Validating Your New Stainer According to CAP and CLIA Requirements

Validating Your New Stainer According to CAP and CLIA Requirements

Congratulations on purchasing a new autostainer! There are three parts to the start-up process:  Instrument Verification, Stain Protocol Optimization, and Validation of the Staining Protocols.

Instrument Verification see CAP All Common Checklist (06/04/2020) COM.40350

Optimally, the company you purchase the instrument from should provide an operator manual; provide verbal procedural instructions; and test the stainer to verify its proper operation. You should also be provided signed documentation confirming that it operates as intended.

If you are in a large hospital system, the Clinical Engineering department will do a safety, operational, and functional inspection. Obtain a copy of the Clinical Engineering records showing their approval of the instrument and include these documents with the other papers you are putting together for the Verification and Validation.

Stain Protocol Optimization

Prior to staining patient tissue, you will need to test the staining protocol(s) to make certain that the stain quality that the new instrument is providing is acceptable for diagnosis. This needs to be done even if the staining protocol is the same one as used on the old instrument – they are different machines, and will operate differently, thus requiring an independent optimization.

To verify what an acceptable H&E stain looks like, look at the photos in reputable Histotechnology reference books such as Carson, Bancroft and Brown. To improve your knowledge, read the differences between reference-quality H&E staining versus poor quality staining. If you ever found yourself in a legal predicament you really don’t have the excuse to say, “well, this is what my pathologist likes”. Consider what could happen if your lab was sued based on the quality of your work and your employer had to go to court. The prosecuting attorneys would show text book quality photos, as established by ASCP, NSH and/or HistoQIP (proficiency standard for US laboratories) to support their case. In turn, your employer would need to show high quality photos as evidence to win its case.

With this ideal in mind, do some test runs with several different types of tissues. Use tissues that you process frequently, such as breast, skin, GI biopsy, and placenta. Use tissues that are sensitive to the stain expressions of the eosin and hematoxylin chemicals, such as small and large intestine.

To assess the staining quality of eosin you may want to use epithelial, muscle and lymphoid tissue to better visualize nuclei. Use muscle, collagen, epithelial cells, and mucin cells to better visualize connective tissue. Use eosinophils and plasma cells to better visualize various WBCs. 

To document your results, you need to design an H&E Stain Optimization Worksheet. Start with a grid that lists all the elements of the protocol that is being used on your H&E stainer.

Document the following details on the H&E Stain Optimization worksheet:

  • Name of solution/reagent in each container
  • Time that slides spend in each container
  • Temperature, if appropriate, e.g., running water
  • pH of appropriate solutions, e.g., water, hematoxylin, eosin, acid rinse
  • Date and test run number
  • Signature of the Tech that oversaw the run
  • Signature of the Pathologist that assessed the results

Next, have a section on the optimization worksheet where you record your results. Some example questions might be:

Overall Stain Quality (observed using a 10x objective):

  • Is the staining even?Are the nuclei standing out darker than the background?
  • Is there an absence of splotches, e.g., water droplets?

Hematoxylin Staining (observed using a 40x objective):

  • Is the nuclear wall dark and crisp?
  • Is the chromatin pattern stippled, not smudgy?
  • Is the nucleolus, if present, a red to purple color?
  • Are other cells (plasma cells and pancreatic acinar cells) which are expected to stain bluish, doing so?
  • Are the mucin cells clear of color (often seen as a pale blue color if you are using a Gill hematoxylin and/or are not doing a regressive stain)?
  • Are the muscle and connective tissue cells free from a bluish color?

Eosin Staining (observed using a 40x objective):

  • Are RBCs the darkest red?
  • Are eosinophil granules, Paneth cell granules, and zymogen granules as dark, or nearly as dark, as RBCs. Note, if you are using a fixative with acetic acid, these organelles will be lysed and this question does not apply.
  • Is muscle tissue a medium shade of pink, and is collagen a light shade of pink?
  • Can muscle be differentiated from collagen? Observing medium size blood vessels should show this differentiation.

