The H&E Stain: Far From Routine Part 2

This blog is a followup to the previous article “The H&E Stain: Far From Routine”.  In that article, the basics of the routine hematoxylin and eosin (H&E) stain were discussed.  Now we shall discuss how to trouble shoot the routine H&E and how to ensure a high quality stain, once you have worked with your pathologist to determine the best stain result for your laboratory.

 

A standard H&E staining protocol is provided below.  It applies to either a manual or automated staining procedure.  While it may not be ideal for your laboratory, it can be used as a starting point.  The final color rendition of your H&E stain should be determined by working with your pathologists.  This will consequently make their job easier.  Each day, once the H&E stain set up is completed, you should run down one test slide to confirm that the staining is optimal.  This also will help you to document quality control procedures.

 

Hematoxylin and Eosin (H&E) Stain

 

PRINCIPLE: Hematoxylin stains nuclear material a dark blue, while the eosin stains cytoplasm and connective tissue varying shades of pink.

 

SOLUTIONS:

  1. Harris’ hematoxylin
  2. Alcoholic Eosin
  3. Clarifier (i.e. acid alcohol)
  4. Bluing reagent (i.e. ammonium hydroxide)

 

PROCEDURE:

  1. Cut paraffin sections 4-5 microns.  Bake at 60 C for 45 minutes; allow to cool.
SOLUTION TIME NOTES
xylene 2 x 5 min Removes paraffin; this is a minimum time
100% alcohol 2 x 2 min Removes xylene
95% alcohol 1 x 2 min Begins hydration
Running water 2 min Hydrates the sections
Harris’ Hematoxylin 5 min Stains nuclei and tissue
Running water 2 min Removes excess hematoxylin
Clarifier 1 min Removes hematoxylin from tissue
Running water 2 min Stops clarifier action
Bluing reagent 1 min Changes hematoxylin from red to blue with positive ions
Running water 2 min Stops bluing action
95% alcohol 1 min Readies for eosin
Eosin 1 min Stains cytoplasm pink
95% alcohol 15 seconds Differentiates eosin into 3 shades
100% alcohol 3 x 2 min Dehydrates
Xylene 2 x 5 min Readies for coverslip

Each step in the H&E stain procedure is essential, and if not managed correctly, any of them can cause suboptimal slides.  We shall begin discussion here and continue through the next blog, identifying potential problems and solutions along the way.

 

  1. Deparaffinization and dehydration. Many H&E and special stain protocols begin with step #1 as simply “Deparaffinize and hydrate to water”. In fact, nothing could be more complicated.  The sections to be stained are surrounded by a matrix of paraffin wax, which is insoluble in water.  All of the paraffin must be removed to ensure optimal staining.  Two changes of xylene (or xylene substitute) for 5 minutes each may accomplish this in a low volume laboratory; however a high volume laboratory may need to use three changes of xylene / xylene substitute to completely remove all paraffin.  In automated H&E stainers, it is important to adjust the height of xylene (and all reagent) containers to ensure that the sections next to the label end of the slide are being deparaffinized sufficiently.

 

  1. Harris’ hematoxylin is recommended for histology. Gill’s hematoxylin is generally used for cytology, and as a counterstain in many special stains. Harris’ hematoxylin continues to oxidize as it sits in the original container, and in the stain containers; hence, it should be filtered through Whatman #1 filter paper prior to use for the days’ staining runs.  This will prevent precipitate from adhering to the microscope slides.

 

  1. Clarifier. Harris’ hematoxylin is a regressive stain.  A clarifier solution must be used to remove excess hematoxylin staining from both the nuclei and other tissue elements.  The original formulation  is 1% hydrochloric acid in 70% ethanol, for “a few quick dips”.  However, with the current use of automated stainers, “a few quick dips” needs to be quantified into seconds or minutes. This usually requires a dilution of the original clarifier solution, with some experiments to determine the exact time.  The recommended starting point for clarifier on an automated stainer is to use the following for 30 seconds:

 

100% ethanol                            700 ml

tap water                                  2600 ml

hydrochloric acid (concentrated)  2.0 ml

 

Once made, the resulting intensity of the nuclear stain can be adjusted by changing the time of the clarifier incubation.  Alternatively, one may adjust the amount of acid added to the alcohol (i.e. decrease from 2.0 ml to 1.5 ml).  These adjustments will allow you to obtain a final stain with the required nuclear staining intensity.

