College of American Pathologists
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  Histologic preparations: first program, now book


CAP Today




June 2009
Feature Story

The CAP released last month its new how-to guide to good histologic slide preparation, edited by Richard W. Brown, MD, of Memorial Hermann Healthcare System, Houston, and immediate past chair of the CAP/NSH Histotechnology Committee.

The National Society for Histotechnology/CAP HistoQIP program got its start in 2003 and was the first in the nation in which labs were able to obtain an objective evaluation of the quality of their histologic sections, special stains, and immunohistochemical preparations. The aim at that time of the CAP/NSH Histotechnology Committee—the group behind the program—was to give feedback to labs about the problems seen in their slides. Now, the CAP has a new book, developed in conjunction with the NSH, that’s an extension of the program; it’s titled Histologic Preparations: Common Problems and Their Solutions. Dr. Brown says in the preface that the book is an expanded version of the program’s educational critiques.

Of the book’s similarly organized chapters, he writes: “After a brief presentation of the technique or stain, including the underlying biochemistry where applicable, there is a list of common problems encountered by laboratories using that stain or technique and a number of possible solutions. The numerous illustrations ... include examples of excellent-quality preparations and of the numerous problems that can occur.”

The book consists of 16 chapters on the following: fixation and processing, microtomy, frozen sections, H&E, Gram stain, mycobacteria, H. pylori, spirochetes, fungi, trichrome stains, reticulin, elastin stains, basement membrane, mucin stains, amyloid, and immunohistochemistry. It’s for histotechnologists and histotechnicians, histotechnology students, pathology residents, and pathologists serving as medical directors of histology labs. To order ($40 for members, $50 for nonmembers), call 800-323-4040 option 1.

Here, as a preview of what’s between the covers, is the chapter on amyloid by Vinnie Della Spera­nza, MS, HTL(ASCP)HT, manager for anatomic pathol­ogy services, Medical University of South Carolina, Charleston

Vinnie Della Speranza

The term amyloid, meaning starch-like, is a misnomer coined by Virchow when he observed that amyloid deposits would stain blue with the iodine reaction, suggesting the presence of starch or cellulose. Despite a striking morphologic uniformity in virtually all cases, amyloid is now known to encompass a spectrum of secondary protein structure diseases.1 The only common denominator is the tendency of these proteins, under specific circumstances, to form β-pleated sheets of antiparallel fibrils, regard­less of the source protein’s original primary structure or function.2

Amyloid deposits occur insidiously and may be found in many organs, often in the walls of small blood vessels. Definitive diagnosis requires the morphologic identification of amyloid deposits in biopsies of the affected organs, which may be surprisingly difficult. Amyloid deposits have a characteristic amorphous hyaline or glassy appearance, but otherwise they stain no differently from other proteins in a routine hematoxylin and eosin stain.

The progressive accumulation of extracellular protein fibrils can lead to cellular atrophy, ischemia, necrosis, and, ultimately, organ failure due to the detrimental effect of these accumulated fibrils on blood supply and normal cellular function. Therefore, correct recognition of amyloid deposits is of considerable importance in diagnostic surgical pathology.3 The clinician will have many questions for the pathologist: Is the amyloid systemic or localized? Is the amyloidosis primary, secondary, or inherited? The answers to these questions are important because patients with localized disease do not require systemic therapy, and their long-term prognosis is excellent. The prognosis and treatment modalities for the three forms of systemic amyloidosis differ significantly, making accurate characterization essential.4

All amyloid deposits share the following physical properties:

  • Appearance of amorphous, eosinophilic deposits under light microscopy after hematoxylin and eosin staining.
  • Bright green birefringence under polarized light after staining with the cotton-wool dye Congo red.
  • Regular fibrillary structure observed with electron microscopy.
  • β-pleated-sheet structure demonstrated with x-ray diffraction.5

By definition, any protein deposits staining with Congo red and exhibiting green birefringence when viewed with polarized light are amyloid.4 Although the identification of amyloid deposits in tissues without characterization of the source protein is not especially helpful to the clinician, the use of tinctorial stains remains an important screening tool for the initial diagnosis of amyloidosis.