If the results are not satisfactory, follow up to find out what is causing the problem. Then make changes to the H&E protocol, run another rack of test slides, and then record the results. Continue doing this until you finally get good quality H&E staining. This is now your optimized H&E Stain Protocol.

Now, you are required to do a control run every day, which can tell you when you need to rotate or change solutions/reagents. Or, you may want to do a control run every 200 or 400 slides.

Validation of the Staining Protocol

Before the stainer is used for any patient slides, you should validate the staining program(s).

According to the new CAP standards which were published in June 2020, Validation of the staining protocols is no longer required! COM.40350 – see NOTE 8: This checklist requirement (validation) does not apply to LDTs that employ the following methods: Manual microscopy (eg, histopathologic and cytologic interpretation, microscopic examination of blood or body fluids, Gram stains)”

HOWEVER, all US labs are licensed by CLIA, and CLIA does require stain protocol validation. You could still be inspected by a CLIA inspector even if you are CAP accredited, and if you did not do validations you would be cited.

Each different H&E program must be separately validated. This means that if you use one program with more delicate staining for your biopsies, and a different program for routine surgical specimens, both staining programs must be separately validated. To do this, stain 20 different slides of differing common tissue types according to your optimized protocol. Twenty is the commonly accepted number of test runs for most laboratory validations.

Design your Staining Protocol Validation Worksheet with the following details:

  • A header that includes the make, model, and serial number of the instrument.
  • The body to record the following information:
  • Accession/ID number for the 20 slides
  • Tissue type
  • Review approval/non-approval
  • Comments
  • A footer with the following information:
  • A statement which says: “This protocol has been validated and is approved for patient use”.
  • A signature sign-off for the Medical Director and the date
  • The lab name and address

You are required to keep the records of the Instrument Verification and the Staining Protocol Validations for the years you own the instrument plus two years. Optimization records are not required to be archived.

Re-validation of the instrument is required if:

  • The staining protocols are changed
  • The solutions/reagents are changed
  • The instrument is moved to a different location, within or outside of your lab
  • The instrument has had any major repairs


  1. Peggy A. Wenk, BA, BS, HTL(ASCP)SLS, Former Program Director, Beaumont School of Histotechnology
  2. Beth A. Cox, HTL/SCT(ASCP)QIHC, 11/09/2020
  3. Robert G. Rankin, MSM, SM(ASCP), 11/09/2020
  4. CAP All Common Checklist COM.40350, 06.04.2020

Small Specimens

Small Specimens

Some specimens may be very tiny; on the order of less than 0.1 cm.  Some methods employ the use of mesh cassettes, “tea bag” biopsy pouches, sponges, wrapping paper, etc. in order 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 has to be handled yet again, possibly resulting in additional fragmentation of the tissue, or possibly complete tissue loss.

A proper method of using wrapping papers is described by Dr. Izak B Dimenstein in his Technical Note published in the Journal of Histotechnology (2016.  Vol. 39, No. 3, pages 76-80).  This method works very well for fragile tissues such as prostate and breast needle biopsies.  Dr. Dimenstein notes that 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.

With regard to 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.

With regard to embedding, 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 has a new product called “embedding gel” that can be used in conjunction with their Paraform cassettes.  This material can be used to hold small specimens in place, in proper orientation.

An alternative method is to use a product called “HistoGel”.  HistoGel is a liquid at 55 C, and has a gelatin consistency at room temperature.  The idea is simply to surround the tiny tissue fragment(s) with liquefied HistoGel, allow it to cool to room temperature (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.)  During embedding, the button is simply removed from the cassette and embedded as usual.  Additional slides may have to be taken in order to reach the fragment; however, the tissue cannot be lost in processing.  This fact far outweighs the extra effort involved during the surgical grossing procedure.

Figure 1A shows a black ink dot, marking the presence of a small group of cells that represent a tiny fragment submitted in Histogel.  Figure 1B is a high power micrograph of the specimen stained with PAS.  Fungi are clearly demonstrated.  The specimen was so small that it had to be serially sectioned.  Slides #5, 6, and 7 out of 30 slides showed the tissue fragment.



  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
  4. Dermatopathology Laboratory Techniques.  CM Chapman, I Dimenstein.  In press.