#7 Tissue Processing Artefacts

#7 Tissue Processing Artefacts

The most noticeable tissue processing artefacts are the wrinkles and tears in the tissue sections which are evident even at low power (Figure 1).  Incomplete fixation would not cause such artefacts, as the cellular histology is acceptable (i.e. nuclear and cytoplasmic staining is within quality control limits).  Improper embedding would also not be the cause of these artefacts. When skin specimens are mis-embedded, usually tearing and wrinkles would be localized within the epidermis, and within the dermal-epidermal junction.

The most likely cause of the artefacts seen in “Figure 1” is due to incomplete dehydration, resulting in incomplete paraffin infiltration.  When tissue is not completely dehydrated, excess water is left in the tissue. If xylene is used as the clearing agent, it can dissolve up to 2% water – but no more.  Also, if xylene substitutes are used, they are completely intolerant of any residual water. If water is present in the tissue after dehydration and clearing, paraffin cannot infiltrate the tissue completely.  Without complete paraffin infiltration, the tissue can tear and fold during microtomy. If this affects all tissues in the run, the tissues may have to be reprocessed by melting down the blocks and placing the tissues back into the cassettes, with new cassette tops. The cassettes can now be run back through changes of xylene and 100% alcohol.  After rinsing in 95% alcohol, the tissues can be put back into the tissue processor to begin processing at the formalin step. Finally, they can be run back through xylene and into paraffin. This procedure should remedy the problem.

Another cause of tissue processing artefacts is referred to as “nuclear streaming” (Figures 2A and 2B). Nuclear streaming is a result of three possible circumstances; incomplete dehydration, too rapid dehydration, or processing solutions that are overused and in need of changing out.  When water in the tissue moves out too quickly, resulting in “squeezing” the cell, it leaves a final appearance of the nuclei in a “streaming” configuration.

 

Other questions to ask when experiencing tissue artefacts are:

Did the tissue dry out during transport?  If so, the client should be contacted to see if they used an empty specimen bottle, with no formalin in it.  Also, the specimen bottle should be inspected to see if it was cracked and the formalin leaked out.

Are the tissue processor reagents clean / exhausted?  The tissue processor maintenance logs should be checked to see when the last time the reagents were changed out.  A hydrometer can be used to determine the actual percentages of the alcohol reagents.

What about the tissue processor reagent bottles?  Did the tissue processor reagents get swapped? The hydrometer can be used to determine the answer.  Also, do all of the reagent bottles contain fluids that measure up to the “fill” mark? There must be enough reagent in the bottles to fill the processor retort.

 

If you are wrapping small tissues, are the reagents getting full penetration into and through the tissue?  There should be only one layer of paper surrounding the tissue, not several layers.

If you are using sponges, are reagents carrying over?  Sponges are notorious for getting saturated with reagent, and then holding onto it.  If enough of the reagent carries over into the next reagent bottle, it can change the concentration and/or contaminate the subsequent reagent.

If you are using biopsy cassettes, are the holes too small, which may cause what’s called “air lock”?  Some biopsy cassettes have micro-holes in them. You must be sure that the holes are not so small that they prevent the proper movement of reagents through them.

So, when you see tissue artefacts on your slides, troubleshooting using the above techniques involving theory, chemistry, and considering the physics of the consumable products being used, you should be able to resolve most issues when they occur in your laboratory.

References:

  1. Chapman, C.M. (2017). The Histology Handbook: Amazon CreateSpace Independent Publishing Platform
  2. Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform
  3. Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory.  Submitted to J Histotechnol
#6 Understanding Tissue Processor Procedures

#6 Understanding Tissue Processor Procedures

Tissue Processing

Standard tissue processing may be carried out on any number of open and closed tissue processors, although closed processors are preferred due to safety concerns, both for the tissues and laboratory personnel.  Closed system processors are “smart’ enough to prevent tissues from drying out in the event of a power failure, and the chemical fumes are kept inside the processor; released through filters and/or vented to the outside of the laboratory space.

Another tissue processing option is the use of microwave assisted processors that use conventional heat and microwaves to adjust and maintain temperature control during processing.  Specimens are dehydrated through ethanol and isopropanol.  Then, after vacuum vaporization, specimens are infiltrated with molten paraffin.  The specimens are then ready for embedding.  There are also ancillary units that will perform automated embedding of the tissues, if the proper cassettes are used.

A major advantage of microwave assisted tissue processors is the rapidity of processing of biopsy specimens.  Small tissue biopsies of skin, prostate and gastrointestinal tissue can be processed into paraffin in approximately one hour.  This is extremely valuable for cases requiring “rush” status.  Another advantage is that, since no xylene is used, the tissues are generally much softer in the paraffin block, and therefore much easier to cut during microtomy, resulting in fewer cutting artefacts.

