(use: Working solution for Gomori Basement Membrane Stain.)

(use: Sulfated Alcian Blue procedure for Amyloid stain.)

SOLUTION:

	125 ml	500 ml
Safranin O Stain 1%, Aqueous	Part 1350A	Part 1350B

 

Additionally Needed:

Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Hematoxylin Stain Set, Weigert Iron	Part 1409
Fast Green Stain 2.5%, Aqueous	Part 10852
Acetic Acid 1%, Aqueous	Part 10012

 

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

 

APPLICATION:

Newcomer Supply Safranin O Stain for Cartilage procedure is to stain cartilage, mucin, and mast cell granules in tissue sections and assist in demonstrating articular cartilage loss in arthritic and other articular disease processes.

Safranin O is a basic dye that stains growth plate cartilage and articular cartilage (proteoglycans, chondrocytes and type II collagen) varying shades of red. The intensity of Safranin O staining is proportional to the proteoglycan content in the cartilage tissue. Fast Green counterstains the non-collagen sites and provides clear contrast to Safranin O staining.

If tissue is decalcified, decalcifying agents such as EDTA, nitric acid and hydrochloric acid may extract proteoglycans and result in weak Safranin O cartilage staining.

 

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.

 

STAINING PROCEDURE:

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. Wash well in running tap water; rinse in distilled water.
   3. Prepare fresh Weigert Iron Hematoxylin (Part 1409); mix well.
      1. 1. Solution A: Ferric Chloride, Acidfied 20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic 20 ml
   4. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   5. Wash in running tap water for 10 minutes; rinse in distilled water.
   6. Prepare 0.25% Fast Green Stain, Aqueous; combine and mix well.
      1. 1. Fast Green Stain 2.5%, Aqueous (Part 10852) 5 ml
         2. Distilled Water 45 ml
   7. Stain in 0.25% Fast Green Stain, Aqueous for 5 minutes.
   8. Rinse directly in Acetic Acid 1%, Aqueous (Part 10012); 10-15 seconds.
   9. Place directly in Safranin O Stain 1%, Aqueous for 5 minutes.
   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:

Cartilage	Red to orange
Mucin and mast cell granules	Red to orange
Bone, connective tissue and cytoplasm	Green
Nuclei	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. 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. 346-347.
   2. Callis, Gayle and Diane Sterchi. “Decalcification of Bone: Literature Review and Practical Study of Various Decalcifying Agents, Methods, and Their Effects on Bone Histology.” The Journal of Histotechnology 21.1 (1998): 49-58.
   3. Chevrier, Anik, Evgeny Rossomacha, Michael D. Buschmann and Caroline D. Hoemann. “Optimization of Histoprocessing Methods to Detect Glycosaminoglycan, Collagen Type II, and Collagen Type I in Decalcified Rabbit Osteochondral Sections.” The Journal of Histotechnology 28.3 (2005): 165-175.
   4. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 256.
   5. Modifications developed by Newcomer Supply Laboratory.

(use: Stock solution for Gomori Basement Membrane & GMS; Working solution for VVG Elastic, Gordon & Sweets and Mercury pigment removal.)

SOLUTION:

SOLUTION:

	1 Liter
Zamboni Fixative	Part 1459A

 

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

 

APPLICATION:

Newcomer Supply Zamboni Fixative is a ready-to-use phosphate buffered picric acid-formaldehyde (PAF) fixative with applications for light and electron microscopy.  Zamboni fixative provides general fixation with rapid penetration, optimal preservation and stabilization of cellular proteins.

 

METHOD:

Fixation:

• • Small Biopsies: Minimum of 1 hour
  • Larger Biopsies: Minimum of 4 hours

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

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue in Zamboni Fixative after surgical excision.
      1. 1. See Procedure Note #1.
   2. Hold tissue specimens in Zamboni Fixative until ready to process.
      1. 1. See Procedure Note #2.
   3. Rinse Zamboni fixed tissue thoroughly in running tap water followed by Phosphate Buffered Saline 0.1M, pH 7.4 (Part 133104) for a minimum of 15 minutes prior to processing.
   4. Processing:
      1. 1. Light microscopy: place on tissue processor starting in either Formalin 10%, Phosphate Buffered (Part 1090) fixation step or first dehydration station.
         2. Electron microscopy: a secondary osmium tetroxide fixation is recommended. Refer to protocol for electron microscopy processing.

 

PROCEDURE NOTES:

1. 1. For electron microscopy studies, fix tissues within 15 minutes after excision. Mince into 1mm cubes for expedient fixative infiltration.
   2. Tissue can be held indefinitely in Zamboni Fixative at room temperature without compromising preservation.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 21, 334, 336.
   2. Dapson, Janet Crookham and Richard Dapson. Hazardous Materials in the Histopathology Laboratory: Regulations, Risks, Handling, and Disposal. 4th ed. Battle Creek, MI: Anatech, 2005. 150, 265-266.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 48, 328-330.
   4. Zamboni, Luciano and Cesare De Martino. “Buffered Picric Acid Formaldehyde: A New Rapid Fixative for Electron Microscopy”. Journal of Cell Biology (1967) 35: 148.
   5. Modifications developed by Newcomer Supply Laboratory.

 

(use: Fite Stain for Leprosy & Nocardia.)

 

• Acetone, Alcohols, and Xylene do not have expiration dates.
• Shelf life is 2 yrs once opened.

 

• Acetone, Alcohols, and Xylene do not have expiration dates.
• Shelf life is 3 years from date of manufacture.

CI 50240

• Shelf Life is 4 years from date of manufacture.

