60 Minutes: 20 Histology Tips
When was the last time that your pathologist brought you a slide of decalcified bone, and said it was the best she ever saw? Ever wonder why your PAS stain is not staining the basement membrane the way it should? These questions and 18 others will be discussed and answered during the one hour Webinar: “60 minutes: 20 Histology Tips”. Issues related to routine histology, special stains, immunohistochemistry, histology equipment and other subjects will be the focus. All participants are invited to bring their histology “pet peeve” questions along for discussion and resolution to your histology issues.
- Effectively manage the various different specimen types received in the histology laboratory, with regard to proper processing and embedding.
- Troubleshoot the routine hematoxylin and eosin (H&E) stain.
- Troubleshoot immunohistochemistry stains.
- Effectively utilize automated staining platforms in the histology laboratory.
- Utilize novel methods to solve histology quality issues.
MR. CLIF CHAPMAN, HTL (ASCP), QIHC (ASCP):
Today's learning objectives, which we will accomplish without question, are to see examples of technical issues in the histology laboratory which require troubleshooting and problem resolution. We will look at them, and I will help you understand how the issues are resolved, and hopefully, that learning process will lead to your ability to troubleshoot new issues, which will arise in your histology laboratory and learn how to resolve those issues. And we will accomplish these objectives by looking at 20 histology tips in 60 minutes.
Tip #1 -- Histology Overview
So, I'd like to begin with a histology overview. I think it's good to kind of begin at the beginning. And in the late 1700s and into the 1800s is when generally everyone agrees that this was the birth of histology. In 1665, Robert Hooke published a book called Micrographia, and he coined the term "cell" because the plant materials he was looking at under crude lenses resembled the small uniform cells that monks lived in in the monasteries of that time. A little bit later, in the mid-1700s, Anton van Leeuwenhoek learned to grind lenses and make quality microscopes, and this was really the beginning of histology.
And then soon thereafter, there was a need for stained specimens, because whether you were hand-cutting specimens with a razor blade, or however you obtained your sections, you needed some contrast, so there was a need for stained specimens, and the hematoxylin and eosin stain, or what we refer to as the H and E stain, was developed and became the mainstay of pathology diagnosis. The H and E stain, along with special stains, are tinctorial stains, which impart color to the specimen under study. And for 150 to 200 years after that, it was the main stain used in the pathology laboratory.
In the 1960s and into the 1970s, really a revolution took place, where the immunohistochemistry methods were worked out, and IHC, as it is referred to, utilized specific antibodies against proteins in order to localize these proteins within tissues and cells. So now, in addition to the tinctorial stains, pathologists could investigate what proteins that cells were making, and most importantly, this aided them in making the diagnosis of cancer.
So, kind of fast-forward to today, in the 2000s, we've got a similar revolution going on. In the early 2000s was the development of the FISH technique, so the fluorescent in situ hybridization technique, and that's the use of probes, which are labeled with fluorescence molecules to be incubated onto tissue sections, and they seek out and bind to analogous portions within the DNA. So now, pathologists can investigate cells which seem normal with an H and E stain, and even with special stains, but may contain genetic changes that can cause disease. And the most recent development is the ability to stain messenger RNA, again, using specific probes, and that's the machinery that makes protein. So now, a cell could look normal, it may not even be ready to accumulate proteins, but messenger RNA is available and being made, and in this case, we're able to label that and see that, and that's a very exciting development, indeed.
Tip #2 -- Chemistry
However, in order to understand these new techniques and new information, we are going to have to backtrack a little bit and do a little bit of chemistry. I know it was everybody's favorite subject in high school and college. I know it was mine, without question. So, no fear here. If it wasn't your favorite subject, we'll go easy on you. But the idea is we need to understand chemistry in order to fully understand histology.
So, we need to understand specifically the biochemicals in life, and basically here we are talking about the elements of carbon, hydrogen, and oxygen, and the way that they bind. And just a quick refresher, that a single atom has a nucleus composed of protons and neutrons around which electrons reside. It used to be thought that they kind of spun around kind of like the planets circle the solar system, but now they use a probability diagram with an electron cloud, but suffice it to say that this is the glue that holds molecules together, the sharing of electrons.
You'll see that the chemicals that we use in histology are very similar to what we just saw. Here's formaldehyde with its carbon, hydrogen, oxygen. Here's xylene with its carbon and hydrogen, and its mixture of double bonds and single bonds. And here's ethyl alcohol, or ethanol as we know it, showing this ethanol group, which is an oxygen and a hydrogen.
