate setting, then select set and then done.

Select open, go to your folder, then new folder, then add date.

Select file, then file name, and change sample id.

Set control settings, to find peak by placing control under arm, then acquire. Use Fl2-A and FL2-W, and SSC-H and FSC-H for acquisition plots, and Counts and FSC-HFls-A for histogram. When ready to run uncheck setup. Run FACS on appropriate tubes. Once complete, open new, and select gate plot and add twice, then select histogram and select file, then copy into powerpoint and change file for all samples, save and quit. Follow

warm-down procedure on FACS, by running Conrad solution for 5 minutes and without arm for 1 minute, then remove waste and shut down machine.

According to FACscan, use appropriate fraction for FISH, by setting the concentration of cells, preparing slides (like DNA fiber), and putting on the probes according to FISH protocol with the following exceptions;

After final Alexa-Strepavidin incubation and wash continue as follows (All washes are in coplin jar under yellow light without cover slip. Always use new cover slip for incubations):

Add Anti-rat CldU(Anti-BrdU, Accurate Chemical, #OBT0030) to 0.5% BSA (1 to 100 dilution), then add 150 l to each slide for 1hr at 37°C, followed by 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Alexa 594 Anti-Rat IgG to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed by 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Anti-mouse IdU (Anti-BrdU, BD, #347580) 0.5%BSA (1 to 50), then add 150 l to each slide for 1 hr at 37°C, followed 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Anti-Idu (Alexa Mouse 647, for far red) to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Alternative DAPI staining. Take DAPI (.25mg/ml) and dilute 1:1 with PBS and add 100 l to slides and cover with cover glass for 10 min at room temp followed by 2x PBS wash for 3 min. Then add 20 l Mounting Medium to slides, cover with cover slip and store at 4°C. Allow Mounting Medium to set for 3 hr or over night.

If using Alexa 350 for blue, do not add DAPI.

Cell Viability Assay:

Prepare cells in 2 ml plates at 4x105 concentration in the following labeled drug concentrations 0, 0.1, 1, 10 g/ml APH. Incubate overnight.

Remove cells from incubator and add appropriate concentrations of drug for 2 hrs.

After treatment, trypsin the cells and collect with 1 ml medium. Set concentration for 1050 total cells divided by your cell count for 300 cells per 2 ml of medium.

Add cells to 7 ml of medium for each sample.

Mix and add 1 ml at a time into 6 well and incubate for 10 days.

After time, remove medium and add 2 ml of fixation solution (50% Methanol, 5% Acetic Acid, made fresh from RT materials) for 1 hr at room temperature.

Remove fixation solution and add 2 ml of Wright’s Geisma Stain and allow to incubate for 1 hr at room temperature.

Count colonies manually!

Fluorescent In Situ Hybridization with Idu and CldU:

(From BioPrime Labeling kit #18094-011)To make probe dissolve 100 ng/ml DNA in 5 l TE buffer.

On ice, add 20 l 2.5x Random Primers.

Denature by heating for 5 min. in a water bath. Immediately cool on ice.

On ice, add 5 l 10x DNTP Mixture, then add water to total volume 49 l.

Mix briefly, and quickly add 1 l Klenow Fragment. Mix gently, but thoroughly, and centrifuge for 15-30 sec.

Incubate at 37°C for 1 hr.

Add 5 l stop buffer.

Store probe in -20°C.

Prepare and fix cells on slide using DNA fiber protocol:

Add Idu for 20mM into cells and incubate for desired time at 37°C.

Remove tube and spin down at 1,000 RPM for 5 min.

Remove medium and wash 1x with PBS (-MgCl –CaCl) (resuspend pellet) at 1,000 RPM for 5 min.

Remove PBS and add fresh medium to each tube and add CldU for final conc. 20 m for desired time at 37°C. (For us, we normally will do Idu 10 min, CldU 20 min)

Remove tube and spin down at 1,000 RPM for 5 min.

