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Post Electrophoretic Analysis

Method for Western Blotting


Prepare and run an SDS PAGE gel. Select a gel percent which will give the best resolution for the size of antigen being analyzed (if known). If the size is not known, a 12% gel is a good starting point. Load enough protein to provide 0.1-1ng of antigen per well. It is generally advantageous to load serial dilutions of sample, so as to ensure that one lane at least will fall in the optimal range for detection. If desired, use prestained markers, such as National Diagnostics' ProtoMarkers, to monitor transfer efficiency.


Upon completion of the run, the gel must be transferred onto a blotting membrane. This can be accomplished by semi-dry transfer or wet transfer. Both are electrophoretic transfers, and require equipment designed for that purpose. The procedures outlined below are intended as general outlines. For best results and to ensure safety, follow the equipment manufacturer's instructions for this phase.


  1. Rinse electrode plates with deionized water.
  2. Cut six sheets of Whatman 3MM paper and one sheet of blotting membrane to the size of the gel (or slightly smaller).
  3. Wet the membrane. Soak nitrocellulose in deionized H2O for 3 minutes. Soak PVDF in methanol for 1 minute. Transfer membrane to transfer buffer (composed of 0.02M tris base, 0.15M glycine and 20% MeOH) and soak for 3 minutes.


  1. On the lower plate (positive, red lead), place these items in the following order:
    • Three sheets of 3MM (precut and wetted with transfer buffer)
    • Transfer membrane
    • Gel
    • Three sheets 3MM (precut and wetted with transfer buffer)

    Remove any air bubbles by rolling over each layer as placed with a 10ml pipette. Be careful not to disturb the stack, or to let the gel stick to the pipette. Roll gel after placing first upper layer of paper, if desired. Note that bubbles trapped in the stack will distort current flow, leading to lateral band displacements and failure of bands to transfer.

  2. Check to be sure that no portion of the upper paper stack contacts the paper or the electrode underneath the gel. Contacts between upper and lower stacks will short circuit the current, distorting the transfer. In some systems, parafilm or plastic wrap may be arranged around the gel to prevent this short circuiting from occurring. Check the instructions.
  3. Place the upper (negative, black lead) electrode plate on top of the stack, and apply current. Consult apparatus instructions; typical conditions are 1 hour at 0.8 mA/cm2. Over-transfer may dry the gel and drive proteins through a Nitrocellulose membrane.


  1. Cut two sheets of Whatman 3MM paper and one sheet of transfer membrane to the size of the gel.
  2. Wet the membrane by soaking nitrocellulose in water for 2 minutes. Soak PVDF in methanol for 2 minutes.
  3. Place membrane in transfer buffer (see "Semi-Dry Blotting" above).
  4. Assemble transfer "sandwich" by placing the following items in the following order:
    • Filter paper sheet
    • Gel
    • Membrane
    • Filter paper sheet
  5. Assemble sandwich clamps and support pads per the equipment manufacturer's instructions.
  6. Place sandwich in transfer tank with membrane side closest to the positive electrode (Red lead).
  7. Add cold transfer buffer, and initiate cooling procedure.
  8. Apply voltage. This parameter is entirely dependent upon the apparatus used. Large format gels may require 50 - 75 V for 5 - 15 hours, while minigels can be blotted in one hour at 50 - 100 V in some systems.


It is prudent to mark the blot in a permanent way for orientation, by notching or clipping a corner. If prestained markers were used, their positions should be marked in pencil or an alcohol indelible ink. Often well positions can be distinguished and marked immediately after transfer.

Notes: Nitrocellulose membranes may be air dried prior to further processing. It has been reported that this improves protein retention on the blot. After transfer, Coomassie staining of the gel can give information about the efficiency of transfer.


  1. Stain the blot and mark the positions of well and markers. If prestained markers were used, this step may be skipped.
  2. At this point it is very helpful to spot diluted primary and secondary antibody on an unused area on the blot. This can be invaluable for troubleshooting. If a blot fails (no bands are detected) the antibody spots can be interpreted as follows:
    1° & 2°: 2° antibody functioning well: label okay, 1° antibody may have failed.
    2° only: 2° antibody failed to bind 1°
    No spots: Label enzyme denatured - remake 2° antibody dilution.


A variety of blocking reagents are available. It is worthwhile to optimize blocking procedures, as this step determines the background level of the blot, and hence the detection limit. The most universal blocking agents contain mixtures of proteins and surfactants. This combination provides good to excellent blocking on most membranes.


