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Processing Fatty Specimens

PACE credits are no longer available for webinars more than 6 months old.

Overview

One of the most critical steps in histology is fixation, especially when it comes to fatty tissue. If a specimen is not well fixed, the lack of full fixation could affect staining steps and possible diagnosis. A laboratory thoughts of what alcohols to use in dehydration steps and the solutions used in clearing steps in optimum processing of fatty tissue is an important step to get these tissues to ultimate processing and fixation. This presentation will review the process of processing fatty tissue, processors types, and good quality measures for ultimate fixation of fatty tissues.


Learning Objectives:

  1. Review the process of specimen handling before processor.
  2. Review Fatty Specimens and why they are so difficult to process.
  3. Overview of the different technologies for processing and solutions used.
  4. Expand on good quality techniques for processing fatty specimens.

Webinar Transcription

MS. MARI ANN MAILHIOT:  Good morning everybody and thank you for attending.

     First off let’s talk about our objectives this morning.  We’re going to review the process of handling specimens for the processor.  That’s really important how we handle them, what we do with them when we get them into the laboratory, and things like that.  If there’s no formalin we really should add formalin unless the surgeon specifically says don’t do it.  We want to analyze the reasons why fat is so difficult to process, and we also would like to discuss the different technologies that are available to us now to process fat, including the different solutions that are on the market, and we want to also identify good quality control for fatty specimens.

     Of course, we’re going to have to start off with grossing, that is one of the most important things in histology.  If your specimen is not grossed properly you’re not going to get the proper results that you’re looking for.  The next step after grossing is going to be fixation.  Are you going to use 10% buffered formalin?

     Now, in the case of us presenting fat today, I’m thinking more of breast specimens so we do have to follow the CAP regulations about using neutral buffered formalin.  Once we’re done with the fixation of course, we can go to dehydration on our tissue processor, it starts there and we have graded alcohols that will help us with that dehydration, and then we're going to talk about clearing reagents.  We all know that xylene is one of the best reagents in histology, but we also know that there are some affects to our health from using xylene, so now the world of histology is thinking about maybe we should be xylene-free, and that’s not a bad idea from someone who suffers from adult-onset asthma as being exposed to xylene in the workforce long before we had proper air exchanges, and we didn’t even have air conditioning at University of Chicago, we had to open up the windows.  You can understand how important this is to me to know that we’re going in that direction.

     Last, but not least, we have infiltration with paraffin and the thing with paraffin we have to remember is you have to use the paraffin that’s the best one for you and your lab.  There are so many opinions about paraffin in the histology world that I could not begin to say use this one, this one, this one.  It’s entirely up to you and what works the best for you.  There's no way you want to be frustrated sitting and cutting at a microtome because your paraffin is not the best.  Last, but not least, is we have protocols for processing fatty specimens on some of the higher end tissue processors today.

Grossing

     We get our specimens into histology and to the lab and let’s hope there's proper identification on those specimens, let’s hope the surgeon has given us information about where that specimen was taken from, and possibly a little bit of history about that patient.  Once that’s done and we’ve checked everything, we start our grossing process.  Most cases if it is a breast we’re going to have to ink margins.  Now we do inking in other types of histology to skin when we have an elliptical piece of skin, and I’m mentioning skin also because it does have a fat pad on it and sometimes it’s really hard to process skin specimens when you don’t get that fat fixed properly on that specimen.

     The selection of the most represented pieces you’ve got this big breast specimen or this big skin specimen and you have to look at it and say which do I think is going to be the best piece for me to take and put into the cassette.  Sometimes that’s a thinking process as you look at that specimen.  Then you want to get the specimen down to an ideal size.  Optimally we have 1 to 2 millimeter biopsies for rapid processing and we have 3 to 5 millimeters for routine overnight processing.  We have to really think about this process ahead of time to make sure we get our specimen to the optimal size so that means no more stomping on the cassette to close it. 

