Rosen lab website http://www.bcm.edu/rosenlab last modified August 31, 2005

Rosen Lab Immunostaining Tutorial By Sandy Grimm, PhD

Place your slides in a glass or metal slide holder, pictured at right. The glass holder will fit 10 slides and the metal holder will fit 30 slides. You will also need at least three staining dishes to wash/rinse the slides during later steps. We have a dedicated hood set up with all the reagents needed to deparaffinize, rehydrate, dehydrate, or stain for eosin and hematoxylin (H&E), as pictured below.

The first step for immunostaining is to remove the paraffin from the sections on the slide, by immersing the slides in three changes of Xylene for 3 minutes each. Blot any excess Xylene from the slide holder using paper towels, and place the slides in three changes of 100% ethanol for 3 minutes each. Next, rehydrate the sections by placing the slides in 95%, 80% and 70% ethanol for 2-3 minutes each. Finally, place the slides in 1X PBS for ~5 min. The next step for immunostaining is to perform antigen retrieval to reverse the cross-links formed by fixation (we fix our tissues 2 hours in 4% paraformaldehyde). The most common method we use is to microwave the slides in a solution of 10 mM sodium citrate pH 6 for 20 minutes. With the slides in a glass slide holder, place the whole holder in a 2L glass beaker containing 600 ml of 10 mM sodium citrate pH 6. Be sure to remove the metal handle. If you have less than 10 slides, fill the extra space with blank slides. This is to ensure even heat distribution for every experiment. Mark a line at the top of the buffer before microwaving. Heat the beaker in the microwave for 10 minutes and replace any evaporated water (filling to the line with a squirt bottle of water). Microwave for another 10 minutes. Remove the beaker from the microwave and place on your bench to cool for 20 minutes.

Rosen lab website http://www.bcm.edu/rosenlab last modified August 31, 2005

Rosen Lab Immunostaining Tutorial, p.2 Wash the slides in 3-4 four changes of distilled water for 5 minutes each. Wash once with 1X PBS for 5 minutes. An optional step is to incubate the sections in 3% hydrogen peroxide to block endogenous peroxidases, which could give a false positive signal when incubated with DAB. The 30% stock of hydrogen peroxide can be diluted with either 1X PBS or methanol. Place the slides in a glass slide holder and incubate 10-15 minutes in a staining dish containing ~300 ml of 3% H2O2. Wash three times with 1X PBS for 5 minutes each. Circle each tissue section using a Pap pen, which will leave a slight green cast. This is done so different antibodies can be added to adjacent sections and to keep the volume of antibodies as small as possible.

Immediately after circling each section, add blocking buffer to the sections. Usually 50 µl of solution is enough to cover the sections. The blocking buffer you use depends on the antibody, but the most common ones we use are:

1. 5% BSA + 0.5% Tween-20 in PBS 2. 5-10% normal goat serum in PBS (or whichever species the secondary antibody was raised in) 3. The M.O.M. kit blocking reagent (Mouse on Mouse kit for monoclonal antibodies)

Place the slides in a humid chamber (to prevent evaporation) and block for anywhere from 1 to 5+ hours. We purchased our slide boxes (which hold 10 slides) from Evergreen Scientific. http://www.evergreensci.com/ch/index.htm Aspirate or decant the block solution from the slides and add ~ 50 µl of primary antibody, diluted in the appropriate diluent, to each section. Place in the humid chamber overnight at room temperature or 4ºC. Aspirate or decant the primary antibody from the slides and place the slides in a glass slide holder. Using three staining dishes containing ~250 ml of liquid, wash three times with 1X PBS for 10 minutes each. The secondary antibody used will depend on the primary antibody. Most of our antibodies are rabbit polyclonals, so we use a biotinylated goat α-rabbit secondary antibody. Dilute the secondary antibody anywhere from 1:200 to 1:1000 in the same diluent used for blocking and/or primary antibody. Carefully wipe any excess PBS from slides and add ~ 50 µl of secondary antibody to each section. Incubate in the humid chamber for 1 hour at room temperature. Aspirate or decant the secondary antibody from the slides and place the slides in a glass slide holder. Use the same three staining dishes containing 1X PBS to wash the slides for 3 times 10 minutes each.