 

SOLUTION:

	100 ml
Poly-L-Lysine Adhesive Stock	Part 1339A

 

Additionally Needed:

Non-Adhesive Slides	Part 6215 (Frosted End)	Part 6206 (Colored End)	Part 6210 (Plain)
EasyDip™ Slide Staining Jar	Part 5300
EasyDip™ Slide Staining Rack                    OR EasyDip™ Slide Staining Kit	Part 5300RK        OR Part 5300KIT

 

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

 

APPLICATION:

Newcomer Supply Poly-L-Lysine Adhesive Stock when diluted to a working solution, provides a strong adhesive coating to non-adhesive microscopic slides.  This applied coating enhances bonding of tissue sections to microscopic slides for use in histological, microwave and immunohistochemistry (IHC) staining procedures, leaving minimal or no background staining.

One liter of Poly-L-Lysine Working Solution will coat approximately 900 slides. Exceeding 900 slides per liter of working solution may affect product performance.

 

METHOD:

Technique:  Paraffin or frozen sections
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. 1. Fill slide rack with clean and dry non-adhesive slides.
   2. Dilute Poly-L-Lysine Adhesive Stock to a working solution; combine and mix well.
      1. 1. Poly-L-Lysine Adhesive Stock 10 ml
         2. Distilled Water                 90 ml
         3. See Procedure Note #1.
   3. Pour Poly-L-Lysine Working Solution into a plastic staining dish, using sufficient solution to cover racked slides. Keep solution covered to avoid evaporation and contamination.
      1. 1. EasyDip™ Slide Staining Jars (Part 5300) and Racks (Part 5300RK) and EasyDip™ Slide Staining Kit (Part 5300KIT) each hold 80 ml of solution with a 12-slide capacity.
   4. Soak in Poly-L-Lysine Adhesive Working Solution for 5 minutes.
      1. 1. Increased soaking time does not improve performance.
         2. See Procedure Notes #2 and #3.
   5. Drain slides. Blot and tap excess solution from slides/rack.
   6. Dry racked slides in a 60°C oven for 1 hour or overnight at room temperature in a dust-free environment.
   7. Store dried treated slides in a clean slide box at room temperature and low humidity.
      1. 1. Slides will adhere together if not thoroughly dried before storing.
   8. Wash emptied slide racks, plasticware and glassware after use to ensure all adhesive is removed.

 

PROCEDURE NOTES:

1. 1. Poly-L-Lysine Adhesive Stock and working solutions will react and leave deposits on glassware. The use of plastic containers and graduated cylinders when mixing, storing solutions and coating slides is recommended.
   2. Store used Poly-L-Lysine Working Solution at 2°- 8°C in a plastic bottle for up to three months. Discard solution if turbidity develops.
   3. Filter diluted Poly-L-Lysine Working Solution between uses.
   4. Do not add to or mix fresh solution with used diluted solution.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 70.
   2. Huang, W.M, S.J. Gibson, P. Facer, J. Gu and J.M. Polak. “Improved Section Adhesion for Immunocytochemistry Using High Molecular Weight Polymers of L-Lysine as a Slide Coating.” Histochemistry 77.2 (1983): 275-279.
   3. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	500 ml	1 Liter	1 Gallon
Wright Stain	Part 1420A	Part 1420B	Part 1420C

 

Additionally Needed:

Alcohol, Methanol Anhydrous, ACS	Part 12236
Wright Stain Buffer, pH 6.8	Part 1430

 

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

 

APPLICATION:

Newcomer Supply Wright Stain for Smears is an unbuffered Wright staining solution used for differential staining of cell types in peripheral blood smears as well as bone marrow smears/films.

 

METHOD:

Technique: Flat staining rack method
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

 

PRESTAINING PREPARATION:

1. 1. Prepare a well-made blood smear or bone marrow with a focus on uniform cell distribution.
   2. Allow slides to thoroughly air-dry prior to staining.
   3. Filter Wright Stain Solution prior to use with quality filter paper.
      1. 1. Filter sufficient stain to allow 1 ml of stain per slide.

 

STAINING PROCEDURE:

1. 4. Place slides on flat staining rack suspended over sink.
   5. Fix by flooding slides with Methanol (Part 12236) for 10-30 seconds.
   6. Drain off Methanol.
   7. Flood each slide with 1 ml of filtered Wright Stain for 3-5 minutes.
      1. 1. See Procedure Notes #1 and #2.
   8. Retain Wright Stain on slides.
   9. Directly add 1 ml of Wright Stain Buffer, pH 6.8 (Part 1430) to each slide; agitate gently to mix with Wright Stain.
   10. Stain for an additional 6-10 minutes.
   11. Wash well in distilled water; rinse thoroughly in running tap water.
   12. Air-dry slides in a vertical position; examine microscopically.
   13. If coverslip is preferred, air-dry slides and coverslip with compatible mounting medium.

 

RESULTS:

Erythrocytes	Pink
Neutrophils	Granules – Purple
Eosinophils	Granules – Pink
White blood cells	Chromatin – Purple
Lymphocytes	Cytoplasm – Blue
Monocytes	Cytoplasm – Blue
Bacteria	Deep Blue

 

PROCEDURE NOTES:

1. 1. Timings are suggested ranges. Optimal staining times will depend upon staining intensity preference.
   2. Smears containing primarily normal cell populations require minimum staining time; immature cells and bone marrow smears/films may require longer staining time.
   3. The color range of stained cells may vary depending on buffer pH and pH of rinse water.
      1. 1. Alkalinity is indicated by red blood cells being blue-grey and white blood cells only blue.
         2. Acidity is indicated by red blood cells being bright red or pink and lack of proper staining in white blood cells.
         3. If necessary adjust buffer pH accordingly to 6.8 +/- 0.2.

 

REFERENCES:

1. 1. Lillie, R. D. and Harold Fullmer. Histopathologic Technic and Practical Histochemistry. 4th ed. New York: McGraw-Hill, 1976. 747-748.
   2. McPherson, Richard and Matthew Pincus. Henry’s Clinical Diagnosis and Management by Laboratory Methods. 22nd ed. Philadelphia: Elsevier Saunders, 2011. 522-532.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 154-155.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Wolbach Giemsa.)

(use: Gordon & Sweets Reticulum Stain & stock solution for melanin bleaching.)

