The H&E Stain – A Lesson on Consistency and Reproducibility
Producing an H&E stained slide is a process. It starts way before the slide is loaded on a stainer or moved down through containers by hand. Producing quality, consistent and reproducible H&E stained slides is a process as well as a lesson in standardization. In this presentation, we will look at the process and learn the standardization necessary to have consistent and reproducible H&E slides.
- Review the many factors that influence the quality of an H&E stain.
- Describe how an H&E works.
- Explain what makes a regressive H&E stain different from a progressive one.
- Review what experts suggest in order to maintain consistent and reproducible H&Es.
So in this lesson, we will review the many factors that influence the quality of an H&E stain, and we’ll see what we can do to reduce the number of those factors. We will describe how an H&E works so that we will not make errors based on lack of knowledge. We will explain what makes a regressive H&E stain different from a progressive one. We’ll look at the potential caveats of each, and maybe determine which one might work best for you. We will review what experts suggest in order to maintain consistent and reproducible H&Es and perhaps take away some good suggestions for reaching each of our goals of consistency and reproducibility.
Just to enlighten you as to where my experiences and knowledge were gained over the years, I made this little map so that the red indicates states and/or countries in North America where I have been in hospitals either training, setting up staining solutions, or troubleshooting staining. As you can see, I’ve covered the continental USA, and each of those red dots, and in each of those red states, I may have been in as many as a dozen places, or maybe more than a dozen places. But the last 13 years I lived on an airplane.
Key statements regarding H&E
In my past 40 years interacting with pathologists and technologists all over North America, whether it would be in clinical, research, or the industry, I’ve been told many times that there are some general statements that can be made regarding the H&E. These are not the only statements, by far, that could be made, but they seem to be the most common.
First, there’s a statement that I think every one of us will agree with; the H&E is by far the most often-requested stain by a pathologist. Second, I think most of us would agree that there are many factors that influence the quality of the H&E, even before the slide reaches the staining platform. Will you personally agree that the H&E is possibly the most misunderstood and abused stain in histology?
Well, there are a lot of people who do, and what they believe is that it stems from lack of experience or knowledge of the chemistry involved in an H&E. Last but not least, the dreaded one, the difference between a good and a bad H&E could simply be the preferences of the pathologist looking at it, and that is by no means putting the pathologists down at all, because after all, the education they’ve had, they really deserve to have a slide to look at that they like looking at. However, there may be some compromises that have to be made.
So let’s look at each one of these statements a little closer. The H&E is by far the most often-requested stain by a pathologist. Basically all this slide is saying to us is that there’s a lot of hospitals in the U.S. Not all of them do H&E stains, but most of them do. But this cites a study that was done jointly by the College of American Pathologists and NSH, looked at different sizes of hospitals and independent laboratories, and basically came up with a number, 1.8 H&E slides are cut per block. So again, I don’t think any of us will disagree that there’s a lot of H&Es out there.
Again looking at this, many factors that influence the quality of an H&E stain. Some happen before they get to the staining platform, and it actually starts with the removal of tissue from the body. There’s some things that we can control, and other things that we have no control over. For instance, if you have a surgeon that crushes the specimen with forceps; if the tissue is allowed to lay out and dry before it’s put in a fixative; there’s cauterized samples; necrotic material that wasn’t even viable to begin with.
So once we get the tissue, hopefully we are getting it in formalin or some other fixative. Fixation is important, really important, in the process to quickly stop all cellular activity, including deterioration. When done properly, the tissue should be resistant to all processing that follows. There are important factors to consider in fixation. You need to select a proper fixative. You need adequate time in the fixative. You need a correct ratio of fixative volume relative to the size of the tissue. In other words, you don’t want to put a whole uterus and cervix in 5 ml of formula and expect it to fix properly.
If you look at this work of art here, you will see that it represents good fixation before dehydration. You will see that we have free water represented by the black circles, and bound water represented by the red circles. You can see that there is a strong matrix that’s present being formed by these hydrophobic bonds, creating a strong architecture of the tissue. This is what you want.
Considerations regarding dehydration
After fixation it’s important that processing protocols are critical to the size and nature of tissue relative to their size and nature. In other words, gastric biopsies probably don’t want to be processed with your large, fatty specimens like breast tissue and colon tissue. Proper dehydration works by diffusion, which is the inward passage of alcohol and outward passage of water. At some point we’re going to achieve equilibrium and we’re only going to have a little bit of free water in the tissue. We’re going to have our bound water still in place, creating those bonds that are going to keep our matrix strong. If we under-dehydrate, there’s water left in the tissue. If we over-dehydrate and remove the bound water, then we might end up with something that I like to call “crispy critters.”
