#1 Introduction to Immunohistochemistry

#1 Introduction to Immunohistochemistry

Rankin Biomedical Logo

14515 Mackey Rd,
Holly, MI 48442
248.826.2412
sales@rankinbiomed.com

Anthony van Leeuwenhoek is credited with being the first person to use a microscope to view tiny “animacules” in 1674. At approximately the same time, Robert Hooke used a similar microscope to view thin slices of cork. The structures that he observed resembled the tiny cells that monks lived in at the local monastery, so he named the structures “cells”.

From that time through the 1800’s and early 1900’s, laboratorians experimented with various dyes to impart contrast to sections on microscope slides in order to see these cells. Joseph von Gerlach is credited with developing the first histology stain by using a solution of carmine in 1858 to stain brain cells. Several years later, in 1896, the hematoxylin and eosin (H&E) stain combination was worked out by Paul Mayer. The H&E stain has since remained the cornerstone of pathological diagnosis to this day.

The mid 1900’s saw the rapid development of many special stains. These stains are also used in present day histology laboratories to detect microorganisms, cell products and tissue structures. Similar to the H&E stain, these special stains use colored dye solutions to impart color and contrast to the desired structures. However, many times cells under study appear normal with these stains, even though they are later proved by other methods to display pathology.

In the 1960’s and 1970’s a novel method was developed which helped pathologists to diagnose cancer cells. The method was based on using antibodies to specifically bind to protein antigens within the cells and tissue elements of a tissue section on a microscope slide. This was followed by using a “detection chemistry” to visualize the binding, such that it could be seen using a light microscope. The method is called immunohistochemistry, or “IHC” for short.

Since that time many thousands of antibodies have been developed for use in IHC. Each one will specifically bind with only one protein site. Through experimentation and research, pathologists now have a large menu of different proteins which can be detected and visualized within the specimens on the microscope slides. The presence or absence of these proteins provides information from which to form a diagnosis.

The IHC method relies on the ability to develop and use antibodies that are directed to, and bind only with, very specific proteins. Early literature described the binding as a “lock and key” arrangement. However, it was soon discovered that the binding of an antibody to its protein antigen counterpart is a very complex, three-dimensional event.
Mammals have an immune system that makes antibodies (Ab) to foreign proteins, for example, bacteria and viruses. If you inject mice, rabbits, or other animal subjects with pure protein preparations, their immune system will make antibodies to the protein. These proteins are also referred to as antigens (Ag). The antibodies are highly specific – each will bind to only one antigen.

For example, if you are exposed to the chicken pox virus, you most likely will contract the chicken pox disease. For 2-4 weeks from the exposure time, your immune system makes antibodies to the chicken pox virus. From that point on, any time you are exposed to the chicken pox virus, the antibodies circulating in your blood protect you from the disease by binding with and inactivating the chicken pox virus particles.

In histology, we can use pure antibody preparations to specifically bind to protein antigens in tissue sections. Since antibodies themselves are too small to be seen under a microscope, we can “label” them with colored tags which can be observed under the microscope. The proteins can be localized within cells, outside of cells, or even within cell nuclei.

Future segments of the educational series will delve into the methodology of this fascinating technique.

Figure 1. IHC staining for the proteins of keratin in a microscopic section of skin. Keratin is a protein that is produced by epidermal cells and in this preparation is stained brown. Cell nuclei are visualized by a blue hematoxylin counterstain. Original magnification X 400.

References:

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

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

3. https://www.nature.com/milestones/milelight/full/milelight02.html

4. Von Gerlach, J. (1858). Mikroskopische Studien aus dem Gebiet der menschlichen Morphologie (Enke)

5. Sternberger, L.A., Hardy, P.H. Jr, Cuculis, J.J. & Meyer, H. G (1970). The unlabeled antibody enzyme method of immunohistochemistry: preparation and properties of soluble antigen-antibody complex (horseradish peroxidase-antihorseradish peroxidase) and its use in identification of spirochetes. J. Histochem. Cytochem. 18, 315–333

Article Categories

#7 Tissue Processing Artefacts

#7 Tissue Processing Artefacts

Rankin Biomedical Logo

14515 Mackey Rd,
Holly, MI 48442
248.826.2412
sales@rankinbiomed.com

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

Article Categories

#6 Understanding Tissue Processor Procedures

#6 Understanding Tissue Processor Procedures

Rankin Biomedical Logo

14515 Mackey Rd,
Holly, MI 48442
248.826.2412
sales@rankinbiomed.com

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

Article Categories

#5 Troubleshooting in the Histology Laboratory

#5 Troubleshooting in the Histology Laboratory

Rankin Biomedical Logo

14515 Mackey Rd,
Holly, MI 48442
248.826.2412
sales@rankinbiomed.com

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

Article Categories

#4 Alternative Methods for Preparing and Embedding Specimens

#4 Alternative Methods for Preparing and Embedding Specimens

Rankin Biomedical Logo

14515 Mackey Rd,
Holly, MI 48442
248.826.2412
sales@rankinbiomed.com

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

Article Categories