Processing procedures using microwave assisted tissue processors must be clearly and accurately defined, with much attention during the validation process.  The fixation and dehydration steps must be complete to ensure proper infiltration with molten paraffin.  Like routine tissue processors, the basic stages of tissue processing must accomplish:

  1. Fixation of tissue to stabilize proteins and harden the tissue
  2. Dehydration of tissue to remove all unbound water
  3. Clearing of tissues to remove the dehydrant
  4. Infiltration of tissue with molten paraffin, to ensure the embedding process is successful

 

There are many factors involved in tissue processing, which provide many opportunities for things to go awry.  Carry over of fixative into the processing alcohol can inhibit subsequent dehydration.  If the absolute alcohol stations prior to the clearing stations contain water, this will result in incomplete dehydration as well.  When water is left in the specimen, it cannot be removed by the clearant, and becomes trapped within the tissue during paraffin infiltration.  The resulting paraffin blocks will be soft and difficult to cut during microtomy.

Conversely, tissue can become “over dehydrated” if the processing times in alcohol are too long.  Tissues contain an amount of molecular “bound water’ within the nuclei and some other tissue elements.  If this water is removed during extended dehydration, the resulting paraffin blocks may be dry, scratchy, and hard to cut during microtomy.  Soaking in cold ice water, once the block is faced off, may sometimes be used as a remedy.  This is common in laboratories that use only one tissue processor to process all of their tissues, regardless of size and type.  In this case, smaller tissues (i.e. biopsies) may become dry and brittle for cutting.

The next blog will discuss specific tissue processing artefacts that are observed in the microscope slide.  Now that you have a background in the chemistry and rationale of tissue processing, you will be able to understand how you can troubleshoot and remedy these all too common processing artefacts.

 

References:

  1. Chapman, C.M. (2017). The Histology Handbook: Amazon CreateSpace Independent Publishing Platform
  2. Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform
  3. Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory.  Submitted to J Histotechnol
#5 Troubleshooting in the Histology Laboratory

#5 Troubleshooting in the Histology Laboratory

The first four blogs of the troubleshooting series focused on being proactive with regard to the prevention of sub-optimal events in the histology laboratory.  Unfortunately, we are not able to predict every single potential issue that may cause a sub-optimal event in the laboratory.  As a result, another strategy is required.  This next series of blogs explains the chemistry and theory of fixation and processing.  Learning these concepts will form a basis of knowledge that will allow the histologist to troubleshoot sub-optimal events that may occur in the histology laboratory during these phases of specimen preparation.

Living tissues are made up primarily of carbon, hydrogen and oxygen and are known as the elements of biochemistry.  Histologists need to know the chemistry of fixation, processing and staining.  Histology involves using formaldehyde to chemically “fix” dynamic, living tissue into a static “snapshot” of cellular activity.  The cells in your body are currently metabolizing energy sources and performing chemical reactions to ensure that all of your bodily functions continue, and you stay alive.  When tissue is removed from the body (i.e. surgery or biopsy), the cells no longer receive oxygen from the blood, and the cells begin to die, and autolyze (i.e. break down).  The fixation process “fixes” the tissue, and stops the autolysis process, thereby preserving the cellular structure and tissue architecture, for subsequent processing into a paraffin block.

At the molecular level, formaldehyde is a simple molecule, consisting of one carbon atom joined to two hydrogen atoms with a single bond, and one oxygen atom with a double bond (Figure 1A).  Carbon is stable when it forms a total of four bonds.  A double bond contains a lot of energy – similar to compressing a spring.  The bond wants to “spring apart” to release the energy.  It does this by “springing apart” the double bond, to provide two single bonds, which immediately bind two other molecules.  This is what is meant by the term “cross linking” fixation, as it relates to formaldehyde.  The formaldehyde molecule cross links molecules within the protein structure of the cells.  Optimal fixation is the basis of optimal processing and results in an optimal microscope slide (Figure 2A).  Suboptimal fixation cannot be remedied after the slide has been made (Figure 2B).  Even though there are procedures for “running back” specimens to formalin and then reprocessing them, the result will always remain sub-optimal.  Therefore, it is of paramount importance to ensure that specimens are completely fixed prior to processing.

