Lessons From the Laboratory: Your Tissue Processing Questions, Answered

Thank you, Kaylee, for that warm introduction, and thanks to all of you that have taken the time out to be a part of this webinar. Today's webinar about tissue processing artifacts is entitled, Lessons from the Laboratory, Your Tissue Processing Questions Answered. The objectives today, at the conclusion of this session, will include the three conditions as to why pre-analytic conditions impact downstream results, identify the three critical steps to ensure optimal tissue processing, and discuss the effects of over and under fixation on downstream results.

Today's presentation is a continuation of last year's webinar that was hugely successful, thanks to many of you that attended. We had so many questions to follow up on, and we could not wait to have this session to do just that.

Let's do a little bit of review. Going back to the webinar, let's refresh what good looks like. Regardless of what your PI or pathologist prefers regarding stain intensity, we all want to see architecture that is intact and well-preserved cellular structures. On the left, we can see a section of skin with keratinocytes staining purple and the dermis staining mostly pink, and red due to the collagen, elastic, and loose connective tissue that's present. On the right is a nice section of GI tissue that shows the nucleus staining purple-blue, the mucin, a light purple, and the cytoplasm, nice pink. In the center, we can also appreciate supporting structures that also include, of course, the vascular supply.

What causes dark nuclei H&E staining?

Let's start with the questions that were submitted from last year's webinar. The first question comes in asking sometimes in H&E staining, the nuclei stain is very dark. What is the cause? There can be several causes. A couple of things that come to mind is what type of hematoxylin is being used? You may know or may not that there are several formulations and types of hematoxylin. The regressive Harris hematoxylin that is most widely used for staining, while in cytology, the progressive Gill's hematoxylin is used most often. There are certainly others, such as Meyers and Weickert's, still used today in special stains.

The intensity of the hematoxylin may be due in part to the cell type. Whether the cell is in a cell cycle, apoptosis, euchromatin, heterochromatin will certainly sometimes affect the intensity of what we're seeing under the scope. Also, too, going back to the Harris hematoxylin, if you are using the regressive Harris, it is a stain that overstains structures such as the nucleus and the cell membrane. And we use an acidic solution, some labs will make up a very weak acidic solution, or you can certainly buy pre-made from vendors that may be called define or something like that they label it with. But it's usually just a weak acid solution to kind of clean up the slide and pull back on some of that heavy staining.

We are always getting under-stained hematoxylin and histopathology staining. We are using Harris hematoxylin followed by bluing and destaining. Can any steps of tissue processing result in this problem, though eosin staining and tissue morphology are fine. There's several things, of course, that come to mind. First thing in the question I did notice is that they're using Harris hematoxylin followed by bluing and destaining. And I was curious, I'm hoping that they're doing the destaining first because Harris is a regressive stain, and then following that with the bluing. I'm not sure if they meant to put those in the opposite order, but just checking.

Some of the things that can cause understaining, is how long has the tissue been in formalin? Tissue that's been in formalin for longer than a couple of weeks, I think we don't realize that formalin continues to fix tissue even after we're done with it, whether you're in a clinical or research setting. If it's in there for weeks, you're continually making those covalent bonds, building those methylene bridges. And so even though we are accustomed to using antigen retrieval when we're doing IHC to open sites back up, if you leave tissue in formalin for extended periods of time, weeks or months, then you can actually tie up those sites beyond what AR can actually retrieve. So that might be some of the cause. If it hasn't been in a long time, then you might be able to antigen retrieve the tissue and pop those sites back open for H&E staining. We I had done that before when we had autopsy tissue that was maybe sitting in formalin for, you know, longer than what clinical specimens do.

And then is the hematoxylin, you know, old? Is it expired? How long is your slide sitting in the hematoxylin? You know, from vendor to vendor, there are different formulations. And so, the times may vary a little bit. If you're not, you may purchase it commercially, but if you're making it in-house, which I'm not sure if I know of anybody that's doing that very often, it's quite laborious. It does have to, especially with Harris, you make up the Harris and then you have to let it ripen before it's ready to use.