(use: PTAH Stain & working solution for melanin bleaching.)

(use: Gomori Mod. Iron Stain & stock for Colloidal Iron.)

(use: Oil Red O in Propylene Glycol, Fat Stain.)

SOLUTION:

	250 ml	500 ml
Trichrome Stain, Gomori One-Step, Aniline Blue	Part 1403C	Part 1403B

 

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
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 Trichrome Stain, Gomori One-Step, Aniline Blue 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 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. 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.

1. 9. Stain in Trichrome Stain, Gomori One-Step, Aniline Blue for 20 minutes.
   10. Differentiate in Acetic Acid 0.5%, Aqueous (Part 100121); 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: Oil Red O in Propylene Glycol, Fat Stain.)

(use: Steiner Silver Impregnation Methods.)

(use: Schmorl Melanin Stain.)

(use: IHC wash solution; dilute 1:10 before use.) Store at 2-8°C.)

 

Tech Memo 2: Trichrome Stain, Gomori One-Step, Light Green for Frozen Muscle Biopsies

 

SOLUTION:                                                                                                                                               

(use: Alternative to Bouin as mordant for the liver Trichrome, and as differentiator for Sulfated Alcian Blue for Myocardial Amyloid.)

 

 

(use: Stock sol’n for muscle biopsies.)

(use: Instead of keeping the hazardous powder to make-up Bouin, etc.)

 

 

SOLUTION:

	250 ml	500 ml
Trichrome Stain, Gomori One-Step, Fast Green	Part 14021A	Part 14021B

 

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
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 Trichrome Stain, Gomori One-Step, Fast Green 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 procedures are designed to be used with Coplin jars filled to 40 ml following the 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 1 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. Stain in Trichrome Stain, Gomori One-Step, Fast Green for 20 minutes.
   10. Differentiate in Acetic Acid 0.5%, Aqueous (Part 100121) for 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: Counterstain for Mucicarmine.)

(use: Gram Stains, eg. Brown Brenn.)

 

(use: Brown & Hopps.)

 

 

(use: Trichrome, Masson Light Green.)

(use: Trichrome, McLetchie.)

(use: With Gomori One-Step Trichrome for Decolorization if needed.)

(use: Masson Trichrome Stain.)

(use: Klatskin Trichrome for liver.)

(use: With Giemsa Stain.)

(use: IHC wash and rinse.)

(use: IHC diluent, rinse and wash.)

(use: Buffer for Glycogen Digestion; Does not include enzyme.)

 

Molded from acetal, the patented Micromesh Biopsy Tissue Processing & Embedding Cassettes keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents. The Micromesh Biopsy Tissue Processing & Embedding cassettes ensure efficient fluid exchange and drainage.

 

PACKAGING OF THE MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 250 cassettes/box; 1,000 cassettes/case with covers assembled.

 

MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 1676 square openings (0.38mm) allowing for a greatly improved fluid exchange without having to use biopsy pads.
• Large anterior and posterior slots in both cassette and cover ensure that the cassette will sink rapidly.
• One large square compartment measuring 27mm is perfect even for needle biopsies.
• Recessed cover allowing more cassettes to be stacked in automatic labeling machines and tissue processors.
• The one-piece integral lid eliminates the need for separate steel lids.
• Anterior writing area is at a 45° angle.
• Cassettes also available in QuickLoad™ Sleeves (Part 5123).

(use: Rinsing unfixed tissues and diluting antibodies, eg. in IHC.)

(use: IHC diluent, rinse and wash.)

 

Made of acetal, the Microsette Biopsy Tissue Processing & Embedding Cassettes can hold up to six tissue specimens, each one placed in it’s own 7×12 mm compartment, numbered from 1 to 6.  The Micromesh biopsy tissue processing & embedding cassettes keep specimens safely submerged in liquid and are resistant to most histological solvents.

 

PACKAGING OF THE MICROSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES (6 Compartments):

• 250 cassettes/box; 1,000 cassettes/case with covers assembled.

 

MICROSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES (6 Compartments):

• Cover and base have over 2,000 (0.26mm) square openings to maximize fluid exchange and ensure proper drainage.
• No biopsy pads are necessary.
• Six separate 7x12mm compartments, numbered from 1 to 6.
• Approximately 170 holes (each having a diameter of 0.26 mm) per compartment.
• Anterior writing area is at a 45° angle.
• Lid is pre-attached to base

(use: PAS for glycogen or amoeba and in Gomori Basement Membrane Stain.)

 

These Slimsette tissue processing & embedding cassettes are suitable for hoppers accepting plastic sleeves such as Thermo Fisher printers.  They load in cassette labeling instruments in one simple operation!  The transparent sleeve allows viewing of the Slimsette tissue processing & embedding cassettes in order to confirm there are no tissue cassette jams in the sleeve during the printing process. Molded from acetal, these cassettes keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents.

PACKAGING OF THE SLIMSETTE TISSUE PROCESSING & EMBEDDING CASSETTES IN QUICKLOAD SLEEVES:

• 75 cassettes/sleeve; 10 sleeves/case; 750 cassettes/case.

 

SLIMSETTE TISSUE PROCESSING & EMBEDDING CASSETTES IN QUICKLOAD SLEEVES:

• Convenient plastic dispensing sleeve compatible with Thermo Fisher printers.
• 114 openings each measuring 1 x 5 mm allowing for efficient fluid exchange and drainage.
• A unique recessed cover which is a great space saving feature allowing more cassettes to be stacked in automatic labeling machines and in storage cabinets.
• One-piece integral lid eliminates the need for separate steel lids.
• Lids can be opened and closed as often as necessary and they always relock securely without danger of specimen loss.
• Anterior writing area is at a 45° angle.
• Cassettes also available without QuickLoad sleeves (Part 5126).

(use: Can be diluted to 0.5% for PAS or PAMM.)

(use: Cell flow fixation.)

