SOLUTION:

	1 Liter	1 Gallon	20 Liters
Hydrochloric Acid 5%, Aqueous	Part 12086A	Part 12086B	Part 12086C

 

For storage requirements and expiration date refer to individual bottle label.

 

APPLICATION:

Newcomer Supply Hydrochloric Acid 5%, Aqueous is a ready-to-use solution for acid cleaning glassware and instrumental in ensuring laboratory glassware/plasticware is properly acid cleaned prior to solution preparation and performing silver, iron or calcium staining procedures.

 

METHOD:

Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. 1. Before proceeding with acid cleaning, glassware should be cleaned with a laboratory grade glassware detergent, washed and rinsed.
      1. 1. Most new glass is slightly alkaline and should be washed before initial use.
   2. Using safety precautions, pour sufficient Hydrochloric Acid 5%, Aqueous into glassware; gently swirl to thoroughly coat all surfaces/sides/edges with the solution.
   3. Repeat this process for all glassware/plasticware to be used in procedures that requires acid cleaning, including bottles, beakers, flasks, Coplin jars, graduated cylinders, stir rods, pipettes, thermometers and lids.
      1. 1. See Procedure Note #1.
   4. Rinse all glassware/plasticware with a minimum of four changes of distilled water, ensuring all surfaces/sides/edges are well rinsed.
      1. 1. It is essential that final rinses are distilled water or chlorine-free water.
         2. Tap water rinses may leave surface contaminates on glassware resulting in potential staining issues.
   5. Dry glassware in a designated area for acid cleaned glassware.

 

PROCEDURE NOTE:

1. 1. Use fresh Hydrochloric Acid 5%, Aqueous with every batch of glassware acid cleaned. The solution can be reused on multiple pieces of glassware/plasticware acid cleaned at the same time.

 

REFERENCES:

1. 1. aceglass.com. “What Are Good Cleaning Techniques for Laboratory Glassware?,” December 20, 2016.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 134.
   3. Modifications developed by Newcomer Supply Laboratory.

 

• Shelf Life is 2 years from date of manufacture.

 

(use: Alcian Blue 1.0 pH Sulfated Mucosubstances.)

(Vented cap allows for release of pressure. Condensation inside shipping bag may occur.)

STAIN SOLUTION:

	Part 1202A	Part 1202B	Part 1202C
Hematoxylin Stain, Mayer Modified	500ml	1 Liter	4 Liters

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Lithium Carbonate, Saturated Aqueous                 OR Scott Tap Water Substitute	Part 12215     OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844
Eosin Y Working Solution                OR Eosin Y-Phloxine B Working Solution	Part 1072     OR Part 1083

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Hematoxylin Stain, Mayer Modified is a ready to use progressive hematoxylin that does not require filtering and does not contain chloral hydrate or alcohol. Due to the progressive staining nature of Mayer hematoxylin, over-staining is less likely and a differentiation step is not required.  Mayer hematoxylin is an excellent choice as a counterstain in immunohistochemistry (IHC), amyloid and copper staining procedures.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material; nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  Longevity of hematoxylin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

H&E STAINING PROCEDURE WITH MAYER MODIFIED:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Stain with Hematoxylin Stain, Mayer Modified, 10 to 20 minutes, depending on preference of nuclear stain intensity.
      1. 1. See Procedure Note #3.
   4. Wash well in tap water for 3 minutes.
   5. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   6. Wash in three changes of tap water; rinse in distilled water.
   7. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   8. Counterstain in Eosin Y Working Solution (Part 1072) or Eosin Y-Phloxine B Working Solution (Part 1083) for 30 seconds to 3 minutes, depending on preference of intensity.
   9. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.  Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Bright pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Special stains or IHC nuclear counterstain may require less time.
   4. Store hematoxylin at room temperature in tightly capped container away from direct light to minimize oxidation and extend shelf life.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 112-115.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Patholog 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 36-38.
   3. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 127-130.
   4. Modifications developed by Newcomer Supply Laboratory.

STAIN SOLUTION:  

	Part 1201A	Part 1201B	Part 1201C
Hematoxylin Stain, Harris Modified	500ml	1 Liter	1 Gallon

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                  OR Scott Tap Water Substitute	Part 12215     OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844
Eosin Y Working Solution               OR Eosin Y-Phloxine B Working Solution	Part 1072      OR Part 1083

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Hematoxylin Stain, Harris Modified is a ready to use regressive hematoxylin that does not require filtering, is mercury-free and can be used in either manual or automated staining platforms. This modified Harris formulation contains glacial acetic acid for more precise nuclear staining and ethylene glycol to increase solution stability and reduce surface precipitate.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material; nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  Longevity of hematoxylin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

H&E STAINING PROCEDURE WITH HARRIS MODIFIED:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Stain with Hematoxylin Stain, Harris Modified, 1 to 5 minutes, depending on preference of nuclear stain intensity.
   4. Wash well in three changes of tap water.
   5. Differentiate quickly in Acid Alcohol 1% (Part 10011).
      1. 1. Nuclei should be distinct and background very light to colorless.
   6. Rinse well in three changes of tap water.
   7. Blue in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   8. Wash in three changes of tap water; rinse in distilled water.
   9. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   10. Counterstain in Eosin Y Working Solution (Part 1072) or Eosin Y-Phloxine B Working Solution (Part 1083) for 30 seconds to 3 minutes, depending on preference of intensity.
   11. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Bright pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Store hematoxylin at room temperature in tightly capped container away from direct light to minimize oxidation and extend shelf life.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 112-115.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 142-144, 153-154.
   3. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 127-130.
   4. Modifications developed by Newcomer Supply Laboratory.

STAIN SOLUTION:

	Part 12013A	Part 12013B	Part 12013C
Hematoxylin Stain, Harris	500ml	1 Liter	1 Gallon

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                    OR Scott Tap Water Substitute	Part 12215      OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844
Eosin Y Working Solution                OR Eosin Y-Phloxine B Working Solution	Part 1072      OR Part 1083

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Hematoxylin Stain, Harris is a ready to use regressive hematoxylin that does not require filtering, is mercury-free and does not contain glacial acetic acid or ethylene glycol.   This Harris formulation is designed for either manual or automated staining.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material; nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of  tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.   Longevity of hematoxylin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

H&E STAINING PROCEDURE WITH HEMATOXYLIN, HARRIS:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Stain with Hematoxylin Stain, Harris, 1 to 5 minutes, depending on preference of nuclear stain intensity.
   4. Wash well in three changes of tap water.
   5. Differentiate quickly in Acid Alcohol 1% (Part 10011).
      1. 1. Nuclei should be distinct and background very light to colorless.
   6. Rinse well in three changes of tap water.
   7. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   8. Wash in three changes of tap water; rinse in distilled water.
   9. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   10. Counterstain in Eosin Y Working Solution (Part 1072) or Eosin Y-Phloxine B Working Solution (Part 1083) for 30 seconds to 3 minutes, depending on preference of intensity.
   11. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.  Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Bright pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Store hematoxylin at room temperature in tightly capped container away from direct light to minimize oxidation and extend shelf life.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 112-115.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 142-144, 153-154.
   3. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 127-130.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Elastic-Masson Trichrome combination stain with myocardial applications.)

(use: Verhoeff-Van Gieson Elastic Stain.)

CI 75290

• Shelf Life is 4 years from date of manufacture.

 

SOLUTION:

	500 ml
Haupt Gelatin Adhesive	Part 1151A

 

For storage requirements and expiration date refer to individual bottle label.

 

APPLICATION:

Newcomer Supply Haupt Gelatin Adhesive is a blended solution of high-quality gelatin, glycerin and phenol with a variety of procedural uses including subbed slide/direct slide coating applications.  Haupt Gelatin Adhesive works to create a strong adhesive bond between tissue sections and microscopic slides to prevent or reduce the loss of sections due to the nature of the tissue and tissue treatments, such as:

• • Thick sections
  • Harsh staining procedures
  • Animal tissues
  • Plant preparations
  • Bone specimens
  • Resin, plastic/methyl methacrylate (MMA) sections

 

METHOD:

Technique:  Frozen, paraffin or resin, plastic/MMA sections
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURES:

Haupt Gelatin Adhesive Subbed Slide Preparation:

1. 1. Use only clean and dry microscopic slides.
   2. Place a large drop of Haupt Gelatin Adhesive on slides, spread evenly over surface creating a thin film.
      1. 1. Allow a minimum of 30 minutes drying time.
         2. Background staining may occur with thicker films.
         3. See Procedure Note #1.

 

Water Bath Method for Sections on Subbed Slides:

1. 3. Fill water bath with distilled water; maintain temperature at 5°- 10°C below melting point of embedding medium.
      1. 1. See Procedure Note #2.
   4. Float paraffin tissue sections onto Haupt subbed slides.
   5. Warm slide slightly on edge of water bath to straighten section.
   6. Drain and dry slides.

 

Vapor Fixation Method for Sections on Subbed Slides:

1. 7. Mount paraffin or frozen sections on Haupt subbed slides; dry 1 minute.
   8. Under fume hood, add 2-4 ml of concentrated formaldehyde to bottom of a Coplin jar.
   9. Place slides in Coplin jar; formaldehyde should not be in direct contact with tissue sections.
   10. Tightly cover and place in 60°C oven for 30 minutes to 1 hour.
       1. 1. See Procedure Note #3.
   11. Remove slides from Coplin jar; drain and dry.

 

Method for Resin, Plastic/MMA Sections on Subbed Slides:

1. 12. Place droplets of filtered/processed water on Haupt subbed slides.
   13. Transfer resin or plastic/MMA sections to water droplets.
   14. Air dry or use low heat.

 

Method for Resin, Plastic/MMA Sections on Non-Subbed Slides:

1. 15. Place droplets of Haupt Gelatin Adhesive on clean, dry slides.
   16. Transfer resin or plastic/MMA sections directly to Haupt droplets.
   17. Add 1-2 drops of 50% ethanol atop sections.
   18. Manipulate sections by teasing or stretching to remove any folds.
   19. Air dry or use low heat.

 

PROCEDURE NOTES:

1. 1. Dry slides in dust-free environment and store dried subbed slides indefinitely in a slide box at room temperature and low humidity.
   2. Clean interior/exterior of water bath on a daily basis to deter contaminates and residual gelatin adhesive build-up.
   3. Formalin vapor renders gelatin insoluble, strongly affixing tissue sections to subbed slides.

 

REFERENCES:

1. 1. Haupt, Arthur W. “A Gelatin Fixative for Paraffin Sections.” Stain Technology 5.3 (1930): 97-98.
   2. Huang, Bing Quan, Michael John Sumner, Claudio Stasolla and Edward C. Yeung. Plant Microtechniques and Protocols. Springer, 2015. 88, 94.
   3. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 581.
   4. Presnell, Janice, Martin Schreibman and Gretchen Humason. Humason’s Animal Tissue Techniques. 5th ed. Baltimore: Johns Hopkins University Press, 1997. 468.
   5. Modifications developed by Newcomer Supply Laboratory.

Holds 100 slides

• Rust-resistant clasp & hinge

• Stackable

• Cork lined

• Heavy Duty

 

 

(use: Buffer for digestion in Alcian Blue pH 2.5. Does not include enzyme.)