Remove medium and wash 1x with PBS (resuspend pellet) at 1,000 RPM for 5 min.

Check pellet, remove supernatant, and resuspend with appropriate amount PBS for specific conc (1x106 cells/ml). Can be stored at 4°C for a few days.

Add 2.5 l of sample with 7.5 l of SDS lysis buffer(0.5% SDS in 200 mM Tris-HCl, pH 7.4, 50 mM EDTA) (pipette up & down 3-5 times), and place total solution on uncoated glass slide, which should be placed at an angle to allow for solution to run down, for 8 min. covered with foil. (We use a tip box)

After 8 min., fix cells in 3:1 methanol/acetic acid (prepare fresh) for 5 minutes in coplin jar at RT.

Remove and allow to dry on tip box covered with foil for 8 min. and store in 70% Ethanol at 4°C overnight.

Store slides in 4°C.

Prepare probe solution for 1 slide as follows. Multiply as necessary. Items should be on ice.

Probe (45ng): 2.2 lSsDNA (Salmon Sperm): 0.8 l Cot-1: 2 lWater: (Add to 22 l) 17 l

Add 2.2 l 3M Sodium Acetate per slide at RT. Multiply as necessary.

Add 51 l 100% Ethanol per slide at RT. Multiply as necessary.

Place in -70°C for 30 min., then centrifuge at 14,000 rpm for 15 min., remove supernatant.

Add 200 l 80% Ethanol, wash but do not resuspend.

Centrifuge at 14,000 rpm for 5 min., and remove supernatant completely.

Add 6l water for each slide/probe in tube with pellet.

Create Hybrid Mix as follows (this is enough for 2 slides, multiply as necessary) :

Formamide: 37.5 l50% Dextram : 15 l20x SSC: 6.25 lWater: 3.75 l

Add 25 l per slide into tube. If tube contains 3 slides worth of probe add 75 l of hybrid mix.

Vortex, and place in 80°C for 10 min.

Immediately place on ice for 5 min. followed by incubation for at least 30 min at 37°C.

Prepare slides for denaturation as follows:

Remove slides from 4°C, and remove Ethanol.

In 50 ml tube add as follows:

4N NaOH: 700 l100% Ethanol: 12 mlWater: to 40 ml

Place slides (up to 2, face out) in the solution for 3 min.

Then using cold Ethanol place slides in tubes as follows:

70% Ethanol for 3 min.

80% Ethanol for 3 min. 90% Ethanol for 3 min.

100% Ethanol for 3 min.

Allow to dry for 8 min. at angle, covered with foil.

Once dry, add probe on slide and seal coverslip with rubber cement. Usually probe amount 25-32 l, depending on amount of probe solution available and number of slides.

Place in wet-box in 37°C incubator over night.

After overnight, remove cement, coverslip, and place slides in 2x SSC while preparing the following in 50 ml tubes (3-50%Formamide/2xSSC(cold 4°C) & 3-2xSSC and place in water bath set to 45°C:

50% Formamide/2x SSC 3 min. x 3 times

2x SSC 3 min. x 3 times

5% BSA, 200 l covered by cover slip for 30 min. in 37°C. Use wet-box for BSA and for all following incubation steps. (All washes are in coplin jar under yellow light without cover slip. Always use new cover slip for incubations)

Quick wash PBS

Alexa 488-Strepavidin for one hour in 37°C created as followed for one slide:Alexa 488-Strep 1 l5% BSA 13.3 lPBS 188.7 lUse 200 l per slide and use cover slip.

Wash with 0.1% Triton-PBS 3x 3-5 min., followed by quick was PBS

Biotin-Strepavidin (Vector, BA-0500) for one hour in 37°C created as followed for one slide:

Biotin-Strep 2.4 l5% BSA 13.3 lPBS 184.3 lUse 200 l per slide and use cover slip.

Wash with 0.1% Triton-PBS 3x 3-5 min., followed by quick was PBS

Alexa 488-Strepavidin for one hour in 37°C created as followed for one slide: Alexa 488-Strep 1 l

5% BSA 13.3 lPBS 188.7 lUse 200 l per slide and use cover slip.