We also need to understand the concept of the pH scale. So, to be a successful histologist, you need to understand this concept. pH stands for potential hydrogen, which is really a measure of the acid-alkaline balance of a particular solution. So, here we have the middle of this scale. It's a scale that goes from 0 to 14. The middle of the scale is 7, and this is the neutral solution, things like water. Milk is right around neutral, as well. As we go to the left, the numbers get lower, and these solutions are called acids. As we go to the right, the numbers between 7 and 14 refer to basic solutions, and you should remember that this is a logarithmic scale, which means that each value is 10 times the previous value.
So, why is this important? It's important for two reasons as histologists, because the first reason is you need to understand the concept of acids and bases. It's the mechanism by which staining works within tissue sections. It's the method by which you will make up stain solutions, or if you don't make them up and buy them, you should understand how they work.
And secondarily, you want to be a safe histologist, and that is a requirement to understand pH so that this does not happen to you. This is an example of a chemical burn. This was caused by a spill of acid. The effects are immediate and destructive. This would look exactly the same were it a spill of strong base. This example that I've used here, while a little bit shocking, is nothing like the ones that I turned down to not show you, so if you're having trouble staying awake at night, you need to stay up late because you're going to watch the game on TV, and it doesn't start till 10. So, you don't want to fall asleep. So, you go on your computer or your handheld device and just google chemical burns to the hand, and you will see some examples that will definitely keep you awake, perhaps for too long over the evening.
Tip #3 -- Fixation and Processing
So, back to fixation and processing, you need to understand. So, we've talked about the chemicals, the molecules involved. We've talked about the concept of PH. Now, this is very important, because when we have artifacts in the final slide, many of them originate in the fixation and processing part of tissue processing. So, you can see that, again, formaldehyde has this double bond that it likes to break and to help it cross-link proteins. Xylene has what's called the benzene ring. It likes to mix with organic solutions. And alcohol has this OH group over here on the side. So that, if we look at a fixation, you've probably heard of formaldehyde being a cross-linking fixative, which is very interesting, because that's exactly what it does. Here's a protein, and it interacts with formaldehyde. This double bond breaks, as we spoke, and some chemistry magic happens, and you wind up with this protein being cross-linked with -- I can't make the arrow go the other way -- but this one down on this end. And that's how we cross-link proteins. You should also know that alcohol fixes tissue by denaturization of proteins, which is a different chemical process, and we'll talk about that a little bit later. And fixation, of course, results in killing cells and stabilizing the proteins for subsequent processing in our histology laboratories.
Dehydration is done because tissues, when they are received, are 94% water, and this water needs to be removed and replaced with paraffin wax. Water and wax do not mix. One of the cardinal rules, the first rules in chemistry, is like dissolves like. So, in this case, we have an aqueous solution and an organic solution. Those do not mix. That's like taking your vinegar and your oil, and you have to shake it up before you put it on your salad, because it doesn't naturally mix. You need to shake it up and make a suspension. Leave it alone, it'll separate later on. They absolutely don't mix. So, we want to take the tissue that was originally water-based and put it into a waxed substance, and the way we do that is we've got to dehydrate it, get the water out. So, our friend alcohol is down here with this OH group on the end, which combines with water, H2O, to result in the tissue being denatured, these molecules are then denatured, and then the H3O strips off the hydrogen. That removes the water.
So, now we're almost there. We've fixed the tissue, we've gotten the water out. Now, we need to clear the tissue, which is basically we want to get the final tissue into paraffin wax, but ethanol does not mix with paraffin. So, we need xylene or a xylene substitute, which is an intermediate chemical, which connects with both ethanol and paraffin. And again, since we reviewed our chemistry, we see that xylene has these little methyl groups on the end, CH3, which kind of look like ethanol, so it'll mix with ethanol, but it also has a benzene ring, which looks like these little guys over here, which are in paraffin, and so this part will mix with paraffin wax. So, xylene or a xylene substitute -- you can use aliphatic hydrocarbons, some other chemicals that we can use as a xylene substitute -- then those will act as the intermediate chemical.
Tip #4 -- Processing Artefacts
And again, why are talking about this? Why did I just torture you with five or ten minutes' worth of chemistry? Because you need to understand organic chemistry and the chemistry of fixation and processing, because in the final slide, you may see these kinds of artefacts.