     [Slide 6 00:06:31] You can see that I have a picture here of a cassette with a piece of tissue inside and there’s actually two pieces of tissue.  There’s enough space in that cassette for fluid penetration throughout the tissue.  That’s really, really important and throughout the cassette you want the fluid to get into the cassette and get into the tissue.  If that cassette was totally filled there would be no room for a proper fluid penetration.

Fixation

Our body is made up of all kinds of proteins and lipids, carbohydrates, and inorganic salts.  What we need to do in fixation is to coagulate everything in our body including the proteins.  That’s really, really important.  Once we do that then we stop autolysis which is the self-destruction of the tissue.  Stopping autolysis does prevent the tissue from acting upon itself and that’s really important.  We break down those intracellular proteins and we also prevent decomposition.

     It probably would be important for me to bring up something I think is important.  I have pleasure last summer of working with someone in a histology lab that does regular histology but they also have been asked by the medical examiner in their area to take on their specimens and process them for them.  They did not have a regular histology lab.  She called me because she was really frustrated about the fact that her specimens were not processing properly.  She could not get any proper sections and she was extremely upset about that.

     Basically, her and I worked through all of the details of her processing and I have to say she really was on top of everything, she did all of the things that are required of an excellent tech, and we whittled it down to the fact that the medical examiner was offsite so they put their specimens in a bucket of formalin.  One of the things that she told them is she wanted the formalin changed once before it comes to the lab.  Probably that was not done at that lab.

     I asked her when she received the specimens in her lab to change out the formalin.  When we started doing that, her specimens increased so much, everything went so much clearer and much better on the slide so there’s a lot of things that we have to remember with fixation and one of them is to make sure we have clean reagents, we change our formalin on a more normal basis, so if you start grossing at 8:00 in the morning maybe at 12:00 or 1:00 you dump that formalin out and you either add more specimens to that tray or you start up another tray, but make sure you empty out your formalin.  It is a big concern of mine.

     The other thing that we have to remember, and I can tell you, you can see this in your H and E slides your hematoxylin will be a grey color and your eosin will be very pale.  That’s because the fixation started in formalin but it finished in alcohol.  You know that you did not have enough time in your fixative. 

     What are some of the properties of a good fixative?  It should penetrate that cell very rapidly, it should also, starts from the outside, goes to the inside of that specimen and it takes a while to reach the area that it needs to fix.  What am I saying about this?  I’m talking about big specimens.  You better cut them open and make sure that it has two or three or four points of entry into that specimen, so the middle can get fixed and so can the outer edges get fixed and hopefully the two will meet at some point.

     When you bisect something you’ll always come back, especially if it’s a big specimen, and then take representative specimens from that.  Columns need to be pinned on a board.  They have fat also and if they’re not pinned and opened properly they will not fix property and we know that for a fact when we sit down to cut specimens.

     Here is something that I thought would be a good reminder of what happens when we don’t cut a large specimen open.  You take a look at the picture on the right-hand side and there’s a little blue box around it and a red box.  There is a pink area at the top of the uterus that is not fixed.  The next day you come into the lab and the pathologist brings you the slide, Mari Ann everything was so beautiful on these slides except this one particular area.  I don’t understand.  There’s a lot of distortion and it doesn’t look like it was fixed well but I don’t know how that happened because everything else is so well.  It happened because we did not open up that specimen and submit that specimen to formalin and that’s very, very important.

     If your fixative has a job to do that job is going to be stabilize the tissue.  Why would you want that stabilized tissue?  Because we’re going to subject that its use to really rough tissue processing.  Going through alcohols and going into a clearing agent is tough on the tissue and then going into paraffin so we have to make sure that we have taken care of toughening up that specimen for the rigors of processing.

     First thing we can talk about here is my choice of fixative.  We know that formalin is our fixative.  We’re going to use this for fixing all specimens and I did find some interesting information when I was preparing the presentation.  Back in the later 1900s there was a gentleman by the name of Frederick Blum and he discovered formalin but used it as a disinfectant.  I’m glad I wasn’t back there when he was disinfecting the laboratory because that stuff is really bad.  Also, in 1987, the government said we’re going to standardize fixation and embalming and we’re going to use 10% buffered formalin.  That’s just a little bit of information for you.  Those of you that have been in histology for a long time, I’ve seen changes in histology that I just can’t imagine.  I’ll be honest with you.  It’s all for the positive, so we’ve really improved how we can process specimens and how we can get the best results to our patient.  That’s really, really important.