Once the tissue is fixed in formalin, the proteins within are cross linked and stabilized.   The tissue is in a solution of 4% formaldehyde in 96% water – similar to the natural water content of the human body.  In routine histology, the goal is to embed the tissue in a paraffin wax block.  Water and wax do not mix.  To be able to infiltrate the tissue with wax, and embed it in a paraffin block, the water must be removed; the tissue must be dehydrated.

Dehydration is usually accomplished by using a graded series of alcohols to remove the water and replace it with 100% alcohol.  Alcohol and wax do not mix.  Therefore, histologists can use an “intermediate substance”, to bridge the gap between alcohol and wax.  For most laboratories, this substance is xylene – although now there are xylene-substitutes that can be used as well.

The molecular structure of xylene is shown in Figure 1B.  You can see that it is a “hybrid” molecule.  The center is a “ring’ of carbon atoms, with alternating single and double bonds.  The exterior is made up of single bonds to hydrogen.  This unique structure allows xylene to mix with both alcohol and paraffin.  This brings us to the first rule of chemistry: “like dissolves like”.  The middle ring of xylene is described as “organic”, which is like the organic ring structure of paraffin.  The exterior is a straight chain “inorganic” structure, which is like the structure of alcohol.

The principles above form the basis of understanding the chemical basis of fixation and processing of tissues in histology.  Once you understand them, you can troubleshoot fixation and processing issues that will occur in your laboratory, as we will see in the next blog.

Fig 1A – Formaldehyde molecule
Fig 1A – Formaldehyde molecule
Fig 1B – Xylene molecule
Fig 1B – Xylene molecule
Fig 2A – Optimal fixation. Note nuclear detail. Original magnification x 60.
Fig 2A – Optimal fixation. Note nuclear detail. Original magnification x 60.
Fig 2B – Suboptimal fixation. Note poor nuclear detail. Original magnification x 60.
Fig 2B – Suboptimal fixation. Note poor nuclear detail. Original magnification x 60.

References:

Chapman, C.M. (2017). The Histology Handbook: Amazon CreateSpace Independent Publishing Platform

Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform

Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory.  Submitted to J Histotechnol

#4 Alternative Methods for Preparing and Embedding Specimens

#4 Alternative Methods for Preparing and Embedding Specimens

Some specimens may be very tiny; on the order of less than 0.1 cm. Some preparation methods employ the use of mesh cassettes, “tea bag” biopsy pouches, sponges, wrapping paper, etc. to contain the specimen and prevent it from escaping the tissue processing cassette. A disadvantage of the above methods is that upon embedding, the specimen must be handled yet again, possibly resulting in additional fragmentation of the tissue, or possibly complete tissue loss. These are among the most difficult specimens to troubleshoot, as the most common suboptimal event is: where is the tissue that is supposed to be on the slide? Did it survive processing? Did it escape the processing cassette? Did it fragment during embedding? This blog will examine several methods of dealing with tiny specimens, such that each and every one received in your lab ends up on a microscope slide.

A proper method of using wrapping papers is described by Dr. IB Dimenstein. This method works very well for fragile tissues such as prostate and breast needle biopsies. Dr. Dimenstein notes that the use of sponges/polyester pads may result in a ‘compression artefact”, which can occur during processing. Specifically, tissue needle biopsies may compress and narrow, comparable to the size of the pad mesh holes. One solution is to lay out and wrap the core in lens paper, and then sandwich the wrapped specimen between two sponges, which have been pre-soaked in formalin.

Regarding core fragments remaining in the specimen bottle, Dr. Dimenstein recommends filtration through a porous paper, such as the internal layer of a Kimberly-Clark protective mask. This is more reliable than trying to remove fragments with a pipette, where mucous or tiny fragments may stick to the inside of the pipette. He does not recommend filtration through nylon mesh bags, as the tiny fragments are difficult to retrieve during embedding.

When embedding these specimens, Dr. Dimenstein recommends the use of a tamper to flatten the core, to keep it parallel to the block face. If two cores are present, they should be embedded parallel to each other, and to the horizontal axis of the block. Filtration specimens should be embedded in the manner similar to embedding any aggregate of tissue, carefully grouping the fragments into the middle of the mold.

In addition to the above method, Sakura Finetek makes a product called Tissue-Tek Paraform Tissue Orientation Gels that can be used in conjunction with their Paraform cassettes. This material can be used to hold small specimens in place and in proper orientation, from grossing through embedding.