Other cotton dyes have been explored for amyloid staining with varying success, most notably Sirius red. This dye has failed to gain popularity, however, because it does not exhibit the polarizing characteristics considered the hallmark of amyloid staining with Congo red. The presence of carbohydrate moieties in amyloid fibrils has encouraged some investigators to use carbohydrate stains, such as periodic acid–Schiff or Alcian blue, to demonstrate amyloid deposits; however, dye uptake is variable and generally poor with these methods. Likewise, metachromatic stains, such as crystal violet or methyl violet, attempt to capitalize on the carbohydrate content of amyloid fibrils, but their staining is not very specific, and the low sensitivity of these methods has caused them to fall in disfavor.

Staining with fluorochrome thioflavin T is an alternative method to improve the sensitivity of amyloid detection; however, the stained sections are not permanent, and tissue com­ponents other than amyloid, including fibrinoid, keratin, intestinal muciphages, Paneth cells, zymogen granules, and juxtaglomerular apparatus, all stain with thioflavin T.6 Furthermore, the appropriate excitation and barrier filters required for fluorescence microscopy may not be available in most laboratories. Therefore, Congo red, specifically the alkaline Congo red method of Puchtler, et al.,7 remains the gold standard for the demonstration of amyloid in tissue sections.

With the Congo red stain, false-positive or false-negative results are usually related to (1) the staining technique, (2) the microscope equipment, or (3) the presence of small amounts of amyloid.2 Congo red in aqueous solution will bind nonspecifically to many tissue structures, including collagen and elastin. Excessive differentiation may lead to decolorization of the amyloid, while collagen remains stained. The Benhold Congo red technique does not yield reproducible results and should be avoided.2

The use of alcoholic solutions, high salt content, and high pH, as in the Puchtler Congo red method, greatly increase staining specificity for amyloid. A saturated salt solution in alcohol at alkaline pH is used in both the dye solution and as a pretreatment of the tissue sections just before staining. High salt content and alkaline pH are believed to depress dye ionization and electrostatic binding to nonamyloid structures. Saturation of the salt and dye solutions is very important, and the instructions must be followed exactly.3

The Congo red stain can be applied to tissues fixed in a variety of fixative solutions, including Bouin, Helly, Zenker, ethanol, and formalin; this technique is also effective for frozen sections and cytology smears. This versatility makes the Congo red stain suitable for the examination of tissues in many settings, including the examination of archival tissues in paraffin blocks. Tissues that have been stored for prolonged periods in formalin will have diminished staining; therefore, fixation for inordinately long periods should be avoided.2,6 One report indicates that Carnoy fixative confers a greater risk of false-positive staining8 and should be avoided.

Virtually any tissue containing known deposits of amyloid may be used as a suitable control. However, as previously stated, fixation for prolonged periods may diminish staining; therefore, tissues identified as positive for amyloid should be processed immediately into paraffin blocks. Large deposits of amyloid (presumably older deposits) frequently show less intense staining and may not exhibit green birefringence with polarized light.9 Smaller deposits, thought to represent more recent amyloid deposits, can often be found in blood vessel walls and tend to make better controls.

One author has reported that sections that have been precut and stored will lose staining reactivity over time. Therefore, it is recommended that control sections be relatively freshly cut. Sections of 6- to 8-µm thickness are most desirable, because this thickness increases the likelihood of discovering small amyloid deposits. Thinner sections may fail to exhibit birefringence.3

What should be seen in a good Congo red stain
In a section that has been well stained with Congo red, the staining should be limited to amyloid deposits present in the tissue section. In general, amyloid will appear a muted red or pink color, the intensity of which is largely dependent on the size, density, and age of the amyloid deposit (Fig. 15.1).

The Congo red-stained section will also show green birefringence with polarizing microscopy; this finding is mandatory for diagnosing amyloid (Fig. 15.2). It should be noted that other tissue elements, including collagen, smooth muscle, and striated muscle, will also display birefringence of varying colors that may be mistaken for amyloid.