     The other thing I want to talk about is tissue thickness.  I made the comment about no longer stepping on the cassette to close it.  That is very true.  Back in the day the specimen was falling out the sides of the cassette and that just is not acceptable.  The other thing too is postmortem storage.  I can address that a little bit because if come from a big teaching hospital and we stored everything from the 1920s on up and some of it was never thrown out.  You know that some of those specimens that were stored in the early twenties the formalin was never changed out so now we have formalin acid and really and truly that specimen isn’t that useful if we wanted to any research on it.

     The other thing to remember once you open up the bottle of formalin it starts crosslinking to itself.  Second of all once you’re all done with a specimen and it’s properly fixed and you want to store it and you put it back in the bottle and store it, that formalin is going to continue to crosslink that specimen.  I don’t think I really thought too [Slide 12 00:15:38] much about that crosslinking myself.  I did later on when I was doing IHC and stuff like that, but I really did not give it that much of a thought.  Good information for you to thinking about definitely when you’re storing your tissue.

     What about fixation and desirable properties of a fixative the thing that I like the most is that it prevents shrinkage of tissue.  We don’t want our tissue to shrink.  The other thing is it provides the hardening of the tissue for intracellular components to be retained during processing.  That is important.  That’s what I was talking about the rigors of tissue processing.  It activates the enzymes so they don’t destroy the protein.  That’s really important.  We’re going to always make a reference to staining properly.  You want it to enhance staining.  You do enhance staining if it’s properly fixed.

     Formalin crosslinks proteins creating a gel and that stops that protein from acting on its job in our body.  That’s really what that means.  Once that action stops then decomposition is going to stop and some other things that go on if we don’t have that specimen in formalin.

     The other thing that I think I want to stress here is there’s a relatively short time for fixation.  Again, I have to bring up CAP because it’s a minimum of 6 hours to 48 hours for breast specimens if it’s a HER2 new.  I’m going to keep on bringing that part up but generally in some cases formalin is a relatively fast fixative.  You’re not going to have 48 hours if you’re going to open up your specimen and expose it to formalin.  That’s also something to think about.  I think we’re doing really well in histology now because I think we’re stepping up and saying no, we can’t do this anymore; we have follow the guidelines, and of course, when we’re being asked by CAP inspectors to show documentation of everything so that really, really is important.

     The other thing I can probably mention too is that sometimes we all use a different fixative and the ones that comes to mind is Bouin’s for specific things that we want to enhance in staining but the application here for fat is not necessarily that important because we don’t use Bouin’s.  A long time ago when I was in University of Chicago some of the pathologists decided they would ink their specimens and then put them in Bouin’s, put them in the oven and then take them back out again and that would help the inking process.  I don’t know exactly what that did to the whole process with the breast.  There were some complaints about staining in ICH from that.  We have to be really, really careful now in histology, we have to follow the guidelines.

     Universal fixative is formalin.  I told you that in 1987 government said we’re going to have it as a universal fixative and an embalming solution.  To be effective the specimen has to be in 20 times its own volume so you can’t have this big piece of tissue in a teeny tiny bottle with a very little bit of formalin inside nor is it ridiculous to put a biopsy in a big container.  You have to pay attention and especially if a lot of your specimens come from the outside so you're an outreach lab and that lab sends you specimens to process for them.  They’re not necessarily histologists and I certainly don’t want to put any body down for not doing their job, but the theory and the philosophy is not there.  They don’t understand so a lot of times specimen bottles are dumped over and sometimes formalin is not put back in that bottle.  Sometimes they, like I said, put a big piece in this tiny bottle.  Once you get the specimen you have to take it out and hopefully there’s not a whole lot of damage.  If you get the specimens in 24 hours, 24 hours is almost too late.  If you get them within the same day you might be able to salvage something.  There has to be some kind of training by you if you are doing an outreach to doctors’ offices.