This picture represents, in the top left-hand corner, it represents good dehydration. We’ve got that little bit of freestanding water, and we’ve got all of our bound water, and if you look to the right, what you have is a nice, clean view of the tissue section. If we remove the bound water then what we’re going to see is that our matrix has broken down, and now we’re going to get a crushed view of the same sample.
This is an example of over-dehydration of a tissue sample. This looks like it’s a little gastric biopsy, some kind of little biopsy, and if you look, you can see the cracks are evident there. How many of you out there would be taking this block and putting it on a block of ice or some cold water, letting it soak for a little bit, and actually the first several hundred microns, maybe, are going to soak up some water, and if you carefully put your block back in your microtome and sneak up on it and capture that first four or five ribbons, then you are likely to get maybe a halfway decent representation of this block. However, you are going to end up with probably thick and thin sections, because as you soak that block the tissue expands, and you may have five micron sections, and you may have ten micron sections on the same slide.
The other thing that this slide indicates is a situation called basophilia, and that is when the slide tends to take up more of the hemotoxin than it normally would. You end up with this purple hue all over the slide. But once we have dehydration properly achieved, then proper clearing needs to be performed. Up to this point, if you look through a thin slice of tissue as it comes out of the alcohol or the dehydrant, it will be opaque; you cannot see through it. After a few minutes in xylene or another clearing agent, you can hold the tissue up and the tissue will be transparent.
Proper clearing is intermediate step between dehydration and paraffin, in other words, we can’t go from alcohol, normally speaking, to our infiltrating medium paraffin and have infiltration to occur. So the clearing removes the alcohol, it renders the tissue transparent, and also hardens it. Keep that in your mind when you’re thinking about those pieces of cervix that you have that you just can’t get a section on. It may be that they are hard to begin with, for sure, but it also may be that they’ve been left in xylene a little bit too long. If the tissue is under-dehydrated and water remains, then proper clearing cannot occur. Therefore, good infiltration will not happen and you will end up with soft and slushy tissue that will need partial reprocessing.
Processing protocols critical to size and nature is extremely important, as we saw that tissue that was cracked. If all steps previous to infiltration are done properly, then you’re going to get good infiltration, you’ll have a strong matrix for sectioning, and I always like to kind of mention the thumb test. If you take your thumb and you place it on top of a block and you press lightly on it, if it’s well-processed, well-infiltrated and ready to cut, then it’s going to keep your thumb out of it and it’s going to feel firm. But if it’s that big piece of breast that was processed on a gastric biopsy protocol and you put your finger through it, then you’re going to sink right down into the tissue and it’s going to be kind of squishy and moist, and you are definitely going to want to reprocess that.
But you don’t have to take it all the way back to formalin; there’s no reason. Oftentimes this is mistaken for under-fixed or not-fixed, that’s what a lot of folks like to say at the embedding table. They will say, this tissue is not fixed; however, the tissue is not dehydrated. You’re not going to know if it’s fixed properly until you look at it under the microscope and see if you can see the nuclear detail well. But normally just taking it back, getting rid of the wax, go back through some alcohol, start it in some fresh 100 and bring it forward, there’s really no reason to take it all the way back to aqueous formalin and start all over again trying to remove all that water.
Considerations regarding microtomy
Now if we leave processing, which is super-important, as I think you can probably already see, for achieving consistent H&Es, and good H&Es. If we go to microtomy, we can have things like compression, chattering, blade marks, thick and thin. Thick and thin we talked about on that slide, with the one that we sneak up on after we soak it. Compression can be processing, under-infiltrated; it can be your room is too warm; a number of things. It could be your knife blade is dull. Chattering can be from over-hardened in xylene, or it can just be from a loose microtome. But as you’re looking there you can see that was my first microtome, and I am really hoping that you've got something that looks better than that.
This slide shows you what a well-prepared embedded tissue sample should look like. It may not be green, but it should look like this. It should sit in there, it should be the size it was when we started, and there it goes. We can do our thumb test and it’s firm. Slide A here represents a good section from the block. We should be able to take Slide A, place it over the top of that embedded tissue sample, and they should match identically in size. There you know that you are giving the pathologist the best representation you can of that piece of tissue. Slide B reflects either poorly processed tissue or perhaps some sectioning issue, because if you hold it over the well-embedded tissue sample, it’s going to be about half its size and you’re going to get that squished look, like we looked at earlier.