How can I overcome zonal fixation?

Moving on to the next question. How can I overcome zonal fixation? Great question. With large specimens or even with hollow type specimens that like a gallbladder or maybe a kidney, uteri is a very common bladder that is considered kind of hollow organs. The thing I would certainly respond with is prep, prep, prep. Large hollow organs need to be open and prep large solid organs like liver need to be prepped by taking it, trying to keep some orientation and making anywhere from a centimeter to a centimeter and a half full thickness cuts through without actually detaching those sections so that formalin can get access and start making those covalent bonds, building those methylene bridges to fix tissue to preserve not only architecture, but certainly structures. And then we'll talk a little bit about molecular impact.

One of the things that might help, especially if you're dealing, if you if you have the facility and accommodations to do this, is if you have containers that can accommodate these type of specimens, any kind of stir bar at the bottom of a container helped to move that formalin around, certainly helps too. Just a little trick that we used to use.

Speaking of perfusion, someone asked, “can you explain what it is?” In histology, it's the mechanical, of course, technique in moving fluid into cavities and channels of tissue that we incur. It's more than just letting it sit in the formalin and let passive diffusion happen, right? We're going to move it in. This is kind of maybe an archaic example of actually a mechanical pump that's going to move that fluid through organs and vessels that's needed maybe for those types of studies. We don't normally do a lot of perfusion where I come from in the clinical world, but I know in research it is very common. So yeah, it is really a great procedure in preserving delicate structures that need to be examined.

What can cause perfusion artifacts?

Great question to follow up with is what can cause perfusion artifacts? When you look at this image on the right, you can see, I've got two images here. The one on the left is, you can see the architecture is just almost perfect. You see on the right some disruption and that is where the fluid that is being moved into this organ to preserve is being too much pressure and too much volume maybe that it disrupts some of the fine delicate structures that we see in lung.

How long can tissue be kept in neutral buffer formalin before tissue processing or histopathology?

Someone asked, how long can tissue be kept in neutral buffer formalin before tissue processing or histopathology? I think you have to take in consideration what type of testing you're going to be doing. It's forgiving when it comes to H&E staining and a lot of antibodies. We do have recommendations from Leica, who I work for, that we would like to see a minimum of 6 hours. We're finding as increased paraffin, tissue that's formalin fixed tissue that's embedded in paraffin, is being examined in the research world. We're finding that there are some considerations that need to be considered. So, if you're doing just normal tissue processing, you need to certainly put it in for six hours. But if you're going to be looking at structures, DNA and RNA, there's from what I have information I've gathered, there are there’s recommendations that you need at least, you know, 12 to 24 hours. If you go beyond the 24 hours, I don't know the chemistry behind it, but the formula actually starts to break up the RNA in shorter segments that you probably want to study. So just take note of that.

If we can fix a tissue for 24 hours, how long after that can the tissue be in the 70% ethanol or in PBS?

Next question is, if we can fix a tissue for 24 hours, how long after that can the tissue be in the 70% ethanol or in PBS? And so, if you'll see down at the bottom right-hand corner of this slide, you'll see two links that will connect you to two sites that I went to that made recommendations for this. It's very common, just kind of the short answer is that it's very common to store tissue in 70%, three to four weeks. Of course, this is after you've already fixed the tissue in formalin and you've rinsed it right, you're going to move it into the 70%. If you need PBS, use cold PBS only for two to three days.

How do I prevent ice crystals?

Next question. How do I prevent ice crystals? Obviously, the person asking this question comes from a laboratory that I think would probably be using #1 fresh tissue and would be using some type of cryostat. If you are getting fresh tissue from your necropsy, I would think, you're going to transfer it. Try not to transfer it in a jar that is full of saline, because that tissue is going to absorb some of that saline and make it very wet. You're trying to freeze tissue, number one, without a lot of water in it, and you want to freeze it quickly to keep the rapid crystal formation from happening. So, if you do get a sample, and maybe it was in saline, pat it dry if you can before freezing it. If you have liquid nitrogen, it's great, but a lot of labs may not be able to have that on site. So just taking note of not having it in a container full, soaked in saline, or at least pat it dry before you freeze it, and that should help once you get it mounted and ready to go.