The first one we'll talk about is one of over-processing. You can see in this micrograph of an H&E slide, we've got a tissue fold here. We've got another tissue fold here. We have some tearing in the dermis, and we have an actual hole in the dermis right here, and this, I can tell you, is due to over-processing of the tissue in which the tissue dries out. Now, please be aware that while we want to remove water from the tissue, there is a certain amount of bound water which exists in the tissue. It helps hold the DNA together. It helps hold some of the basic molecules of the collagen together, so we really don't want to -- when we dehydrate the tissue, we don't want to dehydrate it completely. We want to leave a little bit of water, and a little bit means less than probably 1/100th of a percent, but we do want to leave some. If we don't leave it, then the tissue gets dry, brittle, scratchy, it's difficult to cut, and if you're a histologist, you know exactly what I'm talking about. So, if you have this issue, you have to go back to your processing schedule, and you've got to shorten the times in the alcohol, decrease the times, so that you leave the bound water in the sections.
Another artefact that you may see in your finished slide are these vacuoles, open spaces, inside cells. In this case, this is a piece of epidermis here, and you can also see some down here in some of the dermal cells. There are two causes, at least, and maybe more, but we'll talk about two causes.
The first is if you're using microwave processing, and the heat gets too high, or you have heat for too long, then what can happen is gases inside the tissue can vaporize and get trapped, and this is exactly what you'll see. So again, the solution to this is rather easy. You cut back on the temperature, you cut back on the time, and then it should make this go away.
If you don't use microwave processing and you see this, it may be, if you live in the Boston area or New England or northern climes, as I might refer to it, where the temperature goes below freezing, this is a classic freezing artefact in tissue. So that, to be proactive about this, if your laboratory is indeed located in these northern climates, and you see this artefact, you should think about distributing formaldehyde fixative, which has a percentage of ethanol in it. Up to 10% is usually recommended. This will decrease the freezing point of that solution, from 32 degrees Fahrenheit down to between 10 and 15 degrees. And as I look out the window now, boy, the wind's really blowing now. Right now, it's probably 20 degrees out, so we'd be all set, but tonight the temperature is expected to go down to 0, so that if there were any specimen bottles in, like, a clinician's office in a lockbox and that happens to be outside -- by the way, Strata has gone around and made sure that all those lockboxes are inside, at least in a lobby. We don't leave anything outside anymore. The other thing to worry about is, if it's a Friday and your couriers are driving around picking up specimens, and someone doesn't get back in time, and they take the car home and leave it in the driveway, that's going to get below freezing as well, in addition to the interior of airplanes, FedEx planes, and things like that, so it's something to think about, and this is kind of a classic freezing artefact.
Continuing with processing artefacts, I think everyone, if you've been a histologist, you have seen this thing called chatter. It's kind of a vibration of the tissue as it passes over the knife edge. This is a micrograph of a prostate needle biopsy, and you can see light striations in it. This is a classic venetian blind effect caused by chatter. And not too long ago, there was an article written by Choi and his group on the effects of fixation on GI biopsies, and of course, this may occur in any small biopsy tissue: Any kind of small needles, GI biopsies, little aggregates, any of that. This artefact can be affected by the fixative type, the time of fixation. Their work showed that alcohol-containing fixatives seemed to be a little better, with a maximum time of six to eight hours of fixation time. Over-processing was to be avoided, because that can lead to chatter as well, and I've noticed even in our laboratory, the speed of cutting, of microtomy, with the advent of automated microtomes, where the operator can set the cutting speed, and the laboratory management team wants everyone to kind of work faster and get more slides out, so if the dial goes from zero to ten, well, you know, ten would be better, right? It goes faster, it gets done quicker, and that's exactly wrong, because a cutting speed that's too fast can result in exactly this kind of artefact. As a point of practice, I advise people in our laboratory to keep their speeds between five and six, somewhere around there, certainly no higher than seven.
Tip #5 -- Special Cases: Alopecia
Okay, we'd like to move on from processing artefacts into artefacts that you may see or may be caused by special cases, and this puts us now into -- instead of being in a reactive mode, looking at that final side, saying, "Gee, what the heck happened to this slide?" Now, we need to kind of be proactive, look ahead. We need to identify what kinds of specimens can come into your laboratory that aren't the routine specimens, if you will, whether you're a hospital laboratory, whether you're a private dermatopathology laboratory, what to be on the lookout for.
Being in a dermatopathology laboratory, one of the things we look out for are special cases of alopecia. Why do we look for that? Because this is the only skin specimen that we receive that is not cut perpendicular to the dermal-epidermal junction. Instead, it's cut parallel to the epidermis. So, you see that we cut it parallel to the epidermis. Again, these are specimens for alopecia, which is hair loss, balding, not related to male pattern baldness, which my father and grandfather so kindly gave to me. Thanks, guys. No, not that, but ones that have a pathological cause. So, we want cross-sections of the hair follicle. Here are one, two, three hair follicles very up-close to the surface of the epidermis where the hairs are thin, and here's the same follicles, one, two, three. You can see how they're bigger, they're fatter, they're more plump, because that's the hair bulb, which is further down in the dermis.