 

  1. Bluing reagent. The recommended starting point for bluing reagent is:

 

Tap water                                                    3500 ml

Ammonium hydroxide (concentrated)               1.5 ml

 

As with the clarifier solution, results can be obtained by adjusting the time of incubation and/or concentration of the solution.  Alternatively, one can use lithium carbonate as a bluing reagent, or simply use a running water step to “blue” the nuclei.

 

  1. Eosin.  Working eosin solution is the most stable reagent of the H&E stain.  Hundreds of slides can be stained with the same batch of eosin on board an automated stainer.  The incubation time is usually kept short, as eosin penetrates and stains rapidly and reproducibly (i.e. 1-2 minutes).  The variable step in eosin staining is the subsequent 95% alcohol incubation, which produces the final “three shades of pink” within the tissue sections.  The number of 95% stations (i.e. one or two), and the incubation times (i.e. 30 seconds to 2 minutes) will determine the final quality of eosin staining.  Finally, it is important that the final 100% alcohol stations after the eosin staining remain uncontaminated with water.  Any water left in the sections after the coverslip is applied may cause the eosin to “bleed out” of the section.

 

In summary, the information contained herein will hopefully help you to manage and maintain consistent high quality of the H&E slides produced in your laboratory.

 

REFERENCES:

  1. Theory and Practice of Histological Techniques. JD Bancroft, A Stevens ed. Churchill Livingstone, NY.  Fourth edition. 1996
  1. Theory and Practice of Histotechnology.  DC Sheehan, BB Hrapchak.  CV Mosby Company, St. Louis. First edition. 1980.
  2. Luna L.  AFIP. Manual of Histologic Staining Methods.  Third Edition. McGraw-Hill.

p39. 1968.  As modified by CM Chapman

  1. Dermatopathology Laboratory Techniques.  CM Chapman, I Dimenstein.  In press.
  2. The H&E Stain: Far from Routine.  CM Chapman.  Advance for Laboratory Professionals.  April 8, 2002.  Vol. 14 No.8

The H&E Stain: Far From Routine Part 1

[Editor’s note:  Segments of following blog are taken directly from an original article “The H&E Stain: Far from Routine” published by the author in Advance for Medical Professionals in April 2002.]

 

What exactly is a routine “H&E”?  And what makes it routine?  The first question is easy.  “H” stands for ”hematoxylin” and “E” stands for “eosin”.  Both are dyes used to stain tissue sections in histology.  However, the procedure for correctly applying this combination of stains to tissue sections is far from routine.

Since both dyes are water soluble, the first step is to completely remove the paraffin that is present in the tissue sections from the microscope slides.  Soaking in xylene, or a xylene substitute, followed by 100% and 95% alcohol, allow the slides to then be immersed in running water, which hydrates the sections.

The next step is to place the slides in a solution of hematoxylin, a natural dye obtained from the logwood tree Haematoxylon campechianum that will stain the nuclei of cells blue/black.  The staining is enhanced by the addition of an aluminum, iron or lead salt, which acts as a mordant for the hematoxyin to bind to the tissue sites.  This type of hematoxylin is referred to as “Harris” type, and is a “regressive’ stain.  That is: the tissues are overstained with the hematoxylin, which stains all tissue elements, and then differentiated with an acid alcohol solution (i.e. “clarifier”) to remove excess hematoxylin, leaving only the nuclei of the cells stained.

The composition of the clarifier varies.  The original procedure written by Lee Luna in the Armed Forces Institute of Pathology (AFIP) Manual specifies differentiation of the Harris hematoxylin solution with a “1% acid alcohol” solution for a time duration of “a few quick dips”.  This solution is 1% hydrochloric acid in 70% ethanol.  However, with the current use of automated stainers, “a few quick dips” needs to be quantified into seconds or minutes. This usually requires a dilution of the original clarifier solution, with some experiments to determine the exact time.

Harris’ hematoxylin can be compared to “Gill’s” or “Mayer’s” hematoxylin, which are used “progressively”.  The longer the sections are left in these solutions, the darker the nuclei stain.