(use: Steiner-Steiner & Steiner-Chapman Modified Silver Stain.)

(use: Brown-Brenn & Brown-Hopps Gram Stains and mercury pigment removal.)

SET INCLUDES:

		Part 1131A
Solution A:	Methenamine 3%, Aqueous	250 ml
Solution B:	Silver Nitrate 5%, Aqueous	50 ml
Solution C:	Sodium Borate 5%, Aqueous	50 ml

Individual stain solutions may be available for purchase under separate part numbers.

 

Additionally Needed For Jones Basement Membrane Stain:

Basement Membrane Control Slides	Part 4055
Hydrochloric Acid 5%, Aqueous	Part 12086 (for acid cleaning glassware)
Periodic Acid 1%, Aqueous	Part 13304
Gold Chloride 0.25%, Aqueous	Part 11287
Sodium Thiosulfate 2.5%, Aqueous	Part 13889
Light Green SF Yellowish Stain 0.1%, Aqueous	Part 12203
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

The Newcomer Supply Jones Basement Membrane Silver Set provides the silver staining solutions for the Jones Basement Membrane Stain for identification of glomerular and tubular basement membranes in renal tissue.  This procedure is also referred to as the Periodic Acid-Methenamine Method (PAMM).

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique: Paraffin sections cut at 4 microns
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Sets are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the set may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.
   3. Prepare Silver-Methenamine Working Solution and mix well:
      1. 1. Solution A: Methenamine 3%, Aqueous      40 ml
         2. Solution B: Silver Nitrate 5%, Aqueous 2 ml
         3. Solution C: Sodium Borate 5%, Aqueous 4 ml
   4. Preheat Silver-Methenamine Working Solution to 45°-60°C in a water bath 20-30 minutes before use.
      1. 1. Maintain solution between 45°-60°C to minimize precipitate.
         2. Do not preheat solution if using Microwave Modification.

 

STAINING PROCEDURE:

1. 5. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #3 and #4.
   6. Place in Periodic Acid 1%, Aqueous (Part 13304) for 15 minutes.
   7. Wash in tap water for 5 minutes; rinse in distilled water.

1. 8. Incubate slides in preheated Silver-Methenamine Working Solution (Step #4) at 45°-60°C or at room temperature for 12-18 minutes until sections appear paper-bag brown.
   9. Periodically remove control, rinse in warm distilled water, check microscopically for adequate silver impregnation. Basement membranes should be dark brown. If tissue structures are not sufficiently dark, return slides to warm silver solution.  Recheck at 2-3 minute intervals until desired intensity is achieved.
      1. 1. Staining at room temperature will require longer incubation.
   10. Microwave Modification: See Procedure Note #5.
       1. 1. Place sides in a plastic Coplin jar (Part 5184) with prepared Silver-Methenamine Working Solution (Step #3). Microwave for 3 minutes at 70°C.
          2. Check microscopically for adequate development.
          3. If additional incubation is required, return slides to heated silver solution and recheck at regular intervals.
   11. Rinse in three changes of distilled water.
   12. Tone in Gold Chloride 0.25%, Aqueous (Part 11287) for 1 minute.
   13. Rinse well in three changes of distilled water.
   14. Place in Sodium Thiosulfate 2.5%, Aqueous (Part 13889) for 2 minutes.
   15. Wash in tap water for 5 minutes; rinse in distilled water.
   16. Counterstain in Light Green SF Yellowish Stain 0.1%, Aqueous (Part 12203) for 1 minute.
   17. Quickly rinse slides in two changes of distilled water.
   18. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Kidney glomerular basement membranes	Black
Intra-glomerular deposits	Black
Reticular fibers	Black
Nuclei	Outlined in black
Cytoplasm	Light green

 

PROCEDURE NOTES:

1. 1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
   2. No metals of any kind should come in contact with silver solutions to prevent precipitation of silver salts.  Use plastic forceps (Part 5500) or paraffin coated metal forceps.
   3. Drain slides after each step to prevent solution carry over.
   4. Do not allow sections to dry out at any point during procedure.
   5. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   6. If using a xylene substitute, follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Jones, David B. “Nephrotic Glomerulonephritis,” American Journal of Pathology 33.2 (1957): 313–329.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 97-99.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 187-188.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Gomori Basement Membrane Stain.)

(use: Gordon & Sweets Reticulum, Fontana-Masson Argentaffin and GMS Stains.)

 

EASYDIP SLIDE STAINING JAR SPECIFICATIONS:

• • Made of acetal polymer
  • Available in 5 colors
  • Use individually or link together as many as you need
  • Resistant to most staining agents including alcohol and xylene*
  • Dimensions: 2 ½ x 3 x 3 5/8 in. H
  • Temperature range: -80°C to +30°C
  • Uses just 80 ml reagent for 12 slides
  • Jars all have attached hinged lids
  • Ideal for special stains, frozen sections and other processes
  • Can be used with the EasyDip Slide Staining Rack (Part 5300RK) (sold separately)

 

*Jars are not resistant to: phenol, iodine, ferric chloride and are not autoclavable.  Also not recommended for Gram stain.

 

 

EASYDIP SLIDE STAINING RACK SPECIFICATIONS:

• • Made of acetal polymer
  • Holds up to 12 slides vertically
  • Slides fit into individual slots for free passage and rapid drainage of staining fluids
  • The lid completely covers the EasyDip Slide Staining Jar to minimize spill and evaporation
  • The handle is permanently attached to the rack for easy insertion and removal of slides without your fingers touching the solution
  • Available in dark gray only
  • Temperature range: -80°C to +30°C
  • Do not use in microwave
  • Dimensions: 2 ¼ x 2 ½ x 3 ¾ in. H
  • Can be used with the EasyDip Slide Staining Jar (Part 5300) (sold separately)

 

Avoid using in picric acid and phenol for prolonged periods of time.

 

The EasyDip Slide Staining Kit Contains: 5 jars (one of each color) & 1 rack

EASYDIP SLIDE STAINING JARS SPECIFICATIONS:

• • Made of acetal polymer
  • Available in 5 colors
  • Use individually or link together as many as you need
  • Resistant to most staining agents including alcohol and xylene*
  • Dimensions: 2 ½ x 3 x 3 5/8 in. H
  • Temperature range: -80°C to +30°C
  • Uses just 80 ml reagent for 12 slides
  • Jars all have attached hinged lids
  • Ideal for special stains, frozen sections and other processes
  • Can be used with the EasyDip Slide Staining Rack (Part 5300RK) (sold separately)

 

*Jars are not resistant to: phenol, iodine, ferric chloride and are not autoclavable.  Also not recommended for Gram stain.

 

EASYDIP SLIDE STAINING RACK SPECIFICATIONS:

• • Made of acetal polymer
  • Holds up to 12 slides vertically
  • Slides fit into individual slots for free passage and rapid drainage of staining fluids
  • The lid completely covers the EasyDip Slide Staining Jar to minimize spill and evaporation
  • The handle is permanently attached to the rack for easy insertion and removal of slides without your fingers touching the solution
  • Available in dark gray only
  • Temperature range: -80°C to +30°C
  • Do not use in microwave
  • Dimensions: 2 ¼ x 2 ½ x 3 ¾ in. H
  • Can be used with the EasyDip Slide Staining Jar (Part 5300) (sold separately)

 

Avoid using in picric acid and phenol for prolonged periods of time.

 

EASYDIP ANODIZED ALUMINUM JAR RACK HOLDER SPECIFICATIONS:

• • Made of anodized aluminum to resist rust, corrosion and abrasion
  • Securely holds up to six Easy Dip Slide Staining Jars
  • Helps prevent tipping
  • Provides easy mobility of the stationed jars, if needed
  • Dimensions: 16 ¾ x 4 x 1 ½ in H
  • Temperature range: -80°C to +30°C
  • To be used with EasyDip Slide Staining Jars (Part 5300) and EasyDip Slide Staining Racks (Part 5300RK) both sold separately

(use: With Millonig Buffer pH 7.0, #12442 for EM.)

(use: Wolbach & May-Grunwald mod. Giemsa Stain.)

CI 15510

• Shelf Life is 4 years from date of manufacture.

 

(use: Brown-Hopps modified Gram Stain.)

 

 

IPC Blue™ Tissue Marking Dyes are a proprietary formula made with Toluidine Blue biopsy marking dye in a 10% neutral buffered formalin.

• Economical – one drop per biopsy.
• Convenient dispenser bottles prevent product from spilling or drying out.
• Continues fixation while grossing.
• Can be added directly to your tissue or on the tissue processor.

 

BENEFITS OF THE IPC BLUE™ TISSUE MARKING DYES:

• Dark blue color allows better visualization of biopsy during embedding & cutting.
• Do not interfere with any other staining.  The IPC Blue™ Dye is totally replaced when the slide is stained.  (When Eosin is used as the marking dye, the tissue section will remain stained with Eosin).
• IPC Blue™ Tissue Marking Dyes do not evaporate as quickly as Eosin, because it is formalin based instead of alcohol based.

 

IPC BLUE™ TISSUE MARKING DYES DIRECTIONS FOR USE:

At the Grossing Station: One to two drops are placed on a biopsy.

On the processor:  1 oz. of IPC Blue™ Tissue Marking Dye is added to the first formalin.  (It is important to note that Eosin CANNOT be used on the tissue processor, as this will remove and replace IPC Blue™ Tissue Marking Dye in the specimen).

The recommended application is to add 1 oz. to the first formalin and then rotate it to the second formalin before changing.

PRACTICAL APPLICATION OF FORMIC ACID 96%, ACS:

Formic Acid 96%, ACS can be used in histology/autopsy as a additional step after fixation to reduce CJD infectivity.

 

TECHNICAL NOTES FOR USING FORMIC ACID 96%, ACS:

Do not reuse open bottle.  Use fresh bottle each time

 

KEY COMPONENT:

>95% FORMIC ACID

 

RESEARCH:

• Abstract on inactivating virus infectivity in formalin-fixed tissue samples from patients with Creutzfeldt-Jakob disease.
• WHO infection control guidelines for transmissible spongiform encephalopathies.

 

REFERENCES:

1. Brown, P., Wolff, A., Gadusek, D.C. 1990.  A simple and effective method for inactivating virus infectivity in formalin-fixed tissue samples from patients with Creutzfeldt-Jakob disease. Neurology 40:887-890.
2. CLSI M29-A3 Protection of Laboratory Workers from Occupationally Acquired Infections; Approved Guideline – Third Edition p 47.

SOLUTION:

	4 X 1 Gallon	20 Liter Cube
Lymph Node Grossing Aid	Part 1093A	Part 1093B

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Lymph Node Grossing Aid is a ready-to-use fixative that serves as either a primary or secondary fixative to enhance the visibility and yield of lymph nodes in specimens in which lymph node examination is critical, such as; breast, colon, lung, radical neck dissection, small intestine and omentum.

The combination of formaldehyde, ethanol and acetic acid in the Lymph Node Grossing Aid solution allows for rapid tissue penetration.  Following sufficient immersion, lymph nodes will be visibly grayish in color, standing out from surrounding tissues and readily identifiable.