Common problems associated with H&E staining
We go to the third statement that H&E is possibly the most misunderstood and abused stain in histology. Again, you may be one of these folks that don’t believe that. Maybe you think that the Dieterle is the most misunderstood and abused stain in histology, but when we finish I’m hoping to show you what discord is out there as far as how to really get consistent in H&E and reproducible H&E.
Remember the red areas that I showed you on the map? Well, professionals all over that area have given me, over time, problems that are associated with H&E staining, and I made a short list of them. Here are some of them, and as you’re looking at these and thinking about these, imagine or remember if you have this problem, or have had this problem, and as we go through the rest of the presentation, I’m going to try to point out some of the causes of these problems and the remedies. Red or brown nuclei; hazy nuclei; nuclear ghosting, which means about the same thing as nuclear bubbling, or I can’t see anything in my nucleus; poor contrast of nucleus and cytoplasm; eosin leaching out of the tissue after you take it to the pathologist, of course; and the famous, the slides are always better toward the beginning of the week.
Here is more. After staining, there are unstained areas in the tissue; there are areas with eosin staining, but no hematoxylin staining. This a really cool one that I kind of want to stop and talk a little bit about, because I had this happen to me. There were these little round dots on my slide that I thought, what is this? I could see eosin staining in them, but I couldn’t see hematoxylin. I turned to a professional, an expert in this field, and I said, what is this? What it was, was the fact that paraffin molecules are round, and if you leave paraffin molecules in your slide, the hematoxylin cannot get through the paraffin because it’s an aqueous solution. However, the eosin can get through the paraffin because it’s an alcoholic solution. So if you ever see these perfectly round pieces of unstained except for eosin in your tissue, you know you've got paraffin on your slide.
Then there’s the pale nuclei; nuclei too dark, diffuse staining, or basophilia, like we saw in the over-processed, over-dehydrated slides; and the bottom sections on the slide are darker than the top ones; that too could come from that overly dehydrated biopsy.
Even after seeing the previous list of all the problems that can happen, plus there’s many more, more laboratories will say the H&E process is fairly straightforward, and I think we’ll all agree with that. You remove the paraffin, you get the tissue back to a hydrated state, because we know that in the staining process like environment likes like environment, and if we’re going into an aqueous hematoxylin, then we want to rehydrate the tissue to get to that aqueous hematoxylin so it will stain as good as it can stain. Staining the tissue nuclei with the primary stain, hematoxylin, it may involve differentiation, it may involve a bluing step. It probably will involve counter-staining, dehydration, and clearing.
Consistency of staining
The H&E process, fairly straightforward, yet so hard to maintain quality slide after slide. To get a better understanding, and I know most of you are probably experts in how an H&E works, but just in case there’s one or two people out there like I used to be that didn’t. Maybe you missed this chapter at school, but we’re going to review it to help us all, hopefully.
There’s a little general information that we’re all aware of, that after we cut our slides we’re going to put them in a staining rack, and we’re going to place them in an oven, preferably at 65-70 degrees centigrade. We’re going to dry them for about 15-20 minutes; this drying will stick the tissue down to the slide better, keep it from falling off during staining; it will also start the melting of the paraffin in and around the tissue sample.
One thing I would like to warn you about, is if you like to go to higher temperatures to possibly get your slides out of the dryer quicker, then you’ve really got to remember to drain your slides first, because if you saw that nuclear bubbling, nuclear ghosting, this can be where it happens. It’s usually caused by heat, and if there’s water under the tissue itself, then once it gets into that hot dryer it can start steaming and do damage to your tissue.
We’ll continue with deparaffinization in xylene, because we don’t finish the deparaffinization in the oven, we start it. Xylene is a popular substance to use. There are xylene substitutes. Really the only big difference is that xylene seems to be more robust than most of the substitutes, so you can get away with 2-3 changes, 3 if you can, because the first one’s going to get really loaded with paraffin. The second one will have a little carryover. The third one will be nice and clean to make sure that all the paraffin has been removed. If you do decide to use a substitute, then please maybe put an extra one in, maybe leave it a little bit longer, and for sure change it a little more often than you would your xylene.
Xylene removes any remaining paraffin left on the slides or in the tissue. At this point in time, the sections, if you hold the piece of tissue up and look at it, or hold the slide up, you should see that the tissue is actually transparent; you can hardly even see anything is on it.
Next comes hydration. We got our paraffin off of our slide, and we’ve got to work it down to water. How many of you know that xylene is a petroleum-based substance, it’s like an oil, and oil and water do not mix. Have you ever gone by your hematoxylin station and seen that there’s like an oil slick on top of it? That’s because you did not remove all the xylene.