SOLUTIONS:

	500 ml	1 Gallon
Papanicolaou Stain, OG-6	Part 1330A	Part 1330B
Papanicolaou Stain, EA-50, Fast Green	Part 1310A	Part 1310B
Papanicolaou Stain, EA-65, Fast Green	Part 1320A	Part 1320B
Papanicolaou Stain, EA-50, Light Green	Part 1312A	Part 1312B
Hematoxylin Stain, Gill I (single strength)	Part 1180A	Part 1180C
Hematoxylin Stain, Gill II (double strength)	Part 1180D	Part 1180F
Hematoxylin Stain, Gill III (triple strength)	Part 1180G	Part 1180I

 

Additionally Needed:

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

 

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

 

APPLICATION:

Newcomer Supply Papanicolaou (Pap) Stain provides classic staining solutions for cytology preparations, allowing for crisp, distinct nuclear detail and differentially stained cytoplasm. Gill Hematoxylin is the optimal nuclear stain and the two counterstains, Orange Gelb (OG) and Eosin Azure (EA) provide the subtle range of green, blue, and pink hues to the cellular cytoplasm.

Papanicolaou Stain, EA, is comprised of a combination of two dyes: Eosin Y and Fast Green SF or Light Green.  EA-50 and EA-65 denote the varying proportions of dyes in each solution. The stains are suitable for both Gyn (EA-50) and Non-Gyn (EA-50, EA-65) specimens.  Formula of choice will depend upon staining preference.

 

METHOD:

Technique/Fixation:  Preparation and choice of fixative is dependent on specimen types.
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. Fix in 95% Ethyl Alcohol for 5-10 minutes.
      1. 1. Fixation times may vary depending on specimen type.
   2. Rinse in running distilled water for 1 minute.
   3. Stain in Hematoxylin Stain, Gill I, Gill II or Gill III for 1 to 6 minutes, depending on specimen and preference of nuclear stain intensity.
   4. Wash in distilled water until clear.
   5. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 30 seconds.
   6. Wash in running distilled water for 30 seconds.
   7. Dehydrate in 70% Ethyl Alcohol: 10 dips.
   8. Dehydrate in 95% Ethyl Alcohol: 10 dips.
   9. Stain in Papanicolaou Stain, OG-6 for 1-2 minutes, depending on specimen type and preferred stain intensity.
   10. Rinse in two changes of 95% Ethyl Alcohol; 10 dips each.
   11. Stain in Papanicolaou Stain, EA-50 or EA-65 for 3-5 minutes, depending on specimen type and preferred stain intensity.
   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:

Chromatin	Blue
Keratin	Orange
Squamous cells	Shades of pink
RBCs, nucleoli, cilia	Shades of pink
Cytoplasm	Shades of blue-green

 

PROCEDURE NOTES:

1. 1. Pap stains can be implemented for either manual or automated staining. Timings may vary depending on staining platform used.
   2. Solutions should be filtered or replaced daily to prevent cross-contamination and maintain optimal staining.
   3. If using a xylene substitute, follow manufacturer’s recommendation for the clearing step.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 127-128.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 326-327.
   3. Koss, Leopold G. Diagnostic Cytology and Its Histopathologic Bases. 3rd ed. Philadelphia: Lippincott, 1979. 1218.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTIONS:

	500 ml	1 Gallon
Papanicolaou Stain, OG-6	Part 1330A	Part 1330B
Papanicolaou Stain, EA-50, Fast Green	Part 1310A	Part 1310B
Papanicolaou Stain, EA-65, Fast Green	Part 1320A	Part 1320B
Papanicolaou Stain, EA-50, Light Green	Part 1312A	Part 1312B
Hematoxylin Stain, Gill I (single strength)	Part 1180A	Part 1180C
Hematoxylin Stain, Gill II (double strength)	Part 1180D	Part 1180F
Hematoxylin Stain, Gill III (triple strength)	Part 1180G	Part 1180I

 

Additionally Needed:

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

 

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

 

APPLICATION:

Newcomer Supply Papanicolaou (Pap) Stain provides classic staining solutions for cytology preparations, allowing for crisp, distinct nuclear detail and differentially stained cytoplasm. Gill Hematoxylin is the optimal nuclear stain and the two counterstains, Orange Gelb (OG) and Eosin Azure (EA) provide the subtle range of green, blue, and pink hues to the cellular cytoplasm.

Papanicolaou Stain, EA, is comprised of a combination of two dyes: Eosin Y and Fast Green SF or Light Green.  EA-50 and EA-65 denote the varying proportions of dyes in each solution. The stains are suitable for both Gyn (EA-50) and Non-Gyn (EA-50, EA-65) specimens.  Formula of choice will depend upon staining preference.

 

METHOD:

Technique/Fixation:  Preparation and choice of fixative is dependent on specimen types.
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. Fix in 95% Ethyl Alcohol for 5-10 minutes.
      1. 1. Fixation times may vary depending on specimen type.
   2. Rinse in running distilled water for 1 minute.
   3. Stain in Hematoxylin Stain, Gill I, Gill II or Gill III for 1 to 6 minutes, depending on specimen and preference of nuclear stain intensity.
   4. Wash in distilled water until clear.
   5. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 30 seconds.
   6. Wash in running distilled water for 30 seconds.
   7. Dehydrate in 70% Ethyl Alcohol: 10 dips.
   8. Dehydrate in 95% Ethyl Alcohol: 10 dips.
   9. Stain in Papanicolaou Stain, OG-6 for 1-2 minutes, depending on specimen type and preferred stain intensity.
   10. Rinse in two changes of 95% Ethyl Alcohol; 10 dips each.
   11. Stain in Papanicolaou Stain, EA-50 or EA-65 for 3-5 minutes, depending on specimen type and preferred stain intensity.
   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:

Chromatin	Blue
Keratin	Orange
Squamous cells	Shades of pink
RBCs, nucleoli, cilia	Shades of pink
Cytoplasm	Shades of blue-green