 

METHOD:

Fixation Recommendations:

Larger Biopsies: A minimum of 10-12 hours.
Small Biopsies:  A minimum of 4 to 6 hours.

Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue in an adequate amount of Lymph Node Grossing Aid solution after surgical excision.
      1. 1. See Procedure Note #1.
   2. Gross tissue specimen in a manner to allow Lymph Node Grossing Aid to adequately penetrate.
      1. 1. Breast tissue: work with entire specimen or remove smaller area with suspected lymph nodes.  Bread loaf tissue to increase surface area and promote fixative penetration.
         2. Colon tissue: cut open, rinse, pin-out and work with either a segment of colon or entire specimen.
         3. Other specimens: gross to increase tissue surface area and promote fixative penetration.
   3. Return specimens to Lymph Node Grossing Aid solution and fix for a sufficient amount of time, depending upon size and type of tissue.
   4. After adequate immersion in Lymph Node Grossing Aid, remove specimen and re-gross.
      1. 1. Remove adipose and palpate for lymph nodes or identify nodes by their visibly grayish color.
   5. Hold tissues in either Lymph Node Grossing Aid or Formalin 10%, Phosphate Buffered (Part 1090) until ready to process.
      1. 1. See Procedure Note #2.

 

PROCEDURE NOTES:

1. 1. A specimen received in Formalin 10%, Phosphate Buffered can remain in fixative until initial grossing occurs, then transferred to Lymph Node Grossing Aid solution as a secondary fixative.
   2. Extended storage in Lymph Node Grossing Aid is not recommended.
      1. 1. After fixation, transfer Lymph Node Grossing Aid fixed tissue to Formalin 10%, Phosphate Buffered for long-term storage purposes.

 

REFERENCES:

1. 1. Koren, Rumelia, Shlomo Kyzer, Adrian Paz, Vladimir Veltman, Baruch Klein and Rivka Gal. “Lymph Node Revealing Solution: A New Method for Detection of Minute Axillary Lymph Nodes in Breast Cancer Specimens.” The American Journal of Surgical Pathology 21.11 (1997): 1387-1390.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 4.
   3. Newell, Ken, Barry Sawka and Brian Rudrick. “An Inexpensive, Simple and Effective Aid for the Retrieval of Lymph Nodes From Colorectal Cancer Resections.” Archives of Pathology Laboratory Medicine 125 (2001): 642-45.
   4. Modifications developed by Newcomer Supply Laboratory.

 

Tissue Microarray (TMA) is a technique enabling tissues from many donor blocks to be arrayed on a single slide.  The array mold is specifically designed to be simple, easy to use and inexpensive.  Tissues can be analyzed in the same conditions enhancing the efficiency of the research.

The mold kit will allow you to perform TMAs faster while giving excellent results.  By using array molds, you can process up to 170 specimens onto one single slide in very little time.

BENEFITS OF THE TISSUE MICROARRAY MOLDS:

• Mold made of silicone
• 45° cut corner on one corner for sample orientation
• View many different samples on the same slide
• Process up to 170 specimens onto one slide
• Easily stored in a drawer
• Can be reused hundreds of times without losing its flexibility
• Withstands temperatures from -100°C to +250°C

Printable pdf version

INSTRUCTIONS FOR USE:                                      

1		Place the mold in an oven for 30 minutes at 70°C to 80°C.
2		Slowly dispense liquid paraffin (60°C to 65°C) until the top of core rods are fully submerged.  If bubbles are formed, remove them with a pair of heated forceps.
3		Position a cassette on the mold.
4		Fill embedding cassette with paraffin.
5		Cool at room temperature or at about 4°C for 30 to 60 minutes.  Warning:  At lower temperatures, cracks may appear in the block.
6		Slowly separate the mold from the embedding cassette.
7		Trim paraffin around the edges of the recipient block.
8		Extract the marked tissue from the donor block by using the appropriate tissue punch. a.  Place the donor block on a horizontal and flat surface. b.  Hold the tissue punch in your hand perpendicularly to the marked position of the donor block. c.  Slowly insert the tissue punch into the donor block at the proper depth of 5mm.  Don’t insert it too quickly and too deep to prevent damaging the donor block and the tissue punch.
9		By slowly pushing on the tissue punch plunger, deliver the extracted tissue into the corresponding hole of the recipient block.  Then, gently push in all the tissue cores to ensure evenness for microtomy.
10		Place the recipient block on a glass slide (facing down) and incubate the block at 37°C to 45°C for 3 hours up to overnight.  The delivered cores will adhere to their respective holes in the recipient block.  Do not pull the slide from the TMA block.
11	With the recipient block still warm and tacky, heat another slide in an oven to around 70°C for approximately 10 minutes.  Then, place it under the slide that is already stuck to the Array block.  The Array block surface should quickly turn to liquid.  Move the two slides around on the Array block to push any surface air bubbles away and to flatten the Array block surface.
12	Now, remove second slide and place Array block with original slide (slide down) on counter for 10 minutes in order to cool down.  Once Array block is at room temperature, place it with the slide on an ice tray (no water) to cool for 20 minutes.  Slide should remove easily from Array block which will now be ready for cutting.

 

Notes:

1. The tissue punches are not intended for use directly on patients.  For lab/research purposes only.
2. If some of the mold cores are not needed, simply fill unwanted holes in the paraffin Array block with blank paraffin cores.
3. If the Array mold has cracked or split, you can still use it by placing a rubber band or tape around it.  This will keep the Array mold together when paraffin is poured into it.

 

 

• Shelf Life is 2 years from date of manufacture.

 

 

• Shelf Life is 2 years from date of manufacture.

 

Kit includes:

• 10 Specimen Chucks with Colored O-rings & Orientation Mark: 6 – 30mm, 2 – 25mm, 2 – 40mm
• Teasing Needle
• Angular Brush

 

 

Features:

• Colored O-rings
• Orientation Mark
• Proper tolerance and soft aluminum

(use: CNS tissue e.g. large animal brains.)

(use: Used in flotation techniques for ova and parasites.)

• Shelf Life is 2 years from date of manufacture.

 

Features:

• Colored O-rings
• Orientation Mark
• Proper tolerance and soft aluminum

 

Along with offering formalin vials in multiple sizes, we also sell bulk volumes of 10% buffered formalin in 4 x 1 gallon cases and 20 liter (5 gallon) cubes.

 

Features:

• Colored O-rings
• Orientation Mark
• Proper tolerance and soft aluminum

 

Highest quality vials & lids.

 

Features:

• Colored O-rings
• Orientation Mark
• Proper tolerance and soft aluminum

• Shelf Life is 2 years from date of manufacture.

 

TRICHROME, MASSON, FAST GREEN STAIN KIT INCLUDES:

		Part 9180A
Solution A:	Bouin Fluid	250 ml
Solution B:	Ferric Chloride, Acidified	125 ml
Solution C:	Hematoxylin 1%, Alcoholic	125 ml
Solution D:	Biebrich Scarlet-Acid Fuchsin Stain, Aqueous	250 ml
Solution E:	Phosphotungstic Acid 5%, Aqueous	250 ml
Solution F:	Fast Green Stain 2.5%, Aqueous	250 ml
Solution G:	Acetic Acid 0.5%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Trichrome, Masson, Fast Green Stain Kit procedure, with included microwave modification, is used to differentially demonstrate connective tissue elements, collagen and muscle fibers.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the staining procedure provided below.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Solution A: Bouin Fluid to 56-60°C in oven or water bath. (Skip if using overnight method or microwave procedure.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Mordant in preheated Solution A: Bouin Fluid (Step #2) for one hour at 56-60°C or overnight at room temperature. Cool at room temperature for 5-10 minutes.
      1. 1. Skip Step #4 if tissue was originally Bouin fixed.

       Microwave Modification:  See Procedure Note #3.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Solution A: Bouin Fluid. Microwave for 5 minutes at 60°C.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin; combine and mix well.
      1. 1. Solution B: Ferric Chloride, Acidified 20 ml
         2. Solution C: Hematoxylin 1%, Alcoholic           20 ml
   7. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   9. Place in Solution D: Biebrich Scarlet-Acid Fuchsin Stain, Aqueous for 2 minutes.
   10. Rinse in distilled water.
   11. Place in Solution E: Phosphotungstic Acid 5%, Aqueous for 5 minutes.
   12. Transfer directly to Solution F: Fast Green Stain 2.5%, Aqueous; 5-6 minutes.
   13. Rinse in distilled water.
   14. Place in Solution G: Acetic Acid 0.5%, Aqueous; 2 quick dips.
   15. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen and mucin	Green
Muscle fibers, cytoplasm and keratin	Red
Nuclei	Blue/black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining will be compromised.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Brown, Richard. Histologic Preparations: Common Problems and Their Solutions. Northfield, Ill.: College of American Pathologists, 2009. 95-101.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 162-165.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 190.
   4. Vacca, Linda L. Laboratory Manual of Histochemistry. New York: Raven Press, 1985. 308-310.
   5. Modifications developed by Newcomer Supply Laboratory.

TRICHROME, MASSON, ANILINE BLUE STAIN KIT INCLUDES:

		Part 9179A	Part 9179B
Solution A:	Bouin Fluid	250 ml	500 ml
Solution B:	Ferric Chloride, Acidified	125 ml	250 ml
Solution C:	Hematoxylin 1%, Alcoholic	125 ml	250 ml
Solution D:	Biebrich Scarlet-Acid Fuchsin Stain, Aqueous	250 ml	500 ml
Solution E:	Phosphomolybdic-Phosphotungstic Acid, Aqueous	250 ml	500 ml
Solution F:	Aniline Blue Stain, Aqueous	250 ml	500 ml
Solution G:	Acetic Acid 0.5%, Aqueous	250 ml	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Trichrome, Masson, Aniline Blue Stain Kit procedure, with included microwave modification, is used to differentially demonstrate connective tissue elements, collagen and muscle fibers.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Solution A: Bouin Fluid to 56-60°C in oven or water bath. (Skip if using overnight method or microwave procedure.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Mordant in preheated Solution A: Bouin Fluid (Step #2) for one hour at 56-60°C or overnight at room temperature. Cool at room temperature for 5-10 minutes.
      1. 1. Skip Step #4 if tissue was originally Bouin fixed.

       Microwave Modification:  See Procedure Note #3.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Solution A: Bouin Fluid. Microwave for 5 minutes at 60°C.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin; combine and mix well.
      1. 1. Solution B: Ferric Chloride, Acidified 20 ml
         2. Solution C: Hematoxylin 1%, Alcoholic         20 ml

1. 7. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   9. Place in Solution D: Biebrich Scarlet-Acid Fuchsin Stain, Aqueous for 2 minutes.
   10. Rinse in distilled water.
   11. Place in Solution E: Phosphomolybdic-Phosphotungstic Acid, Aqueous for 10-15 minutes.
   12. Move directly to Solution F: Aniline Blue Stain, Aqueous; 5 minutes.
   13. Rinse in distilled water.
   14. Place in Solution G: Acetic Acid 0.5%, Aqueous for 3-5 minutes.
   15. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen and mucin	Blue
Muscle fibers, cytoplasm and keratin	Red
Nuclei	Blue/black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining may be compromised.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Brown, Richard. Histologic Preparations: Common Problems and Their Solutions. Northfield, Ill.: College of American Pathologists, 2009. 95-101.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 162-165.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 190.
   4. Vacca, Linda L. Laboratory Manual of Histochemistry. New York: Raven Press, 1985. 308-310.
   5. Modifications developed by Newcomer Supply Laboratory.