Evaluation of Congo red-stained sections with a fluorescence microscope also may be used to identify minute amyloid deposits. Using a fluorescein isothiocyanate filter, amyloid deposits appear yellow-orange, increasing the sensitivity of the technique. However, Congo red fluorescence is not specific, and the presence of characteristic green birefringence with polarizing microscopy should be confirmed in deposits identified with fluorescence microscopy.10

Problems encountered with the Congo red stain

Problem: Weakly stained tissues
Appearance: Faint uptake of Congo red dye throughout the section (Figs. 15.3 and 15.4).

  • Tissue was fixed for a prolonged period in a formaldehyde-containing fixative.
  • Cut sections were stored for a prolonged period.
  • Solutions of Congo red are not stable in the presence of salt and alkali.
  • Avoid storing tissues in formaldehyde-based fixatives for prolonged periods.
  • Cut the sections just before staining, or cut only as many control sections as can be used in a short period.
  • Seal the cut paraffin blocks to help preserve control tissue reactivity.
  • Prepare fresh working solutions just before use.11

Problem: Nonspecific staining
Appearance: Structures other than amyloid bind the Congo red; high background staining may make it difficult to distinguish true amyloid deposits if present in the tissue (Figs. 15.5 and 15.6).

  • Collagen, elastic fibers, and keratin may stain nonspecifically with some fixatives and some procedures.
  • Aqueous Congo red solutions have a tendency to stain structures nonspecifically; this is especially true of the original Benhold formulation.
  • The pH is not sufficiently alkaline.
  • Use polarizing microscopy to help distinguish connective tissue components (grey or silver) from amyloid deposits (green).
  • Rotating the slide on the stage during polarizing microscopy can help distinguish true amyloid deposits from connective tissue. When the slide is rotated, connective tissue fibers will lose the dichroism, while amyloid will not.2
  • Use the Puchtler Congo red method, because the high salt content of the prestain rinse and staining solutions tends to diminish nonspecific staining.
  • After staining, increase time in the alcoholic potassium hydroxide rinse to improve differentiation with the Highman technique.
  • Avoid the use of Canada balsam mounting medium because it will fluoresce.11

Problem: Incorrect color of birefringence
Appearance: Structures may exhibit yellow, red, or white dichroism (Figs. 15.7, 15.8, and 15.9).

  • Section thickness may be incorrect; this artifact is especially likely in thin sections.3
  • Ensure that sections are cut at 6 to 8 mm.
It should be noted that old (large) amyloid deposits will often display diminished birefringence. Smaller deposits in blood vessel walls, for example, may be more likely to demonstrate the characteristic apple-green color.

Problem: Precipitate on tissue
Appearance: A red precipitate is randomly present throughout the tissue (Fig. 15.10).

  • Salt solutions were prepared with isopropyl alcohol.
  • Ensure that the salt solutions are prepared with ethyl alcohol because the salt does not dissolve well in isopropyl alcohol and will deposit crystals on the section.


  1. Kaplan B, Martin BM, Livneh A, Pras M, Gallo GR. Biochemical subtyping of amyloid in formalin-fixed tissue samples confirms and supplements immunohistologic data. Am J Clin Pathol. 2004;121:794–800.
  2. Rocken C, Sletten K. Amyloid in surgical pathology. Virchows Arch. 2003;443:3–16.
  3. Carson FL. Histotechnology: A Self-Instructional Text. 2nd ed. Chicago: American Society of Clinical Pathologists; 1997:125–129.
  4. Gertz MA. The classification and typing of amyloid deposits. Am J Clin Pathol. 2004;121:787–789.
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  9. Geisinger KR, Stanley MW, Raab SS, Silverman JF, Abati A. Modern Cytopathology. London: Churchill Livingstone; 2004.
  10. Fail M, Self S. A novel approach for the demonstration of amyloid in thin (2 micron) sections of kidney. HistoLogic. 2000;32:1–3.
  11. Horobin RW, Bancroft JD. Troubleshooting Histology Stains. London: Churchill Livingstone; 2000.