     I’m going to stress that formalin is definitely the fixative for HER2 new staining.  I apologize for doing it so often, but my point here is that we have to pay attention to this now that it’s so important to CAP and they said this is the way we want it to be.

     A common form of formaldehyde is 10% buffered formalin and formalin as you know, is a gas and to that gas we add water till it’s a 40% formaldehyde and then we bring it into the lab and we do our dilutions to have it 10% formalin.  We do add buffered salts to stabilize the formalin and that makes the fixative keep a lot longer.  The other thing is, buffered formalin if it’s not buffered will leave a pigment behind.  I do know some people do use unbuffered formalin and that is their choice.  They have specific reasons why they do that and they know that they have [Slide 16 00:21:53] to remove that pigment in order to have a good stain.

     Alcoholic formalin.  This is going to be a big controversy.  When it comes to question time at the end of the presentation, if somebody is using alcoholic formalin they might want to spend a little note into Rick and let him know that you are and just offer how well it works for you.  I still have to remind you formalin is first.  You have to fix in formalin.  The minimum is 6 hours.  The max at this point is 48 hours; understand that it may change but I have not heard of the changes yet.

     What about alcoholic formalin?  It can be used as a secondary fixative and the good thing about alcoholic formalin; not only is it a fixative but it’s a fixative that’s been mixed with alcohol so there’s some dehydration going on at this time.  That might help the specimen but again, if you’re going to change anything you must validate all your protocols. 

     I’m going to fix my specimen this long in formalin and then I’m going to switch it over to alcoholic formalin and validate everything.  You have to have a specimen that’s only fixed in formalin and then the same specimen or same patient that you then submit to formalin and then submit to alcoholic formalin.  I’ve heard a lot of positive feedback from alcoholic formalin but again, I tend to worry more about what CAP wants from us.  It is something for you to think about.

     I did put a little blurb on here and I wanted to see if everybody was awake.  It says one to two hours.  That’s just a statement.  It doesn’t mean that you’re going to do that.  That’s too short of a time and I just wanted to make sure that you guys were awake.  It can be several hours afterwards.  If you only fix your specimen for six hours and you want to do the rest of it in alcoholic formalin you know, it’s going to be longer than this.  I put an arbitrary time in there just to make you stop and think about this.

Dehydration

     What about dehydration?  We’re going to start with our alcohols.  Everybody knows in histology ethanol is the best alcohol that you can use for tissue processing but there’s also methanol and isopropyl alcohol.  Isopropyl alcohol I’ve never been fond of it but it can be used in special processes and work very well.

     The other thing is methanol.  Methanol is a wood-based alcohol and our friends in histology that do plant histology process with methanol and they get really good results.  Again, the staining is going to be very different between methanol and isopropanol.  You’re going to have grayer hematoxylin and you’re going to have paler eosin.

     In dehydration, the water slowly is replaced with alcohol and we start with a low volume of alcohol.  Let’s say 70%.  Again, I have to stress 70% is important because it gets rid of the formalin salts.  If you don’t get rid of the formalin salts and you guys have one of the tissue processors on the market, they all use rotary valves now.  The rotary valves will stall because there’s a salt build up on that rotary valve.  Seventy percent alcohol is extremely, extremely important.  Once you get through the 70% you can go to 80 or you can skip 80 and go to 95% and go to 100.  You basically brought your tissue from a water state down to an alcohol state which is really important.  Again, you have diffusion of alcohol and into the specimen and water out of the specimen.

     At the end when we’re done with all of this you need to have 3 to 4% water left in the tissue.  That’s the bound water.  If we don’t have it that’s when we have a difficult time cutting our specimens on the microtome because they’re extremely dry.

     Ethanol is the best for processing but what happens if your hospital does not want to pay for the license for the alcohol.  It is a money problem sometimes in the budget.  You have option.  You have reagent grade alcohol which is 95 parts ethanol divided between methanol and isopropyl alcohol.  They make up the other 5%.  You do not need a license for reagent-grade alcohol so that would help.  The only thing that I could see that would be a difference is probably you’re going to have to change your alcohol more often when you use reagent-grade alcohol.  Don’t feel bad because you can’t have ethanol in your lab.  You can certainly accommodate all of your processing and staining to reagent-grade alcohol.