 

PROCEDURE NOTES:

1. 1. Pap stains can be implemented for either manual or automated staining. Timings may vary depending on staining platform used.
   2. Solutions should be filtered or replaced daily to prevent cross-contamination and maintain optimal staining.
   3. If using a xylene substitute, follow manufacturer’s recommendation for the clearing step.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 127-128.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 326-327.
   3. Koss, Leopold G. Diagnostic Cytology and Its Histopathologic Bases. 3rd ed. Philadelphia: Lippincott, 1979. 1218.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTIONS:

	500 ml	1 Gallon
Papanicolaou Stain, OG-6	Part 1330A	Part 1330B
Papanicolaou Stain, EA-50, Fast Green	Part 1310A	Part 1310B
Papanicolaou Stain, EA-65, Fast Green	Part 1320A	Part 1320B
Papanicolaou Stain, EA-50, Light Green	Part 1312A	Part 1312B
Hematoxylin Stain, Gill I (single strength)	Part 1180A	Part 1180C
Hematoxylin Stain, Gill II (double strength)	Part 1180D	Part 1180F
Hematoxylin Stain, Gill III (triple strength)	Part 1180G	Part 1180I

 

Additionally Needed:

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

 

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

 

APPLICATION:

Newcomer Supply Papanicolaou (Pap) Stain provides classic staining solutions for cytology preparations, allowing for crisp, distinct nuclear detail and differentially stained cytoplasm. Gill Hematoxylin is the optimal nuclear stain and the two counterstains, Orange Gelb (OG) and Eosin Azure (EA) provide the subtle range of green, blue, and pink hues to the cellular cytoplasm.

Papanicolaou Stain, EA, is comprised of a combination of two dyes: Eosin Y and Fast Green SF or Light Green.  EA-50 and EA-65 denote the varying proportions of dyes in each solution. The stains are suitable for both Gyn (EA-50) and Non-Gyn (EA-50, EA-65) specimens.  Formula of choice will depend upon staining preference.

 

METHOD:

Technique/Fixation:  Preparation and choice of fixative is dependent on specimen types.
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. Fix in 95% Ethyl Alcohol for 5-10 minutes.
      1. 1. Fixation times may vary depending on specimen type.
   2. Rinse in running distilled water for 1 minute.
   3. Stain in Hematoxylin Stain, Gill I, Gill II or Gill III for 1 to 6 minutes, depending on specimen and preference of nuclear stain intensity.
   4. Wash in distilled water until clear.
   5. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 30 seconds.
   6. Wash in running distilled water for 30 seconds.
   7. Dehydrate in 70% Ethyl Alcohol: 10 dips.
   8. Dehydrate in 95% Ethyl Alcohol: 10 dips.
   9. Stain in Papanicolaou Stain, OG-6 for 1-2 minutes, depending on specimen type and preferred stain intensity.
   10. Rinse in two changes of 95% Ethyl Alcohol; 10 dips each.
   11. Stain in Papanicolaou Stain, EA-50 or EA-65 for 3-5 minutes, depending on specimen type and preferred stain intensity.
   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:

Chromatin	Blue
Keratin	Orange
Squamous cells	Shades of pink
RBCs, nucleoli, cilia	Shades of pink
Cytoplasm	Shades of blue-green

 

PROCEDURE NOTES:

1. 1. Pap stains can be implemented for either manual or automated staining. Timings may vary depending on staining platform used.
   2. Solutions should be filtered or replaced daily to prevent cross-contamination and maintain optimal staining.
   3. If using a xylene substitute, follow manufacturer’s recommendation for the clearing step.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 127-128.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 326-327.
   3. Koss, Leopold G. Diagnostic Cytology and Its Histopathologic Bases. 3rd ed. Philadelphia: Lippincott, 1979. 1218.
   4. Modifications developed by Newcomer Supply Laboratory.

 

The Slimsette Biopsy Tissue Processing & Embedding Cassettes are more compact, easier to use and more efficient than ever. Made of acetal, the Slimsette biopsy processing & tissue embedding cassettes keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents.

 

PACKAGING OF THE SLIMSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 500 cassettes/box, 1,500 cassettes/case with covers assembled.

 

SLIMSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• Designed for biopsy specimens.
• Efficient fluid exchange and drainage with 392 openings each measuring 1 x 1 mm.
• A unique recessed cover, a great space saving feature, allowing more cassettes to be stacked in automatic labeling machines and in storage cabinets.
• One-piece integral lid eliminates the need for separate steel lids.
• Lids can be opened and closed as often as necessary and relock securely without danger of specimen loss.
• Large labeling areas for easy identification.
• The anterior writing area is slanted at a 45° angle.
• Also available in QuickLoad™ Sleeves (Part 5122).

 

DIMENSIONS OF THE SLIMSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 1 5/8″ x 1 1/8″ x 1/4″ H

SOLUTIONS:

	500 ml	1 Gallon
Papanicolaou Stain, OG-6	Part 1330A	Part 1330B
Papanicolaou Stain, EA-50, Fast Green	Part 1310A	Part 1310B
Papanicolaou Stain, EA-65, Fast Green	Part 1320A	Part 1320B
Papanicolaou Stain, EA-50, Light Green	Part 1312A	Part 1312B
Hematoxylin Stain, Gill I (single strength)	Part 1180A	Part 1180C
Hematoxylin Stain, Gill II (double strength)	Part 1180D	Part 1180F
Hematoxylin Stain, Gill III (triple strength)	Part 1180G	Part 1180I

 

Additionally Needed:

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

 

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

 

APPLICATION:

Newcomer Supply Papanicolaou (Pap) Stain provides classic staining solutions for cytology preparations, allowing for crisp, distinct nuclear detail and differentially stained cytoplasm. Gill Hematoxylin is the optimal nuclear stain and the two counterstains, Orange Gelb (OG) and Eosin Azure (EA) provide the subtle range of green, blue, and pink hues to the cellular cytoplasm.