(use: Weigert, Lillie & Mallory Iron Hematoxylins: Elastic and Schmorl Melanin stains. Can be diluted to 5% & 2%.)

TRICHROME, McLETCHIE, ANILINE BLUE STAIN KIT INCLUDES:

		Part 9177A
Solution A:	Biebrich Scarlet-Acid Fuchsin Stain, Aqueous	250 ml
Solution B:	Iodine, Lugol’s, Aqueous	250 ml
Solution C:	Phosphotungstic Acid 2%, Alcoholic	250 ml
Solution D:	Aniline Blue Stain, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Trichrome, McLetchie, Aniline Blue Stain Kit  procedure is for the differential demonstration of collagen and muscle fibers.  This modified trichrome protocol provides time efficient results without the use of Bouin Fluid or a hematoxylin nuclear stain.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Place in Solution A: Biebrich Scarlet-Acid Fuchsin Stain, Aqueous for 5 minutes.
   4. Rinse in several changes of distilled water.
   5. Place in Solution B: Iodine, Lugol’s, Aqueous; 2 minutes.
   6. Rinse in several changes of distilled water.
   7. Differentiate one slide at a time in Solution C: Phosphotungstic Acid 2%, Alcoholic, for 15-30 seconds with gentle agitation.
      1. 1. To avoid over-differentiation, do not exceed 30 seconds.
         2. If sections are over-differentiated, wash well in distilled water and repeat Steps #3 through #7.
   8. Rinse quickly in several changes of distilled water.
   9. Stain in Solution D: Aniline Blue Stain, Aqueous for 1-3 minutes.
   10. Rinse in several changes of distilled water.
   11. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen	Blue
Muscle fibers, cytoplasm and keratin	Magenta to red
Nuclei	Dark red

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-instructional Text. 5th ed. Chicago: ASCP Press, 2020. 162-166.
   2. McLetchie, Norman G.B. “Trichrome McLetchie Modification”. Laboratory Procedure: Lakes Region General Healthcare, Laconia, NH.
   3. Modifications developed by Newcomer Supply Laboratory.

TRICHROME, GOMORI ONE-STEP, ANILINE BLUE STAIN KIT INCLUDES:

		Part 9176B	Part 9176A
Solution A:	Bouin Fluid	250 ml	500 ml
Solution B:	Ferric Chloride, Acidified	125 ml	250 ml
Solution C:	Hematoxylin 1%, Alcoholic	125 ml	250 ml
Solution D:	Trichrome Stain, Gomori One-Step, Aniline Blue	250 ml	500 ml
Solution E:	Acetic Acid 0.5%, Aqueous	250 ml	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Trichrome, Gomori One-Step, Aniline Blue Stain Kit procedure, with included microwave modification, uses a one-step solution combining a plasma stain and a connective tissue stain to differentially demonstrate collagen and muscle fibers.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Solution A: Bouin Fluid to 56-60°C in oven or water bath. (Skip if using overnight method or microwave procedure.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Mordant in preheated Solution A: Bouin Fluid (Step #2) for one hour at 56-60°C or overnight at room temperature. Cool at room temperature for 5-10 minutes.
      1. 1. Skip Step #4 if tissue was originally Bouin fixed.

       Microwave Modification:  See Procedure Note #3.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Solution A: Bouin Fluid. Microwave for 5 minutes at 60°C.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin; combine and mix well.
      1. 1. Solution B: Ferric Chloride, Acidified 20 ml
         2. Solution C: Hematoxylin 1%, Alcoholic         20 ml

1. 7. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   9. Stain with Solution D: Trichrome Stain, Gomori One-Step, Aniline Blue for 20 minutes.
   10. Differentiate in Solution E: Acetic Acid 0.5%, Aqueous; 2 minutes.
   11. Rinse quickly in distilled water.
   12. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen and mucin	Blue
Muscle fibers, cytoplasm and keratin	Red
Nuclei	Blue/black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining may be compromised.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Brown, Richard. Histologic Preparations: Common Problems and Their Solutions. Northfield, Ill.: College of American Pathologists, 2009. 95-101.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 165-166.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 191-192.
   4. Vacca, Linda L. Laboratory Manual of Histochemistry. New York: Raven Press, 1985. 308-310.
   5. Modifications developed by Newcomer Supply Laboratory.

(use: Verhoeff Elastic & Hall Bile Stains.)

TRICHROME, GOMORI ONE-STEP, FAST GREEN STAIN KIT INCLUDES:

		Part 9175A
Solution A:	Bouin Fluid	250 ml
Solution B:	Ferric Chloride, Acidified	125 ml
Solution C:	Hematoxylin 1%, Alcoholic	125 ml
Solution D:	Trichrome Stain, Gomori One-Step, Fast Green	250 ml
Solution E:	Acetic Acid 0.5%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Trichrome, Gomori One-Step, Fast Green Stain Kit procedure, with included microwave modification, uses a one-step solution combining a plasma stain and a connective tissue stain to differentially demonstrate collagen and muscle fibers.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the staining provided procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Solution A: Bouin Fluid to 56-60°C in oven or water bath. (Skip if using overnight method or microwave procedure.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Mordant in preheated Solution A: Bouin Fluid (Step #2) for one hour at 56-60°C or overnight at room temperature. Cool at room temperature for 5-10 minutes.
      1. 1. Skip Step #4 if tissue was originally Bouin fixed.

       Microwave Modification:  See Procedure Note #3.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Solution A: Bouin Fluid. Microwave for 5 minutes at 60°C.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin; combine and mix well.
      1. 1. Solution B: Ferric Chloride, Acidified 20 ml
         2. Solution C: Hematoxylin 1%, Alcoholic           20 ml

1. 7. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   9. Stain in Solution D: Trichrome Stain, Gomori One-Step, Fast Green for 20 minutes.
   10. Differentiate in Solution E: Acetic Acid 0.5%, Aqueous; 2 minutes.
   11. Rinse quickly in distilled water.
   12. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen and mucin	Green
Muscle fibers, cytoplasm and keratin	Red
Nuclei	Blue/black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining may be compromised.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Brown, Richard. Histologic Preparations: Common Problems and Their Solutions. Northfield, Ill.: College of American Pathologists, 2009. 95-101.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 165-166.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 191-192.
   4. Vacca, Linda L. Laboratory Manual of Histochemistry. New York: Raven Press, 1985. 308-310.
   5. Modifications developed by Newcomer Supply Laboratory.

(use: Schmorl Melanin Stain.)

JONES BASEMENT MEMBRANE STAIN KIT INCLUDES:

		Part 9167A
Solution A:	Methenamine 3%, Aqueous	250 ml
Solution B:	Silver Nitrate 5%, Aqueous	50 ml
Solution C:	Sodium Borate 5%, Aqueous	50 ml
Solution D:	Periodic Acid 1%, Aqueous	250 ml
Solution E:	Gold Chloride 0.25%, Aqueous	250 ml
Solution F:	Sodium Thiosulfate 2.5%, Aqueous	250 ml
Solution G:	Light Green SF Yellowish Stain 0.1%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Hydrochloric Acid 5%, Aqueous	Part 12086 (for acid cleaning glassware)
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Jones Basement Membrane Stain Kit procedure, with included microwave modification, is a silver technique for identification of glomerular and tubular basement membranes in renal tissue.  A light green counterstain is used to enhance results.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique: Paraffin sections cut at 4 microns
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.
   3. Prepare Silver-Methenamine Working Solution and mix well:
      1. 1. Solution A: Methenamine 3%, Aqueous      40 ml
         2. Solution B: Silver Nitrate 5%, Aqueous 2 ml
         3. Solution C: Sodium Borate 5%, Aqueous 4 ml
   4. Preheat Silver-Methenamine Working Solution to 45°-60°C in a water bath 20-30 minutes before use.
      1. 1. Maintain solution between 45°-60°C to minimize precipitate.
         2. Do not preheat solution if using Microwave Modification.

 

STAINING PROCEDURE:

1. 5. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #3 and #4.
   6. Place in Solution D: Periodic Acid 1%, Aqueous for 15 minutes.
   7. Wash in tap water for 5 minutes; rinse in distilled water.

1. 8. Incubate slides in preheated Silver-Methenamine Working Solution (Step #4) at 45°-60°C or at room temperature for 12-18 minutes until sections appear paper-bag brown.
   9. Periodically remove control, rinse in warm distilled water, check microscopically for adequate silver impregnation. Basement membranes should be dark brown. If tissue structures are not sufficiently dark, return slides to warm silver solution.  Recheck at 2-3 minute intervals until desired intensity is achieved.
      1. 1. Staining at room temperature will require longer incubation.
   10. Microwave Modification: See Procedure Note #5 (page 2).
       1. 1. Place slides in a plastic Coplin jar with prepared Silver-Methenamine Working Solution (Step #3). Microwave for 3 minutes at 70°C.
          2. Check microscopically for adequate development.
          3. If additional incubation is required, return slides to heated silver solution and recheck at regular intervals.
   11. Rinse in three changes of distilled water.
   12. Tone in Solution E: Gold Chloride 0.25%, Aqueous for 1 minute.
   13. Rinse well in three changes of distilled water.
   14. Place in Solution F: Sodium Thiosulfate 2.5%, Aqueous; 2 minutes.
   15. Wash in tap water for 5 minutes; rinse in distilled water.
   16. Counterstain in Solution G: Light Green SF Yellowish Stain 0.1%, Aqueous for 1 minute.
   17. Quickly rinse slides in two changes of distilled water.
   18. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Kidney glomerular basement membranes	Black
Intra-glomerular deposits	Black
Reticular fibers	Black
Nuclei	Outlined in black
Cytoplasm	Light green

 

PROCEDURE NOTES:

1. 1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
   2. No metals of any kind should come in contact with silver solutions to prevent precipitation of silver salts.  Use plastic forceps (Part 5500) or paraffin coated metal forceps.
   3. Drain slides after each step to prevent solution carry over.
   4. Do not allow sections to dry out at any point during procedure.
   5. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   6. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Jones, David B. “Nephrotic Glomerulonephritis,” American Journal of Pathology2 (1957): 313–329.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 97-99.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 187-188.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Working solution for VVG Elastic Stain.)

SOLUTION:     

	250 ml	500 ml
Fast Green Stain 2.5%, Aqueous	Part 10852A	Part 10852B

 

Additionally Needed:

Trichrome, Liver Control Slides                    OR Trichrome, Multi-Tissue Control Slides	Part 4690     OR Part 4693
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Bouin Fluid	Part 1020
Hematoxylin Stain Set, Weigert Iron	Part 1409
Biebrich Scarlet-Acid Fuchsin Stain, Aqueous	Part 10161
Phosphotungstic Acid 5%, Aqueous	Part 13345
Acetic Acid 0.5%, Aqueous	Part 100121
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Fast Green Stain 2.5%, Aqueous, a component of the Trichrome Stain, Masson, Fast Green procedure, is used to differentially demonstrate connective tissue elements, collagen and muscle fibers.   The microwave modification for this stain is included.