An alternative method is to use a product called “HistoGel”. HistoGel is a liquid at 55℃ and has a gelatin consistency at room temperature. The idea is simply to surround the tiny tissue fragment(s) with liquefied HistoGel and allow it to cool to room temperature (approximately two minutes), thereby trapping the tissue fragment in the HistoGel, much the same way fruit is embedded in the gelatin of a Jell-O fruit mold. The resulting “button” of HistoGel containing the tissue is placed into a tissue processing cassette and processed as usual. (Note: do not use microwave processing, or “Rush” processing.) During embedding, the button is simply removed from the cassette and embedded as usual. Additional slides may have to be taken to reach the fragment embedded in the HistoGel; however, the tissue cannot be lost in processing (Figure 1A, 1B). This process far outweighs the extra effort involved during the surgical grossing procedure.

Troubleshooting in the Histology Laboratory

In summary, it is excellent practice to identify all specimens that may be received by your laboratory for “non-routine” histology. Once identified, procedures should be developed for proper receipt and handling of the specimens. This is the only way to ensure the highest patient care quality and to avoid later instances of troubleshooting sub-optimal events.

REFERENCES:

  1. Chapman, C.M. (2017). The Histology Handbook: In Search of the Perfect Microslide. Amazon CreateSpace Independent Publishing Platform
  2. Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform
  3. Dimenstein, I.B. (March 2016). Technical Note. Submitted to J Histotechnol (Vol.39(3): 76-80)
  4. Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory. Submitted to J Histotechnol
#3 Specimen Identification & Optimal Embedding

#3 Specimen Identification & Optimal Embedding

No matter what type of histology laboratory you work in – hospital, research, reference, teaching facility – there will be times when you receive specimens that you do not normally receive. It is important to identify these specimens upon receipt, so that they can be handled correctly to avoid any sub-optimal event that may compromise the specimen and the resulting microscope slide.

Fingernail or toenail specimens are such specimens. Usually, the clinician suspects a fungal infection, however, in some cases, a malignant melanoma may be suspected. In either case, the nail must first be fixed thoroughly in formaldehyde. Upon receipt, it may be prudent to have the surgical grossing team use the “Nail Map’ (devised by Renig, et. Al.) shown in the “Handling of Nail Specimens” procedure contained herein, especially for nail bed specimens. Use of this map, along with proper inking, will help to maintain all three axes of orientation; epidermis-dermis, medial-lateral and proximal-distal.

After fixation, the nail should be held overnight in nail softening solution, which will soften and continue to fix the nail (5% Tween in 10% neutral buffered formalin). Decalcification solutions should not be used, as the hardness of the nail specimen is caused by keratin protein, not calcium. The following day, the specimen may be processed routinely into a paraffin block. It is important to use a “large side” processing protocol, not a “small” or “rush” protocol to ensure proper processing.

During cutting, additional unstained slides should be made for any special or immunohistochemical stains that the pathologist may want with the H&E stain. The use of the nail softening solution will make the microtomy go easier and result in optimal sections.

After cutting, the sections should be picked up on gelatin coated slides (or some other coated slide that works in your laboratory). This insures that the sections will adhere to the slide during H&E, periodic acid Schiff’s (PAS) and immunohistochemical staining. This will also help you avoid troubleshooting a “section fall off” event during staining.

Please notice that the literature describes the use of ammonium hydroxide solutions, “Nair”, and acid solutions for softening nails. Based on experience, these solutions are not recommended. The use of a 5% solution of Tween 85 (in formaldehyde) is recommended. Nail with an attached bone is a surgical pathology specimen and will require decalcification at some point.

The optimal mode of microtomy for nail specimens starts with optimal embedding. Nail specimens should be embedded on edge, at a 45 degree angle to the microtome knife edge. Hand wheel rotation should be slow, to avoid shattering the tissue. If nails continue to be “hard’ during microtomy, once the block face is opened and the nail tissue exposed, the entire block can be soaked for 15 to 30 minutes in the Tween solution described above, or in a simple detergent solution. Detergents act on the keratin proteins to soften them. Once sections are floated out, they should be picked up on gelatin coated slides. Extra slides should be picked up at this time for special stains requests.

Since the majority of nail specimens are taken for demonstration of possible fungi, the Periodic Acid Schiff’s (PAS) stain and the Gomori’s Methenamine silver (GMS) stain will most likely be requested. Nail specimens received for assessment of melanoma will require a Fontana-Masson stain along with other assorted immunohistochemical (IHC) stains (i.e. S-100, Melan-A, MITF). These unstained slides should be prepared at the time of initial microtomy to shorten turnaround times.

As with the previous blogs in this series, the best way to troubleshoot suboptimal events is to recognize exactly where in your laboratory these events could occur, and to address them “up front”. That is the case with nail specimens. In addition, it is important to be proactive in the embedding and microtomy tasks associated with any unique specimens that your laboratory may receive, which will further decrease chances for a suboptimal event to take place.