Wash with 0.1% Triton-PBS 3x 3-5 min., followed by quick wash PBS

After final Alexa-Strepavidin incubation and wash continue as follows:

Add Anti-rat CldU (Anti-BrdU, Accurate Chemical, #OBT0030) to 0.5% BSA (1 to 100 dilution), then add 150 l to each slide for 1hr at 37°C, followed by 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Alexa 594 Anti-Rat IgG to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed by 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Anti-mouse IdU (Anti-BrdU, BD, #347580) 0.5%BSA (1 to 50), then add 150 l to each slide for 1 hr at 37°C, followed 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Add Anti-Idu (Alexa Mouse 647, for far red) to 0.5%BSA (1 to 100), then add 150 l to each slide for 1 hr at 37°C, followed 3x 0.1% Triton in PBS wash (3-5 min.), then quick wash PBS.

Alternative DAPI staining. Take DAPI (.25mg/ml) and dilute 1:1 with PBS and add 100 l to slides and cover with cover glass for 10 min at room temp followed by 2x PBS wash for 3 min. Then add 20 l Mounting Medium to slides, cover with cover slip and store at 4°C. Allow Mounting Medium to set for 3 hr or over night.

Add 20 l Mounting Medium (no DAPI) IF USING 350 to slides, cover with cover slip and store at 4°C.

Troubleshooting:

Adjusting the initial concentration of cells per slide may help. I usually use between 6000-10000 cells per 2.5 l.

The steps to making the probe are very important. They should be followed closely and performed quickly.

Alternative DAPI staining. Take DAPI (.25mg/ml) and dilute 1:1 with PBS and add 100 l to slides and cover with cover glass for 10 min at room temp followed by 2x PBS wash for 3 min. Then add 20 l Mounting Medium to slides, cover with cover slip and store at 4°C.

Protocol for IdU and P-BLM staining. ( Protein can be substituted as needed)

For double immunostaining with P-BLM and IdU, cells were incubated with 20 uM IdU (4ul) for 5 minutes before treatment with 0.5ug/ml(10ul)APH.

After treatment with the indicated drugs, cells were washed with PBS, exposed to a hypotonic lysis solution (10 mM Tris-HCl pH 7.4, 2.5 mM MgCl2, 1 mM phenylmethylsulfonyl fluoride, and 0.5% Nonidet P-40) for 8 minutes on ice. Then quick wash PBS

Cells were fixed in 4% paraformaldehyde in PBS for 10 minutes, washed in PBS 2x 5min

Incubate in 100% methanol at -20°C for 15 minutes, and then washed in PBS 2x 5min.

Block with PBS containing 0.5% BSA for 30 minutes, then quick wash PBS

Cells were incubated with 100ul of first antibody, P-BLM (Rabbit) 1:2000, make 400 ul(so, 400 ul rabbit serum and .2ul P-BLM) for 50 min. at 37C, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS 5 min, then PBS quick wash.

Add 100 ul second antibody (Alexa 488 anti-rabbit) at 1:100 in rabbit serum ) for 50 min. at 37, followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS 5 min, then PBS quick wash.

Samples were then fixed again with 4% paraformaldehyde for 10 minutes at room temperature and washed with (PBS for 5 minutes x2).

DNA was denatured in 500 ul 1.5 N HCl for 5 minutes at 37C and washed with (PBS for 5 minutes x2).

Add 100 ul anti-IdU (Anti-BrdU, BD, #347580) antibody diluted 1:50 in PBS with 0.5% BSA for 50 min. at 37C followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS 5 min, then PBS quick wash.

Add 100 ul secondary antibody Cy3 diluted 1:100 in PBS with 0.5% BSA for 50 min. at 37C followed by quick wash 0.1% Triton in PBS, 0.1% Triton in PBS 5 min, then PBS quick wash.