Here's a photo micrograph of kind of a typical alopecia case where you see a degenerating hair follicle, another degenerating hair follicle. This hair follicle is almost all gone, and then at the same there's this infiltrate of inflammatory cells, so something is happening is here, something is causing this hair loss. One of the things that the pathologist can do with this preparation as well is they can count hair follicles. In order to prepare these specimens, they are almost always a 3 or a 4 mm diameter punch specimen, so we cut them in the middle, or actually 2 to 3 mm below the epidermis. Here's the cut surface. Now you've got two pieces. You want to ink the cut surface red. Ink this cut surface red. This goes in cassette A, this goes in cassette B, and the grossing personnel will know to embed the ink surface down to get a cross-section. So, grossing personnel have to be alert. They have to check the requisition. They need to look for terms like alopecia, hair loss, traction, baldness, and then make sure to identify the cassette so that the embedding personnel will do it properly.
In the final slide for -- this is Headington procedure, where the slide is prepared in such a manner. You can see with a PAS there is a thickened basement membrane. You can see with the elastic tissue there are scarred follicles. More importantly for us, which is very obvious, is one can actually go and count hair follicles: 1, 2, 3, 4, 5, 6, 7, let's say that's per a high-power field. Then the pathologist can look up on the chart, look up the patient demographic, for example. You look up my demographic: Male, Caucasian, over 60. How many hairs should they have? If they don't have male pattern baldness, they should have 20, and there's only 3, then you know something is going on with pathology to cause that hair loss.
Tip #6 -- Special Cases: Immunofluorescence
Another special case -- and again, whether you're a hospital laboratory or a reference laboratory, you may receive specimens for immunofluorescence. Now, these might not even be specimens that you're going to handle. They may show up in your lab as a pass-through specimen, maybe being routed someplace else. We do receive these specimens, and you can see that the cardinal rule here is any specimen submitted for immunofluorescence has to be submitted in a special immunofluorescence transport medium, sometimes referred to as Michel's medium, named after Dr. Michel, who invented the medium, or Zeus transport medium, or simply immuno transport medium. It cannot go into formaldehyde. If these specimens go into formalin, and they're fixed in formalin, even for three minutes, then it cross-links the proteins, it destroys the antigenicity, so that this particular stain cannot be done. Here's the immunofluorescence. This result is a basement membrane staining. This result is intercellular staining. So, what we do in our laboratory, see the big red arrow here? That's the immuno bottle. There's the regular fixation bottle, and you can see they look very different. The big one's got the big yellow label, says formalin in big letters. This one's smaller because usually the specimens are smaller. As I pointed out, they're 3 or 4 mm punch specimens, but very visually different, so that all handling personnel and triage personnel know this guy does not go into the surgical grossing room to be put into formaldehyde; this guy goes into the immunohistochemistry laboratory. Very important to be proactive about that, because the laboratory does not want to receive a specimen and then mishandle it into the wrong fixative so that the requested procedure can't be done.
Tip #7 -- Minute Specimens
Okay, continuing along, these might not be special cases, but you certainly receive minute specimens. I have been doing histology for -- it used to be over 30 years. I'm afraid now it's over 40 years, but anyway, I don't think it's me, and my eyesight is getting worse, but I think the specimens, generally speaking, are getting smaller.
When I first started at Massachusetts General Hospital in the surgical pathology laboratory, you got nice big, whopping big pieces of tissue, and if something bad happened, you could always go back to the formalin jar and get more tissue. But now, with the advent of fine needle aspirates, small needles, skinny needle biopsies, those days are gone. So, over and over again, we get smaller and smaller pieces of tissue. So, how to handle those: Because you don't want them, you don't want to compromise them. In different laboratories that I've worked in and different people I've talked with, the choices are to use histo-wraps or lens paper to wrap the tissues. The mesh bags, you can use those. The mesh cassettes, you could those to trap and keep those tiny fragments in. The blue sponges, they're not the best because they carry over some solutions, and if it's something mucinous, the specimen will actually kind of melt into the sponge during processing and be really difficult to get out. And in addition, all of these do require that you open the cassette after processing and then poke them more and dig them out and perhaps break them to embed them.
