Casein Blocking Buffer Protocol for Southern or Western Blotting

Southern Blot Procedure
Western Blot Procedure

To specifically detect an antigen immobilized on a solid support, the unoccupied sites on the support must be blocked. Blocking of the non-specific sites may be accomplished by a variety of protein and detergent solutions; however, the blocking solution must be compatible with the detection system. Our Casein Blocking Buffer is compatible with a variety of detection systems, including fluorescein and DIG detection systems.

Reagents Required

• Phosphate-Buffered Saline (PBS) with 0.05% (v/v) TWEEN^® 20 (PBS-T), (Product No. P3563)
• Protein probe or antibody
• 0.1 M Tris, 0.1 M NaCl, pH 9.5 (for Southern blotting)

Preparation

Dissolve the contents of one container of the casein blocking buffer in 800 mL deionized water. Once the contents are dissolved, add deionized water to 1,000 mL and stir to mix.

Storage/Stability

Store powder at room temperature. After reconstitution, store the casein blocking buffer at 2-8 °C to avoid bacterial contamination. Solutions may be kept up to one week at 2-8 °C after reconstitution.

Procedure

Southern blot blocking

1. After transfer and cross-linking of labeled nucleic acid to a nitrocellulose, nylon, or positively charged nylon membrane, incubate the membrane with casein blocking buffer for 60 minutes (0.6 mL/cm²) at ambient temperature or for 30 minutes at 37 °C with gentle agitation. Also, note that blocking can be accomplished overnight at 2-8 °C.
   Note: Casein blocking buffer is not suitable for blocking PVDF membranes.
   
   
2. The membrane may now be probed with the DNA fragment according to your specific protocol.
   
   

Western blot blocking, probing, and detection procedure

1. After protein transfer to a nitrocellulose membrane, incubate the membrane with casein blocking buffer for 10 minutes (0.6 mL/cm²) at ambient temperature or for 30 minutes at 37 °C with gentle agitation. Alternatively, blocking can be accomplished overnight at 2-8 °C.
   
   
2. Dilute the primary antibody in casein blocking buffer. A common dilution for primary antibodies is 1:1,000, but may vary. Dilutions may vary from 1:100 to 1:100,000 or higher. The researcher must determine the optimum dilution factor.
   
   
3. Incubate the membrane with the primary antibody (0.6 mL/cm²) for 1-16 hours at 2-8 °C with gentle agitation.
   
   
4. Wash the membrane, with gentle agitation, 3-5 times for 5 minutes each with PBS-T.
   
   
5. Dilute the enzyme-antibody conjugate in casein blocking buffer and incubate the membrane for 30-120 minutes. After the incubation, wash the membrane with gentle agitation, 5-6 times for 5 minutes each with PBS-T.
   
   
6. The membrane may now be exposed to chromogenic or chemiluminescent substrate as per the manufacturer's instructions.

Suggestions for colorimetric detection:

1. The membrane should be exposed to the colorimetric substrate until a positive signal results, but as background begins to develop, the reaction should be stopped. The membrane should be exposed to substrate for no longer than 60 minutes.
   
   
2. For colorimetric peroxidase substrates, the reaction may be stopped by removal of substrate and transfer of the membrane to a solution of 0.1% sodium azide with 1% SDS in either PBS or TBS (Tris-buffered saline).
   
   For alkaline-phosphatase substrates, the reaction maybe stopped by removal of substrate and transfer of the membrane to a solution of 0.3 M sodium phosphate, pH 5.5.

Suggestions for chemiluminescent detection:

1. Following exposure to substrate, the excess substrate should be blotted off and the membrane transferred to a solid support. Transferring the membrane to a "page protector," slightly larger than the membrane itself, is recommended. The supported membrane should then be placed within a heat sealable bag. Using gentle pressure, smooth out air bubbles between the membrane and the plastic bag by rolling a glass test tube or pipet over the contained membrane. Seal the bag and wipe off any excess substrate from the outside of the bag. Place the contained membrane into a film cassette and expose to film.
   
   
2. Initially, an exposure of 1 minute should be used; however, if no signal results, expose the membrane to film for longer. If there is excess signal, use as short an exposure as technically possible. See the troubleshooting guide for other hints.

Suggestions for Western blot detection:

1. Dilutions of the enzyme-antibody conjugate depend on the substrate used for subsequent detection and should be optimized by the researcher. General guidelines for dilutions are 1:5,000 for chromogenic substrates and 1:50,000 for chemiluminescent substrates. The dilution ratios are based on an initial concentration of ~1 mg/mL enzyme-antibody conjugate.
   
   
2. In sensitive systems, fingerprints will show up. Use powder-free gloves at all times. Avoid the use of forceps with ridges, as these also tend to show up.
   