Why does my paraffin sometimes crumble when I'm cutting? 

Why does my paraffin sometimes crumble when I'm cutting? Well, the first thing I would think about... is the quality of paraffin that you're using. There's so many different types out there, from so many vendors. So, it could be just a formulation that doesn't fit for the laboratory work you're doing. Is it crumbling within the tissue sample? I'd kind of like to know some of, you know, knowing where the crumbling is happening would kind of help us determine if it's crumbling right after you embed it and you're cutting, or are you using free spray and maybe overusing it where it creates the cracks in the paraffin in the tissue? That could certainly be a reason. And for labs that can't use the free spray or don't purchase free spray, they find a workaround by putting the blocks in a freezer for a short time but then forget about it. And of course, by the time they pull it out, there's a lot of cracks in the paraffin. So, I hope that helps.

Can paraffin expire?

I'm so glad somebody asked this question. Can paraffin expire? And if so, can it be used if it's expired? Paraffin from a laboratory standpoint is very inert and takes time to degrade. So, paraffin, there's formulations in there too, we introduce plastics and other chemicals to keep it stable. If there is a degradation of the material past the expiration date, it would show in the product to be more brittle and cracking more because it's breaking down those smaller aliphatic carbon chains. You can see this is a great example of expired paraffin and it cracked, not because it was put in the freezer and not because it was over-sprayed, but because it's just old paraffin.

Can the crush artifact come from a tech pressing down on tissue too hard with forceps during embedding?

Another great question, can the crush artifact come from a tech pressing down on tissue too hard with forceps during embedding? Yes, absolutely. Tissues like maybe uterus or liver are a little bit more, you know, resistant to that pushing down because you do need to put some pressure on the section because you want to capture the entire section, right? You don't want one side lifting up or curling around. And so, a lot of times people will buy what we call tampers, to make sure everything is pushed down on the same plane, especially if it's small. It's critical because you put in a piece of tissue, you want to see that whole piece make it to the slide. And so yes, nuclear streaming is probably going to be absent because that seems to be a pre-analytical impact or a technique. Sometimes when people are handling fresh tissue, they can crush it in an area and cause, you know, nuclear stream. So, you probably won't see it after processing. It's usually with the embedding like this image on the left is kind of a good example of sometimes somebody pushing down too hard with forceps and just making an indentation. So, it can happen.

Would you happen to have tips for better slicing?

Next question. I'm currently struggling to slice in the microtome. My tissue gets folded with holes, which results in terrible staining. Would you happen to have tips for better slicing? Thank you. You're welcome. I hope I can offer you a few answers or at least tips or maybe things to consider that would help you resolve this issue. And then I can say you're welcome, maybe. There may be more than just one issue going on here.

One thing I do want to mention throughout this presentation, maybe more than once, is that we have 17 application specialists that just support core histology across the United States. We also have application specialists in Canada. I don't know the number of people they have up there, but you're always encouraged to reach out to your application people if your instrument is under warranty or under a service contract, because that service for you, for us to come in and troubleshoot if it's related to your tissue processor or your stainer, even your microtone, we offer those services. So you can call us and set up an appointment that we can come in and maybe, you know, sometimes on-site is very valuable on determining the root cause of a problem and finding resolution with, you know, talking to staff that's there on site.

One of the things I want to offer too is, is suboptimal staining due to folds or holes? Or is it the morphology of suboptimal? I wasn't sure exactly where this was coming from. When we look at tissue that has holes, when you're cutting at the microtome, it's usually a processing issue. So, I would want to look, maybe number one, I want to look at your tissue size and the protocol you're using. Because if you're using a very short protocol and your tissue is rather large, you can see where there might be a problem with under-processing. If it might be just the opposite too, you could certainly have a very long protocol for a small specimen, and that would certainly create artifacts, but not like where there's holes right when you're sectioning. Embedding practices might be another thing to consider, or the type of paraffin. Like I said, paraffin comes in a lot of flavors, some with very little plastic, some paraffins have a lot of plastic. And so that might be something I would want to look at.