In either case, the resulting hematein-mordant stains nuclei a reddish color.  Subsequent treatment in a weak basic aqueous solution, using either lithium carbonate or ammonium hydroxide, changes the dye molecules from red to blue; hence the term “bluing reagent”.  The resulting blue color is dependent upon the freshness, type and age of the mordant.  It is important that the pH of the bluing reagent is not too high, as section loss may occur during staining.  Additionally, warm running tap water may be used as the “bluing reagent”, as it contains positive ions.  The final blue color will be decided upon by the pathologist and histologist, who should agree what is optimal for the laboratory.

Eosin, the second dye in the H&E, is used to demonstrate the general histology of the tissue architecture.  When used correctly, eosin should stain both cytoplasmic and tissue elements in three shades of pink.  There are several types of eosin dye available, however Eosin Y is widely used for routine staining, as it is soluble in both water and alcohol.  Usually a 0.5 % to 1.0 % solution of Eosin Y is made up in 80% alcohol, with a small amount of acetic acid added.  Additionally, phloxine may be added to provide a more intense reddish color.

After staining in eosin for one to three minutes, the final shades of pink are determined by the differentiation steps following this staining.  The differentiation can be carried out in running water, 95% or 100% alcohol.  In any case the slides must be dehydrated completely after differentiation in several changes of 100% alcohol and xylene prior to coverslipping.  Failure to do so may cause “bleeding” of the eosin dye in the final microscope slide and/or variable eosin staining.

Complete removal of paraffin is an essential first step in any H&E staining procedure.  If xylene substitutes are being used, it may be necessary to adjust and lengthen times to guarantee paraffin removal.  In addition, xylene substitutes do not tolerate traces of water, as xylene does.  Thus, it is important also in the dehydration steps prior to coverslipping that the slides move through fresh changes of alcohol and xylene substitute.  If any water remains in the sections during coverslipping, the final slides will appear cloudy.  Additionally, the eosin stain may bleed out of the sections, as mentioned above.

This article has provided the basics of the H&E “routine” stain.  Please keep an eye open for the next blog, which will provide additional information regarding how to maintain the quality of your H&E stain, once you have optimized it.

 

REFERENCES:

  1. Theory and Practice of Histological Techniques.  JD Bancroft, A Stevens ed.  Churchill   Livingstone, NY.  Fourth edition. 1996.
  2. Theory and Practice of Histotechnology.  DC Sheehan, BB Hrapchak.  CV Mosby Company, St.     Louis. First edition. 1980.
  3. Luna L.  AFIP. Manual of Histologic Staining Methods.  Third Edition. McGraw-Hill.

p39. 1968.  As modified by CM Chapman.

  1. Dermatopathology Laboratory Techniques.  CM Chapman, I Dimenstein.  In press.
  2. The H&E Stain: Far from Routine.  CM Chapman.  Advance for Laboratory Professionals.  April 8, 2002. Vol. 14 No.8.

Silver Stains

In the histology world, the mere mention of a “silver stain” may be the cause of panic and uncertainty with regard to the performance of the stain, and the quality of the final resulting microscope slide.  All other special stains, with few exceptions, are relatively easy and straightforward to perform; not so with silver stains.

Silver stains can be categorized into (a) stains to visualize substances, such as calcium, melanin and reticulin and (b) stains for microorganisms, such as fungi and spirochetes.  The goal of all silver stains is the same: to get metallic silver to precipitate out at the staining site, and then replace it with gold to provide the final, stabilized, black reaction product.

The specific procedures for these stains are outside of the scope of this blog.  However, the following stain procedure outline explains the steps used.

Oxidation enhances subsequent staining by the silver solution.  Oxidizers include phosphomolybdic acid, potassium permanganate and periodic acid.

Sensitization usually employs a metal salt to help bind silver from the silver solution.  The original sensitizer for Wilder’s silver technique and the Steiner and Steiner stain is 1% uranyl nitrate.  However, Margeson and Chapman pioneered the substitution of zinc formalin for the original radioactive uranyl nitrate solution, which is also a strong oxidizer.

Silver impregnation solution contains metallic silver in a solution.  The idea is to have the silver carrying solution composed such that the silver ions will move from the solution, bind to the tissue section and then precipitate out in metallic form.

The reduction step in the reaction involves providing electrons, in the form of substances such as hydroquinone and formaldehyde, to chemically make the silver ions precipitate out into visible metallic silver.  This allows the structures in question to be visualized in dark black staining.