   
3. If using a biotin-avidin system for detection, the blocking reagents should be free of biotin. Casein blocking buffer is not recommended for use in these systems since milk contains large and variable amounts of biotin.
   
   
4. Casein blocking buffer is not recommended for use in systems detecting phosphoproteins, due to the presence of phosphorylated proteins. Gelatin blocking buffer (Product No. G7663) is recommended for those systems.
   
   
5. To verify the quality of the secondary antibody, perform a "blank" membrane, in which the primary antibody is omitted. If background is present without the primary antibody, either use a different blocking reagent or dilute the conjugate further.
   
   
6. The antibodies that are used for the detection of antigens can usually be removed by immersion of the membrane in a buffer containing 100 mM glycine, pH 2.3 for 30 minutes with agitation. This is not a universally applicable procedure; some antigens are dissociated from the membrane, so subsequent probing will be faint or non-existent. Many researchers prefer to prepare membranes in parallel if possible, thereby avoiding the uncertainty of this "stripping" step.

Troubleshooting

Table 1

Problem	Possible Cause	Remedy
No Sigmal	No target present	If the labeled nucleic acid is not present on the membrane, it cannot be detected. Include a positive control with the next assay.
	No target protein present	Verify transfer by visualizing proteins on the membrane using a Ponceau S solution (P7170). If possible, a positive control should always be run to ensure components are functioning.
	Overblocking	Masking of a signal can occur if the blocking reagent is used at too high a concentration. A dilution of 1:1 to 1:3 may be done to decrease the concentration. If the problem persists, different blocking reagents should be tried.
	Interference from contaminants in blocking reagents (e.g. biotin systems)	If the blocking reagent is contaminated with biotin, the free biotin will bind the streptavidin conjugate so that it will not be available for detection.
	Interference from contaminants in blocking reagents (e.g. phosphoprotein systems)	If the blocking reagent contains phosphoproteins, the phosphate sites will bind the alkaline phosphatase conjugate so that it will not be available for detection.
	Inadequate exposure time using chemiluminescence system	First exposure should be 1 minute. If there is no signal, expose for longer. For example, try exposing for 5 and 10 minutes. If there is excess signal, try as short an exposure as practical (down to 1 second) without using a cassette.
	Enzyme conjugate may have lost enzymatic activity	Determine if the enzyme conjugate is active.
High Background	Too much antibody or protein conjugate	Perform a titer of the antibody or protein conjugate until an acceptable signal-to-noise ratio is obtained.
	Inappropriate blocking reagent	Increase the concentration of the blocking reagent by preparing the reagent with one-half the recommended volume of water. In addition, some antibodies may crossreact with certain blocking reagents. To test for this possibility, prepare a “blank” membrane that does not contain the primary antibody.
	Inappropriate blocking protocol	Increase the blocking time and increase the blocking temperature to 37 °C.
	Inappropriate wash protocol	Increase the number of washes.
	Overincubation in colorimetric substrate solution	Decrease the staining time. The membrane should be exposed to the colorimetric substrate until a positive signal is obtained, but as the background begins to develop, the reaction should be stopped. For colorimetric substrate: Incubate for 5-10 minutes or whenever bands are visible. The time required may be increased or decreased, but should not be longer than 60 minutes. To stop reactions: For alkaline-phosphatase substrates, use a solution of 0.3 M sodium phosphate, pH 5.5. For horseradish-peroxidase substrates, wash the membrane with 0.1% sodium azide with 1% SDS in either TBS (Tris-buffered saline) or PBS (phosphate-buffered saline).
	Inappropriate film	Switch to film designated for chemiluminescent detection such as Kodak® Biomax® Light, MS, and MR film.
Extraneous spots	Aggregated protein or antibody conjugate	Filter the conjugate through a 0.2 micron cellulose acetate filter or centrifuge the conjugate solution at 10,000 x g for 10 minutes and use the supernatant.

References for Southern Blotting

1. Sambrook J, et al.. 1989. Molecular Cloning: A Laboratory Manual. Second Edition. Cold Spring Harbor Laboratory Press, p. 9.47-9.50.

References for Western Blotting

1. Bjerrum O, Heegaard N. 1988. CRC Handbook of Immunoblotting of Proteins. Second Edition. CRC Press, p. 229-236.

2. Dunba B. 1994. Protein Blotting: A Practical Approach. NY: IRL Press, p. 67-70.

3. Fortin A, Berkova N, Tremblay S, Côté F, Rousseau F, Khandjian E. 1994. A 56- to 54-kilodalton non grata signal in immunoblot analysis using the horseradish peroxidase chemiluminescence system. Biochem. Cell Biol.. 72(5-6):239-243. https://doi.org/10.1139/o94-034