Papanicolaou Stain, EA, is comprised of a combination of two dyes: Eosin Y and Fast Green SF or Light Green.  EA-50 and EA-65 denote the varying proportions of dyes in each solution. The stains are suitable for both Gyn (EA-50) and Non-Gyn (EA-50, EA-65) specimens.  Formula of choice will depend upon staining preference.

 

METHOD:

Technique/Fixation:  Preparation and choice of fixative is dependent on specimen types.
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. Fix in 95% Ethyl Alcohol for 5-10 minutes.
      1. 1. Fixation times may vary depending on specimen type.
   2. Rinse in running distilled water for 1 minute.
   3. Stain in Hematoxylin Stain, Gill I, Gill II or Gill III for 1 to 6 minutes, depending on specimen and preference of nuclear stain intensity.
   4. Wash in distilled water until clear.
   5. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 30 seconds.
   6. Wash in running distilled water for 30 seconds.
   7. Dehydrate in 70% Ethyl Alcohol: 10 dips.
   8. Dehydrate in 95% Ethyl Alcohol: 10 dips.
   9. Stain in Papanicolaou Stain, OG-6 for 1-2 minutes, depending on specimen type and preferred stain intensity.
   10. Rinse in two changes of 95% Ethyl Alcohol; 10 dips each.
   11. Stain in Papanicolaou Stain, EA-50 or EA-65 for 3-5 minutes, depending on specimen type and preferred stain intensity.
   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:

Chromatin	Blue
Keratin	Orange
Squamous cells	Shades of pink
RBCs, nucleoli, cilia	Shades of pink
Cytoplasm	Shades of blue-green

 

PROCEDURE NOTES:

1. 1. Pap stains can be implemented for either manual or automated staining. Timings may vary depending on staining platform used.
   2. Solutions should be filtered or replaced daily to prevent cross-contamination and maintain optimal staining.
   3. If using a xylene substitute, follow manufacturer’s recommendation for the clearing step.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 127-128.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 326-327.
   3. Koss, Leopold G. Diagnostic Cytology and Its Histopathologic Bases. 3rd ed. Philadelphia: Lippincott, 1979. 1218.
   4. Modifications developed by Newcomer Supply Laboratory.

 

The Slimsette Tissue Processing & Embedding Cassettes are more compact, easier to use and more efficient than ever. Made of acetal, the Slimsette tissue processing & embedding cassettes keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents.

 

PACKAGING OF THE SLIMSETTE TISSUE PROCESSING & EMBEDDING CASSETTES:

• 500 cassettes/box, 1,500 cassettes/case with covers assembled.

 

SLIMSETTE TISSUE PROCESSING & EMBEDDING CASSETTES:

• Efficient fluid exchange and drainage with 114 openings each measuring 1 x 5 mm.
• A unique recessed cover which is a great space saving feature allowing more cassettes to be stacked in automatic labeling machines and in storage cabinets.
• One-piece integral lid eliminates the need for separate steel lids.
• Lids can be opened and closed as often as necessary and relock securely without danger of specimen loss.
• Large labeling areas for easy identification.
• The anterior writing area is slanted at a 45° angle.
• Also available in QuickLoad™ Sleeves (Part 5121).

 

DIMENSIONS OF THE SLIMSETTE TISSUE PROCESSING & EMBEDDING CASSETTES:

• 1 5/8″ x 1 1/8″ x 1/4″ H

 

These Micromesh tissue processing & embedding biopsy cassettes are suitable for tissue cassette labelers with hoppers that accept plastic sleeves.  The cassettes will load in a cassette labeling instrument in one simple operation!  The transparent sleeve allows viewing of the Micromesh tissue processing & embedding biopsy cassettes in order to confirm there are no tissue cassette jams in the sleeve during the printing process.

 

PACKAGING OF THE MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 75 biopsy cassettes/sleeve; 10 sleeves/case; 750 cassettes/case.

 

MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• Convenient plastic dispensing sleeve compatible with Thermo Fisher printers.
• 1,676 (0.38mm) square openings, and large anterior slots allowing for increased fluid exchange and faster sinking in liquids.
• Large square 27mm compartment, perfect for needle biopsies.
• No biopsy pads are necessary.
• A unique recessed cover which is a great space saving feature allowing more cassettes to be stacked in automatic labeling machines and in storage cabinets.
• Lids are pre-mounted on cassettes and can be open and closed as often as necessary and always relock securely without danger of specimen loss.
• Anterior writing area is at a 45° angle.
• Cassettes also available without sleeves (Part 5120).

 

These Slimsette tissue processing and embedding, biopsy cassettes are suitable for hoppers that accept plastic sleeves such as Thermo Fisher printers.  They load in cassette labeling instruments in one simple operation!  The transparent sleeve allows viewing of Slimsette tissue processing & embedding, biopsy cassettes in order to confirm there are no tissue cassette jams in the sleeve during the printing process. Molded from acetal, these cassettes keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents.

 

PACKAGING OF THE SLIMSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES IN QUICKLOAD SLEEVES:

• 75 biopsy cassettes/sleeve; 10 sleeves/case; 750 cassettes/case.

 

SLIMBSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• • Convenient plastic dispensing sleeve compatible with Thermo Fisher printers.
  • Cassettes designed for biopsy specimens.
  • 392 openings each measuring 1 x 1 mm allowing for efficient fluid exchange and drainage.
  • A unique recessed cover which is a great space saving feature allowing more cassettes to be stacked in automatic labeling machines and in storage cabinets.
  • One-piece integral lid eliminates the need for separate steel lids.
  • Lids can be opened and closed as often as necessary and they always relock securely without danger of specimen loss.
  • Anterior writing area is at a 45° angle.
  • Cassettes also available without QuickLoad sleeves (Part 5127).

(use: Bleach out melanin or phosphotungstic acid in the PTAH Stain & is stock for Gordon & Sweets Reticulum.)

