Antibody Protocols

Protocol:

1. Block membrane by incubating 1 hour at room temperature with shaking in Blocking Solution (5% nonfat milk in TBST (50mM Tris, 100mM NaCl, 0.05% Tween-20, pH 7.6).

Note: Use 5% BSA in Blocking Solution for phospho specific antibodies.

2. Dilute primary antibody at the appropriate dilution in Blocking Solution.

3. Incubate the membrane with diluted primary antibody overnight at 40C with agitation.

4. Remove antibody solution. Wash the membrane 3 times for 5-10 minutes each time at room temperature in TBST (50mM Tris, 100mM NaCl, 0.05% Tween-20, pH 7.6) with shaking.

Note: Increasing the concentration of Tween-20 to 0.1% reduces the background and increases the specificity, but it will reduce the sensitivity.

5. Incubate membrane with secondary AP or HRP conjugate diluted (according to manufacturer’s instructions) in Blocking Solution for 1 hour at room temperature with shaking.

6. Wash the membrane as in Step 4.

7. Wash membrane with TBS for 2-5 minutes before proceeding Chemiluminescent Reaction.

8. Prepare and use the Chemiluminescent substrate (for AP or HRP) according to the manufacturer’s instructions.

9. Immediately wrap the membrane and expose to X-ray films for 10 second to 1 hour period. The exposure time may vary according to the mount of antibody and antigen. Or place into imager and adjust accordingly.

Deparaffinization

1. Incubate slide at 60 C for 60 minutes.

2. Deparaffinize in Xylene for 10 minutes and repeat one more times.

3. Hydrate in 100% alcohol for 5 minutes, next in 95% alcohol for 5 minutes, next in 85% alcohol for 5 minutes, next in 75% alcohol for 5 minutes.

4. Dip into distilled water for 5 minutes

5. Dip into TBS (50 mM Tris, 100 mM NaCl, pH 7.6), leave for 5 minutes, and repeat two times.

Antigen retrieval

1. Bring 500-2000 ml 10 mM citrate buffer (pH6.0) to the boil in a stainless steel pressure cooker.

2. Put the slide into staining rack and lower into pressure cooker ensuring the slide is well immersed in citrate buffer.

3. When the pressure indicator valve has risen after 3-4 minutes, incubate for 1 minute.

4. Cool the slide naturally to room temperature.

5. Dip into distilled water, leave for 5 minutes, and repeat two times.

6. Dip the slide in TBS for 5 minutes and repeat two times.

7. Immerse slides in 3% H2O2 (in fresh methanol) for 15 minutes at room temperature.

8. Wash with distilled water two times, 5 minutes each time.

9. Wash with TBS (pH 7.6) two times, 5 minutes each time.

Staining with Primary Antibody

1. Dilute primary antibody with 3% BSA in TBS. Cover the tissue section on the slide with diluted primary antibody (use 50-150 μl for each slide).

2. Incubate at 37C for 30 minutes or at room temperature for 60 minutes (The optimal incubation time, incubation temperature, and antibody dilution should be determined by the individual laboratory).

3. Wash with TBS two times, 5 minutes each time.

Staining with Secondary Antibody

1. Incubate with 100-200μl Polymer Enhancer Incubate 30 minutes at 37C

2. Wash with TBS for 3 times, 5 minutes each time.

3. Incubate with 100 200μl Polymerized HRP and incubate 30 minutes at 37C

4. Wash with TBS for 3 times, 5 minutes each time.

5. Add dAb solution and incubate 3-10 minutes (The reaction progress and the optimal time should be determined accordingly with microscope).

6. Wash with distilled water for 2 times, 5 minutes each time.

7. Counterstain sections in hematoxylin if required,wash with distilled water. Immerse slides in 0.1% HCl ethanol for 1-10 seconds, wash with distilled water.

8. Dehydrate through 95% ethanol for 1 minute, 100% ethanol for 2×3min, Xylene for 2×3min, and coverslip with mounting medium.