What do you exactly mean by chatter when cutting?

Next question, what do you exactly mean by chatter when cutting? Well, chatter can have certainly more than one root cause and is the most common artifact that we see for over-processed tissue. Also, for tissue that tends to be dry anyway, like liver is probably the first thing that comes to mind for me. And if you're cutting really fast, because it has a tendency to already be an organ that's a little on the dry side, and you're over-processing it on top of that, I can see where chatter would have a tendency to occur. But chatter is a common term used in histology. You're going to have these elongated, you can see the image here, you're going to have elongated thin and thick areas that run parallel to the knife blade. or may even fragment within the section. Dense or calcified tissue structures are also prone to these thick and thin. It's just nature, it's very difficult to perfectly process structures like bone or dense cartilaginous tissue. And so, I think the tip I would have there is even though you've processed this tissue as best you can, I think making sure you're not cutting too fast to create that chatter would help. Of course, uterus is also a very dense, drier organ that we see that can see a lot of chatter with and blood. Bone marrows, clots and placentas always create havoc in the histology lab when it comes to these types of specimens. And we, you know, even though we can have an optimal processing protocol, you may have to actually maybe have a little water on top of your ice that you could soak it a little bit and then take your time in cutting a nice section right off, you know, maybe just maybe one or two sections in because usually that first section off the block is going to be a little wet. And so maybe the next one will take that and it should be a lot better.

How should I approach fixing chatter on glass slides?

How should I approach fixing chatter on glass slides? Well, even after, say, 42 years of histology experience, I don't know that I know of a way to reverse this artifact, even if you were able to get the section back off the slide, which is difficult to do in itself. So, my only answer for this slide would be just to recut it. Next question. I personally have trouble with the parts of bone like sternum, femur, chunking out of the block during microtomy, even with the surface decal in addition to initial decalcification with formic acid. Can you help me? I would love to. Well, as I had mentioned in the previous slide, these type of specimens are tough. Even if your section is thin, no more than three millimeters, and it doesn't fill the cassette so fluids have access to actually process, and you've needled down that processing protocol where it's just perfect, it still seems that sometimes there's going to be areas where most of it's decal, but one area is not. Or it's tissue that you didn't expect to have, you know, calcifications in.

The only thing I would want to look at is what's the type of tissue and maybe understand why the under decalcification occurred. You know, with bone marrow specimens, we have to decal those, but we must take into consideration that we just can't use the decals come in very different flavors. There's formic acid, right, hydrochloric, there's blends, and there's EDTA, right? So, you must think about If I'm doing a femur, I might be able just to use formic acid. If doing bone marrow and I'm really concerned about the hematopoietic structures in that sample, I may want to take the EDTA approach. It does take a longer time, but it's more, it's better, it's more controlled, and it's not as harsh, especially when you're going to be doing IHC. So just think about that, especially Kappa Lambda. Kappa Lambda is very sensitive to acids, and so you really want to use the EDTA. Take those things into consideration. Maybe you didn't decal long enough and certainly that resolves a lot of microtomy issues.

You can, at the microtomy station, surface decal. Maybe you thought you had a decal like you needed to. You get it on the microtome and oh you're just having problems like you see in this picture here where there's you know open spots. You can see that it's obviously a decal problem and you can just lay that block on maybe a four-by-four piece gauze that's been soaked with your decal solution and just do a surface decal. Make sure you rinse all that decal off before you put it on your metallic microtome, so it doesn't degrade anything. And then just take right off the top. I hope that helps.

What would be an effective way to decalcify arteries?

Moving on to the next question. What would be an effective way to decalcify arteries? Great question. Again, formic acid is the most popular in the laboratories I've worked in. There are ones that are certainly a little bit harsher, like hydrochloric. And I'll just list EDTA as another decal, but it certainly takes a long time. It's usually used for bone marrows. And usually with arteries, you're not going to be doing a whole, I don't know that there's a lot of cellular structure that would be as sensitive as bone marrows, but you could certainly use the EDTA. That was an easy one.