Add 8 ul DAPI in mounting medium to slides and store in 4C.

Allow DAPI to set for 3 hr or overnight.

Trouble shooting: The fixation steps and denature steps can be adjusted as necessary.

Cell Pellet Harvest (Minna Lab)

Grow cells to 90% confluency

In hood, remove medium and wash with (RT) PBS, then remove.

Add 2 ml of Trypsin and incubate long enough to detach cells.

Add 4 ml of Trypsin Neutralizing Solution and collect cells in 15 ml tube.

Spin at 1,000 RPM for 5 mins. After spin remove medium.

Add 3 ml PBS (RT) and resuspend pellet.

Spin at 1,000 RPM for 5 mins. After spin remove PBS.

Add 1ml PBS per 1.5 ml tube (ex. 3ml for 3 tubes) and resuspend.

Place in 1.5 ml tube, and spin at 4,000 RPM for 5 mins.

Remove PBS and store in -80C.

Reagents:

To make 50% Formamide/2X SSC for FISH:

Add 500 ml Formamide with 100 ml 20xSSC

Add water to 1 L

To make DAPI, take stock DAPI (.25 mg/ml in PBS), and add to mounting medium at 1:25:

For 1 ml, 40 l stock to 960 l mounting medium.

To make hypotonic lysis solution:

5 ml of 1 M H-CL Tris, Ph 7.5 (10 mM conc.)1.25 ml of 1 M MgCl2 (2.5 mM conc.)25 ml of 10% N-P40 in water (.5% conc.)87 mg of PMSF

Add water to 500 ml

To make 0.l% PBS-Triton:

Add 1ml Triton X-100 to 1 L PBS

To Thaw and Freeze Cells:

To freeze, collect 500 l of cells in DMSO/20% FBS and place in tube. Cover in Styrofoam box, wrapped overnight in -80.

After overnight, remove from Styrofoam and place in cell box.

To thaw, remove tube and warm with water. Do not completely dissolve.

Place in 9ml of medium and centrifuge at 1,000 RPM for 5 min.

Remove supernatant to 200 l and place cells in 10ml of medium in plate or flask.

To use Epifluorescent Microscope:

Make sure the computer is turned on, then turn on the following:

Mercury Lamp-Make sure the burner light is steadyCamera Power SupplyLudl Box

Turn on iVision. Familiarize yourself with the functions. In order to view with your eyes set the following:

Eyepiece knob set to the eyeFilter wheel set to the slot without tapeUsing the F keys will allow you to switch between channels

In order to capture a 3-color image use the F11 key (Triple Auto), and set the microscope as follows:

Eyepiece knob set to SPFilter wheel set to first slot with tape, clockwise from the no-tape slot

Adjust the exposure as necessary, click apply, then done to capture the image in the particular channel

Turning off: The mercury lamp should stay on if someone is using within 30 minutes. Refrain from turning on/off the lamp frequently, turn off the camera power supply, and the Ludl box

To use confocal:

Make sure the computer is turned on, and then turn on the following:Fans-leave on for at least 10 minutes before turning on the lasersLaser-red & greenBoard to laser-This looks like a computer towerMercury lamp-ignite once you are ready to use the microscopeRT Power SupplyNikon boxNikon remote focus

Once everything is set, open SimplePCIIn order to view with the microscope set the following:Eyepiece knob set to AOil trap out-underneath the channel control box(this should remain out during the use of the microscope, whether eye or camera)Eyepiece control pin set to BINO

You can switch between channels with the channel control box. The left is off, then FITC, TRITC, and a Triple. The confocal is not able to capture images in DAPI

To set the microscope to capture images, set the following:Eyepiece knob set to CEyepiece control pin set to PHOTOChannel box set to off

In Simple PCI, click on the camera icon, and set/adjust the following:Capture mode: 4x IntegrationFast Scan mode: 2x Quadrant ScanClick Fast Scan Icon to Adjust PMT black, Gain, Exposure settings

When satisfied with adjustments, click stop to stop fast scan, then click capture icon(not capture1), once image is captured(wait a few seconds), click stop, and once stop icon returns to capture, on the image window, click the disk icon in order to save the components into the folder of your choice.