 

The Swingsette Tissue Processing & Embedding Cassettes hold tissue specimens during processing and embedding and can be used for storage.  Molded from acetal, they keep specimens safely submerged in liquid and are resistant to the chemical action of most histological solvents.  The efficient flow-through slots maximize fluid exchange and ensure proper drainage.

 

PACKAGING OF THE  SWINGSETTE TISSUE PROCESSING & EMBEDDING CASSETTES:

• 500 cassettes/box, 1,500 cassettes/case

 

SWINGSETTE TISSUE PROCESSING & EMBEDDING CASSETTES:

• Made of acetal.
• Efficient flow-through slots maximize fluid exchange and ensure proper drainage.
• Special hinge allows cassettes to be opened and closed as often as necessary.
• Large tab for convenient and easy opening of lid.
• The anterior writing area is at a 45º angle.
• Cassettes also available in QuickLoad™ Sleeves (Part 5129).

(use: Mod. of sperm vitality stain.)

 

The Swingsette Biopsy Tissue Processing & Embedding Cassettes are designed to hold biopsy specimens during processing and embedding and can be used for storage.

 

PACKAGING OF THE SWINGSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• 500 cassettes/box; 1,500 cassettes/case.

 

SWINGSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES:

• Designed to hold biopsy specimens.
• Made of acetal.
• 1mm square openings to maximize fluid exchange.
• Anterior writing area is at a 45° angle.
• Large tab for convenient and easy opening of lid.
• Easy to remove pre attached cover with special hinge.
• Cassettes also available in QuickLoad™ Sleeves (Part 51291).

 

• Shelf Life is 2 years from date of manufacture.

 

SOLUTION:  

	250 ml	500 ml	1 Liter
Nuclear Fast Red Stain, Kernechtrot	Part 1255A	Part 1255C	Part 1255B

 

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 Nuclear Fast Red Stain, Kernechtrot is a versatile stain and counterstain that combines nuclear fast red dye with an aluminum sulfate mordant to selectively stain nuclear chromatin red and provide nonspecific background tissue staining in shades of pink.

Nuclear Fast Red (NFR) is also known by its Germanic origin name of Kernechtrot. Kernechtrot and Nuclear Fast Red are interchangeable terms for the dye and solution.

Nuclear Fast Red Stain, Kernechtrot is used in a variety of staining procedures that include:

• • Alcian Blue 1%, pH 2.5 Stain
  • Alcian Blue 1%, pH 1.0 Stain
  • Fontana Masson Stain
  • Iron, Gomori Prussian Blue Stain
  • Reticulum, Gordon & Sweets Stain
  • Schmorl Melanin Stain
  • Sudan Black B Stain
  • Victoria Blue Stain
  • Von Kossa Calcium 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.

 

STAINING PROCEDURE:

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. Proceed with staining procedure of choice.
   3. Counterstain in Nuclear Fast Red Stain, Kernechtrot for 5 minutes.
      1. 1. Shake solution well before use; do not filter.
         2. See Procedure Note #3.
   4. Rinse well in distilled water.
      1. 1. See Procedure Note #4.
   5. 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:

Nuclei	Pink-red
Cytoplasm	Pale pink
Other tissue components	Dependent on stain procedure used

 

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. Precipitate will normally settle out of Nuclear Fast Red Stain, Kernechtrot, and can be redistributed by shaking the solution well before each use. Do not filter precipitate out.
   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. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 145-146.
   2. Kiernan, John. Histological & Histochemical Methods. 3rd ed. New York: Oxford University Press, 2003.96,114.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980.183.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	125 ml	500 ml	1 Liter
Mucicarmine Stock Stain, Mayer	Part 1250A	Part 1250B	Part 1250C

 

Additionally Needed For Mucin, Mayer Mucicarmine Stain:

Mucin Mucicarmine Control Slides                   OR Cryptococcus Control Slides	Part 4455     OR Part 4135
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Hematoxylin Stain Set, Weigert Iron	Part 1409
Metanil Yellow Stain, Aqueous                  OR Tartrazine Stain 0.25%, Acetic Aqueous	Part 12235     OR Part 14016
Coplin Jar, Plastic	Part 5184 (for microwave modification)

                                     

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

 

APPLICATION:

Newcomer Supply Mucicarmine Stock Stain, Mayer a component of the Mucin, Mayer Mucicarmine Stain procedure, is used to stain acid epithelial mucin (sialomucin, sulfomucin) and also useful for the demonstration of the encapsulated yeast Cryptococcus neoformans. Metanil Yellow or Tartrazine Stain provides a light yellow counterstain to connective tissue and cytoplasmic elements.

 

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.

 

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 (Part 1409) 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 Weigert Iron Hematoxylin Working Solution; 7 minutes.
   5. Rinse in running tap water for 10 minutes.
   6. Prepare Mayer Mucicarmine Working Solution; combine, mix well.
      1. 1. Mucicarmine Stock Stain, Mayer                 10 ml  
         2. Tap Water (do not use distilled water) 30 ml      
   7. Stain in 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 Mayer Mucicarmine Working Solution. Microwave 10 minutes at 70°C.

1. 8. Rinse in several changes of tap water.
   9. Counterstain with preferred method:

• • 1. 1. Metanil Yellow Stain, Aqueous; stain 30 to 60 seconds. Proceed directly to Step #10.
       2. Tartrazine Stain 0.25%, Acetic Aqueous; 5 quick dips, rinse 30 seconds in distilled water. Proceed to Step #10.

1. 10. Dehydrate quickly through 95% and 100% ethyl alcohols. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

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. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 161-162.
   4. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 168-169.
   5. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	120 ml	120 ml X 6
Mold Release Concentrate	Part 1246A	Part 1246A

 

Additionally Needed:

Alcohol, Ethyl Denatured, 100%                   OR Alcohol, Isopropyl ACS, 100%	Part 10841     OR Part 12094
Base Molds, Stainless Steel, Heavy Duty	Part 5103
Base Molds, Stainless Steel	Part 5104
Base Molds, Disposable	Part 5105

 

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

 

APPLICATION:

Newcomer Supply Mold Release Concentrate, when diluted to a working mold release solution, facilitates the clean release of cooled, solidified paraffin embedded tissue blocks from tissue embedding base molds.