Would you use vacuum on your fixation, ethanol and xylene steps or just your wax embedding steps?

Moving on to the next question. Would you use vacuum on your fixation, ethanol and xylene steps or just your wax embedding steps? So, this opens up, certainly, I love this question because this is where I live and breathe a lot of times. And it's certainly open to interpretation, preference, maybe where you were trained at on what they taught you about processing. And maybe that's the only way is from the facility maybe you worked at or trained at. But it's open. There are so many different protocols, and you can create so many variations. It's probably unlimited. But the main goal of tissue processing is just to remove your free water. infiltrate, be able to infiltrate with a medium that will allow you to suction. That's in a nutshell, right? Protocols vary and they need to be customized for the tissue that's being processed.

It's not uncommon to see, especially if you go back to your textbooks, even to see pressure and vacuum, what we call cycling on a lot of the reagent steps, because it's kind of a push and pull effect, especially if your specimens are dense or if somebody that's not, you know, prosecting the way they should and they've maybe put more in the cassette that needs to be. It's thicker than it needs to be. PV is kind of tool we use to kind of overcome those barriers that would normally create under processing. So, it's not uncommon to see PNV on the reagent steps. I don't like to see it for me. I don't like to see it on the paraffin because that's the last segment of your processing. And maybe you could do it on the usually know there's three paraffin steps and you could do it may be on that first one. But if you're pushing and pulling and there's a little bit of residual xylene in that dense specimen, you're going to get some artifacts once you stain, or even when you're cutting maybe, if it's enough in there, that's going to cause some issues, right? And so I like to put just vacuum on the paraffin steps so that I'm sure that pulling in one direction, I'm going to get rid of any residual xylene or whatever clearant you're using that can cause the morphology, especially after staining, look appear like hazy.

Take note that once you get xylene trapped in tissue, it's hard to get that out. Once you've completed your processing, there are cassettes. I want to make note, too, with this processing. They were talking about pressure vacuum. If you don't have an instrument that has a pressure vacuum option, I believe most do, unless it's the instrument similar to the old technicons. Then you can purchase cassettes that actually have side vents that kind of can sort of take the place, not as well, but they have side vents that will help with the fluid moving into the cassette. And so that if you don't have that option, you can certainly, you can even use those types of cassettes with the side vents on protocols that do have even pressure and vacuum. That helps overcome some of the barriers that people don't prosect correctly too. I hope that helps. That was kind of a long answer, wasn't it?

During dehydration and rehydration, is there a major effect if we do it overnight for each ethanol concentration?

Moving on to the next question. During dehydration and rehydration, is there a major effect if we do it overnight for each ethanol concentration? I think this would be reasonable to do. I think, though, I would want to know the size, of course, the size of the tissue samples, because we're talking about a long protocol, I would think. You know, the goal in processing is to remove free water, like I had mentioned before. But one thing I didn’t mention was that we want to preserve, we don't want to remove the bound water in tissue. And there's not a lot of articles on bound water, but there is increasing research on it, because it's certainly, as you can see here, I have a picture here of a protein structure with bound water. And so, they're finding out that the importance of bound water, so more attention is certainly being garnered. We want to remove all the free water, keep the bound water. I would want to know the size of the tissue, right? We want to make sure that your alcohols are graded. We don't want to remove that water too rapidly or too harshly. So, we want a nice gradient, maybe a 70 and 80 and 90, if you're going to do something like that overnight. I think the whole goal of this is trying to avoid over-processing and drying out the tissue far too much.

What is the best practice to reprocess a specimen?

Leica has a great article or it’s a nice PDF of the different methods in reprocessing. Some I am familiar with; some I am not as they come from other countries. With, and I'm based here in the US of course, But before reprocessing, this is the one I am probably the most familiar with, is direct reprocessing. You see a picture here of cassettes. You're going to take those cassettes out of the tissue processor, try to let as much paraffin drain off those cassettes when you're done. And then before you reprocess, you're going to kind of blot the excess paraffin off in new cassettes, place the cassettes back into the tissue processor, and then you can just actually do your run again.

