Ari Berman – 12/18/2007
Three-color Western Blot Protocol
This protocol assumes that you have a standard western blot protocol in place in your
laboratory. The steps provided here are intended to modify that protocol to incorporate
three-color fluorescence into your gels. This procedure can also be used with a single or
two colors just as easily. I’ve found this system to work much more reliably (and less
stressfully) than the standard ECL methods and it removes the need for stripping or
reprobing your membranes.
Materials needed:
 Millipore Immobilon-FL PVDF membrane (for fluorescence; Cat: IPFL00010)
 PBS or TBS
 Tween-20
 Bovine Serum Albumin (BSA)
 Invitrogen Novex Sharp protein ladder (shows up under fluorescence)
 Three Primary antibodies – Rabbit, Mouse, Sheep hosts (or other combination)
 Donkey Anti-rabbit IgG – AlexaFluor 488, Donkey Anti-mouse IgG – AlexaFlour
555, Donkey Anti-sheep IgG – AlexaFlour 633 (or appropriate combination for
primaries, the key is 488, 555, and 633 excitation wavelengths. See Figure 1 at
the end of the protocol for the spectra analysis)
Procedure:
* - Note: all volumes listed are for a single membrane. Scale up appropriately for more
membranes.
1.
2.
3.
4.
5.
6.
7.
Collect cell lysate, load and run gel as specified in laboratory protocol.
Cut appropriate sized piece of Immobilon-FL membrane for gel.
Place membrane in methanol for 10 seconds to activate, avoid bubbles.
Equilibrate membrane in transfer buffer for at least 10 minutes.
Equilibrate completed gel in transfer buffer for 5 minutes.
Transfer according to lab protocol.
Remove transferred membrane from transfer apparatus and place immediately in
PBS.
8. Rinse 2x quickly (manual agitation, ~15 sec) with PBS to remove methanol from
membrane.
9. Make blocking buffer as follows:
 10mL/membrane
 1xPBS, 0.1% Tween-20, 5% BSA (w/v)
o Pre-mix PBS and tween until in solution
o Add powdered BSA to pre-measured volume of PBS-Tween
o Vortex at full speed until last traces of BSA are no longer visible
(solution should be yellowish, but clear)
o Push blocking buffer through syringe filter (0.20um) into clean tube
o Use mixture for duration of experiment, store at 4°C when not in use
Ari Berman – 12/18/2007
10. Add 5mL blocking buffer to membrane and block at room temperature rocking at
40rpm for 1 hour.
11. Mix primary antibodies with 2mL of blocking buffer for incubation.
o Make sure that each of your primary antibodies are from different hosts and
that you have secondary antibodies to match those hosts. With proper
blocking, the same host as the host tissue may be used without much trouble.
o Be sure to vortex antibody mixture thoroughly.
12. Place blocked membrane in heat-sealed polypropylene pouch.
13. Cut one corner from the pouch and add the 2mL of antibody mixture to the pouch.
14. Remove any large bubbles from the pouch and re-seal cut corner.
15. Incubate overnight at 4°C rocking at 40rpm.
16. Cut a corner from the pouch and drain antibody mixture into a tube for storage, mix
with 0.05% sodium azide. Can reuse antibody mixture 3x.
17. Cut the membrane out of the pouch and place in 4-6mL 1x PBS, 0.1% Tween-20
(wash buffer).
18. Manually agitate the membrane, then lift it out of the wash buffer with forceps and
drain the liquid from the washing container.
19. Replace the membrane and wash 3x10min in wash buffer rocking at room
temperature.
20. Remove blocking buffer from 4°C storage and equilibrate to room temperature.
21. During the final wash, mix 4mL blocking buffer and appropriate secondary antibodies
at 1:1000 dilution (must be AlexaFluor 488, 555, and 633 to work well with
Typhoon).
22. Vortex thoroughly, store in the dark until use (wrap tube in foil).
23. Drain the final wash from the membrane container.
24. Add the 4mL of secondary antibody mixture to the membrane and wrap the container
in aluminum foil in a light-proof manner to avoid photobleaching of fluorescent dyes.
25. Incubate membrane at room temperature for 2 hours rocking.
26. Wash the membrane as in steps 22-23, re-wrapping the container in foil during each
wash (it is vital to minimize the membranes exposure to light once the fluorophores
are in contact with the membrane through the end of the experiment).
27. After the final wash, rinse briefly in PBS and place membrane on a piece of Whatman
paper protein side up, in a drawer (out of the light) to dry for 1 hour.
28. Wrap the membrane carefully in foil and take the dried membrane to the Typhoon
9810 in Bldg 1, rm 117.
o Note: You must be trained and checked out by Christina Ferrell in order to use
the Typhoon.
29. Open the typhoon control system and allow the lasers to warm up.
30. Wipe down the platen of the Typhoon with the dust cloth located on top of the
machine.
31. Place the dried membrane protein side down onto the platen with the top (wells)
facing the left side of the platen.
32. Close the lid and select the proper area to be scanned
33. Select Fluorescence as the acquisition mode, then click Setup.
34. Select the 520 BP 40 emission filter from the drop-down list and change the PMT to
300. Make sure the selected laser is Blue2 (488).
Ari Berman – 12/18/2007
35. Click the second checkbox and select 580 BP 30, set PMT to 300. Make sure the
532nm laser is selected.
36. Click the third checkbox and select 670 BP 40, set PMT to 300. Make sure the 633nm
laser is selected.
37. Click OK.
38. Select the third “R” from the left on the bottom row from the orientation menu.
39. Set the Typhoon to “press sample” and to scan at 200um.
40. Select “FlourSep” from the image analysis menu below the user comment field.
41. Once the typhoon reads ready, click SCAN.
o The Typhoon will scan your blot three times, once at each wavelength
42. When the scanning is complete, your image will be transferred to the fluorsep
program.
43. Right-click on your image and select “Actual Size” to zoom out.
44. Change the view to “side-by-side” view (seventh button from left on top toolbar).
45. Click on “F1” box and resize it to fit the average band size on your gel. All boxes will
match the size change.
46. Move the F1 box to one of the bands in channel 1.
47. Do the same with the F2 box in channel 2, and the same for F3 in channel 3.
48. Click “Perform Separation”.
49. Exit Fluorsep and load up ImageQuant.
50. Select your file from inside the correctly named folder that the images were saved
under from the scanning program.
51. Right-click on the image and select “Gray/Color adjust”, adjust the image to
minimize background and brighten the bands of interest. Do this for each channel.
52. Click OK. You should see your finalized gel with three colors in front of you.
53. You can either quantify using ImageQuant, or export the images for use with other
programs.
54. To export the color image, go to Edit->Copy Image.
o Load Microsoft Powerpoint.
o Paste the image into powerpoint.
o Either save as a powerpoint file, or export it as a tif file.
55. To export the individual channels for quantification, change to side-by-side view as
was done in FluorSep.
56. Click once on the channel 1 box, then go to File->Save As and select tif from the
“Save as type” dropdown menu. Repeat the procedure for each of the channels.
57. Save the images to a USB drive or to the shared network drive if you have access to
that system
58. Quantify as you would a normal gel.
Ari Berman – 12/18/2007
Figure 1 – Spectra analysis with Typhoon lasers and filters for AlexaFluors called
for in this protocol
Figure 2 – An example of a successfully completed three-color western blot
Please email Ari Berman at ari@bermanism.com with any questions about this protocol.