Hi, I am Victor Chi and I'm a graduate student in the Chandy Lab in the Department of Physiology and Biophysics at the University of California Irvine. Today I'm gonna demonstrate how to stain paraffin embedded sections using the A b, C kit from Vector Laboratories. This procedure involves de waxing and hydrating paraffin embedded sections, performing heat induced antigen retrieval to unmask antigens, block peroxidase, and non-specific protein binding, incubating with CD four and tyrosine hydroxylase and secondary antibodies, and developing and mounting the slide.
So let's get started. Since the slides are coded in paraffin, the first step is to dewax the slides. This is done usually in a fume hood using three jars of xylene after three washes consisting of five minutes for each wash.
The slides are then put into a hundred percent ethanol. You'll notice that the wax on the slides are gone, but be careful not to handle the slides with gloves because the xylene will dissolve the rubber of the gloves. From there, you carry over the slides in the a hundred percent ethanol back into the lab where they will undergo a hydration gradient.
So here I'm gonna talk to you about the hydration gradient that will be putting our slides through. It first starts with two baths of a hundred percent ethanol. From there, we transfer our slides to two baths consisting of 95%ethanol.
Then we transfer our slides to one bath of 70%ethanol, and from there we'll transfer our slides to a bath of 50%ethanol. After that, we transfer our slides to 30%ethanol, and two minutes later we'll transfer our slides to double distilled water. And upon treatment with double distilled water, we will let our slide sit in PBS.
So now I'm gonna show you how to carry the slides from one bath to the next in the carriages. So the first step is to using your carriage holder, remove the slides and scrape gently to remove excess liquid so that you're not carrying too much liquid on over to your next bath. From there, submerge it in and wait two minutes till your next gradient change.
So many times when the tissues fixed, the antigen becomes masked so that the antibody can't bind to it. So what I'm gonna do now is show you a method of heat induced antigen retrieval using sodium citrate. So now we're here at the microwave where I'm going to unmask my CD eight antigen, of which I will try to detect using immunohistochemistry.
And here I'm going to put my solution in and microwave it for 15 minutes. So we'll boil our sodium citrate with our slides for 15 minutes to make sure that our slides don't dry out. We'll be watching our slides so that upon boil over we'll refill with sodium citrate to make sure the slides remain hydrated.
So before I put my slides in the sodium citrate, I've already prewarm my solution for 45 seconds. Then I've put my slides into the sodium citrate, and now I'm heating my sodium citrate for 15 minutes. With the slides in it, usually the solution will begin to boil over at about two to two and a half minutes into the boiling process.
At that point, I will stop and replace the sodium citrate lost with fresh sodium citrate. After boiling the sodium citrate for 15 minutes, we wanna remove our our slides and solution and cool them. And so the best way to do this is to cap your slide holder, pour off the excess liquid, and just run it over a stream of cold water until the solution inside the jar is cooled to room temperature.
And this usually takes about 10 to 15 minutes. So after removing our slides from the sink, we need to block the endogenous peroxidase activity in the tissues. So by, so what we're gonna do is make a solution of hydrogen peroxide utilizing 30%hydrogen peroxide and making up a 1%hydrogen peroxide solution.
And from here, I just transfer the slides directly into the hydrogen peroxide solution. And now we'll incubate our slides in the hydrogen peroxide for 20 minutes. Now, after 20 minutes have passed, after blocking the endogenous peroxidase, I'm gonna now wash the hydrogen peroxide off the slides using washes of PBS.
So I'm gonna transfer my slides to the carriage and wash it with three washes consisting of five minutes each. So these slides have been placed back to back to avoid contact with the other slides that may damage the tissue. So I'm merely sliding off the slides and I am arranging the slides so that the tissue has no possibility of contacting the glass.
There we transfer our slides to PBS. So after washing our slides with PBS, I'm now gonna block the slides for non-specific binding of proteins with a solution of 5%BSA 5%goat serum and 0.1%sodium azide. So you want to change the goat serum to another species if you're working with goat antibodies.
So right now, I'm just gonna put the slides into our slide holder and fill it with our solution. So normally I just fill it halfway so that the volume of of the slides will just make sure that the tissues are submerged. So one thing you want to make sure of when putting the slides in the container is to avoid these tissue from touching the sides of the container.
So for the last slide, rotate it inward so that the tissue doesn't touch the container. So now that I've blocked my slides overnight, I'm gonna wash off the excess blocking solution with three washes of five, consisting of five minutes in PBS. So at this point, we want to draw a circle using liquid blocker around our tissue so that we would minimize the amount of antibody used.
If you're not worried about conserving antibody, then you can always incubate the slides in the same casings that we did the blocking step in. So from this point on, I'm gonna show you how we draw with liquid blocker around our tissue. So first we remove blood off the excess liquid, careful not to touch the tissue.
So just wipe around at the corners to remove excess liquid. Again, being very, very careful to not touch the tissue. Take the liquid blocking pin, press down gently get the flow going, and from there, encircle the tissue with your pen.
Allow the liquid blocker to dry for about 15 seconds. At this point, we want to add the primary antibody. And so using a pipette, make sure you cover the tissue with your primary antibody and transfer it to your humidity controlled box.
So we use a Kim wipe. Wed wedded with some di water in this box to prevent the slide from drying out as it's incubating with the primary antibody. Now that our primary antibody has been incubating for an hour, we're gonna remove our slides and wash off the excess antibody with PBS.
So again, just allow the liquid to drain and insert it into your PBS wash. And again, wash three times for five minutes. So after our three five minute washes, we're gonna add our secondary antibody to our tissue in the same fashion that we added our primary antibody.