So, we will use HistoGel for very small specimens -- when I say very small, 0.1 mm or smaller -- because HistoGel is a liquid at 55, but then at room temperature becomes gel-like, so imagine a Jell-O mold with all the fruit trapped inside, so HistoGel is a mini-Jell-O mold. You can trap the tissue fragments inside. Here it is being dripped out of a pipette onto the specimen. You'll get this dome of HistoGel, and when it solidifies, you can put that in the cassette, and away you go. And then the next morning, for embedding, you simply just lift out the little blob of HistoGel and embed it. If there's orientation involved, however, you'll have to do it at the time of embedding. You can trim one end to get a flat side down or something, but you cannot do orientation.
However, that's the bad news, but the good news is when you have a very tiny fragment, it can't go anywhere. It's in that drop of HistoGel. Here's an example of a case we actually did have. Here's the original fragment. That's a photo micrograph under, I think, just 2x power, the ink dot showing where the fragment is. So, we cut initial sections on this and didn't see anything, so we went through and cut 30 serial sections, and out of 30, slides #5, #6, and #7 had sections of the tissue, and we took one of those and did a PAS stain on that, and you can see that it's positive for fungi. So, this is a method that's a little labor-intensive. You can't orient the tissue at the time of use of HistoGel, but the bottom line is you can't lose the specimen, and you won't be damaging it the next morning trying to embed it.
Tip #8 -- Bone Specimens
I'll take some time with this. I was surprised at the previous webinar. We had quite a number of questions about bone, about decalcification. Bone has its own issues, of course, and whether you're a hospital laboratory or a private lab, you're going to get pieces of bone for some reason. The first decision is, which decalcification solution should you use? In our laboratory, we're a reference laboratory. We have some time. We want good histology, so we use 10% formic acid and 10% formalin, which is not buffered. You want that solution to be acidic, because the acid in the de-cal will pull the calcium out of the bone, because the calcium is what makes the bone hard.
The good news is the histology is great. The bad news is it takes some days to decalcify. There was an article in The Histologic of 2011 that said you can use microwaves to speed up decalcification. I guess you can experiment with that if you want. I stay away from heat during any of the processing or decalcification. As we saw before, tissues are basically protein, and the more that you heat them, the harder they get. You want the soft-boiled egg, you just go a little bit of heat; if you want that thing really hard all the way through, then you give it more heat. So, I like to stay away from heat during processing. It makes the tissue really hard.
No matter what decalcification solution you do use, it's a good idea to use a big volume of de-cal solution. Here, we have a little schematic. Here's a stir plate with the stirrer on. The heat is off. Here's a little stir bar. Here's your decalcification solution. Here's the cassette, and I put a little fishing line on there to hold it over the edge. This is probably a 2L beaker. Fill it up with de-cal, get the spinner going, put a top on it to seal it or some parafilm, and then leave it 24 hours. Come back, use your fishing line, pull up the specimen, check it, and see if it's softer. If it's not, then dump out the de-cal solution and put in fresh, because the decalcification solution will become expended. It will get filled up with calcium, and then it won't be as efficient pulling out the calcium. When you do this every day, you'll get to a point where you'll be done. You can judge the endpoint by feeling it manually, but the best way is to use the chemical endpoint determination.
Now, back to which decalcification to use: Obviously, if you're in a hospital and you are receiving bone marrow biopsies, you don't have two or three days to do it, so use a rapid de-cal, which contains nitric or hydrochloric acid. The good news is it's fast. The bad news is the histology is not so good, because those are very strong acid and don't bode well for a cellular structure, but again, it will get it done quickly. You can use Bouin solution with some picric acid. That will do the decalcification for you. If you're in a research facility and you want to preserve antigens, then you can use EDTA. That takes weeks for a complete decalcification, but the histology is excellent, and the antigens for IHC are preserved. These formic acid de-cal specimens have excellent histology, as you can see here. We also use a gelatin-coated slide to pick up these sections. It helps the de-cal to adhere to the glass slide.
Tip #9 -- Specimens for Gout/Crystal Identification
Continuing along with special cases, you may receive cases for crystal identifications, specifically uric crystals for gout. These specimens should be submitted in 100% alcohol. When they're received, they should go on to a tissue processor with a program that starts in 100% alcohol, doesn't go back, doesn't start in formalin, and that's because uric crystals will dissolve in water, so you want to use 100% alcohol. If it's processed correctly, you'll be able to make an H&E. Then, you'll be able to put the H&E under a polarizing scope and see these nice uric crystals underneath the polarized light.