Once finished, you need to do the following:

Close SimplePCIClose the oil catchSet the channel box to offTurn off everything in reverse order. Allow for the fans to stay on at least 10 minutes before turning off the lasers.If someone is signed up within 30 minutes, leave the mercury lamp on.

To use the Pathway:

Log onto the computer, and turn on the following:Power to the PathwayLamp ALamp B

Once they are ready, a green light indicator will turn on by the chamber for the slides.If you do not have a user, one can be created by opening the AttoVision Config shortcut on the desktop. From here you can create a new user and clone the setting from other users into your username. A password is not required.

All slides are place down into the slide holder which sits on top of the objective. This can be control by the joystick, which also have a knob for fine and coarse adjustment.

Open AttoVision from the desktop.Once open, you need to open the following windows which can be located in the setup menu, but also have icons as well on the main window for AttoVision:

Dye set up (blue drop icon)XYZ PositionSet up Macro (Blue down arrow)Assay Launch (can be found in capture)

Whatever macro you want to run can be found in the assay launch box or can be viewed/edited manually in the set up macro window. If there is not a macro for the dye combination you want, you can create one by first setting up a probe cycle as follows:

Click probe cycle icon (blue drop with circled arrow), name the combination, then select the dyes from the drop-down list and insert them in the order you want in the probe cycle. Once finished click save. Then select the set up macro. From here name the Macro then from the list select your probe cycle and add it to the macro. Select (1 data point), and add other steps in the macro as desired. If you only want the macro to capture the specific dye images then the 1 data point is fine. If you want to use your montage settings, select use active montage mode.

In order to set up your montage settings, select the set up menu, then montage capture.Name your montage or select one already available. Adjust the height and width and then save. We normally use a 4x4 montage for immunofluorescence. For intensity quantification, we want about 300 cells per sample.

In the Dye set up, you can select each dye and set the exposure. It is a good idea to set the exposure of DAPI to around 1000 or 1500. You can see the channel intensity by moving the mouse over the cells. In order to view a live image, select the camera with green arrow icon in the Dye set up window. If other channels, such as Alexa 488 you want your positive signals to be around 1500 or 2000 intensity, so adjust exposure as necessary. This is something you need to adjust on your own.

When you are ready to capture the montage, depending on your macro, you may have a start position. You can set this with the XYZ position. When you have an area that you want to capture with a montage, select read, then save in the XYZ position window.

Click the run macro icon (green triangle), then create you data name and select run. You can repeat this as many times as necessary for your different samples.

Once you are finished you can select the Anaylze menu, then re-analyze experiment.

From here you can select your data, and then select the dyes you wish to analyze, then click next to go to Wells, then next to Processing, you may want to select or de-select the flat-field and other processing, then select next to reach segmentation.

In segmentation, select re-segment, then select system-rois, then select “polygonts” for nuclei then select the auto button. Make sure DAPI is selected as your image reference. Here you can threshold manually and by pixels. You can test and it will show you how it selects the rois. You want to get this as good as possible, but it may be impossible for the software to completely segment the nuclei.

Once you are done, click close, then select next dye threshold. Here can adjust the threshold according to dye and not shape like segmentation. Once finished, click next and then select what you want to measure and apply it to the dyes you want. Then click run. Once finished you can view the data, which is located here:

C: drive, then AttoVision, then users, then your name, then data. Select the folder, and then scroll down to find a .txt file called ROI summary re-analysis. This can be exported to Excel. Your pictures are here as well.

In order to map the 384- well plate to your slide for fiber analysis, perform the following.

In the C: drive, then Attovision, there will be two .ini files called AttoImageSlide.ini, then Attoimage.ini. Rename the AttoImage.ini to AttoImageoriginal.ini, and then rename the AttoImageSlide.ini to AttoImage.ini. After you image capture, do the reverse to have the original renamed to AttoImage.ini, and the slide rename to AttoImageSlide.ini.THESE STEPS ARE CRUCIAL!