Fast Green Stain 2.5%, Aqueous is also used as a counterstain in the Safranin O Stain for Cartilage.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply stain procedures are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.

 

PRESTAINING PREPARATION: 

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Bouin Fluid (Part 1020) to 56-60°C in oven or water bath. (Skip if using overnight method or microwave procedure.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Mordant in preheated Bouin Fluid (Step #2) for one hour at 56-60°C or overnight at room temperature. Cool at room temperature for 5-10 minutes.
      1. 1. Skip Step #4 if tissue was originally Bouin fixed.

       Microwave Modification:  See Procedure Note #3.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Bouin Fluid. Microwave for 5 minutes at 60°C.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin (Part 1409); combine, mix well.
      1. 1. Solution A: Ferric Chloride, Acidified 20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic 20 ml

1. 7. Stain slides in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   9. Place in Biebrich Scarlet-Acid Fuchsin Stain, Aqueous (Part 10161) for 2 minutes.
   10. Rinse in distilled water.
   11. Place in Phosphotungstic Acid 5%, Aqueous (Part 13345); 5 minutes.
   12. Transfer directly to Fast Green Stain 2.5%, Aqueous; 5-6 minutes
   13. Rinse in distilled water.
   14. Place in Acetic Acid 0.5%, Aqueous (Part 100121); 2 quick dips.
   15. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Collagen and mucin	Green
Muscle fibers, cytoplasm and keratin	Red
Nuclei	Blue/black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining will be compromised.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Brown, Richard. Histologic Preparations: Common Problems and Their Solutions. Northfield, Ill.: College of American Pathologists, 2009. 95-101.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 162-165.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 191-192.
   4. Vacca, Linda L. Laboratory Manual of Histochemistry. New York: Raven Press, 1985. 308-310.
   5. Modifications developed by Newcomer Supply Laboratory.

(FYI: Higher % Isopropyl Alcohol & contains ketone.)

See also Alcohol Denatured ACS.

MUCIN, MAYER MUCICARMINE STAIN KIT INCLUDES:

		Part 9151A	Part 9151B
Solution A:	Ferric Chloride, Acidified	125 ml	250 ml
Solution B:	Hematoxylin 1%, Alcoholic	125 ml	250 ml
Solution C:	Mucicarmine Stock Stain, Mayer	125 ml	125 ml
Solution D:	Metanil Yellow Stain, Aqueous	250 ml	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual bottle labels. 

 

APPLICATION:

Newcomer Supply Mucin, Mayer Mucicarmine Stain Kit procedure, with included microwave modification, stains acid epithelial mucin (sialomucin, sulfomucin) and useful for the demonstration of the encapsulated yeast Cryptococcus neoformans.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Prepare fresh Weigert Iron Hematoxylin Working Solution directly before use; combine and mix well.
      1. 1. Solution A: Ferric Chloride, Acidified   20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic             20 ml
   4. Stain in fresh Weigert Iron Hematoxylin Working Solution for 7 minutes.
   5. Rinse in running tap water for 10 minutes.
   6. Prepare fresh Mayer Mucicarmine Working Solution; combine and mix well.
      1. 1. Solution C: Mucicarmine Stock Stain, Mayer   10 ml   
         2. Tap Water (do not use distilled water)   30 ml    
   7. Stain in fresh Mayer Mucicarmine Working Solution for 60 minutes or longer if a more intense stain is desired.

Microwave Modification:  See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar containing fresh Mayer Mucicarmine Working Solution. Microwave for 10 minutes at 70°C.

8. 8. Rinse in several changes of tap water.
   9. Counterstain in Solution D: Metanil Yellow Stain, Aqueous for 30 to 60 seconds.
   10. Dehydrate quickly through 95% and 100% ethyl alcohols. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

1.                                                                                                                                                                              

RESULTS:

Acid epithelial mucin	Deep rose to red
Capsule of Cryptococcus neoformans	Deep rose to red
Nuclei	Black
Other tissue elements	Yellow

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 174-175.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 142-144.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 168-169.
   4. Modifications developed by Newcomer Supply Laboratory.

MOVAT-RUSSELL MODIFIED PENTACHROME STAIN KIT INCLUDES:

(FYI: Higher % Isopropyl Alcohol & contains ketone.)
See also Alcohol Denatured ACS.

IRON, GOMORI PRUSSIAN BLUE STAIN KIT INCLUDES:

		Part 9136A	Part 9136B
Solution A:	Hydrochloric Acid 20%, Aqueous	125 ml	250 ml
Solution B:	Potassium Ferrocyanide 10%, Aqueous	125 ml	250 ml
Solution C:	Nuclear Fast Red Stain, Kernechtrot	250 ml	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Hydrochloric Acid 5%, Aqueous	Part 12086 (for acid cleaning glassware)
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Iron, Gomori Prussian Blue Stain Kit procedure detects loosely bound ferric iron in tissue sections. This histochemical reaction is sensitive enough to demonstrate minute amounts of iron deposits in blood cells, bone marrow and spleen.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Acid clean glassware prior to use to avoid residual iron staining.
      1. 1. See Procedure Note #1.

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #2 and #3.
   4. Prepare fresh Ferrocyanide Working Solution directly before use; combine and mix well.
      1. 1. Solution A: Hydrochloric Acid 20%, Aqueous        20 ml
         2. Solution B: Potassium Ferrocyanide 10%, Aqueous 20 ml
   5. Place in fresh Ferrocyanide Working Solution for 20 minutes.
   6. Rinse in three changes of tap water; rinse in distilled water.
   7. Place in Solution C: Nuclear Fast Red Stain, Kernechtrot for 5 minutes.
      1. 1. Shake solution well before use; do not filter.
   8. Rinse well in distilled water.
      1. 1. See Procedure Note #4.
   9. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Ferric iron deposits	Bright blue
Nuclei	Red
Cytoplasm	Pink

 

PROCEDURE NOTES:

1. 1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
   2. Drain slides after each step to prevent solution carry over.
   3. Do not allow sections to dry out at any point during procedure.
   4. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps

 

REFERENCES:

1. 1. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 179-184.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 217-218.
   3. Modifications developed by Newcomer Supply Laboratory.

 

(FYI: Higher % Isopropyl Alcohol & contains ketone.)

See also Alcohol Denatured ACS.

FUNGUS, PAS/LIGHT GREEN STAIN KIT INCLUDES:

		Part 9122A
Solution A:	Periodic Acid 0.5%, Aqueous	250 ml
Solution B:	Schiff Reagent, McManus	250 ml
Solution C:	Light Green SF Yellowish Stain 0.1%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Fungus, PAS/Light Green Stain Kit procedure identifies fungus and glycoproteins in tissue sections. Polysaccharides present in fungal cell walls are oxidized by periodic acid to aldehydes; aldehydes react with Schiff Reagent, McManus to yield magenta colored fungi.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Place in Solution A: Periodic Acid 0.5%, Aqueous for 5 minutes.
   4. Wash in three changes of tap water; rinse in distilled water.
   5. Drain slides of excess water and stain in Solution B: Schiff Reagent, McManus for 20 minutes.
   6. Wash gently in lukewarm tap water for 10 minutes, allowing pink color to develop.
   7. Counterstain in Solution C: Light Green SF Yellowish Stain 0.1%, Aqueous for 5 seconds.
      1. 1. See Procedure Note #3.
   8. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Fungal cell walls and glycogen	Red to magenta
Background	Pale green

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Increase or decrease staining time in Light Green SF Yellowish Stain 0.1%, Aqueous for preference of counterstain intensity.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 321-323.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 245.
   3. Modifications developed by Newcomer Supply Laboratory.

 

SET INCLUDES:

		Part 1082A
Solution A:	Eosin Y Stock Stain 1%, Aqueous	1000 ml
Solution B:	Phloxine B Stock Stain 1%, Aqueous	100 ml

 

Additionally Needed For Eosin-Phloxine Stain Set:

Alcohol, Ethyl Denatured, 95%	Part 10842
Acetic Acid, Glacial, ACS	Part 10010

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Hematoxylin Stain, Harris Modified                   OR Hematoxylin Stain, Harris	Part 1201     OR Part 12013
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                  OR Scott Tap Water Substitute	Part 12215     OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844

 

For storage requirements and expiration date refer to individual product labels. 

 

APPLICATION:

Newcomer Supply Eosin-Phloxine Stain Set solutions are aqueous based, provide an enhanced counterstain to hematoxylin and eosin stains and can be used in either manual or automated staining platforms.  With the addition of the phloxine B component, differentiation of muscle, connective tissue and epithelial elements tend to be sharper and better demonstrated than with the traditional eosin Y solution.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material: nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  The longevity of eosin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

Standard Working Solution:

Solution A:	Eosin Y Stock Stain 1%, Aqueous	100 ml
Solution B:	Phloxine B Stock Stain 1%, Aqueous	10 ml
Alcohol, Ethyl Denatured, 95%		780 ml
Acetic Acid, Glacial, ACS		4 ml

Combine all solutions and mix well.  Store at room temperature for up to one year.

 

H&E STAINING PROCEDURE WITH EOSIN-PHLOXINE:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Stain with Hematoxylin Stain, Harris Modified (Part 1201) or Hematoxylin Stain, Harris (Part 12013) 1-5 minutes, depending on preference of nuclear stain intensity.
   3. Wash well in three changes of tap water.
   4. Differentiate quickly in Acid Alcohol 1%.
      1. 1. Nuclei should be distinct and background very light to colorless.
   5. Rinse well in three changes of tap water.
   6. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   7. Wash in three changes of tap water; rinse in distilled water.
   8. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   9. Counterstain in Eosin-Phloxine Standard Working Solution for 30 seconds to 3 minutes, depending on preference of intensity.
   10. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Bright pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 119-120.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 86-87, 91-92.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 143-144, 153-154.
   4. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 126-130.
   5. Modifications developed by Newcomer Supply Laboratory.

 

STAIN SOLUTION: 

	500 ml	1 Liter	1 Gallon
Eosin Y Stock Stain 1%, Aqueous	Part 1080B	Part 1080C	Part 1080D

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Acetic Acid, Glacial, ACS	Part 10010
Hematoxylin Stain, Harris Modified                  OR Hematoxylin Stain, Harris	Part 1201     OR Part 12013
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                 OR Scott Tap Water Substitute	Part 12215     OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Eosin Y Stock Stain 1%, Aqueous provides the key component of an Eosin Y Working Solution in the hematoxylin and eosin stain and can be used in either manual or automated staining platforms.  Eosin’s value is in its ability to distinguish between the cytoplasm of different types of cells by staining cytoplasmic components differing shades and intensities of pink to red.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material: nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  The longevity of eosin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

Standard Eosin Y Working Solution:

Eosin Y Stock Stain 1%, Aqueous	200 ml
Alcohol, Ethyl Denatured, 95%	600 ml
Acetic Acid, Glacial, ACS	4 ml

Combine all solutions and mix well.  Store at room temperature for up to one year.