Tip #10 -- Nail Specimens
Nail specimens are fairly common now. There are really two reasons that they arrive in your laboratory. Reason #1 is they will be fingernail clippings, looking for fungal infection, fingernail or toenail, to rule out athlete's foot, severe cases, things like that, and the second reason is you may receive a nail specimen from a nail bed in which they are looking for a melanoma. Here's a final slide from that, some melanoma slides. Here is a slide of a nail specimen that is loaded with fungi with a PAS stain.
What we recommend here, and we use them in the laboratory, is to, upon receipt of nail specimens, to use a 5% Tween detergent made up in 10% neutral buffered formalin. Soak this overnight as a minimum; it should be 24 hours. Process on the large side processor, no short processing. During microtomy, cut extra slides for special stains and IHC, should the pathologist need them. We also use gelatin-coated slides to pick up these specimens, so for de-cal bone and for keratin specimens, we need to use gelatin slides. That helps to keep the sections on the slide.
Tip #11 -- Hair Histology
You can do that with single strands of hair. Sometimes, the pathologist is looking for visible changes, indicating genetic disease such as this. Other times, they will be looking for fungal infection. Here's an unstained hair under high power. Here's the PAS stain showing fungi, and in order to do that, you can use Scotch tape to affix hair strands to a slide for automated H&E or automated special stains, or you can keep the hair strands in a small glass vial or beaker and pour the staining solutions on and off, the PAS stain solutions, and that works quite well.
Tip #12 -- Periodic Acid Schiff's Stain - PAS
You need to understand the PAS and PAS plus D. PAS stains glycogen, which is a sugar molecule. Periodic acid will break those rings, make aldehyde groups. Schiff's reagent binds and makes the color reaction product, which is a magenta color. If the serial slide next to it is incubated with diastase prior to the PAS, there will be no stain, so this is a PAS, and a PAS plus D, looking for glycogen.
We use an alpha-amylase from Sigma. Again, I'm not endorsing the company; I'm just saying this is what works in our laboratory, because we're able to do the diastase digestion 20 minutes at room temperature instead of 60 degrees heated. We make sure to make up the periodic acid fresh, no more than a week old. We use a warm tap water rinse after the Schiff's reagent to make the slides go pink, and we use a Gill hematoxylin for the counter stain, and no clarifier or bluing is needed for that. Here's a final result of stain on fungi on the surface of a skin specimen, and inside the keratin of a nail specimen.
Tip #13 -- Special Stains for Infection
The tip here is that each one is a little bit complicated. We've had AFB, Fite stains, spirochetes. I will give you one example of one that we had to troubleshoot. This is the stain for Gram positive and Gram negative organisms. We use the Brown and Brenn stain, and we have found that in the first half of the stain, staining Gram positive, to make up the crystal violet and sodium bicarbonate solution fresh for each stain, and then to use the Gallego's solution to lock in basic fuchsin in the second half of this stain. Specifically, we were getting a lot of stains coming through that these red organisms were very pale, so once we started to use Gallego's solution, we had success.
Tip #14 -- Silver Stains
Again, if you're a histologist, you've been in the field for a while. You want to make sure you use acid-cleaned glassware, plastic forceps instead of metal forceps so the silver does not precipitate. Use fresh silver solutions and precise temperature and time control, and that will help you to pre-emptively strike and keep away from any issues.
Tip #15 -- Light Microscopy: How to Set Up Your Microscope
The stains are no good unless you can look at them. I'm surprised when I go to people's laboratories how many people don't know how to use a microscope, don't know what the parts are, so it is part of your histologist training. You should make sure that people understand that the oculars are here, the objectives are here, the staged slide goes onto the stage, there is a condenser to focus the light, and the light intensity adjustment. And there's a whole procedure that you should sit, adjust the interocular distance, adjust the focus of the focus ring, and then adjust the condenser using the illumination ring, such that you can center the light coming through, and not only should it be centered, but it should be focused sharply through the condenser. You can also calculate final magnifications using the magnification of the ocular times objective, and you can also use the microscope during microtomy. You can pick up sections, look at them unstained. You can rack the condenser down, and you can see here I've got a nice 2 mm punch photographed unstained, and it shows perfect orientation, epidermis, dermis. And here it is stained with an H&E. Its perfect orientation, and you can check and make sure that you've got a complete section.