Open Attovision, then open the dye set up, XYZ position, and macro set up. You will need to find or create the macro you want for the dye combination.

From here go to set up, then configuration, then specimen matrix. From here follow the prompts for 384 well plate, and map the slide according to prompts. The slide should be faced down. Once that is finished, restart AttoVision.

Once restarted, check to make sure the slide is mapped by selecting the Plate Icon (six green circles), and then navigate. Make adjustments if necessary. Select your macro, and then select overwrite to add the new points (i.e. H1, H2, G1, G2 etc.) to your macro so that it will navigate to these points.

Go to montage capture, and set up your montage to your specifications. Once set, make sure you macro is using the active montage mode and settings.

Adjust your exposure in the Dye set up box, and when satisfied, select the run macro button.

Once finished, DO NOT forget to rename the .ini files to return the mapping to normal.

List of Primary antibody dilutions (have worked for me):

PCNA-rabbit 1:100PCNA-mouse 1:100RPA-mouse 1:100Polymerase Alpha 1:100Polymerase Epsilon 1:100P-ATM 1:100P-Chk1 1:100Gamma H2AX-mouse1:1000Gamma H2AX-rabbit 1:800P-BLM 1:500

Unless stated, all secondaries should be 1:100, with the exception of Gamma H2AX-mouse, it is 1:400. Also, I prefer Goat and Rabbit species to raise secondaries.

Note: Cy3 is for mouse, and is from sheep. The stock Cy3 is kept at -20.Also of note is that these numbers are a good place to start, but are by no means absolute.

Preparation of Cell Lysates:

Place samples on ice

Prepare lysis buffer by adding the following for 500 ul lysis buffer:

20 ul 50x Complete (40 ul/1 ml)- store at -2020 ul 50x PPI (40 ul/ 1ml)- store at -20410 lysis buffer- store at 4

Add specific amount to cells (usually 30-60), and pipette up & down about 20 times.

May quick spin, and place on ice for 20 mins.

Label 1.5 ml tubes to transfer protein in, and label tubes for protein assay. 8 tubes for standard, 1-8.

After 20 min incubation, spin samples at 14,000 RPM (4c)

When close to finished, about 5 mins make the following:

For 15 samples add 1.5 ml Reagent A to 30 ul Reagent S, mix.

Add 100 ul to sample and standard tube.

After spin, carefully place on ice and transfer supernatant to new tube.

Add 2 ul lysis buffer to standards.

Add 2 ul sample to each tube.

Add BSA (2 mg/ml) as follows in tube:

1 2 3 4 5 6 7 80ul 1ul 2ul 3ul 4ul 5ul 6ul 7ul

Add 800 ul Reagent B & mix gently. Incubate 20 mins @ RT.

Turn on Spectrophotometer, and set wave for 750.

After 20 mins, place standard 1 in white cuvette, and press set reference.

Vacuum solution, and run through standard and samples, record numbers and plug into

Excel.Western Blot Protocol (Minna Lab):

Make 10% Acrylamide Gel (Use Table 18.3): Resolving and Stacking. Do not add TEMED till ready.

Clean apparatus and place glass in holder.

Place grey pads (soft) into holder, and carefully place glass plus green holder into cast holder. Gently pour the resolving gel. After checking for leaks, place 200 ul of water onto resolving to help polymerize it. Let incubate for 15-20 mins.

Vacuum out water, and add TEMED to stacking gel. Make double the amount for the number of gels. Place comb (16 teeth) very slowly and let sit for 20 mins.

During this time, have samples sitting on ice, and can prepare by adding SB buffer plus beta-Mercaptol (1 ul beta-merc per 20 ul 4x SB, down to 1 x, then add protein plus water). Well will hold about 14 ul.

Once made, boil for 4 mins.