 

H&E STAINING PROCEDURE WITH EOSIN Y:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Stain with Hematoxylin Stain, Harris Modified (Part 1201) or Hematoxylin Stain, Harris (Part 12013) 1-5 minutes, depending on preference of nuclear stain intensity.
   3. Wash well in three changes of tap water.
   4. Differentiate quickly in Acid Alcohol 1%.
      1. 1. Nuclei should be distinct and background very light to colorless.
   5. Rinse well in three changes of tap water.
   6. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   7. Wash in three changes of tap water; rinse in distilled water.
   8. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   9. Counterstain in Eosin Y Working Solution for 30 seconds to 3 minutes, depending on preference of intensity.
   10. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 119-120.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 86-87, 91-92.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 143-144, 153-154.
   4. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 126-130.
   5. Modifications developed by Newcomer Supply Laboratory.

ELASTIC, VERHOEFF STAIN KIT INCLUDES:

COPPER, RHODANINE STAIN KIT INCLUDES:

		Part 9113A
Solution A:	Rhodanine Stock Stain 0.2%, Alcoholic	50 ml
Solution B:	Hematoxylin Stain, Mayer Modified	250 ml
Solution C:	Sodium Borate 0.5%, Aqueous	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Copper, Rhodanine Stain Kit procedure, with included microwave modification, is for the detection of copper and copper-associated protein (CAP). Abnormal copper accumulations are predominantly found in liver tissue, most notably Wilson’s disease.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare Working Rhodanine Solution; combine and mix well.
      1. 1. Shake Solution A: Rhodanine Stock Stain 0.2%, Alcoholic well before each use.
         2. Solution A: Rhodanine Stock Stain 0.2%, Alcoholic  3 ml
         3. Distilled Water    47 ml

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Stain in Working Rhodanine Solution (Step #2) at 60°C for 1-2 hours or at 37°C for 18 hours.

        Microwave Modification:  See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar containing Working Rhodanine Solution. Microwave for 6 minutes at 70°C.

1. 5. At the end of incubation (for both oven and microwave), to avoid unwanted slide precipitate, pour off warm Working Rhodanine Solution into a second Coplin jar; reserve and set aside.
   6. Rinse slides well in several changes of distilled water.
   7. Check positive control slide microscopically to determine adequate copper/reddish brown development.
      1. 1. Return slides to reserved Working Rhodanine Solution if additional incubation is required.

1. 8. Prepare dilute Mayer Hematoxylin Stain directly before use; combine and mix well.
      1. 1. Solution B: Hematoxylin Stain, Mayer Modified    20 ml
         2. Distilled Water                      20 ml
   9. Stain in dilute Mayer Hematoxylin Stain for 10 minutes.
   10. Rinse in distilled water.
   11. Rinse in Solution C: Sodium Borate 0.5%, Aqueous; 2-3 quick dips.
   12. Rinse well in distilled water.
   13. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Copper	Copper/reddish brown
Nuclei	Light blue

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 251.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 258-260.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 230.
   4. Modifications developed by Newcomer Supply Laboratory.

Tech Memo 1: Differential Stain Kit for Smears & Touch Imprints

 

DIFFERENTIAL STAIN KIT FOR SMEARS & TOUCH IMPRINTS INCLUDES:

		Part 9112B
Solution A:	Xanthene Stain	500ml
Solution B:	Thiazine Stain	500ml
Solution C:	Fixative	500ml

Individual stain solutions may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS

Part 1445

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Differential Stain Kit, a modification of the Wright Giemsa Stain, uses a methanol fixative and aqueous based stains to provide a rapid 3-step process for differential assessment of:  peripheral blood smears, touch imprints, fine needle aspirations (FNA), bone marrow biopsy aspirations, and detecting microorganisms.

 

METHOD:

Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

                                                                                                                                                                                             

STAINING PROCEDURE: 

1. 1. Prepare within an accepted time frame, a well-made blood smear, touch imprint, FNA smear or bone marrow aspiration smear/film per your laboratories protocol, focusing on uniform cell distribution.
   2. Allow slides to thoroughly air-dry prior to staining.
   3. Dip dried slides in Solution C: Fixative 5-10 times, one second per dip. Allow excess fixative to drain.
   4. Dip in Solution A: Xanthene Stain 5 times, one second per dip. Allow excess solution to drain.
      1. 1. See Procedure Notes #1, #2 and #3.
   5. Quickly rinse slides with distilled water.
   6. Dip slides in Solution B: Thiazine Stain 5 times, one second per dip. Allow excess solution to drain.
   7. Rinse slides quickly in distilled water.
   8. Allow slides to air-dry, then examine microscopically.
   9. If coverslip is preferred, allow slides to air-dry; dip dried slides in xylene and coverslip with compatible mounting medium.

 

RESULTS:

Erythrocytes:	Pink to yellowish-red
Platelets:	Violet or purple granules

 

Granulocytes

Neutrophils:	Nucleus – Dark blue to violet
	Cytoplasm – Pale pink
	Granules – Purple to lilac
Eosinophils:	Nucleus – Blue
	Cytoplasm – Blue
	Granules – Red to red-orange
Basophils:	Nucleus – Purple or dark blue
	Granules – Dark purple

  

 

Mononuclear Cells

Monocytes:	Nucleus – Violet
	Cytoplasm – Sky blue
Lymphocytes:	Nucleus – Violet
	Cytoplasm – Dark blue
Bacteria/microorganisms:	Deep blue in varying shapes

 

Muscle and collagen	Pale Pink
Nuclei	Blue/violet
Cytoplasm	Varying shades of light blue

 

PROCEDURE NOTES:

1. 1. The division of stains in this kit provides the advantage of varying dips in Solutions A and B to produce degrees of shading and intensity. Never use fewer than three dips of one full second each.
   2. For more intense stain, increase dips in Solutions A and B.
      1. 1. To increase eosinophilic staining; increase dips in Solution A.
         2. To increase basophilic staining; increase dips in Solution B.
   3. For a paler stain; decrease dips in Solutions A and B.
   4. If using a xylene substitute, follow manufacturer’s recommendation for coverslipping application.

 

REFERENCES:

1. 1. Bain, B.J. “Bone Marrow Aspiration”. Journal of Clinical Pathology 54 (2001): 657-663.
   2. Cox, Charles. “Accuracy of Intraoperative Imprint Cytology for Sentinel Lymph Node Evaluation in the Treatment of Breast Carcinoma.” Cancer Cytopathology1 (2005): 13-20.
   3. “Guidelines of the Papanicolaou Society for Fine-Needle Aspiration Procedure and Reporting.” Diagnostic Cytopathology 17 (1997): 239-247.
   4. McPherson, Richard and Matthew Pincus. Henry’s Clinical Diagnosis and Management by Laboratory Methods. 22nd ed. Philadelphia: Elsevier Saunders, 2011. 522-535.
   5. Thompson, Samuel Wisley and Ronald D. Hunt. Selected Histochemical and Histopathological Methods. 2nd ed. Springfield, IL: Thomas, 1966. 756-762.
   6. Modifications developed by Newcomer Supply Laboratory.

 

Tech Memo 2: Differential Stain, Helicobacter Pylori sp. in Tissue Sections

 

DIFFERENTIAL STAIN, HELICOBACTER PYLORI, SP IN TISSUE SECTION INCLUDES:

		500 ml	1 Gallon
Solution A:	Xanthene Stain	Part 10521A	Part 10521B
Solution B:	Thiazine Stain	Part 10522A	Part 10522B

 

Additionally Needed:

Helicobacter sp., Artificial Control Slides	Part 4275
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Differential Stain procedure, a modification of the Wright Giemsa Stain, provides a rapid staining method for demonstration of Helicobacter pylori sp. in gastrointestinal tissue sections. Procedures for both monochromatic and polychromatic versions of the Differential Stain are provided.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. Proceed with either the monochromatic or polychromatic staining method.

 

Monochromatic Staining Method:  See Procedure Note #1.

1. 1. Place slides in Solution B: Thiazine Stain for 1-4 minutes.
   2. Rinse quickly in distilled water to remove excess stain.
   3. Allow slides to air-dry in a vertical position.
   4. Dip dried slides in xylene and coverslip with compatible mounting medium.
      1. 1. See Procedure Note #2.

 

RESULTS:

Helicobacter pylori sp.	Dark blue
Collagen and muscle	Blue
Nuclei	Blue
Cytoplasm	Varying shades of light blue

 

Polychromatic Staining Method: See Procedure Note #1.

1. 1. Place slides in Solution A: Xanthene Stain for 3-5 minutes.
   2. Drain slides briefly; go directly into Solution B: Thiazine Stain for 1-4 minutes.
   3. Rinse well in distilled water.
   4. Allow slides to air-dry in a vertical position.
   5. Dip dried slides in xylene and coverslip with compatible mounting medium.
      1. 1. See Procedure Note #2.

 

RESULTS:

Helicobacter pylori sp.	Dark blue
Collagen and muscle	Pale pink
Nuclei	Blue/violet
Cytoplasm	Varying shades of light blue

                                                                                                                                                                                             

PROCEDURE NOTES:

1. 1. Provided timings are suggested ranges. Optimal staining times will depend upon staining intensity preference.
   2. The elimination of dehydration steps is necessary to retain the dark stain of the organism.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and coverslipping steps.

 

REFERENCES:

1. 1. Potvin, Carol. “A Modified Diff-Quik Stain for Helicobacter pylori in Gastrointestinal Biopsies.” Laboratory Medicine6 (1994): 389-391.
   2. Skipper, Ray and Don DeStephano. “A Rapid Stain for Campylobacter pylori in Gastrointestinal Tissue Sections Using Diff-Quik.” The Journal of Histotechnology 12.4 (1989): 303-304.
   3. Modifications developed by Newcomer Supply Laboratory.

PHOSPHOTUNGSTIC ACID HEMATOXYLIN (PTAH) STAIN KIT INCLUDES:

		Part 9111A
Solution A:	Zenker Fixative, Modified, Zinc Chloride	250 ml
Solution B:	Acetic Acid, Glacial, ACS	25 ml
Solution C:	Potassium Permanganate 0.25%, Aqueous	250 ml
Solution D:	Oxalic Acid 5%, Aqueous	250 ml
Solution E:	Phosphotungstic Acid Hematoxylin (PTAH) Stain	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

The Newcomer Supply Phosphotungstic Acid Hematoxylin (PTAH) Stain Kit procedure, with included microwave modification, is for the  demonstration of collagen, muscle striations and central nervous system (CNS) structures.

 

METHOD:

Fixation: 10% Phosphate Buffered Formalin (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare Zenker Fixative Working Solution; combine and mix well.

Solution A: Zenker Fixative, Modified, Zinc Chloride       38 ml

Solution B: Acetic Acid, Glacial, ACS                              2 ml

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Fix in Zenker Fixative Working Solution (Step #2) at 56°C; 3 hours.

        Microwave Modification:  See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar containing Zenker Fixative Working Solution. Microwave for 5 minutes at 60°C.