Tip #16 -- Hematoxylin and Eosin
It's different in everybody's laboratory. You should have three colors of eosin, three shades of pink. Your nuclei should be sharp, well-defined. You need to decide what you're going to use for reagents, Harris or Gill hematoxylin. Eosin, do you want to add phloxine to it? Which bluing solution are you going to use? Or you could just use running tap water. Make sure it's completely deparaffinized, and watch out for xylene substitutes; some are very water-intolerant. You need to make sure the alcohols on each side of that are nice and clean. Eosin is usually 1% alcoholic solution to give you three colors. Be careful with your bluing reagents, that the pH is not too high, and again, remove all the paraffin before staining and get all the water out before cover-slipping. Even if a little water is left, you can get eosin bleed after the slide has been cover-slipped and put into file.
Tip #17 -- Safety
Now, we're going to come down the home stretch here and wind up with just a couple of safety things. Please make sure everyone in your laboratory is using their personal protective equipment. Make sure they know where the shower is and how to use it. I don't require everyone to stand under it as I did, but I think a picture is worth a thousand words, isn't it? Plus, it's me, so it's funny.
Tip #18 -- Picric Acid and Mercury
Picric acid in your laboratory: As the Germans would say, das ist verboten. Do not have that in your laboratory. If it dries out, it can explode. Similarly, you should not be having any mercury compounds in your laboratory. There's no need for mercury in a histology laboratory anymore. So, no dry picric acid. You can keep some Bouin's in there, because Bouin's is in solution, so you have to be careful with it. No mercury solutions. There are plenty of mercury substitute stains now. You don't need to use those anymore.
Tip #19 -- Chemical Compatibility
Please, in your laboratory, plan your chemical storage with regard to compatibility. You don't want any chemicals to be incompatible. You want to make sure that everything fits with everything else, so predefine your waste streams. We keep stains waste separate from alcohol and xylene waste; separate from gold and silver waste, which are heavy metals; separate from picric acid waste, which is an oxidizer; separate from cyanide waste, which is an out-and-out poison.
And if you choose to not pay attention to compatibilities, something like this could happen to your laboratory. It could be as severe as this. This happened one year ago, January 2016. A chemical called trimethyl aluminum, people were handling it in the hood, and it's one of those chemicals that explodes on contact with water, so it got in contact with water in the air, humidity, whatever, and blew the laboratory to pieces. No one was killed that day. However, three years before that, the same chemical exploded under the same situation, and it did kill a person. So, please pay attention to the safety of yourself and your workers while you are in the laboratory.
Tip #20 -- Two Stains on One Slide
This happens in our laboratory. The pathologist wants two stains on one slide, and the stain has already been done. So, here comes the selected H&E slide. It has four sections on it. These two, the pathologist wanted an AFB; these two, they want the PAS plus D. So, first thing is make sure the pathologist picks which stains they want on what sections, so you don't finish the project and they say, "Oh, I didn't want that." And then once determined, you should remove the cover slip. Once the cover slip is removed, you're going to need a diamond-point pencil. You will hold it firmly in your hand or lay the slide out on a flat benchtop, and then carefully and steadily draw a line and score the glass right between the sections that you have determined you want to separate. Once you're done that, you can apply even pressure with your thumbs right behind the score, and then once it breaks, you're all set. You can take those two pieces, and you can use Super Glue gel to attach those pieces onto a full microscope backing slide. You can run it down, decolorize the section, and then stain as requested. But remember: Once you glue the piece of slide onto the backing slide, it will never come off. There is no kind of anti-glue that's going to take that off.
So, my final tip for today is that while you may wear many hats in your laboratory, there's only one hat that matters: That's your team hat, whichever one you support. If you have more questions, my contact information, these two e-mail addresses: firstname.lastname@example.org and email@example.com. Additionally, you should know that the book we just got through authoring and came out just this past fall is called Dermatopathology Laboratory Techniques. It has many of the procedures that we've discussed today in it. It is available on amazon.com. Simply go into the book section, put in "Chapman and derm," and it should magically appear. So, at this point in time, I would like to thank you all for listening and paying attention in today's webinar, and I would like to stay here, and I'm happy to answer any questions that you may have at this time, and turn back to Rick.
Questions and Answers
Can you please point out again why do we want to have just a little percentage of water to maintain DNA on its place?
Okay. DNA, you probably remember from high school biology. It's the double helix. The nucleotide bases all fit together as an exact jigsaw puzzle, and it's twisted and turned on each other, so think spaghetti, but think of it as being somewhat movable, but it could become quite brittle. There are water molecules embedded in there that give it a little bit of wiggle room. You don't want it to be like a piece of stone. You want it to be maybe like celery, I guess. Leave a little bit of water there. Same thing with the -- and I'm talking about water on the molecular level, not water you can see, not water you can measure, but between the molecules of DNA, you want a little bit of water just to keep those molecules moving enough so that when you go to section them, when you put the torque forces of a microtome blade to the tissue, that it's not too hard: You're not trying to cut a rock with the blade, you can cut a celery. So, that's the idea behind it.