     Dehydration.  Isopropanol is a new, how can I say this, dehydrant and clearant in this histology world.  I would have never thought this 25 years ago.  I never would have anticipated this but it really does work in the world of xylene-free processing. 

     First off as a clearant it is gentle and penetrates slowly.  It is compatible with xylene so that’s a positive.  For those of you that recycle, it recycles only back to 88%.  In its purest form, it’s only 99% so there is 1% water left in ethanol.  As I said, there will be some recommendations for reagent changes on your tissue processor. [Slide 20 00:28:25] The best way to know what those changes should be is to get together with your vendor and work with them to decide.

     Let’s talk about dehydration and xylene-free and isopropyl alcohol.  For your xylene-free protocols you can have two steps of 82/20 ethanol/isopropanol mix.  What are we doing here?  We’re using most ethanol, or in the case if you’re reagent-grade, reagent-grade you add your isopropanol to that and it should be about 80 parts of the alcohol to 20 parts of the isopropanol.  Then once you have two steps of this, you’re slowly introducing the tissue to IPA you’re then going to have three stations of IPA.  Now the specimen is ready to receive that isopropyl alcohol.  Time in end station can vary, even the heat on the steps can vary.  It depends on the size of the tissue always and your larger specimens need more time and can have a longer time in heat.

     You will get rapid turnaround time when you use xylene-free protocols and they can only be used on high ended processors that are made specifically for that.  Your low end processor really cannot process very well xylene-free.  You can use the xylene substitute but you cannot use isopropanol effectively on the lower end tissue processor.

Clearing

What about your clearing set?  This is where the tissue is infiltrated with some sort of clearing agent.  You have to make sure that your last step, which was 100% alcohol, is compatible with that clearing reagent.  The other thing is it doesn’t over harden the tissue.  Xylene does over harden the tissue but when I started in histology we didn’t use xylene; we used toluene.  Now everybody knows they’re probably oh my gosh, because of the carcinogenic effects of it, but it’s a gentler reagent and it’s softer on tissue.  It doesn’t harden it as much.  We do have people that still use toluene in the lab.  They feel it’s a better clearing agent in the lab and I take my hat off to them.  They do have to have proper air exchanges and they have to make sure they’re monitored for any exposure.  What else would I like to say?  Your clearing reagent is compatible with paraffin always.  If it’s not then it’s not going to work.

     Other things to think about; clearing reagents.  You know my favorite is always xylene.  I’m sorry; I guess I’m an old-school tech.  Xylene substitutes, why should we bother with them?  Because they’re better for our health, they’re better for our tissue, and frankly, some people are going xylene-free.  Maybe they don’t want to go the isopropanol route and do it that way but they definitely want to purchase xylene-free reagents from the different vendors that are out there.  That is a possibility for them.  I’m still going to tell you that it’s okay to do that.

     I’ve processed with xylene-free reagents and I’ve also processed with isopropanol as a clearing reagent and xylene-free the results are phenomenal.  They really are phenomenal.  The sections are beautiful.  You have nothing to worry about in terms of the quality of the tissue.  However, I do like to have xylene in my lab for cleaning the tissue processor.  The other thing to think about is that we do have some xylene substitutes that smell like oranges.  They’re not recommended for use on an automatic tissue processor because there’s orange oil in that mixture and it does gunk up when it gets in with paraffin.  It gets a muddy buildup on your rotary valve and on the valves on the inside of your tissue processor.

     The other thing too is I never was very fond of it because it made me sick, but I do know there are some people out there that really like using them.  Again, it’s a gentle solvent so if we don’t want to expose our tissue to harsh xylene or toluene or whatever we’re using then I guess the orange smelling substitutes are okay to use.  It’s just going to be a matter of opinion I have to tell you folks.  It’s always a matter of opinion in the laboratory.

