Staining cells growing on coverslips
From Diseasepedia.net
No2. Staining cells growing on coverslipsDection limits is approximately 1,000-10,000 molecules/cell if the antigen is localized.
Summary
Rapid and easy
Detects antigen presence and localization
Qualitive to semi-quantitative
Sensitivity dependent on minimal level and localization of antigen
High local concentration of antigen, so lower affinity antibody OK
Caution
Ethanol, FITC, fluroscein, paraformaldehyde
1. Sterilize glass coverslips prior to cell staining. Place round coverslips (grade #1 or #1.5) in 70% ethanol.
Remove individually and flame to sterilize.
2. Aseptically transfer the coverslips into a sterile tissue-culture dish by using a sterile pasteur pipet connected to a vacumn line. A light vacumn flow through the pipet will allow the coverslips to be picked up and moved easily.
3. Plate the cell suspension in the tissue-culture dish at low density. Grow overnight.
4. Remove the cell suspension in the tissue-culture media by aspiration and wash the coverslips once with PBS. Sterility is not needed at this or later steps.
5. Asprate the wash buffer and fix the cells by adding 4% paraformaldehyde. Incubate for 10 minutes at room temperature.
6. Remove the paraformaldehyde by aspiration and wash the cells twice with PBS.
7. Aspirate the last wash buffer and permeabilize the cells by adding 0.2% Triton X-100 in PBS. Incubate for 5 minutes at room temperature.
8. Remove the detergent solution by aspiration. Wash the coverslips in 0.2% Triton X-100 in PBS with three changes over 5 minutes. Drain well but do not allow the specimens to dry. Move each coverslips to the well of a 24-well tissue-culture plate.
9. Add the primary antibody solution to the coverslips. Use 25 ul to cover the entire coverslip. Make sure the coverslips does not touch the side of the well.
Monoclonal antibodies are most often used as undiluted tissue-culture supernatants. Ascites fluids, purified monoclonal and polyclonal antibodies, and crude polyclonal sera need to be checked for the proper dilution prior to use. To determine the proper titration of the antibody to use, test 1/10, 1/100, 1/1000, and 1/10,000 dilutions of the starting antibody solution. All dilutions should be done in 0.2% Triton X-100 in PBS containing 3% BSA.
Each assay should include three controls: (1) An irrelevant antibody from the same species and type as the primary antibody to determine the specificity of the staining. (2) A sample with no primary antibody to test for the background of the labeled secondary antibody. (3) If possible, a known control against a positive.
Incubatefor 60 minutes at room temperature.
10. Wash the coverslips in three changes of 0.2% Triton X-100 in PBS over 5 minutes. After the last wash, drain well but do not allow the specimens to dry.
11. Apply the labeled secondary reagent. Add 25ul.
A useful secondary reagent is anti-immunoglobulin antibodies conjugated to FITC. All labeled secondary reagents need to be checked for the proper dilution prior to use. To determine the proper titration of these reagents to use, test 1/10, 1/100, 1/1000, and 1.10,000 dilutions of the starting material. All dilutions should be done in 0.2% Triton X-100 in PBS containing 3% BSA.
Incubate for 20 minutes at room temperature.
12. Wash the coverslips in three changes of 0.2% Triton X-100 in PBS over 5 minutes. Drain well.
13. On a clean and labeled microscope slide place a drop (approximately 50 ul) of mounting medium.
Remove the coverslips from the dish using fine-tipped forceps and drain the last of the wash buffer by touching the edge of the coverslip to a clean paper towel. Invert the coverslip and place on the dropr of mounting medium with the cell side down. Gently lower the coverslip, touching one edge to the slide next to the drop. Then allow the coverslip to fall on the drop. This will push any bubbles ahead of the falling coverslip. Allow to air dry for at least 30 minutes prior to observing.
14. Observe and photograph under the fluorescence microscope.