And what happens if a little bit of water is left on the slide?
If a little bit of water is left on the slide and you can see it, first of all, it's going to look horrible. Let me differentiate. So, bound water is microscopic water. It's not going to affect staining. It's not going to affect the final appearance of the slide. When you apply the cover slip to a finished slide, if there's any water, any measurable water left, not even that you can see, but measurable water, what will happen is usually the hematoxylin sticks into the nuclei all right. They're usually not affected too much. But it causes the eosin in the section to bleed out, to actually go back in the solution. So, when you file those slides and you pull them even a couple of days later, you'll pull them out and all the eosin will be gone. They won't be pink anymore. You'll just have nuclei to look at.
How long do you keep the bone specimen -- this was during Tip #8, during Bone Specimens --how long do you keep the bone specimen in the decalcification solution?
Okay, that's a great question. The answer is, who knows? That's why you have to check it every day. It depends on the size of the bone. It depends on whether it's compact bone, whether it's trabecular bone. The idea is, get your rotary shaver set up or your stirrer set up. Get your specimens on there, and then every day at the same time, maybe when you first get to work or a certain time of day, go put your gloves on, put your gear on, work in a ventilated hood, take the specimen out. Just squeeze it between your fingers. It should begin to get soft. And if it's a big piece of bone -- let's say it's a 4 mm piece of bone -- when it gets soft enough, you'll be able to work a blade through it, a razor blade through it. Now, you've got two 2 mm thick pieces, and you it back in the de-cal again. So, the idea is to check it every day and change the solution every day. Then, once that piece -- after two days, four days, six days -- when that piece is nice and easy -- you can bend it, you can twist it, and it doesn't break -- it's pretty much done. But I also urge you to go online and look up endpoint decalcification methods, because there's a chemical way that you can determine that all the calcium is out of the bone. The answer is, it's going to depend on which kind of bone it is and what kind of de-cal it is, but you do have to change it every day.
Any recommendations for decalcifying multiple cassettes at once, like 30-40 cassettes of mouse ankles? Just throw them all in a large beaker?
Okay, I have a great suggestion for that, and I think you can still get them. The senior members of the histology crew out there who are listening to this will know that way back in the day, a company called Shandon used to make stainless steel cassettes. They were round, and they measured 5 inches in diameter. If you can get your hands on one of those, you can put the stir bar underneath that, and then put that at the bottom of the beaker. And then you can go ahead and just throw all of your cassettes in there and turn on the stir bar, and it keeps it agitated. You can take a chance. If you put too many cassettes in there with just the stir bar, it tends to flip around and not have a good place to spin around, but if you can get something with holes in it or something to kind of prop up and protect the stir bar, you can simply just fill up a 2L beaker with de-cal and throw all your cassettes right in there, and then the next day dump them off into your waste container and then fill it up again, put the parafilm over the top, and just do it again.
What acid do you recommend to clean glass after silver stains?
So, there's two times to clean glassware: Before the silver stain, you should take all the glassware that you're going to use and lay it out on a nice Kimwipe or something, or a Lab-Sorb, one of those things, and you should use a dilute solution of hydrochloric acid. Conversely, you could also use -- almost everybody has 1% acid alcohol in their lab for the Kinyoun's acid-fast. You can use that as well. So, to clean glassware, fill it up with the acid alcohol. Let it set for a couple of minutes, pour it off, and then rinse seven times with distilled water, and then leave that out. That's acid-cleaned glassware, so that gets you ready to do the stain. When the stain's all done, if you've got silver precipitate, you can use that same solution. Usually, that will take it off. So, 1% acid alcohol. Try that.
I have mouse embryos cleared and stained with alizarin red and alcian blue to distinguish bone and cartilage fixed in ethanol and glycerol. I want to longitudinally section epiphyseal plates, but decalcification destroys the original staining, especially alizarin. I can re-stain with H&E, but it does not distinguish as well. Is there a way to section successfully without destroying the original colors through decalcification?
Wow, that's a tough one. Not that I'm aware of. You may want to think about, as a test, decalcify a specimen completely, process it into paraffin, cut it, and you can attempt to do trichrome stains, but I know what you're trying to do. You're trying to see the difference between the calcified bone and the uncalcified bone, and really, the only way I know of doing that is to run those into glycol methacrylate or methyl methacrylate, which are water-soluble plastics, and then cut sections on that. You've got me stumped on that one.