1. 1. 5. Wash well in three changes of tap water; rinse in distilled water.
      6. Place in Solution C: Potassium Permanganate 0.25%, Aqueous for 10 minutes.
      7. Wash in three changes of tap water; rinse in distilled water.
      8. Place in Solution D: Oxalic Acid 5%, Aqueous for 10 minutes.
      9. Wash in three changes of tap water; rinse in distilled water.
      10. Stain in Solution E: PTAH Stain for 12-24 hours at room temperature, or 2 hours at 56°
          1. See Procedure Note #4.

        Microwave Modification:

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Solution E: PTAH Stain. Microwave for 7 minutes at 70°C.

1. 11. Dehydrate quickly in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.
       1. 1. Dehydrate quickly, alcohol may extract stain.

 

RESULTS:

Collagen, cartilage, elastic fibers	Deep reddish brown
Muscle striations, fibrin, keratin	Dark blue
Glia fibers	Dark blue
Myelin	Lighter blue
Neurons	Salmon/Pink
Nuclei	Blue

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. Adjust staining time according to preference of intensity. Suggested staining time at 37°C is 18 hours.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008.130-131.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015.. 178-180, 201-202.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 193-194.
   4. Modifications developed by Newcomer Supply Laboratory.

COLLOIDAL IRON, MÜLLER-MOWRY STAIN KIT INCLUDES:

		Part 9110A
Solution A:	Acetic Acid 12%, Aqueous	500 ml x 2
Solution B:	Colloidal Iron Stock	125 ml
Solution C:	Acetic Acid, Glacial, ACS	50 ml
Solution D:	Potassium Ferrocyanide 2%, Aqueous	125 ml
Solution E:	Hydrochloric Acid 2%, Aqueous	125 ml
Solution F:	Van Gieson Stain	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Hydrochloric Acid 5%, Aqueous	Part 12086 (for acid cleaning glassware)
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Colloidal Iron, Mϋller-Mowry Stain Kit procedure demonstrates acid epithelial mucin (sialomucin, sulfomucin) and stromal (mesenchymal) mucin. This method is also excellent for the demonstration of the encapsulated yeast Cryptococcus neoformans.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Acid clean glassware prior to use to avoid residual iron staining.
      1. 1. See Procedure Note #1.
   3. Prepare Colloidal Iron Working Solution; combine and mix well.
      1. 1. Solution B: Colloidal Iron Stock                          20 ml
         2. Solution C: Acetic Acid, Glacial, ACS                   5 ml
         3. Distilled Water                                                    15 ml

 

STAINING PROCEDURE:

1. 4. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #2 and #3.
   5. Place in Solution A: Acetic Acid 12%, Aqueous for 30 seconds.
   6. Drain Slides. Do not rinse.
   7. Place in Colloidal Iron Working Solution (Step #3) for 30 minutes.
   8. Rinse in three changes of Solution A: Acetic Acid 12%, Aqueous; 3 minutes each.
   9. Prepare fresh Ferrocyanide-Hydrochloric Acid Solution directly before use; combine and mix well.
      1. 1. Solution D: Potassium Ferrocyanide 2%, Aqueous 20 ml
         2. Solution E: Hydrochloric Acid 2%, Aqueous        20 ml
   10. Place in Ferrocyanide-Hydrochloric Acid Solution for 15 minutes.

1. 11. Wash in running tap water for 1-5 minutes.
   12. Counterstain in Solution F: Van Gieson Stain for 3-5 minutes.
       1. 1. Proceed directly to dehydration step without rinsing.
   13. Dehydrate in two changes of 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Acid epithelial mucin	Blue
Stromal mucin	Blue
Capsule of Cryptococcus neoformans	Blue
Collagen	Red
Muscle and cytoplasm	Yellow

 

PROCEDURE NOTES:

1. 1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
   2. Drain slides after each step to prevent solution carry over.
   3. Do not allow sections to dry out at any point during procedure.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 175-176.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 151-153.
   3. Rekhtman, Natasha and Justin Bishop. Quick Reference Handbook for Surgical Pathologists. Berlin: Springer, 2011. 69.
   4. Modifications developed by Newcomer Supply Laboratory.

STAIN SOLUTION:

	500 ml	1 Liter	1 Gallon
Eosin Y Working Solution	Part 1072A	Part 1072B	Part 1072C

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Hematoxylin Stain, Harris Modified                 OR Hematoxylin Stain, Harris	Part 1201      OR Part 12013
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                 OR Scott Tap Water Substitute	Part 12215      OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Eosin Y Working Solution is a ready to use counterstain in the hematoxylin and eosin stain and can be used in either manual or automated staining platforms.  Eosin’s value is in its ability to distinguish between the cytoplasm of different types of cells by staining cytoplasmic components differing shades and intensities of pink to red.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material: nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of  tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  The longevity of eosin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

H&E STAINING PROCEDURE WITH EOSIN Y:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Stain with Hematoxylin Stain, Harris Modified (Part 1201) or Hematoxylin Stain, Harris (Part 12013) 1 to 5 minutes, depending on preference of nuclear stain intensity.
   3. Wash well in three changes of tap water.
   4. Differentiate quickly in Acid Alcohol 1%.
      1. 1. Nuclei should be distinct and background very light to colorless.
   5. Rinse well in three changes of tap water.
   6. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   7. Wash in three changes of tap water; rinse in distilled water.
   8. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   9. Counterstain in Eosin Y Working Solution for 30 seconds to 3 minutes, depending on preference of intensity.
   10. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 119-120.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 86-87, 91-92.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 143-144, 153-154.
   4. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 126-130.
   5. Modifications developed by Newcomer Supply Laboratory.

AMYLOID, BENNHOLD CONGO RED STAIN KIT INCLUDES:

		Part 9103A
Solution A:	Congo Red Stain 1%, Aqueous	250 ml
Solution B:	Alkaline Alcohol	250 ml
Solution C:	Hematoxylin Stain, Mayer Modified	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

 

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Amyloid, Bennhold Congo Red Stain Kit procedure, with included microwave modification, is used for identifying extraneous protein deposits in amyloidosis. The use of polarizing lenses is essential for visualizing amyloid positive areas or confirm negativity.

 

METHOD: 

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 8 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Place slides in Solution A: Congo Red Stain 1%, Aqueous ; 1 hour.

        Microwave Modification: See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar (Part 5184) containing Solution A: Congo Red Stain 1%, Aqueous. Microwave for 3 minutes at 70°C.

1. 4. Rinse in two to three changes of tap water; rinse in distilled water.
   5. Differentiate in Solution B: Alkaline Alcohol, 5 to 30 seconds, agitating constantly until slide background is cleared of Solution A: Congo Red Stain 1%, Aqueous.
   6. Rinse in two to three changes of tap water; rinse in distilled water.
   7. Counterstain with Solution C: Hematoxylin Stain, Mayer Modified, 3 to 5 minutes.
   8. Wash in running tap water for 5 to 10 minutes.
   9. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Light Field Microscopy:
	Amyloid	Pink to red
	Nuclei	Blue
Polarized Light:
	Amyloid fluorescence	Green, red, orange and yellow

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. For optimal results sections should be cut at 8 microns to provide more intense staining and allow smaller amyloid deposits to be identified.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Buxbaum, Joel N, David S. Eisenberg, Marcus Fändrich, Ellen D. McPhail, Giampaolo Merlini, Maria J. M. Saraiva, Yoshiki Sekijima and Per Westermark, “Amyloid nomenclature 2024: update, novel proteins, and recommendations by the International Society of Amyloidosis (ISA) Nomenclature Committee,” Amyloid 31, no. 4 (Sept 2024): 249-256, DOI: 10.1080/13506129.2024.2405948.
   2. Howie, Alexander J and Owen-Casey Mared P., “Systematic review of accuracy of reporting of Congo red-stained amyloid in 2010-2020 compared with earlier.” Annuls of Medicine 54, no. 1 (Dec 2022): 2511-2516, DOI: 10.1080/07853890.2022.2123558.
   3. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 366-367.
   4. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 177-178.
   5. Modifications developed by Newcomer Supply Laboratory.

ALCIAN BLUE 1%, pH2.5 STAIN KIT INCLUDES:

		Part 9102A	Part 9102B
Solution A:	Acetic Acid 3%, Aqueous	250 ml	500 ml
Solution B:	Alcian Blue Stain 1%, pH 2.5 Aqueous	250 ml	500 ml
Solution C:	Nuclear Fast Red Stain, Kernechtrot	250 ml	500 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Hyaluronidase	For stromal mucin digestion
Hyaluronidase Buffer	Part 1150 (for stromal mucin digestion)

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply Alcian Blue 1%, pH 2.5 Stain Kit procedure, with included hyaluronidase method for stromal mucin digestion, is designed to stain acid epithelial mucin (sialomucin, sulfomucin) as well as stromal (mesenchymal) mucin. The hyaluronidase digestion step is for further differentiation of acid epithelial from stromal mucin.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Digestion Step: Proceed to Step #4 if not running Digestion.
      1. 1. Two control slides and two patient slides are needed.
         2. Label one control slide and one patient slide “with”.
         3. Label the other control slide and patient slide “without”.
         4. Prepare Hyaluronidase Digestion Solution and mix well.

                                Hyaluronidase                                0.025 gm

                                Hyaluronidase Buffer (Part 1150)               50 ml

1. 1. 1. 5. Prepare separate Coplin jar of Hyaluronidase Buffer.
         6. Preheat both solutions from Steps #3d and #3e to 37°C.
         7. Place slides labeled “with” in preheated Hyaluronidase Digestion Solution and slides labeled “without” in preheated Hyaluronidase Buffer.
         8. Incubate both for 2 hours at 37°

1. 4. Wash all slides in running tap water for 5 minutes; rinse in distilled water. Combine slides for remaining steps.
   5. Place slides in Solution A: Acetic Acid 3%, Aqueous for 3 minutes.
   6. Move slides directly into Solution B: Alcian Blue Stain 1%, pH 2.5 Aqueous. Stain for 30 minutes at room temperature or for 15 minutes in a 37°C water bath.
      1. 1. See Procedure Note #3.

1. 7. Wash in running tap water for 10 minutes; rinse in distilled water.
   8. Counterstain in Solution C: Nuclear Fast Red Stain, Kernechtrot for 5 minutes.
      1. 1. Shake solution well before use; do not filter.
   9. Rinse well in distilled water.
      1. 1. See Procedure Note #4
   10. Dehydrate quickly through two changes of 95% ethyl alcohol and two changes of 100% ethyl alcohol. Clear in three xylene changes, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Acid epithelial mucins	Blue
Stromal (mesenchymal) mucin	Blue
Stromal mucin digestion	Marked loss of staining
Nuclei	Pink-red
Cytoplasm	Pale pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. A brief dip in Solution A: Acetic Acid 3%, Aqueous from Step #5 can be added before water rinses to remove excess Alcian Blue Solution if needed.
   4. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
   5. Sigma Hyaluronidase from Bovine Testes (H3506) is the Hyaluronidase used in the digestion step.
   6. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 145-148.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 172-175.
   3. Modifications developed by Newcomer Supply Laboratory.