Toning  of the sections in gold chloride is a chemical reaction whereby the metallic silver is replaced by metallic gold, which is very stable and maintains the black color product.

The use of sodium thiosulfate, or “hypo”, helps to remove any unbound silver remaining from the toning reaction.  This is followed by counterstaining, usually either with nuclear fast red or fast green, for a proper final color rendition.

Thus, with all of these simple, straight forward steps, what could possibly go wrong?  Let’s begin at the beginning and work it through.

Use acid clean glassware for all containers and Coplin jars.  Why?  This will prevent the presence of any unwanted ions on the glass surfaces (including the slides themselves) causing non- specific precipitation of the silver.  You will see this as a black or mirror precipitate on the inside of the Coplin jar, or on the surface of the microscope slide.  If this does happen, the unwanted silver can be removed (see procedure at the end of this article).

Use plastic forceps to handle all slides — not metal.  Metal present in metal forceps may cause the silver to precipitate out.  Back in the old days, before plastic forceps, we simply dipped the tines of metal forceps into paraffin, allowed it to cool, thereby preventing the metal from contacting any of the solutions.

Pay attention to the slides when they are in the silver solution – especially if the solution is a hot, heated solution.  The point at which the silver solution itself may precipitate out is a fine one.  The slides and staining solution should be monitored closely during these incubations.  Also, if you are monitoring the slide under a microscope, make sure to rinse the slide in distilled water before viewing, thereby preventing the foundation of black silver precipitate forming on the microscope stage or slide. Your coworkers especially will appreciate this.

 

References

  1. Sheehan and Hrapchak. Theory and Practice of Histotechnology. Second ed.  CV Mosby Co.  St. Louis. pp 181-183.  1980.
  2. Chapman CM. Dermatopathology: A Guide for the Histologist. Copyright 2003.
  3. Chapman CM and Dimenstein IB. Dermatopathology Laboratory Techniques. Copyright 2015.  In press.
  4. Margeson LM, Chapman CM: The use of zinc formalin as a sensitizer in silver stains for spirochetes. J Histotech, 19:135-138, 1996.
Stains for Microorganisms

Stains for Microorganisms

The staining of microorganisms in histology can be challenging. Filamentous fungi and associated conidia are more easily demonstrated as they are visible under light microscopy when stained with periodic acid Schiff’s (PAS) as in Figure 1. The diameter of fungi filaments is 5-10 microns, which is approximately the same as the diameter of a red blood cell, while their length may be hundreds of microns (Figure 2). Microorganisms are extremely small and are at the limit of resolution of the light microscope. Viruses are even smaller (Figure 3).

Bacteria exist in three different shapes. Cocci are round, bacilli appear as rods and spirochetes are corkscrew shaped. The first level of determination for bacteria is whether they are Gram positive or Gram negative. The original method of Gram’s technique published in 1884 is still used today. Tissue sections are first stained with a crystal violet solution, which stains all of the tissue section. This is followed by Gram’s iodine which locks the crystal violet stain into the cell wall of the bacteria. Subsequent decolorization with acetone removes the stain from everything – except the Gram “positive” bacteria, which remain a dark blue/purple color (Figure 4).

The second phase of the stain involves the application of fuchsin / neutral red solution which stains Gram “negative” bacteria pink/red (Figure 5). This can be the most variable step in the procedure and relies on the correct application and retention of the basic fuchsin within Gram negative bacteria.

There are two additional steps which may result in variable staining as well. The first is an over-decolorization with acetone, which may result in faint staining of Gram positive bacteria. The second is the use and timing of the final counterstain, which may be either picric acid/acetone or tartrazine. While picric acid/acetone results in a more reliable result, the picric acid is a dangerous chemical and requires specific disposal procedures. Tartrazine is safer, but can easily be leached out during the dehydration process prior to coverslipping. The histotechnician performing this stain must rely on experience and precise timing to produce consistent results for this stain.

Actinomycetes are a Gram positive, non acid fast, branching filamentous organism. However, care must be taken when performing the Gram positive stain sequence. It may be necessary to (a) increase the time in crystal violet solution and (b) decrease the time in the acetone decolorization step in order to demonstrate the organism.

