Western Blot

Cell Lysis

For a 24 well plate:

1. Aspirate media.
2. Add 500 uL of cold PBS to each well.
3. Aspirate.
4. Add 200 uL of trypsin.
5. Incubate for 90 seconds.
6. Neutralize with 800 uL of cold complete media.
7. Pipette up and down to disperse clumps.
8. Transfer contents of each well to 2 mL labelled eppendorf tubes.
9. Centrifuge 2500 RPM for 8 minutes.
10. Aspirate supernatant. 
11. Add 500 uL of lysis buffer. 
12. Incubate at 4C with agitation for 30 minutes. 
13. Centrifuge at 12000 RPM for 20 minutes. 
14. Transfer supernatant to new labelled tubes and store at 4C (or on ice). 
15. Discard pellet. 
     

Denaturing Protein 

1. Add 50uL of betamercaptoethanol (reducing agent) to 950uL of Laemmli dye.
2. Add Laemmli dye + betamercaptoethanol to lysate so that it dilutes to 1x (equal volumes of Laemmli and lysate if Laemmli is 2x).  For one gel, this means that for each sample add 10uL dye to 10uL sample.
3. Transfer to heat block set to 98C and boil for 5 minutes. 
4. Store at 4C (reboil if reloading).

BCA Assay (to check protein concentration)

For more details: https://www.piercenet.com/instructions/2160412.pdf

For a sample Excel spreadsheet, see: BCR WB BCA 2014-8-1.xls

1. For a 20mL solution of working reagent, add 10mL Solution A, 9.6mL Solution B, and 0.4mL Solution C to a 50mL tube.

2. In a 96 well plate, make three rows of protein standards.  Add 150uL of protein standard solution and 150uL of working reagent to each well.  To make the protein standards:

   Vial	Volume of Diluent	Volume and Source of BSA	Final BSA Concentration
   A	4.5 mL	0.5 mL of Stock	200ug/mL
   B	8.0 mL	2.0 mL of vial A dilution	40ug/mL
   C	4.0 mL	4.0 mL of vial B dilution	20ug/mL
   D	4.0mL	4.0 mL of vial C dilution	10ug/mL
   E	4.0mL	4.0 mL of vial D dilution	5ug/mL
   F	4.0mL	4.0 mL of vial E dilution	2.5ug/mL
   G	4.8mL	3.2 mL of vial F dilution	1ug/mL
   H	4.0mL	4.0 mL of vial G dilution	0.5ug/mL
   I	8.0mL	0	0ug/mL (blank)

3. Add dilutions of samples to a final volume of 150uL in separate wells, and add 150uL of working reagent to each of those sample dilutions.  Do at least 2 replicates per dilution.  For a seeded 24 well plate, dilutions of 75uL sample + 75uL dH₂O + 150uL working reagent and 25uL sample + 125uL dH₂O + 150uL working reagent tend to result in concentrations that fall within the range of the protein standards.  However, when seeding plates with larger wells one should adjust the dilutions accordingly so that the protein concentrations fall within the range of the protein standards.
4. Cover the plate with a sticky air-tight seal, being sure to close off each well individually so that wells don't contaminate one another.
5. Incubate the plate at 37C, slow shaking, for 2 hours.  Alternately, shake for 30 seconds and then incubate standing for 2 hours at 37C.
6. Cool the plate to room temperature.
7. Use a plate centrifuge (balanced with another plate) to spin condensation/displaced liquid back into the wells.
8. Measure the absorbance at 562nm on a plate reader.
9. BCA assay working reagent requires special disposal.  Make sure to dispose it in a plastic bottle rather than dumping down the drain.
10. Take the average of the replicates.  Subtract from these averages the average blank reading (standard I).
11. Plot a standard curve by plotting the average blank-corrected reading for each standard versus its concentration in ug/mL.  Concentration should be on the x axis and absorbance on the y axis.  Then, fit a polynomial (degree 2) curve to the data.
12. Use the equation for the polynomial curve to calculate back the concentrations for the average blank-corrected absorbances of each sample.  You will need to solve for x in order to find the concentration (y=absorbance, x=concentration).  Make sure to use the correct solution to that equation.  For example, one equation should give higher concentrations for samples with lower absorbances - this is NOT the correct equation to use.  Also, don't just use the equation that Excel gives on the chart because that it is not always precise enough to give accurate concentrations.  Use the LINEST function to find more exact values for the coefficients of the curve fit.
13. Find the concentration of the initial undiluted sample based on the concentrations of the average absorbances for each dilution of sample.