So just using enough volume to cover your tissues, add your secondary antibody, which we're using is goat anti rabbit, and move your slide back into the humidity controlled chamber. And so we're using a secondary antibody, which is biotinylated goat anti rabbit, which is essential when you're using the A BC method. So we transfer our slide into the humidity controlled chamber and allow that to incubate for one out.
So while the secondary antibody is incubating, we need to prepare the A BC reagent. So we're using the vector laboratories, A, B, C reagent from the vector stain kit. So remove reagent a prime it, and allow it to drop three times into 15 milliliters of PBS being cautious not to squeeze the tube because the drops have to be specifically metered and of a specific size.
So after your secondary antibody has been done incubating for about an hour, you take your slides and you put, you wash it again in PBS, just like you did with your primary antibody. So after three washes consisting of five minutes each, you then transfer your slides to this container where you add in your A B, C reagent, which you prepared 30 to 45 minutes ahead of time. And so you just add your A, B, C reagent in here and fill your slides in and allow that to incubate for another 30 to 45 minutes.
So after your slides have been sitting in a, B, C for about 34 to five minutes, we wanna wash off the excess A, B, C solution. So you just remove your slides and add into your PBS solution. And so we wanna repeat the same washing procedure three times in PBS for five minutes each wash.
While the slides are washing, I prepare the DAB peroxidase substrate from vector laboratories. I take the pH 7.5 buffer and I add two drops in. After the pH 7.5 buffer, I take the DAB substrate react reagent, and I add two drops of that in.
Again, the drop size is critical, so don't squeeze the bottle and just allow the drops to fall. Two drops for buffer pH 7.5. So you want to add four drops of the DAB substrate reagent, allowing it to also drop.
You want to also add at the end, you want to add two drops of the hydrogen peroxide substrate reagent. So this is how the DAB substrate reagent mixture should look. Now it's ready to add to the slides, remove your slides, and allow the PBS to just drain off your slides.
So over time, the DAB reagent becomes more brown. So we want to add it to our slides at this point. So take enough volume to cover the tissues and just add it directly onto the, onto the tissue samples.
So just take the appropriate volume and add it directly on top of your tissue samples, and allow the solution to develop as your tissue samples should become more brown after time. After adding your DAB solution, you could see in this brain section where the tissue turns brown here and here. The brown DAB standing reveals the region containing midbrain dopaminergic neurons.
After developing your slides from anywhere from two to 10 minutes, you want to add your, so you want to add your slides to the ice cold water to stop the reaction. So just allow the solution to drain off and add your slides directly into the ice cold water. After immersing your slides in the ice cold water, take your bath to the sink and run cold water into it for approximately five minutes to assure that all the DAB solution is drained out from this bath.
So when you add in the sink, be cautious to not let the stream hit the tissue and have it rather hit one of the corners and allow it to just overflow so that your tissues aren't disturbed by the streaming flow of water. So after removing your slides from the wash, you want to do a counter stain. So why might you want a counter stain?
Well, one reason is that you might wanna see the structure of your tissue. Another is to visualize all the cells. So for our purposes, we're gonna counter stain with hemat toin, which stands for nuclei.
So we want to just take our slides, remove excess liquid, and just dunk it once in the Hemat Hematin solution. One, take it out and add it directly into the water. After adding your slides in the tap water, you want to take your slides and run it under water, just as you did previously with the ice cold water bath.
So for the hemat Toin wash, just allow it to run under running water until the water inside the jar becomes clear. So after removing our counter stained slides from the water, we want to re dehydrate our slides. And so we want to put it back through our ethanol gradient, but in reverse order.
So that would be one wash with deionized water, one wash with 30%ethanol, one wash with 50%ethanol, one wash with 70%ethanol, two washes, and 95%ethanol, and two washes, and 100%ethanol each consisting of two minutes each. So remove your slides, wipe off the excess liquid and add it directly to your 30%ethanol, and allow it to sit for two minutes. So after the last a hundred percent ethanol wash, we want to put the slides back in the xylene.
So take your slides again, scraping the excess liquid off, take the slides and add em into the first xylene for five minutes. So that will be three consecutive xylene washes for five minutes each. And now we're ready to mount our slides.
So what I usually do is one minute prior to the slides being ready, I take my pasture pipette and I draw the perm mount solution, and I allow that to go approximately one minute before my slides are ready to come out After the last wash of xylene, being careful to not submerge your gloves or get your gloves wet with xylene, remove them with a pair of tweezers being careful which side contains the tissue. Gently wipe off the excess of XLE with a Kim wipe, being careful not to touch the tissue, just wipe carefully at the corners and the edges, allowing most of the xylene to evaporate. You can probably work on two slides before one dries out.
So as you're done drying off one slide, the other slide should be ready to work with. Now you have to work with them before they completely dry out. So you wanna take your perimount and draw a thin line next to your tissue, being careful not to touch your tissue.
Remove the perimount taking your cover slip. Apply it at a 45 degree angle and allow it to bend down. And then slide your cover slip, making sure that the perimount covers all the tissues.
Then lay the, lay it down and gently press at the center to remove all excess air bubbles. Allow it to dry while working on your second tissue is the time to look at our slides, not actually. So we need to allow the perma mount to dry overnight before looking at our slides.
Otherwise, under the light of the microscope, the perma mount will bubble, and then you'll ruin your sample. Here's a look at our brain sections after staining with chemicals, rabbit polyclonal antibody directed against tyrosine hydroxylase. Again, the brown DAB stain shows the location of midbrain dopaminergic neurons.
The hematin counter stain turns all of the nuclei in this tissue blue. So I've just shown you how to stain paraffin embedded sections utilizing the A BC kit from Vector Laboratories. Now you'll wanna adjust the conditions according to the tissue types that you'll be staining and the antibodies which you'll be using.
So good luck on your experiments.