I know this, it may not appear to make sense, but there are going to be areas of the tissue that are okay. And we know that because as a histologist, paraffin will adhere to the areas that are okay, and they're processed well. The areas that are under-processed, the paraffin's not able to adhere, and so it will peel off or flake away. And so, in just repeating this run, you can skip the formalin step and start with the first alcohol. The areas that are okay will stay protected by the paraffin that's adhering all the way, of course, to the first xylene, and then it will break, and then it will, of course, melt away. That allows for the tissue that's under-processed to be exposed to those reagents and get the correct processing that it needs. I prefer this method, I like this method, and it's listed in the PDF, and we've provided the link down here at the bottom right-hand corner. I hope that helps.

What is the single most important step to ensure optimal tissue processing occurs within the chosen platform every time?

I think when you look at what your focus of your studies are, your targets that you're going to be staining, whether it's IHC/ISH, I think you have to think about not just pre-analytical variables, but also the time to fixation. So, we use these acronyms now because pre-analytics is becoming a big focus and conversation pieces in pathology. And we use the acronyms PAV for pre-analytical variables and TTF for time to fixation.

We also must take in consideration ischemia time. Time that the organ or whatever you're targeting to pull from your host or animal is the time you ligate to the time it reaches formalin, that ischemic time can actually have an impact on some of the areas of focus you might be studying. I know that if it's not, of course we know if it's not preserved appropriately, you know, we can't fix it, we can't undo it. No modifications can help restore that epitope, you know, that target. Whether they're even lineal or conformational epitopes, I'm not aware of a way to recover those. And so ischemic time using the right protocol with processing, but I think certainly pre-analytical. How long is the specimen now? I know that from some of the articles I've read that just within the first 30 minutes when ischemia begins, you're already starting to have an effects on signaling proteins that deal with phosphorylation right within an hour of that specimen reach in formalin, you can, I had read an article where you can lose, a lot of mitonic figures. And so I think a lot of it up front is thinking about those pre-analytical variables, but then also too, after you're sure that you've done the best in getting that to fixation and making sure it's going to be a minimum of 6 hours, if you're looking at RNA studies, certainly 12 to 24, but not going over that, then you've covered your bases there. And then customizing the protocol for the type of tissue and the size of tissue that you're going to be processing is key to having great results.

So, in that mindset, we'll over fix specimens, then we're going to move to the other side of effect IHC staining. So, we can certainly, we've talked a lot about under-processing and pre-analytical variables. Let's look at the other side of that. And I mentioned that formalin certainly can have adverse effects on tissue that it sits in formalin too long. You know, you're not going to be able to recover those epitope sites. You're not going to be able to break some of those bonds. So, you can certainly over-fix to the point. where your IHC's not going to perform well. You can see here, this is based on an article I pulled. I have the reference down at the bottom right-hand corner that you can take a look at. That depending on the time and the antigen retrieval, you may not be able to open up those antigenic sites that are being targeted. You can see here. I think 1A was like, you know, the correct time, six hours. Then they did, they did pass 12, and then this is the third, like what's labeled 1C, I think is past the 24-hour mark. So even though you're doing antigen retrieval, you may not be able to open those sites back up.

That concludes all the questions we wanted to come back and try to answer. Please visit us at the Leica Biosystems Life Science Portal. I know that my webinar from last year is on there, and I'm hoping this one will be on there too. I think it will be. Read and download. We have a lot of educational content that you can peruse. We also have a fantastic virtual stain gallery that you'll want to just check into. You'll want to bookmark this page in case there's things you want to go back to. Maybe there's, you know, antibodies you're using, and you want to see what that looks like on a virtual stain. We have that for you.

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Thank you so much for taking the time to take this journey of processing, optimal processing with me, answering, hopefully answering these questions the best, to my knowledge, I could do for you. Certainly open to any more questions that you might want to send in, or we would be happy to look at those. Thank you so much. Happy processing.