REFERENCES:

  1. Chapman, C.M. (2017). The Histology Handbook: In Search of the Perfect Microslide. Amazon CreateSpace Independent Publishing Platform
  2. Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform
  3. Reinig et al. (2015) How to submit a nail specimen. Dermatologic Clinics: 33(2)
  4. Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory. Submitted to J Histotechnol
#2 “Non-Routine” Handling

#2 “Non-Routine” Handling

In the previous blog we looked at one way to minimize troubleshooting by being proactive and looking ahead to possible situations and procedures that exist in your laboratory that may cause sub-optimal events. This blog will continue with that same mind set.

Some specimens may be received in formalin into the routine lab; however, they may require “non-routine” handling. Skin specimens submitted for “Slow Mohs” processing are such specimens. These are pieces of skin that are carefully marked with colored inks to indicate exact orientation. Laboratory personnel who receive the specimens, and who perform the surgical grossing techniques must recognize these “up front”. Failure to recognize such specimens may result in changing/destroying the orientation, resulting in the pathologist’s inability to render an accurate diagnosis regarding tumor location and surgical margins.

Similarly, a 2 mm punch specimen of skin may be mishandled by laboratory staff, if not identified “up front” at the time of receipt. These specimens should be identified for the embedding and microtomy personnel. The microtomy personnel must pick up the very first sections and observe the unstained slide under the microscope to ensure correct dermal-epidermal orientation (Figure 1A, B). If the specimen has been mis-embedded, there is enough tissue left to melt down and re-embed correctly. Failure to follow this procedure may result in the production of initial and level slides that are mis-embedded. The result may be that the specimen cannot be diagnosed. This would be a possible troubleshooting event that could not be remedied, with dire consequences.

Troubleshooting in the Histology Laboratory

The procedure of “racking down” the condenser to view unstained sections can be used by the microtomist to determine “full-face” sections, presence of knife marks, tearing of sections and overall section quality prior to staining. These issues are best addressed “up front” when they can be remedied, rather than sending suboptimal slides to the pathologist. Microtomists can now more fully control the quality of the sections that are submitted to the pathologist. This procedure also decreases the chances of a suboptimal event related to tissue section quality.

There are some unique specimens received in the histology laboratory that may cause sub-optimal events for the histologist if not handled properly at the time of surgical grossing. Skin specimens may be taken from the scalp to determine if the patient is suffering from alopecia (hair loss). These are almost always punch specimens and are usually 3-4 mm in diameter. To count hair follicles for comparison, these are the only skin specimens that are not cut perpendicular to the dermal-epidermal junction. Rather, they are cut parallel to it, resulting in cross sections of the hair follicles. This method was developed by Dr. Headington and is referred to as the Headington procedure. Some clinicians may submit two punch specimens: one for the Headington procedure, and the other for routine “vertical” sections.

If a skin punch specimen is sent for the Headington procedure, and is mistakenly prepared for a vertical section, there is a remedy. The two pieces of the split punch in the paraffin block can be melted down, positioned so that the specimen is “back together”, then embedded “on end” with the dermis down. This will provide the pathologist with the proper orientation of the Headington procedure.

Another unique specimen which may be sent to your laboratory for evaluation is a punch specimen of skin for diagnosis of painful sensory neuropathy, which can be caused by any number of diseases that affect the peripheral nerves. Usually, a 3 mm punch biopsy is taken for evaluation of density of intraepidermal nerve fibers (IENF) that are reduced in painful sensory neuropathy. The specimen requires 24 hours fixation in 2% paraformaldehyde (Paraformaldehyde/Lysine/Periodate-PLP) fixative. The procedure requires a special protocol of cryosectioning after embedding in sucrose, but at the grossing level of processing, it is necessary to provide adequate fixation. The principles of grossing are the same in Small Fiber Neuropathy (SFN) tests such as Epidermal Nerve Fiber Density (ENFD) and Sweat Gland Nerve Fiber Density (SGNFD). It is extremely important that these specimens are identified “up front” during accessioning to route them into the proper area of the laboratory for specimen preparation.

REFERENCES:

  1. Chapman, C.M. (2017). The Histology Handbook: In Search of the Perfect Microslide. Amazon CreateSpace Independent Publishing Platform
  2. Chapman, C.M. and Dimenstein, I.B. (2016). Dermatopathology Laboratory Techniques. Amazon CreateSpace Independent Publishing Platform
  3. Chapman, C.M. (2018). Troubleshooting in the Histology Laboratory. Submitted to J Histotechnol