After time, remove glass from holder, and pour water over it to gently remove comb. Clean glass and label wells. Place in running holder. Repeat for other gels.

Vacuum out water, and add fresh running buffer to cover wells.

Add samples according to protocol. Add about 7 ul of ladder.

Run at 100V for 3-5 mins. Allow protein to reach resolving gel.

Run at 200V for ~1 hr or as necessary for good separation.

In tray, add ample cold (4c) transfer buffer, place two sponges, 3 papers, and 1 membrane per gel, and place holder in as well to completely soak. Place 1 sponge on black part of holder.

Remove gel from holder and gently remove smaller glass side. Place paper on top, and turn over, and pull membrane off of glass. Place paper side to sponge, and then place membrane on top, followed by two papers plus sponge. Roll out bubble if necessary and close up transfer holder. Repeat for all gels. Place holder with black facing back.

Add ice tray and cold transfer to top, with stir rod on bottom, and run at 100V for 1 hr stirring gently.

After 1 hr remove gel, and place membrane in try. Add ponceau S dye and incubate rocking for 2-3 mins. Drain excess Ponceau and remove from tray by washing with water.

Cut membrane as necessary, and block with 5% milk in 1% Tween-20 or other mixture for about an hour at RT covered.

Follow with 3x10 mins 1% PBS-Tween wash

Then add appropriate amount of 1st Ab overnight, covered, in cold room or 4c.

Wash 3x10 mins 1% PBS-Tween

Then add 2nd Ab

Wash 3x10 mins 1% PBS-Tween

Then add 1:1 luminol, horseradish peroxidase (regular or super substrate) (ex. 2ml, 2ml), and allow membrane to be covered and rocked vigorously for about 3 mins, turning over half-way. Repeat for all membranes, and place in folder in cassette and develop.

May need to adjust exposure times.

Preparation and infection of lentiviral shRNA constructs

Day 1: Seed 293FT cells at 2.4x106 concentration in 10cm dish per 1 shRNA293FT cells grown in 10 ml DMEM (High Glucose, -Sodium Pyruvate, 10% FBS)

Day 2: Prepare constructs to infect 293FT cells 24 hours after plating in 1.5 eppendorf tube as follows:(Brian has aliquots of helper constructs and controls shGFP, GFP transduction). Thaw before use.

shRNA- 3 mg- volume depends on concentrationVSV-G- 3 mg- volume depends on concentrationDelta8.9- 3 mg- volume depends on concentrationFugene 6- 18 mlDMEM (serum free)- to 300 ml

Before adding Fugene, add all others, quickly vortex, spin down, & add Fugene to middle of tube, and gently mix.

Once complete incubate at room temp for 15 mins.

Add solution to cells (remove from incubator) in a dropwise manner, swirl gently, and place cells back into incubator.

Day 3: (In virus hood): After 24 hours, remove medium (Use bleach to sterilize pipettes after use), and replace medium of 293FT cells with 10 ml medium similar to target medium or appropriate medium. (For Mia PaCa2 we used DMEM normal, serum free, no pen/strep)

If planning to infect the next day after collecting virus, seed target cells to 30-50% confluency. I normally seed 1x105 cells per well in a 6-well plate. If planning to do selection, seed 1 well for selection and another for making lysate.

Day 4: After 24 hours, collected virus supernatant of shRNA as follows:Remove medium in 10 ml syringe and filter in .45 mm filter & collect in 15 ml tube. Tube should have polybrene (10 g/ml) at 2.5 l/5 ml virus. Aliquot in 1 ml and either store at -80c or can use immediately and then store at -80c. You can collect another 24 hours by adding fresh medium if you want, but normally 1 collection is enough. We throw the plates out afterwards.

When planning to infect target cells:

Having seeded cells at 1x105 cells in 2 ml medium 24 hours later prepare for infection with lentiviral shRNA.

If virus frozen, thaw on ice (usually takes about 2 hr) or thaw at room temp. (I usually thaw on ice)

In virus hood, remove medium from target cells and add 1ml virus supernatant to each well needed, and incubate at 37c for 2 hours.