Running an SDS-PAGE gel

1. Use BioRad gel.
2. Load gel into gel running machine.
3. Add 1000 mL of running buffer to both the inside and outside compartments. Make sure the buffer covers the top of the wells.
4. Add 5uL of ladder to the first and last wells of the gel.  
5. Add 16uL of sample to each well.  This should correspond to about 10ug of protein, but down to 2ug can still give some signal.
6. Run gel at 200V for 30 minutes (or until the dye front has reached the bottom of the gel.
7. Remove the gel from the running machine and then remove the gel from its plastic encasement, being careful not to rip the gel.  Cut off the top left corner of the gel.

Transfer

For more details: http://www.bio-rad.com/webroot/web/pdf/lsr/literature/M1703940.pdf

1. Cut two sheets of extra thick blotting paper and one sheet of nitrocellulose membrane to the size of the gel (one set of 2xblotting paper/1xmembrane per gel being blotted).  Cut off the top left corner of the membrane (so that you can orient it relative to the gel).  Also remember that the membrane comes packaged in a protective blue paper - that's not part of the membrane.
2. Soak the blotting paper and in transfer buffer.  Equilibrate the membrane in transfer buffer for 15-30 minutes.  Equilibrate the gel in transfer buffer for 15 minutes.  Don't let the gel equilibrate for too long or the proteins might diffuse out.
3. Put a piece of extra thick blotting paper on the anode and roll a pipette over the surface to get rid of bubbles.  Throughout the loading of blotting paper/membrane/gel try not to get the anode too wet because it makes transfer less efficient.
4. Put the membrane on top of the blotting paper and roll a pipette over it to get rid of bubbles.
5. Put the gel on top of the membrane (make sure that it's aligned in the center).  Roll a pipette over it to get rid of the bubbles and make sure that the gel is lying perfectly flat.
6. Put the second piece of extra thick blotting paper on top of the gel and remove air bubbles with the pipette.
7. Place the cathode and safety cover on the unit.
8. Run at 10V for 15 minutes.
9. Open the transfer machine and dispose of blotting paper and gel.  You can tell that you got successful transfer to the membrane if the ladder now shows up on the membrane.

Incubations

1. Wash the membrane in PBS for 5 minutes, shaking (40mL of PBS required for incubating in a pipette tip box lid).
2. Incubate the membrane with blocking buffer (preferably 5% BSA) for 1 hour, shaking.  5% BSA is made by adding powdered BSA (in Deepak's fridge) to PBST.  Make 100mL (5g BSA to 100mL PBST).  It is difficult to suspend BSA in solution so be sure to vortex and to filter before adding the buffer to the membrane.  Use a vacuum filter for the filtration.
3. Save the blocking buffer (it can keep for up to a week).
4. Wash three times for 5 minutes in PBST.
5. Incubate the membrane with primary antibody solution for 1 hour, shaking.  (Use antibodies diluted in PBST.  Dilute according to manufacturer's recommendations.  Start at the low end of the dilutions.  For polyclonal antibodies like our anti-GFP consider using an even lower dilution.)
6. Save the antibody solution (it can keep for up to a week).
7. Wash three times for 5 minutes in PBST.
8. Incubate the membrane with secondary antibody solution for 1 hour, shaking.  Be sure to cover this because the secondary antibodies are photosensitive.  (Use antibodies diluted in PBST.  Dilute 1/15000.)
9. Save the antibody solution (it can keep for up to a week).
10. Wash three times for 5 minutes in PBST.
11. Image.  Leave the membrane in PBST when taking to image so that it does not dry out (dried out portions show up on the blot reading).