     Paraffin is going to support our tissue once we’re done tissue processing with our xylene.  There are different paraffin types on the market.  There’s different melting points.  Some of the paraffin has plasticizers in it, some of the paraffin has a rubber additive and some have a beeswax additive.  Back a couple of years ago, so I could have a better handle on paraffin, because I had so many questions about is that paraffin too hot for my samples.  I called British Petroleum and I talked to one of their chemical engineers there and he said when paraffin leaves our plant it is 100% paraffin.  There are no additives in that paraffin.  It doesn’t burn until 4 or 500 degrees but what happens when it gets to the companies that purchase it to make paraffin they add their additives.  That’s what brings the temperature for burning or catching on fire of paraffin down much, much lower.

     What I’m going to say about paraffin is it’s the rule of thumb for the lab.  Please everybody, everybody has to sit down and cut, everybody has to be able to work.  You have to find a paraffin that’s going to work for everybody in the lab.  I worked at University of Chicago with 16 tests in the lab.  I can’t say what paraffin we had there at the lab but I can tell you it was optimized to be the best for all of us in the lab.  We had senior techs, we had middle techs, and then we had brand new techs.  Everybody was able to cut. 

Protocols for Fatty Specimens

     Let’s start with our most basic processor.  I’m going to spend a little bit of time, hopefully I didn’t talk too long.  We’re going to have two stations of formalin, but let’s think about this.  Do we have to have two stations of formalin?  I’m hoping that if we’re doing breast specimens or colon specimens that we did our fixing at the grossing station, now I know I’m asking for a lot, but I’m doing my stuff at the grossing station why do I need to have a second formalin on my tissue processor?  Think about taking that off if possible and then you go to a 70%.  Seventy percent is needed for getting rid of the buffered salts in the formalin, then you can have 3-95 stations, 3 95% alcohol stations.  You can also have 100% stations, 2 of them but if you use a xylene substitute, you’re going to have to have 2 stations of 95, 3 stations of 100, and 3 stations of xylene substitute.  You need to make sure that specimen is free and clear of water.

     Last, but not least, is our three steps of paraffin and I get this question all the time; I have to use the same paraffin that I use in my embedding center as I do on my tissue processor.  Absolutely not.  You can process in, there’s infiltration mediums on the market.  You can put in one kind of paraffin and then use another one in your embedding center.  The thing to remember is try to get the melting point of both paraffins to be around the same so if you’re using a paraffin that’s 55 to 60 degrees on the machine and at the embedding center they’ll work better but you can definitely use two different types of paraffin; one for embedding and one on your machine for infiltration. 

     Pressure and vacuum has been overrated and everybody says I need pressure, I need that crutch.  Understand that some studies have been done that it’s not necessary.  It’s only 35 KPA pressure.  That’s not very much.  If I break that down to pounds per square inch it’s about 5 pounds over a square inch.  That’s not that much.  We can get away from the crutch of pressure if we want to.  The other thing that I know pressure does affect is the pump on your tissue processor.  It has a diaphragm that’s made of rubber.  It’s like your lungs; it’s breathes in and out.  That pressure does affect that diaphragm.  It’s something to think about.  I know we all can’t possibly get away from it and I understand because I still tend to stand on my crutches a lot.  The other thing is vacuum is necessary usually in paraffin to help push that paraffin through the tissue and that is important.

     Protocol for a higher end processor.  I’m thinking in my mind that all of you have properly fixed your specimens, so once they’re properly fixed you put them into the retort, and this is the single retort processor.  One step of formalin is all that’s necessary.  Then you have a step of water.  Water is used only to get rid of the salts and rinse the tissue from the formalin.  That’s all it is.  It’s a short time; it’s not a very long time.

     You have six steps of graded ethanol, three steps for the wax, and times in each station can vary, so can the temperature in each station.  Usually what happens is you’d like to have the shortest time in the dirtier solution and the longer time in the cleanest reagent.  In some cases, the machine will have a densitometer, can measure the alcohol and it’ll have autorotation so that densitometer measures the 70% alcohol when it gets to a certain point the machine auto rotates and moves the next alcohol into the position of 70% so that alcohol is going to be somewhere around 70%.  The other alcohol is added at the end.  That is something to think about.  You can do xylene-free processing on this type of tissue processor.  Also, you can run this tissue processor in reagent management mode which means you decide how you want to have your reagents changed.  The machine doesn’t.  You decide.  You put in your variables and things you want.  It has a lot of options for you.