A second determination of bacteria is based on an organisms “acid fastness”. The Kinyoun method for acid fast bacteria, the Fite method for leprosy bacilli and the Ziehl-Neelsen stain for tubercule bacilli are all based on the same stain theory. Tissue sections are first stained with a carbol fuchsin solution, which stains all tissue elements pink/red. Subsequent decolorization in an acid-alcohol solution removes stain from all the tissue elements, except acid fast bacteria. Methylene blue is usually used as a counterstain (Figure 7).

Spirochete bacteria are ubiquitous in nature. They are found everywhere in soil and water. The issue is that some are pathogenic to humans. TheTreponema pallidum spirochete is sexually transmitted and causes syphilis. The Borrelia burgdorferi spirochete is present in certain tick species, and may be transmitted to humans when the tick attaches to the skin to feed. The result is Lyme disease, which if left untreated, can be debilitating to the patient.

Silver stains were developed in the early 1900’s to stain spirochetes. The Dieterle (1927), Warthin-Starry (1920) and Steiner (1944) are still used in histology today to demonstrate spirochetes. Some of the methods have been modified to make them safer to use (Margeson and Chapman, 1996). The stain theory is the same in all methods: pre-treat the sections to make the spirochetes more readily able to pick up and bind the silver solution. The final result is to stain the spirochetes black, against a gray/ brown background (Figure 7). Those who have performed any of these stains are aware of the many staining procedure pitfalls, which can render the spirochetes either over or under stained. As a result, many laboratories currently use immunohistochemistry to stain spirochetes. While both methods can stain spirochetes, neither is able to demonstrate the exact species to which the stained organism belongs.

Both positive and negative control slides should be used when performing stains for microorganisms. The negative control slide ensures that there is no bacterial contamination present in source water or stain solutions.

10271102721027310274102751027610277

 

References

  1. Theory and Practice of Histological Techniques. Chapter 10.       JD Bancroft, A Stevens ed.       Churchill Livingstone, NY.       Fourth edition. 1996
  2. Theory and Practice of Histotechnology. Chapter 9.   DC Sheehan, BB Hrapchak. CV Mosby Company, St. Louis. First edition. 1980.
  3. Margeson LM, Chapman CM: The use of zinc formalin as a sensitizer in silver stains for spirochetes. J Histotech, 19:135-138, 1996.
Staining Fungi

Staining Fungi

Fungi include molds, yeasts and higher fungi. All fungi are eukaryotic and have sterols but not peptidoglycan in their cell membrane. Their cell walls are composed of cellulose; the same building blocks that plants use. Fungi may produce large, reproductive mycelium, called mushrooms, which may be edible, or poisonous. Other naturally occurring fungi may infect humans, one example being “athlete’s foot”.

Fungi are chemoheterotrophs which require organic nutrition and most are aerobic. Many fungi are also saprophytes which live off of dead organic matter, in soil and water and acquire their food by absorption. Fungi may produce sexual and asexual spores. There are over 100,000 species recognized, with over 100 of them known to be infectious agents in humans. Molds are composed of numerous, microscopic, branching hyphae known collectively as a mycelium.

Hyphae growth occurs from the apical tip, and apical vesicles contain materials and enzymes for the formation of new hyphal wall. Older hyphae are less biochemically active and contain many storage vacuoles. In most molds these hyphae have septa, which are cross-walled divisions, but in some there are none and the hyphae are aseptate. A septum is a cross-wall formation which divides one fungal hypha into two cells. These septa may add strength to the hyphae or serve to isolate adjacent parts to allow differentiation, such as during production of the reproductive structures.

Spores are formed from the reproductive mycelium. Asexual spores are produced by the aerial mycelium of a single organism, whereas sexual spores are formed by the fusion of cells and nuclei from opposite mating strains (Figures 1 and 2).

Fungal nail infections are common infections of the fingernails or toenails that can cause the nail to become discolored, thick, and more likely to crack and break. The technical name for a fungal nail infection is “onychomycosis.”

Fungal nail infections can be caused by many different types of fungi (yeasts or molds) that live in the environment. Small cracks in your nail or the surrounding skin can allow these germs to enter your nail and cause an infection. Onychomycosis can be diagnosed by microscopic examination or fungal culture of the nail clipping.

Your histology laboratory may receive skin and nail specimens to be evaluated for presence of fungi. The most common special stains used to visualize fungi are the periodic acid Schiff’s stain (PAS) and the Grocott’s methenamine stain for fungi (GMS).