Rosen lab website http://www.bcm.edu/rosenlab last modified August 31, 2005

Rosen Lab Immunostaining Tutorial, p.3 While the slides are washing, prepare the ABC reagent (Vector Labs). This contains an avidin-conjugated horse radish peroxidase (HRP) enzyme that will bind to the biotinylated secondary antibody. Add one drop of solution A and 1 drop of solution B to 2.5 ml of 1X PBS (in a 15 ml conical tube), vortex to mix well, and let sit at room temperature for at least 30 minutes. Any extra can be stored at 4ºC and used within 1-2 months. You can also buy the Ready-to-use version (RTU) of Vectastain Elite ABC reagent from Vector Labs. Carefully wipe any excess PBS from slides and add ~ 50 µl of ABC solution to each section. Incubate in the humid chamber for 30 minutes at room temperature. Aspirate or decant the ABC reagent from the slides and place the slides in a glass slide holder. Use the same three staining dishes containing 1X PBS to wash the slides for 3 times 5 minutes each. Prepare the DAB solution fresh just before use. DAB, or 3,3´-diaminobenzidine, is a known carcinogen, so use with caution and wear gloves. The DAB reacts with the HRP and forms a brown precipitate. To 2.5 ml of ddH2O, add 1 drop of buffer, 2 drops of DAB, and 1 drop of H2O2 and vortex to mix well. Carefully wipe any excess PBS from slides and add ~ 50 µl of DAB solution to each section. Place the slides on a white piece of paper and incubate at room temperature until the tissue starts turning brown. The reaction maxes out at 10-15 minutes. The staining can also be monitored using a microscope. When I optimize the primary antibody dilution, I try to use the least amount of antibody and let the staining go for the full 10-15 minutes, so there is less guess-work and variability between experiments. Decant the DAB reagent from the slides onto a folded paper towel and, using a glass slide holder, place the slides in a staining dish containing distilled H2O for at least 5 minutes. Place the paper towel, pipet tip and 15 ml conical tube that contain DAB in a small plastic container and add a 10% bleach solution to destroy the DAB. The next step is to counterstain the nuclei with Harris hematoxylin. Since hemotoxylin oxidizes when exposed to air, skim the surface of the solution in the staining dish with a Kimwipe to remove any particulate matter. In addition to the distilled water dish, set up two more dishes containing tap water. Blot any excess water from the slide holder and dip the slides in hematoxylin for 3-5 seconds. Rinse the slides in distilled water and place in tap water for 5 minutes to let the stain develop. Change the water in the distilled water dish. Blot any excess water from the slide holder and dip the slides in acid ethanol (70% ethanol + 0.25% concentrated HCl) for 3-5 seconds. Place the slides in tap water for 2 min, then in distilled water for 2 min. Now the slides are ready to be dehydrated. Note that these ethanols and xylenes are different from the ones used to deparaffinize and rehydrate. Place the slides in 95% ethanol, 2 x 5 min. Next is 3 x 5 min in 100% ethanol. Blot any excess ethanol from the holder and place slides in Xylene for 3 x 15 min each. Now that all the water has been removed from the sections, the slides can be mounted for viewing. If all the water has not been removed, there will be bubbles when you coverslip. We use Permount (Fisher Scientific) to mount the slides and 22x40 mm coverslips. Place some Permount in a small beaker and use a glass rod to place a few drops of Permount on the surface of the slide. Angle a coverslip over the slide and gently and slowly drop on the slide trying to avoid any bubbles. Press the long edges against paper towels to remove any excess Permount. Leave the slides in the hood until the Xylene has evaporated and the slides are now ready to view under the microscope.