For a long time I had no idea what the absolute alcohol did, or 95 really was for, because back in the day we used to go 100%, 100%, 95, 80, 70, 50, and then to water, because we were told in our early training classes that cells don’t like to get upset. If you make the cells upset by throwing them into water too quickly, that could just throw off your entire stain. Basically I believed for a long time that this was just stepping down so I could make my cells happy when they got to the water at the end. However, the real purpose of 100% absolute alcohol is to remove all the xylene. You’ve got to get all the xylene out before you take it down to water.
Any cyto-techs out there, I know that you still use that kind of soft, degrading alcohol down to water, and that’s okay because your cells might be more sensitive than histology cells. But we need to get rid of the xylene, so two changes, at least; one, it’s going to get saturated; the second one should ensure that all the xylene is gone. If we don’t get the xylene off and we get to 95, then we might see a little oil slick, as this representation shows, of oil and water. The 95 actually is the first indication of whether or not you have achieved removing all xylene, because what will happen is, if you have xylene on your slide, especially if you have a slide over 30 or 40 it’s going to multiply the effect.
When you go into your 95 you’re going to see this water, this milky precipitant, and it can go all the way down to water, turning milky. If you see that, you’ve got to back it up, replace your 100% to 95, and bring it back through until you do not see this. Leaving xylene on the slides at this stage can corrupt the stain, so it’s actually like staining over an oil slick, and your doctor may complain of muddy nuclei. When we get to the water and all the xylene is gone, then we want to remove all of the alcohol from the slides also by washing them well in running water.
When we go to our primary staining, the mechanism used for this, and we’re typically talking about hematoxylin because we’re doing H&E, a mechanism is absorption. Absorption just means a solution passing through another substance until they reach equilibrium. If you’ve ever picked up a slide out of hematoxylin and you look at it, you know it kind of looks blue all over; most hematoxylins, anyway. Hematoxylin is absorbed by the cell like spray perfume is absorbed by the skin.
Here’s where a little nomenclature comes into play; hematoxylin is a dye. It is the dye that’s extracted from the logwood tree in South America. If we were to mix that up with some water and put a slide in it, nothing would happen, but when we combine it with what we call mordant, which is a number of metallic salts, and we combine our hematoxylin with a mordant, it because an extremely strong nuclear stain.
I want to explain something to you on this slide. If you look at this Pacman at the top left, you will see it has “hematin” written in it. Truthfully, hematin is the result of hematoxylin dye meeting with aluminum salts, whether it be aluminum, iron, or chromium, and resulting in this great nuclear stain. In effect, our nuclear stain should be called hematin, but we don’t call it that, so I just want you to know that that really should say hematoxylin there. The dye tries to meet the tissue, it can’t, but once we put an aluminum salt mordent between it, look what we have; a nuclear stain that is very strong. It is connected by what we call covalent bonds. Covalent bonds are strong because these two atoms will share each of their electrons, meaning that they both have two, so they now have doubled the number of electrons they had, so it’s a strong connection.
Hematoxylin is a basic dye, carries a positive charge, and loves DNA and RNA because they have “ases” at the end of their name, it is attracted by that. As we’ve already learned, the affinity to be attracted to hematoxylin is called basophilia, and again, they form the covalent bonds.
The mechanism for a counter-stain, first let’s identify what a counterstain is. A counterstain is just a secondary dye, in this case eosin that is used to enhance the primary dye without interfering in it. It gives it contrast, and we all know how important contrast is to our pathologists when they’re looking at low level at the slides. Adsorption, not absorption, is a physical reaction dependent on the charge of the dye and the material to be stained. Molecules stick together and they’re loosely held. What happens when we take our slide to eosin, which should be used at about 5.0 pH? The eosin is adsorbed by the tissue sample, then we generally take it to 95, and we’ll work on that a little bit later. The dehydration process starts, but it also acts to differentiate the cytoplasm, or tone the cytoplasm. If we leave it in 95 long enough we’ll actually lose all of our eosin staining, so we have to be careful with that. But if we want to tone it a little bit, then that’s how we do it.
Eosin is an acid dye, it has a negative charge, it loves the positively charged cytoplasm, and this affinity is called acidophilia. Rather than covalent bonds, it’s held by weak ionic bonds, and what happens here is our atoms don’t share the electrons, they actually give them away, so that one side is left weakened. I want to tell you just for your information, there are two basic types of eosin. Most hematoxylin is aqueous that we use for routine H&E staining, but you do have the choice of using an aqueous or an alcoholic eosin. It’s important to know that, remember, like substances like like substances. If we’re using an aqueous eosin, then we can go from water right to eosin, back to water to rinse it, and then start our alcohols.