Materials

• 10X PBS: To prepare 1L, add 80g NaCl, 2g KCl, 2g KH2PO4 and 28.5g NaHPO4 to 1L water for injection. Adjust pH to 7.4.

• 4% Polyoxymethylene: To prepare 100mL, add 4g polyoxymethylene to 100mL 1×PBS. Adjust pH to 7.4.

• 1×PBS/0.2% Triton X-100(PBS/Triton): To prepare 500mL, add 1mL Triton X-100 to 500mL 1× PBS.

• 1×PBS/3% BSA(PBS/BSA): To prepare 100mL, add 3g BSA to 100mL 1× PBS.

Protocol

     Preparation Fixation permeabilization

1. Rinse cells briefly in PBS.

2. Aspirate PBS, cover cells to a depth of 2-3mm about 200ul with 4% polyoxymethylene.

3. Allow cells to fix for 15 minutes at room temperature.

4. Aspirate fixative, rinse three times in PBS for 5 minutes each.

5. Aspirate PBS, cover cells to a depth of 2-3mm about 200ul with PBS/Triton for 5 minutes at room temperature.

6. Aspirate permeability agent rinse three times in PBS for 5 minutes each.

Immunostaining

1. Gently add 200ul of primary antibody diluted in PBS/BSA to the 24 well plates each well.

2. Incubate 60 minutes at 37℃ or overnight at 4℃.

3. Aspirate diluted primary antibody, then rinse three times in PBS for 5 minutes each.

4. Incubate in fluorochrome-conjugate secondary antibody diluted in PBS/BSA to the 24 well plates 100ul each well for 30 minutes at room temperature in dark.

5. Aspirate diluted fluorochrome-conjugate secondary antibody, then rinse three times in PBS for 5 minutes each.

6. Test under fluorescenct microscope.

Materials:

• Coating Solution: Antigen or antibody is diluted in coating solution for immobilization onto the microplate. Commonly used coating solutions are: 50 mM sodium carbonate, pH 9.6; 20 mM Tris HCL, pH 8.5; or 10 mM pBS, pH 7.2. A protein concentration of 1-10 ug/mL is usually sufficient.

• Blocking Solution: Commonly used blocking agents are: BSA, nonfat dry milk, casein and gelatin. Different assay systems may require different blocking agents.

• Primary/Secondary Antibody Solution: Primary/secondary antibody should be diluted in 1x Blocking solution to prevent non specific binding. It is recommended to dilute antibodies between 1:100 and 1:500. Follow the manufacturer’s advice for secondary antibodies.

• Wash Buffer Solution: Typically 0.1 M phosphate buffered saline or Tris buffered saline (pH 7.4) containing a detergent such as Tween 20 (0.02% 0.05% v/v).

Protocol:

1. Dilute the antigen to 1-2 ug/ml in coating solution

2. Add 100 ul of diluted antigen to appropriate wells. Incubate 2 hours at room temperature or 4 °C overnight.

3. Empty plate and tap out residual liquid.

4. Wash twice with 300 ul Wash solution.

5. Add 300 ul Blocking solution to each well. Incubate 1 hour.

6. Empty plate and tap out residual liquid.

7. Wash twice with 300 ul Wash solution.

8. Add 100 ul diluted primary antibody to each well. Incubate 1 hour at 37 °C or 3 hours at room temperature.

9. Empty plate, tap out residual liquid.

10. Fill each well with Wash solution. Invert plate to empty, tap out residual liquid. Repeat 3 times.

11. Add 100 ul diluted secondary antibody to each well. Incubate 1 hour at room temperature.

12. Empty plate, tap out residual liquid and wash as described in step 10.

13. Give final 5 minutes soak with Wash solution. Tap residual liquid from plate. This washing step is critical to reduce signal background.

14. Fill each well with Wash solution. Invert plate to empty, tap out residual liquid. Repeat 5 times.

15. Dispense 100 ul of substrate (e.g. pNPP, TMB) into each well. Develop the color for 30 minutes.

16. Stop reaction if necessary and read plate with plate reader at the correct wavelength.