     I have to tell you that the xylene-free protocols that come with this particular cannot be edited.  They’re factory protocols.  What does that mean for you if you want to use them?  That means you go ahead and validate your tissue on that xylene-free and validate accordingly and see if you like it.

     The other thing I didn’t stress enough probably is when you get a processor please, please, please validate everything, it’s really, really important as a histotech to know.  I know we have great vendors out there, we have great salespeople out there but the thing of it is in our histology world I’m going to try your processor and see if I like it and I want to try to adapt it to my lab.  Think about that.

     There is a more rapid tissue processor that is on the market and again, one step of formalin, six steps of varying grades of alcohol.  This processor operates by an algorithm.  What does that mean?  Every time you process a program that alcohol is calculated at the end of steps.  You go through a 70% or possibly the name of it will be ethanol.  That ethanol is evaluated and its percentage is dropped down after that step.  This is a very intuitive tissue processor.  It’s a relatively new type of tissue processor.  We’ve never had this before.  It’s been on the market for several years but it is a mathematical process.  There are three xylene steps and there are three wax steps and processing times are definitely variable.  You can do xylene-free.

     The other thing I’ve forgotten is you have two retorts.  What a wonderful thing.  You can process biopsies in one retort and do your larger specimens in another retort. 

     Last, but not least, you also have the high-end processor which can do the xylene-free and it’s still the same process.  You still have your 1 step with 70%, you have an 85% ethanol, then you go with your IPA which is 80 parts of ethanol, 20 parts IPA.  Last, but not least, you have the three steps of wax.

     Again, these particular protocols cannot be edited but there are some high ended tissue processors where you can edit them.  Comes with factory-recommended and tested protocols but I would get together with your vendor to see if it’s necessary for you to adjust any of those protocols.

Summary

First off, let’s talk about the size of our specimens.  Again, 1 to 2 millimeter biopsies, no more than 3 to 5 millimeter for larger specimens.  I keep stressing this but it’s really important because I still get calls from people who have huge, humongous specimens submitted into their tissue processing.  We have to get the word out to everybody as much as we can, and some people are more open to new ideas, and of course, some people are not.  Realizing that is really important to process your fat very well because in terms of breast biopsies we do have to follow what CAP tells us to do.

     We have to remember to change that formalin out in our grosser and that’s really, really important also.  If we don’t do that, just think of it, and it’s really a very odd way to think about it, but if you come from a family of ten and you lived in the 1800s and you only took a bath once a week there was a washtub and the baby went first and the dirtiest kid went last.  You can imagine the dirty kid.  I feel bad for him because he probably had a bath but not like the little baby.  It’s the same thing with our formalin.  If it’s dirty it’s not going to do the job.  Make sure you try to remember to change your formalin out as much as possible.

     We also had an introduction of using IPA as a clearant and as a xylene substitute and that’s really going to probably be the way we’re going to go in the future.  My feeling is I like xylene-free very, very much.  The results are as good as xylene.  The only thing that I feel that’s important is we do have to have xylene in the lab for cleaning of our tissue processor and for some other things that we need too, but it cuts down on how much xylene waste we have if we go xylene-free.  Imagine getting in four gallons and keeping four gallons, or let’s say even eight gallons and keeping them for two or three weeks and that’s all you have to send out to be disposed of.  It’s a big cost savings.

     I also talked about having many types of paraffins in the lab to use and it is a lot of different types of paraffin.  It’s going to be up to you.  I can’t tell you which one to use.  You have to decide what’s best for you and all of your other lab techs.