Both stains are based on the chemistry of the fungal cell wall, which is made of cellulose. Cellulose is composed of glucose molecules, attached together very tightly. The PAS stain uses a primary step of oxidation of the glucose with periodic acid to form aldehyde groups. Once formed, the aldehydes are available to subsequently bind with the Schiff’s reagent, which results in the fungal cell walls being stained pink (Figure 3). Diastase digestion may, or may not be used, as it does not affect the fungal staining; it simply removes any glycogen which may be present.

Some dermatopathologists feel that the GMS stain is a more sensitive stain for detection of fungi in tissue sections. In this stain, chromic acid is used to oxidize the glucose molecules, leaving the aldehyde groups open to bind silver molecules, present in the methenamine silver solution. Subsequent development and toning of the sections renders the fungal cell walls black (Figure 4).

Either the PAS or the GMS method may be utilized to stain fungi in tissue sections.

Previous blogs (Hair Histology) described how to stain fungi that are present on hair shafts.

fungi1fungi3fungi4fungi2

 

References

  1. www.cdc.gov
  2. Scher RK, Rich P, Pariser D, Elewski B. The epidemiology, etiology, and pathophysiology of onychomycosis. Semin Cutan Med Surg. 2013 Jun;32 (2 Suppl 1):S2-4.
  3. http://www.microbiologybytes.com/iandi/6a.html
Special Stains for Mucins

Special Stains for Mucins

When staining sections for the presence of carbohydrates, the two main classes under investigation are glycogen and mucins.  Mucins include substances referred to as mucopolysaccharides, mucosubstances, glycoproteins and glycoconjugates.

Mucins provide an environment that is conducive to molecular diffusion of chemicals, especially those in ionic form.  Mucin also increases the binding between cells, and may help to “shed’ bacteria and viruses.  The demonstration of the presence and absence of mucins within tissue sections is an important tool to assist pathologists in rendering a diagnosis.  The following types of mucins can be distinguished by certain special stains.

Acidic mucins contain sulphur in varying amounts.  Other acidic mucins can have carboxyl groups attached to them, which make them acidic.

Neutral mucins, as the name implies, contain no acidic reactive groups within.  These mucins are epithelial in origin and are commonly found in gastric lining cells.

Alcian blue is the most common stain used for demonstrating the presence of acidic mucins, since the blue dye molecules are cationic (positively charged) and bind to the anionic sulphur and carboxyl groups within the mucin.  Neutral mucins are not usually stained by alcian blue.  The pH of the alcian blue solution can be adjusted to stain different classes of acidic mucins (Figure 1A and 1B).  Alcian blue can also be combined with a PAS stain to show both acidic and neutral mucins (Figure 2).  Mucicarmine is sometimes used to demonstrate acidic mucins (Figure 3).  Mucicarmine also will stain encapsulated fungi such as Cyrptococcus neoformans.

 

007008 001 003
004005 006  

Glycogen is the cellular mode of storing sugar, which is in the form of a six-sided molecule. Glycogen can be demonstrated histologically using the periodic acid – Schiff’s stain (PAS). The glycogen molecules are oxidized by the periodic acid, which breaks the ring and exposes aldehyde groups. Subsequent exposure to Schiff’s reagent causes the aldehyde groups to react with it and form a pink reaction product (Figure 4A).

Presence of glycogen is confirmed by staining a serial section of the tissue with periodic acid – Schiff’s plus diastase (PAS+D). The section is incubated with diastase solution prior to performing the PAS stain – in order to digest all of the glycogen. The end result is the absence of pink stain (Figure 4B).

It is important to remember that the Schiff’s stain is not specific for glycogen. Schiff’s reagent will combine with any aldehyde groups to form pink reaction product. This is evident in using the PAS stain to identify the basement membrane in skin specimens (Figure 5A). Additionally, the PAS stain can be used to stain fungi (Figure 5B). The sugars present in the basement membrane and the fungal cell wall are oxidized and react with the Schiff’s reagent to form the pink reaction product. Digestion with diastase does not remove the staining.

References

  1. Theory and Practice of Histological Techniques. Chapter 10.  JD Bancroft, A Stevens ed.  Churchill Livingstone, NY.  Fourth edition. 1996
  2. Theory and Practice of Histotechnology.  Chapter 9.   DC Sheehan, BB Hrapchak.  CV Mosby Company, St. Louis. First edition. 1980.