  • Blocking Solutions:
    1. ProtoBlock: Dissolve 10g of Reagent B in 170 ml deionized water. Add 20 ml of Reagent A.
    2. Tween/milk: Dissolve 50g nonfat dry milk and 2 g Tween 20 in 1L PBS. If product is to be stored for more than one day, add 0.2 g NaN3 (CAUTION - TOXIC!)
  • Blocking Procedure:
    1. Immerse blot in blocking agent with agitation (i.e. a shaking or rocking table) for 1 - 2 hours at room temperature.
    2. Rinse blot in PBS + 0.2% Tween 20 twice for 5 minutes each.

    Blot is now ready for antibody hybridization.


  1. Dilute 1° and 2° antibodies into PBS + 0.2% Tween 20 (PBST).Note: Including blocking reagent at 0.05 - 0.1X in the antibody solutions will decrease background without significantly lowering band intensity.The optimal dilution must be determined for each antibody. The 2° antibody may be tested on dots of 1° antibody. Dilutions can range from 1:100 to 1:10,000.
  2. Incubate blocked, washed blot with 1° antibody for at least 1 hour at room temperature with agitation.
  3. Wash blot four times for five minutes each with PBST.
  4. Incubate blot with 2° antibody for 30 - 60 minutes.
  5. Wash blot four times for five minutes each with PBST.
  6. Transfer blot to detection reagent.


The most popular antibody labels are isotopic (125I), HRP and alkaline phosphatase. 125I labeling is straightforward, and gives consistent and quantifiable results. Its drawbacks are the hazards and inconvenience which radioactive isotopes bring into the lab. Detection of 125I requires that the blot be placed against X-ray film. Upon development, the film will show bands corresponding to the position and intensity of detected antibody band.

A variety of substrates are available for both alkaline phosphatase and HRP. Protocols are given below for the most commonly used.


Stock solutions (each are stable for up to one year):

Solution A: 0.5g NitroBlue Tetrazolium in 10 ml 70% Dimethylformamide
Solution B: 0.5g BCIP in 10 ml 100% DMF
Solution C: 100mM NaCl, 10mM MgCl2 and 100mM tris (pH 9.5)

  1. To prepare substrate solution, mix 100µl of Solution A with 15ml of Solution C, and then add 50µl of Solution B.
  2. Submerge blot (up to 150 cm2) in 15ml substrate solution. Scale up the amount of solution for larger membranes. Incubate with shaking at room temperature until desired band intensity and contrast is achieved (typically 30 minutes) depending on antibody and label activity.
  3. Stop development in PBS + 20mM EDTA

Note: This stop reagent works for CIP or other eukaryotic antibodies. It does not detect Bacterial Alkaline Phosphatase.


Chromogenic detection with diaminobenzidine (DAB):Note: DAB development with HRP is much more rapid than the alkaline phosphatase/BCIP system. In addition, because the stop solution is simply rinsing away substrate, the reaction may continue for a time after "stopping". Development should be taken only up to the point where bands are acceptable and no background has yet appeared.

  1. Make fresh detection solution with the following items:
    • 9mg DAB Tetrahydrochloride
    • 7ml 100mM Tris pH 7.6
    • 1.5ml 0.3% NiCl2 (CoCl2 may be substituted)
    • 6ml water
  2. Filter through Whatman 1 paper.
  3. Add 15µl 30% H2O2 (or 150µl 3%)
  4. Immerse blot in detection solution (up to 150 cm2/15ml) and shake at room temperature until desired band intensity and contrast are achieved. (Typically less than 10 minutes)
  5. Stop reaction by rinsing blot with agitation in PBS.

Chemiluminescent Detection using National Diagnostics' Protoglow ECL chemiluminescent detection reagent:

NOTES: The extreme sensitivity of Protoglow ECL can lead to high backgrounds, in which the entire blot appears as a black image on the film, even at short (less than a minute) exposures. Background can be lowered by rinsing the blot for one minute in PBST after removal from the detection solution. Light emission is essentially stable for 3-6 hours. Multiple exposures may be taken during this period.

    1. Mix 7.5ml of Reagent A with 7.5ml of Reagent B. Allow combined solution to come to room temperature.
    2. Immerse blot (200 cm2/15ml) in combined A & B reagents at room temperature with shaking 1 minute.
    3. Wrap blot in plastic wrap and place in a film cassette.
    4. Expose blot to X-ray film for 1 - 5 minutes, and develop film as usual.


Western blots may be stripped and reprobed, albeit with some loss of sensitivity. Stripping generally involves the use of reducing agents such as 2-mercaptoethanol to cleave the disulfide bands which hold the antibody probes together.

The stripping solution is formed by mixing the following:

  • 2g SDS
  • 750 µl 2-mercaptoethanol
  • 100ml 65mM Tris HCl pH6.8

Incubate blot in stripping solution 60 minutes at 50°C.

Notes: This solution contains a high concentration of 2-mercaptoethanol; use and heat only in hood. Stripping time and temperature given are typical. Optimal values must be determined for each antibody/antigen combination.