     Fat will process well on a basic tissue processor we know this.  Please, if this is all that you can have in your lab be proud of it, be proud of that fact that you can have that tissue processor.  You can optimize it for use to make the results that you want in your lab.  No one says that you have to step up and get the other types of tissue processors some people just can’t do it.  There’s a money element there so be proud of what type of processor you have.

     The other thing is that we found a mid-road basic processor with still one retort and you could process specimens in that retort, but that high end tissue processor has two retorts which means you can run a biopsy protocol and you can also run an eight or ten-hour protocol in the other retort and have everything get finished in time.  If you’re a 24-hour lab that means the processor finishes, you take the biopsies out, you clean it, you put in another run of biopsies so it does give you great turnaround time.

     Just remember they will do the job but some are faster than others.

     That’s it.

Questions

     We have received many, many questions and please note that I have copied all of them and will be forwarding to Mari Ann if we can’t get to all of them.  Thank you very much for that.

     The first one is from Andrea.  We occasionally have to process breast blocks to be able to cut them.  What does that do to the specimen especially if they need immunostaining?

MS. MAILHIOT:  What you’re saying is they have to reprocess Yolanda?

MS. SANCHEZ:  Correct.

MS. MAILHIOT:  Reprocessing does not really affect everything in IHC.  My suggestion is there’s a reprocessing protocol that was put out a long time ago and you can just take your specimen and block them in the cassette and take and drop them all the way back in formalin.  Never go through the cleaning cycle and never start in alcohol.  Start all the way in formalin.  Purpose being that these specimens are already fixed, the part of the specimen that’s fixed properly has already accepted the paraffin.  Paraffins going to protect that specimen as it goes through.  The other parts of the tissue that did not get infiltrated and dehydrated property will accept that solution and you’ll have less problems with IHC.  This is a suggestion.  You have to do a little bit of research on this.

MS. SANCHEZ:  Thank you so much for that one.  David wants to know what do you think of zinc formalin.  It hardens fat cells.  Does it help.  Then, if it does, does zinc formalin have a vacuum step?

MS. MAILHIOT:  Yes, it can have a vacuum step, that’s number one and it does help but I have heard from some of my friends that they have a lot of problems with zinc formalin.  Number one it has to be zinc sulfate.  It cannot be zinc chloride because the chloride is corrosive to metal parts on the tissue processor.  The other thing is they have a hard time because the zinc deposits on the inside of the tissue processor so you have to make sure with your vendor where you get it how you need to clean that zinc away from the tissue processor.  As far as IHC I’ve not heard anybody complain about anything with IHC.

MS. SANCHEZ:  Great.  Thank you so much.  From Blaze.  If you’re analyzing lipids is there any better or modified protocol for this purpose?

MS. MAILHIOT:  Analyzing lipids, I think I’m a little unclear as to the question.  What does he mean by analyzing lipids?

MS. SANCHEZ:  I’m not sure either.  Sorry.

MS. MAILHIOT:  We can answer Blaze offline if that’s okay because I’d have to look into that.

MS. SANCHEZ:  Absolutely.  Absolutely. 

MS. SANCHEZ:  This one says, can you use 70% ethanol and then continue with IPA?

MS. MAILHIOT:  No, you cannot.  You have to introduce the ethanol in the IPA mixture.

MS. SANCHEZ:  Awesome.  From Elizabeth.  Have you ever used sponges in your cassettes for the formalin step?  Could that lead to possible brittles during sectioning or would the brittle part be improved in the alcohol steps?

MS. MAILHIOT:  The brittle is probably coming from too much time in the alcohol.  Remember the focus here was making sure that we have our specimens properly fixed.  If a specimen is properly fixed chances are exposure to the alcohols are not going to cause the brittleness.  The other thing with sponges is you do have carryover of reagent.

MS. SANCHEZ:  I believe that is all the questions that we can physically ask today but again, I have copied all your questions and will be sending them to Mari Ann in order for her to answer your very great questions.  I apologize we don’t have enough time to go over all of them but she will be answering you within the next couple of days.  Thank you so much.  Rick.

RICK:  Thank you very much Yolanda, and wonderful, wonderful job Mari Ann.  You generated enormous questions, almost a record-setter.  Good for you.

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