After 2 hours, wash gently with PBS (x2).

Add 2ml appropriate medium, and incubate.

If infecting for selection, after 24 hr add appropriate selection drug.Allow for non-selected plate to grow for 72 hours before collecting lysate.

To use L5 Microscrope:

Username for PC and Scheduler: mtorresPassword for PC and Scheduler: mtorres

Make sure power supply, lamp, etc. is turned on. (1,2,3 5 labels) 3 is on microscope.Click on Axiovision. Can use custom toolbar for all changes.Can use Ph2 or Brightfield for capturing light images.DIC used for white light. Select light to eyes. Can do Kohler illumination.Focus image. Turn off field diaphragm, adjust knob until see octagon with light. Place octagon in center of field of view. Adjust then to open the octagon till field of view.Use the DIC to capture images in light.Select live, select to camera, and can measure for exposure.Click on interactive and click on background to increase image quality. When satisfied press snap.Switch to fluorescence. Focus and repeat capturing procedures for white light.

Reagent Recipes:

1 M Tris-HCL pH 6.8 (500 ml) (MW 121.14)400 ml ddH2060.55 g Tris-base

Adjust pH with HCLAdjust H20 to 500 ml

.5 M Tris-HCLDilute equal parts of 1 M Tris-HCL with ddH20

1.5 M Tris pH 8.8 (1 L)181.65 g Tris-base700 ml ddH20

Adjust pH with HCLAdjust H20 to 1000 ml

10 X Running Buffer (2L)60 g Tris-base288 g Glycine20 g SDS

10 X Transfer Buffer (2L)60.6 g Tris-base288 g Glycine

ImmunoPrecipitation

Prepare 10cm dish of 90% confluency of target cells per IP.

On ice:

Place plates on ice and remove medium

Wash with 1x PBS (cold), aspirate PBS. (x2)

Add 1 ml PBS per plate

Scrape cells and collect in 15ml tube

Spin 15ml tube @2,800 RPM for 10 mins. (Prepare lysis buffer at this time 137mM)

After spin, remove PBS, and add 1ml lysis, gently resuspend, and sit at RT for 10 mins

After time, dounce 15 times, and place in 1.5ml tube and spin at 20,000 rcf for 10 mins

Transfer into new tube

For bead conjugation:

Add 1ml lysis buffer to new tube

Add 75l of Ag beads per 4 plates, Add 20l of Sec8 (E-12), wipe tip before adding

Add 1ml lysis to new tube

Add 75l of Ag beads per 4 plates, Add 20l of Igg (control), wipe tip before adding

Rotate for 1 hr at 4C

For preclear:

Add 40l Ag beads to 1ml lysis. Spin at 5,000 rcf for 30s, remove supernatant

Add soluble fraction of IP+ HA Ab (Y-11) (20 l ) to bead tube to pre-clear

Rotate for 2 hr at 4C

For bead conjugation after incubation time:

Spin at 5,000 rcf for 30s, remove supernatant

Add 1ml 1M NaCl IP buffer to each tube, gently resuspend, spin at 5,000 rcf for 30s, remove IP buffer, repeat (x3), 4th time use 137mM IP and follow the same steps.

Place on ice.

For pre-clear after 2 hr incubation:

Spin at 5,000 rcf for 30s, take input (around 50 l)

Add equal volume (usually ~450 l) to each antibody-bead IP, and rotate at 4C for 4 hr

After 4 hrs, spin at 5,000 rcf for 30s, remove supernatant, add 1ml 137mM NaCl buffer to each IP, gently resuspend, and spin at 5,000 rcf for 30s.

Repeat washes with 250mM NaCl, 500 mM NaCl, and 137mM NaCl in the same manner as above.

Add appropriate amount of 2x sample buffer to each sample (Sec8 & Igg)

Add equal amount of 2x sample buffer to input

Boil samples for 10 mins, and then freeze at -20C