If you use an alcoholic or an aqueous eosin it will extend the life of your 95, because you know how it is once you come out of eosin into a clean 95; it’s just totally red. However, when rinsing in water you are kind of diluting your 95 a little bit faster. But if you’re using alcoholic eosin, which I think the highest percentages of laboratories that I’ve ever been into did use alcoholic eosin, which tends to show off the three colors of the eosin much better than the aqueous does. So if you use alcoholic, you go from 95 to eosin, and then either to 95 or 100, depending again on if you’re going to tone your eosin. Now we get back to dehydration. Have you ever noticed how tissue in a histology lab is kind of like rollercoasters? It comes in, it’s wet, it goes through a process of drying it out and getting paraffin in it, and then you want to get the paraffin out, you want to get water back into it, you want to get it stained, then you want to get water out of it and you want to clear it. So it’s like a rollercoaster. Once you know what one solution does somewhere, it does the same thing wherever it is, it just may serve a different purpose.
Here we go to dehydration to remove all the water. We usually start with a 95, 30-60 seconds, and then two ethanols, or you can use isopropanol if you need to, but to be safe you need to have three alcohols after eosin to make sure that there is no water on the slide when it leaves that last 100% alcohol and goes into clearing. Either you can have a 95 and two 100s, or three 100s, depending again whether you want to tone your eosin. Then we go into clearing, and if we recall, clearing is the process here of rendering the cells optically optimal. When we come out of 100% we hold our slide up and it’s opaque. When we put it in xylene and we hold our slide up, we can see through it, it’s clear.
This little slide right here represents, as far as I know, the only true macroscopic test for recognizing whether your slides have been dehydrated or not. This is important whether you go straight to an automatic cover slipper or whether you cover slip by hand. If your pathologist says those slides look pretty muddy today, and by the way, the eosin’s kind of washing out of my slides towards the edges, this is the culprit that causes that problem.
Have you ever thrown mercury on the floor, just fun, playing with it, way back in the day? I know we don’t do it now, but have you ever done that, and you see how the little pieces just fly all over the place, it breaks up and scatters everywhere? Where if you lift a slide out of xylene and it looks like the left or the right side of this windshield, then you’ve got water on the slides and you’re going to see little molecules of water trickle all over the slide. You should not send that to the cover slipper.
What you want to see is the middle of the windshield, which I lovingly refer to as the Rain-X effect. Basically, if you take Rain-X, put it on your windshield and it rains on you, it’s just going to sheet off. You don’t even have to use your windshield wipers. So that’s what you want your slides to look like. If they don’t, we need to work on some rotation. Again, we’re going to cover that when we get to the end of this presentation. The Rain-X effect; just remember that.
Once again we’re looking at our process, which is pretty straightforward, but if you notice, so far we have really not addressed staining of the primary stain that may involve differentiation, and may involve a bluing step. We’ve kind of looked at all the others.
Regressive versus progressive
This leads us to our portion where we’re getting down to the nitty-gritty with the stain, and we’re going to look at the difference between regressive versus progressive. I know there’s a bunch of you out there saying, oh no, Carolyn, I know what this is, and I am proud for you. But I cannot tell you how many times I have been called and said, I’ve got a problem. The first question that I’m going to ask is do you have a regressive or a progressive staining protocol? There was a time when 70-80% of my answer would be, what do you mean what’s the difference? How do I know? So I would go through a few pretty obvious ideas, and then we would come up with which they were staining with.
Let’s look at the most often used kinds of hematoxylin before we go into the difference between the two, because it’s kind of important to know your hematoxylin. Harris is very popular in the U.S., especially in skin labs and some other specialty labs. It continues to oxidize the hematoxylin powder, which we know now is called hematin. It should be filtered every day, so you’ve got to work a little bit on that. It typically uses a regressive stain, which we’re going to cover.
Mayers is fairly common, used in Europe; it also oxidizes and needs filtration. It typically uses a progressive stain. Gills, which we’ve all heard of, oxidizes very slowly, so for the purpose of removing the gold sheen, you don’t have to filter it, but it typically uses a progressive staining process.
Now we’ve got some of these new hybrids out here that are kind of a mix between Gills and Harris, and they typically do not oxidize, so no filtration is needed, and you can use either progressive or regressive, depending on your definition of regressive versus progressive. Yes, there is a real definite definition of regressive and progressive, but there are some gray areas that we will look at.