(use: in Leica Peloris Processor as tint for tissue prior to embedding)

AFB, FITE STAIN KIT INCLUDES:

		Part 91013A
Solution A:	Xylene/Peanut Oil , 2:1	500 ml
Solution B:	Carbol Fuchsin Stain, Ziehl-Neelsen	250 ml
Solution C:	Acid Alcohol 1%	250 ml
Solution D:	Light Green SF Yellowish Stain 0.1%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS

Part 1445

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply AFB, Fite Stain Kit procedure is used to detect the presence of either Nocardia or Mycobacterium leprae (causative agent of leprosy) in tissue sections. Procedural variations are included for detection of either organism.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Filter Solution B: Carbol Fuchsin Stain, Ziehl-Neelsen with high quality filter paper.
   3. If staining for Nocardia, prepare Diluted Acid Alcohol Solution:
      1. 1. Solution C: Acid Alcohol 1%              20 ml
         2. Distilled water 20 ml      

 

STAINING PROCEDURE:

1. 4. Deparaffinize slides in Solution A: Xylene/Peanut Oil, 2:1, two changes for 10-12 minutes each.
      1. 1. See Procedure Note #1
   5. Drain slides, wipe off excess oil, and blot to opacity, removing residual oil.
      1. 1. See Procedure Note #2.
   6. Stain in freshly filtered Solution B: Carbol Fuchsin Stain, Ziehl-Neelsen for 15 minutes at room temperature.
   7. Rinse well in distilled water.
   8. Differentiation for Nocardia:
      1. 1. Differentiate slides individually in Diluted Acid Alcohol Solution (Step #3) until background is pale pink; 10-20 dips.
         2. Quickly rinse in distilled water and check microscopically for correct differentiation.
   9. Differentiation for Mycobacterium leprae:
      1. 1. Differentiate slides individually in Solution C: Acid Alcohol 1% until sections are light pink; 5-10 dips.
   10. Rinse well in distilled water.
   11. Counterstain in Solution D: Light Green SF Yellowish Stain 0.1%, Aqueous; 5-10 dips.
   12. Rinse in distilled water.
   13. Blot excess water from slide and air-dry or oven-dry completely.
   14. Dip dried slides in xylene and coverslip with compatible mounting medium.

 

RESULTS:

Mycobacterium leprae	Red
Nocardia	Red
Other tissue elements	Green

 

PROCEDURE NOTES:

1. 1. Acid-fastness of leprosy organisms is enhanced when the waxy capsule is protected by the mixture of xylene/peanut oil and avoiding dehydrating solutions.
   2. It is important to blot well, residual oil may produce staining artifact.
   3. A small percentage of Nocardia organisms may resist taking the red stain and stain green due to growth phase of the individual organism.
   4. If using a xylene substitute, follow manufacturer’s recommendation for coverslipping step.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-instructional Text. 5th ed. Chicago: ASCP Press, 2020. 215-216.
   2. Fite, George, P.J. Cambre and M.H. Turner. “Procedure for Demonstrating Lepra Bacilli in Paraffin Sections”. Archives of Pathology 43 (1947). 624-625.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 237.
   4. Modifications developed by Newcomer Supply Laboratory.

STAIN SOLUTION:

	500 ml	1 Liter	1 Gallon
Eosin Y Stock Stain 1%, Alcoholic	Part 1070B	Part 1070C	Part 1070D

 

Additionally Needed For H&E Staining:

Hematoxylin and Eosin (H&E) Control Slides	Part 4278
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Acetic Acid, Glacial, ACS	Part 10010
Hematoxylin Stain, Harris Modified                  OR Hematoxylin Stain, Harris	Part 1201     OR Part 12013
Acid Alcohol 1%	Part 10011
Lithium Carbonate, Saturated Aqueous                 OR Scott Tap Water Substitute	Part 12215     OR Part 1380
Alcohol, Ethyl Denatured, 70%	Part 10844

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Eosin Y Stock Stain 1%, Alcoholic provides the key component of an alcohol-based Eosin Y Working Solution in the  hematoxylin and eosin stain and can be used in either manual or automated staining platforms.  Eosin’s value is its ability to distinguish between the cytoplasm of different types of cells by staining cytoplasmic components differing shades and intensities of pink to red.

Hematoxylin and eosin (H&E) staining is used for screening specimens in anatomic pathology, for research, smears, touch preps and other applications.  Its two primary coloring agents stain all cellular material: nuclei (blue), and cytoplasmic elements (pink-red).  Popularity of this stain is due to its simplicity, ability to clearly demonstrate a variety of tissue components, dependability, repeatability, and speed of use.

Quality Control: Since hematoxylin and eosin staining is the foundation of the diagnostic process, maintaining quality is of critical importance. Procedures will vary between laboratories depending upon volume of slides, automation vs manual staining, chemical hygiene and solution integrity.  The longevity of eosin depends upon these factors and stain quality should be regularly screened with an H&E control slide.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

Standard Eosin Y Working Solution, Alcoholic:

Eosin Y Stock Stain 1%, Alcoholic	200 ml
Alcohol, Ethyl Denatured, 95%	600 ml
Acetic Acid, Glacial, ACS	4 ml

Combine all solutions and mix well.  Store at room temperature for up to one year.

 

H&E STAINING PROCEDURE WITH EOSIN Y:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Stain with Hematoxylin Stain, Harris Modified (Part 1201) or Hematoxylin Stain, Harris (Part 12013) 1-5 minutes, depending on preference of nuclear stain intensity.
   3. Wash well in three changes of tap water.
   4. Differentiate quickly in Acid Alcohol 1%.
      1. 1. Nuclei should be distinct and background very light to colorless.

1. 5. Rinse well in three changes of tap water.
   6. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
   7. Wash in three changes of tap water; rinse in distilled water.
   8. Drain excess water; proceed to 70% ethyl alcohol for 10 dips.
   9. Counterstain in Eosin Y Working Solution, Alcoholic for 30 seconds to 3 minutes, depending on preference of intensity.
   10. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue
Erythrocytes and eosinophilic granules	Pink to red
Cytoplasm and other tissue elements	Various shades of pink

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 119-120.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 86-87, 91-92.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 143-144, 153-154.
   4. Suvarna, S. Kim, Christopher Layton and John D. Bancroft. Bancroft’s Theory and Practice of Histological Techniques. 8th ed. Oxford: Elsevier, 2019. 126-130.
   5. Modifications developed by Newcomer Supply Laboratory.

AFB, BASIC FUCHSIN, ELLIS & ZABROWARNY STAIN KIT INCLUDES:

		Part 91012A
Solution A:	Basic Fuchsin Stain 1%, Alcoholic	250 ml
Solution B:	Acid Alcohol 3%	250 ml
Solution C:	Methylene Blue Stain 0.25%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed are two complimentary unstained positive control slides for the initial verification of staining techniques and reagents.  Verification must be documented by running one Newcomer Supply complimentary positive control slide along with your current positive control slide for the first run. Retain the second complimentary control slide for further troubleshooting, if needed.

Individual stain solutions and additional control slides may be available for purchase under separate part numbers.

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual bottle labels.

 

APPLICATION:

Newcomer Supply AFB, Basic Fuchsin, Ellis & Zabrowarny Stain Kit procedure, with included microwave modification, provides a safe method for staining acid-fast bacilli without the use of phenol.  Results are comparable to the AFB, Ziehl-Neelsen method.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply Stain Kits are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Preheat Solution A: Basic Fuchsin Stain 1%, Alcoholic to 60°C in an oven or water bath. (Skip if using Microwave Modification.)

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Place in preheated Solution A: Basic Fuchsin Stain 1%, Alcoholic (Step #2) for 15 minutes.

Microwave Modification:  See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar containing Solution A: Basic Fuchsin Stain 1%, Alcoholic. Microwave for 1 minute at 70°C.

1. 5. Rinse well in running tap water for 2 to 3 minutes.
   6. Differentiate in Solution B: Acid Alcohol 3% until color no longer runs off the slide and sections are pale pink; 3 to 10 rapid dips.
   7. Wash in running tap water for 1 minute.
   8. Counterstain one slide at a time in Solution C: Methylene Blue Stain 0.25%, Aqueous for 15 to 30 seconds.
      1. 1. Dip slides a few times in counterstain; rinse in tap water, followed by a distilled water rinse.
         2. Check microscopically. Sections should be pale blue.
   9. Wash in running tap water for 1 minute; rinse in distilled water.
   10. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Acid-fast bacilli	Red
Nuclei and background	Shades of blue

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Ellis, R C and L A Zabrowarny. “Safer Staining Method for Acid Fast Bacilli.” Journal of Clinical Pathology 46.6 (1993): 559-560.
   2. Modifications developed by Newcomer Supply Laboratory.

CI 45380

• Shelf Life is 4 years from date of manufacture.

For direct slide application.
Store at 2-8°C.

(use: Copper Stain.)

SOLUTION:

	250 ml	500 ml
Rhodanine Stock Stain 0.2%, Alcoholic	Part 10531A	Part 10531B

                                                                                                                                                                              

Additionally Needed, Rhodanine Stain For Copper:

Copper, Animal Control Slides	Part 4130
Hematoxylin Stain, Mayer Modified	Part 1202
Sodium Borate 0.5%, Aqueous	Part 13824
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

For storage requirements and expiration date refer to individual product labels.

 

APPLICATION:

Newcomer Supply Rhodanine Stain for Copper, with included microwave modification, is for the detection of copper and copper-associated protein (CAP). Abnormal copper accumulations are predominantly found in liver tissue, most notably  Wilson’s disease.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

All Newcomer Supply stain procedures are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare Working Rhodanine Solution; combine and mix well.
      1. 1. Shake Rhodanine Stock Stain 0.2%, Alcoholic well before each use.
         2.  Rhodanine Stock Stain 0.2%, Alcoholic                 3 ml
         3. Distilled Water                                47 ml

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Stain in Working Rhodanine Solution (Step #1) at 60°C for 1-2 hours or at 37°C for 18 hours.

        Microwave Modification:  See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar containing Working Rhodanine Solution. Microwave for 6 minutes at 70°C.

1. 5. At the end of incubation (for both oven and microwave), to avoid unwanted slide precipitate, pour off warm Working Rhodanine Solution into a second Coplin jar; reserve and set aside.
   6. Rinse slides well in several changes of distilled water.
   7. Check positive control slide microscopically to determine adequate copper/reddish brown development.
      1. 1. Return slides to reserved Working Rhodanine Solution if additional incubation is required.
   8. Prepare dilute Mayer Hematoxylin Stain directly before use; combine and mix well:
      1. 1. Hematoxylin Stain, Mayer Modified (Part 1202)    20 ml
         2. Distilled Water               20 ml
   9. Stain in dilute Mayer Hematoxylin Stain Solution for 10 minutes.
   10. Rinse in distilled water.
   11. Rinse in Sodium Borate 0.5%, Aqueous (Part 13824); 2-3 quick dips.
   12. Rinse well in distilled water.
   13. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Copper	Copper/reddish brown
Nuclei	Light blue

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The microwave procedure was tested using a laboratory-grade microwave oven. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 251.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 258-260
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 230.
   4. Modifications developed by Newcomer Supply Laboratory.

Ship Ground Only

 

Solution A of the Differential Stain Kit.

• Shelf Life is 2 years from date of manufacture.

 

For Staining Procedure, click here

Click here to view the Differential Stain Kit.