Regressive stain; tissue is intentionally over-stained then rinsed in water. After washing, and this is the one step that clarifies to me if you’re doing a regressive or a progressive stain, it is differentiated in a strong acid solution, most often 0.5 to 1.0% hydrochloric acid alcohol, usually used in 70%. This is a strong acid and it will break the covalent bond and it will remove the hematoxylin. So now I know you’re doing a regressive stain.
Regressive stain; differentiation is the one thing that defines that I am doing a regressive stain, and differentiation means that I’m attacking, remember, the tissue and the mordant that we used, more of the mordant dye. Because if we look at it and it’s held together, one side is holding the dye, the other side is holding the tissue, and the mordant is in the middle. One thing that scientists don’t know, they don’t know which side the acid attacks. Apparently it only attacks one side, so either we’re leaving with big molecules of stain on the left, or big molecules of tissue on the right. Regardless, if it’s timed precisely, it will remove just the excess hematoxylin.
In differentiation, rinsing is crucial to remove all the acid. If you hesitate at this point it could cause removal of too much nuclear stain, thereby producing a light stain. Once we get into this pH environment, our hematoxylin changes. It changes to a red soluble color and the complex has been compromised. On this particular stain we can look at this and actually explain three problems within H&E. One is how we can get a light stain; there are other ways, but one for sure is that we left it in the differentiator too long. Two, for a dark stain, we’ve left it not long enough in differentiator, and then because it turns kind of reddish-brown, if we do not take care of it along the rest of the way, we could present our pathologists with a red-brown nucleus, as was one of the problems.
The bluing is totally necessary after we come out of acid alcohol and rinse, and it stabilizes the nuclear stain, changing that soluble red color back to an insoluble blue, and you’ve got to do it. You can do it in path water because your pH range is usually from 5.4-9.8, but if you’re like the hospitals are in Florida, you never know when they’re going to chlorinate the water, and you’re never going to win, your pH is going to drop or raise. So at this point in time I think it’s more advisable that you use a controlled alkaline solution that’s buffered so it maintains the pH and you know what you're getting every time.
The bluing, we’ve got to wash it really good afterwards. I think you might understand by now that you cannot wash your slides too much. I know it takes away time and it takes you a little longer to get to the finished product, but washing really makes a big difference in how good your slides look. This is a picture of a piece of tissue that was not rinsed well. It went through the alcohol of 95 and went into eosin and still had some alkalinity, and this is what it looks like.
Before we go to progressive, review regressive. Regressive always over-stains, differentiates, and then blues to get our hematoxylin back to an insoluble state, a blue state. The biggest problem, the biggest caveat that I’ve heard all over the world with the regressive stain, is you’ve got to precisely get the timing and the percentage down of your acid alcohol. Really, that seems so easy, but apparently it’s not so easy. Of course, that involves when do you rotate it, et cetera, so that seems to be the biggest problem with a regressive stain.
If we now go into progressive stain, we see that there’s no differentiating necessary because we selectively, as a progressive hematoxylin, we stained just the nuclei. Mayers or Gills is most often used this way. You do need to wash it well and most folks do use a bluing solution so that they can deepen the blue-purple color. Running water, again, will work, but if you have that established, buffered alkaline pH, you know what you’re getting every time.
A common problem of the progressive stain, I saw it lots and lots of times. If you look at the slide on the heft, you can see it. I could have gotten a better slide, but I just wasn’t able to. You can see that it looks more blue-purple than the one on the right. The one on the right represents tissue that has gone through a strong differentiator. The one on the left represents a progressive stain that is staining the mucin and the negative mucin in the goblet cells, and some of the background. I’ve seen slides that just look totally blue after washing because their hematoxylin stuck to the slide so bad. Especially if you use coated slides or even charges slides, you will get that sticking on there. What happens is, the hematoxylin works the same; it loves DNA so it’s using absorption. But in this case, adsorption plays a role, too, and we’ve got some ionic bonding there on the surface of the slide.
I don’t know if anybody else has coined this phrase, but I know I’ve used it a lot; I call it the progressive stain clean up, or the pseudo-regressive stain. Here we’re going to use something, we don’t have to call it a differentiator, we call it an accentuator. It’s mild, it’s made with glacial acetic acid. They’ll have names like clarifier, define, get out the blue, take the blue off, whatever, because this sometimes can be a real problem that will drive you crazy. They’re just strong enough to get rid of the blue that’s on the back of the slides, and that excess in the goblet cells, for instance. Bluing is necessary after this because if you use an acid, even a mild one, you’re going to mess up that complex that you’ve built.
After reviewing the factors that can happen before staining, like processing, microtomy, and actual staining, what are some of the factors that are directly related to staining that cause the H&E to be misunderstood and abused? Granted, we know we’ve got problems before, but we’ve seen how the stains work, and some requirements for getting a nice clean stain.
When I went back and surveyed some professionals, there seems to be just a parable discord in exactly how the reagents should be handled. It’s one of the questions that I’m always asked; how often would you recommend changing this? How often, how often? Here are some of the responses I got from professionals, and the red is what I was thinking in my head. We change everything after 500 slides; you might be wasting reagents. We change everything after 5,000 slides; you might be abusing reagents. We dump everything on Friday and start fresh on Monday; okay, potential waste and abuse. I love this one: we rotate alcohol to xylene daily and change stains every two weeks. Well, I would say, based on what criteria?
We change everything on Monday except hematoxylin and eosin, and we top them off with fresh solution; never know what you’ve got. We look at the solutions and we know when they need changing; if you’ve got 20 techs out there which you all think are really good, wouldn’t that still vary some, tech to tech? My least favorite: when the doctors notice the shift in quality we dump it all and start fresh. I don’t want my slide to be the last one that they look at before they dump it all and start fresh.
Before we finish this part of it, I just want to mention that sometimes a good H&E can just be the difference between what the pathologist wants to look at. Giving the pathologist his credit due, what should he expect? Shouldn’t that H&E be clear so that he can see everything on it? Shouldn’t the slide sections look the same from slide to slide, day to day, and the quality be the same? Shouldn’t the first slide of the day and the last slide of the day look the same? That’s kind of what they’re all looking for, which just totally relates back to consistency and reproducibility.
I asked the population ideas on consistent and reproducible H&Es over the years, and I was astounded that the one thing that they totally agreed on was reagent management of staining reagents. In order to maintain consistency and reproducibility there must be a scientifically logical approach to this, and all processes involved must be standardized and controlled as much as possible. You would not believe this after that list of how people change their solutions, you wouldn’t believe that this was high on their priority, but they make some points regarding solutions.
One is that if you determine that 1 ml is enough to cover one slide in a solution, then if you have 500 ml container, you can do 500 slides. Some hematoxylins, eosins and some of the differentiators, and the buffered bluing, they can far exceed that number. But determining how many slides your lab can successfully stain without changing, I’m told, can be really time-consuming and a real problem.
The second point they made is they said that we’ve learned that manufacturing partners can and do work for you doing the necessary testing, because they’re still in product and they have to validate it. So this takes away the guess work and helps us to standardize this part of the function to its highest degree.
Suggestions for consistency and reproducibility
I asked these professionals for some ideas on how to apply what they knew to actual success in achieving consistency and reproducibility, and I got some suggestions. First, you’ve got standardize every aspect, which we looked over, as much as you can. Collection of specimen, grossing, processing. Processing, you can control; grossing, you can control. Embedding, keep those heats down. Microtomy, use those sharp blades, get some new microtomes.
Suggestion 2: determine what staining platform is best suited to your environment. You can get a close staining system that will take all the guess work out, it’ll tell you when to change the kits, you have to buy from certain manufacturers. You can do that with an open system, and that allows you to use whichever manufacturer you want to choose. Manual hand staining, you’ve still got to have SOPs in place for changing those reagents. It doesn’t matter if you automate it or if you're doing it by hand, you’ve still got to have some standard operating procedures.
Number three: you’ve got to standardize every aspect of the staining process. Use stains with regimented operating procedures for reagent handling. If you can, use pre-made industry partners’ validated kits to take the guess work out, already done the work for you. You can go back to them and fuss at them if it doesn’t meet their standards. Enforce your SOPs.
Number four: choose a source for stains that will offer standardized staining solutions and technical support in setting up your staining process and reagent management. If you’re going to spend a ton of money with a certain vendor, then they should at least tell you how to set your stain up, how long you should have to leave it in each solution, and a reasonable reagent management suggestion. Like on this one, rotate your xylene and alcohol every 500 slides. Change differentiating and blue every 1,000 slides. They advise to change your eosin and hematoxylin after 2,500 slides, or every two weeks, whichever comes first. You’ve also got to understand that there is a shelf life for open containers of reagent.
An important thing, too, let this manufacturer work with you, with all your doctors, you can do a blind study. Put a bunch of slides in front of every doctor; which one do you like the best? But let your manufacturer help you do this, determine what looks the best. If you’ve got 10 pathologists, 6 of them like this, then do what the 6 like and just kindly tell the other 4 that they were outnumbered. It’s called compromise; it’s called respect and compromise.
Number six: choose the source for stains that stand behind their products and offer solutions when you have a problem. I’m hoping that if you maybe had some of those issues, or you exchanged your reagents in some sort of haphazard way, that now after seeing these suggestions, that your plan for rotating reagents might change. Getting down to the bottom line is that reagent management is the key to your reproducibility of your H&E slides from day to day, slide to slide. With all things in front of it, it’s a lot to tackle, but when you get right down to the bitter end of it, your staining reagent management is what’s going to keep you having consistent and reproducible slides.
I think our time is really almost up. Unfortunately, there’s no magic formula for the perfection of an H&E. I wish there was, but there isn’t. Now that we’ve gone through the presentation, I think we might be able to agree that there’s way more to the H&E than meets the eye, that it requires a lot of logic and knowledge, it’s not just a silly thing we do. Perfection can be in the eye of the beholder, which again we respect, but we’ve got to compromis
I know you discussed this topic of dehydration earlier, is over-dehydration of tissue caused by too much time in all of the ethanols, or just too much time in the 100%?
C. DOAN: That’s a really good question. Typically on your processor you’re going to start with a lower-grade alcohol after formalin, because if you don’t you know you’re going to get formalin sediment all over the place. So you’re going to start with 70, probably, or 80, and then go maybe to a 95 and then 100. What I’d say to you is that the 70 and the 95 are going to be kinder to the tissue than the 100% is, so if it’s small samples you want to leave it a little longer in the lower-grade and you don’t have to leave it so long in the higher-graded alcohol, like 100% alcohol. If it’s big tissue, you want to spend a lot of time in the 100% alcohol. I hope that answered it.
Next, and I’m sorry if I’m mangling this, but how often would you recommend changing the defaring (phonetic) station?
C. DOAN: Oh, defaring station. I’m assuming we’re talking about the xylene on the stainer/staining platform. As I kind of alluded to at the end there, it’s dependent on your relationship of how many ml to one slide. So if we take the old adage, which I kind of like, one ml, one slide, and our container holds 750 ml, for instance, then you should be able to do 750 slides and then pull out the first one because it’s going to get the yuckiest first, and then rotate them down and make sure you have a clean one at the end. But I do like the old rule of one slide, one ml, and that’s usually what I would recommend.
What do you mean by the thumb test, Carolyn?
C. DOAN: Okay, so if you can imagine that you’ve got a piece of tissue that you're embedding and you probably--the thumb test is really not necessary, but again, if you are able to crush a specimen with your thumb, sink it down, like have it collapse because there is no matrix formed because it hasn’t been infiltrated, then for sure you need to take that back to alcohol and dehydrate it and bring it forward. It’s loaded with water. Just a funny little nomenclature; nobody really wants to stick their thumb into a block but if you could do that, if you can press that tissue down at all, then you really want to reprocess it. Don’t bother putting it on the microtone because you could lose precious cells by trying to cut it.
Would you please talk a little bit more about the eosin diffusion under the cover slip? What caused it and what are the possible solutions?
C. DOAN: It is almost 100% of the time caused by the fact that you have water on your slide. Something I did not go into but I will tell you now, we have a real problem with when we’re using an automatic stainer, even when we’re hand-staining, and we come out of one of the washes. If you think about your stainer, when you come out of a wash there is water all over the top, all around it; it’s not just on the tissue, the water covers it. So when you go into 95 before eosin, and you go to eosin, then you go to the alcohols after eosin, you most likely have water droplets that are still on the top, because your reagents hardly ever submerge the entire slide. What you need to do to remedy the situation is raise the level of your last alcohols, your 100% and your xylene, to the point that they reach the tippy-top of your slide over it. If you do that and you change those regularly, you will not see that problem, but that is exactly what it’s caused from. You might still be dehydrating pretty good, but if those water droplets are still out there on top, they’ll roll down under that cover slip and you’ll get that diffusion.
How long and in what environment should slides be drained before drying in the oven to prevent nuclear bubble artifacts?
C. DOAN: That’s a really good question, too. I don’t know that I can put a time fix on it, but have you ever seen--maybe some of your co-workers do this, I see it a lot in laboratories, where I go in and you’ve got your water bath, and you’ve got all these slides standing up, and they might even be two or three layers deep, but that’s what those folks are doing, they’re just letting them drain. You can normally visually see the water under your tissue, so what I would do is I would try a little test and put 10 or 20 of them up there, and if they all look like there’s no water on them, then I would go. I don’t think you can put a time on it, because there’s only so much you can do to waste time, because you’ve got to get those slides out. So you can look at it and see if there’s water under it, just to that poi