 

METHOD:

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

 

PROCEDURE:

1. 1. Dilute Mold Release Concentrate to a working solution. Combine solutions in a plastic spray bottle and mix well;
      1. 1. Mold Release Concentrate         5 ml
         2. Ethyl Alcohol or Isopropyl Alcohol       95 ml
         3. Store unused Mold Release Working Solution at room temperature for up to 6 months.
         4. Keep working solution tightly capped when not in use.
   2. Arrange clean and dry stainless steel base molds (Parts 5103, 5104) or disposable base molds (Part 5105) on paper towels or bench paper in a ventilated area.
   3. Spray all embedding base molds thoroughly with Mold Release Working Solution, allowing small beads of solution to form on base mold surfaces that will come in contact with paraffin.
      1. 1. See Procedure Note #1.
   4. Allow embedding base molds to thoroughly air dry prior to use.
      1. 1. Thorough drying of base molds is necessary to avoid residual Mold Release Solution mixing with paraffin during embedding process.
   5. Embed paraffin processed tissue in embedding base molds that have been treated with Mold Release and dried.
   6. Cool paraffin blocks on embedding center cold plate.
   7. After paraffin is solidified, cooled tissue blocks will easily separate from embedding base molds.
      1. 1. See Procedure Note #2.
   8. Repeat treatment of clean and dry base molds with Mold Release Working Solution on a routine basis.

 

PROCEDURE NOTES:

1. 1. Spraying embedding base molds provides a better adherence of Mold Release and preferred over soaking base molds in solution.
   2. If tissue blocks are difficult to remove from embedding base molds, a sharp tap of the base mold on the bench top will facilitate removal.

 

REFERENCES:

1. 1. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 66.
   2. Modifications developed by Newcomer Supply Laboratory.

 

• Shelf Life is 2 years from date of manufacture.

 

SOLUTION:

	1 Gallon	20 Liter Cube
Carson Modified Millonig Formalin	Part 12445A	Part 12445C

 

	30 ml vial, 15 ml fill (100/cs)
Carson Modified Millonig Formalin Vial	Part 12445E

 

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

 

APPLICATION:

Newcomer Supply Carson Modified Millonig Formalin, is a formalin based ready-to-use fixative buffered with sodium monobasic phosphate and sodium hydroxide.  Carson Modified Millonig Formalin promotes fixation with rapid penetration, providing excellent cellular detail and ultrastructural preservation.

This multi-purpose fixative has applications for both light microscopy (LM) and electron microscopy (EM) studies, including immunogold labeling, and can be used in place of standard 10% formalin fixatives.

Carson Modified Millonig Formalin Vials are available in 30 ml vials prefilled with 15 ml of fixative for ease of use, storage and transport.

 

METHOD:

Fixation Recommendations:

• • Small Biopsies: Minimum of 1-2 hours.
  • Larger Specimens: Minimum of 4-6 hours.

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

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue in Carson Modified Millonig Formalin after surgical excision.
      1. 1. See Procedure Notes #1 and #2.
   2. Hold tissue in Carson Modified Millonig Formalin for processing.
      1. 1. See Procedure Note #3.
   3. Processing:
      1. 1. Light Microscopy: place on tissue processor starting in primary fixation step or first dehydration station.
         2. Electron Microscopy: A secondary osmium tetroxide fixation is recommended. Refer to protocol for electron microscopy processing.

 

PROCEDURE NOTES:

1. 1. If received in Formalin 10%, Phosphate Buffered (Part 1090), rinse tissue thoroughly in tap water prior to placing in Carson Modified Millonig Formalin.
   2. For electron microscopy studies, fix tissue within 15 minutes after excision. Mince into 1 mm cubes for expedient fixative infiltration.
   3. Tissue can remain in Carson Modified Millonig Formalin over an extended period of time.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 68.
   2. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 11-12, 335-336.
   3. Carson, Freida L. and James Martin. “Formalin Fixation for Electron Microscopy.” The Journal of Histotechnology 2.2 (1979): 58-60.
   4. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 46.
   5. Modifications developed by Newcomer Supply Laboratory.

(use: AFB Stain, Fite.)

Opaque, plastic back with hinged covers that will fold under to save table space. Clear plastic covers for easy recognition of slide content. Slides lay horizontal and have easy release when pushed on end to pop up. Holds 20 slides (3″ x 1″). Dimensions: 7 9/16″ x 11 1/2″ x 7/16″ H.

Disposable, Individually Wrapped. Rigid plastic handle holds a #11 blade. Don’t expose yourself to potentially infectious specimens while washing instruments. Use and dispose. 50/pkg.

(use: Counterstain for IHC procedures)

(use: Gomori Basement Membrane Stain (PAMM) & Urates.)

 

• Shelf Life is 2 years from date of manufacture.

 

(use: Fixative for blood smears.)

(use: Counterstain for Mucicarmine.)

(use: Elastic-Masson Trichrome & stock solutions for GMS & Gomori Basement Membrane.)

(use: Bluing Reagent)

(use: Stock solution for GMS & Gomori Basement Membrane.)

(use: Counterstain for GMS Fungus.)

SOLUTION:

	250 ml	500 ml
Light Green SF Yellowish Stain 0.1%, Aqueous	Part 12203A	Part 12203B

 

Additionally Needed:

Fungus, PAS, Aspergillus, Artificial Control Slides                             OR Fungus, PAS, Candida, Artificial Control Slides	Part 4232     OR Part 4233
Periodic Acid 0.5%, Aqueous	Part 13308
Schiff Reagent, McManus	Part 1371
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 Light Green SF Yellowish 0.1%, Aqueous provides a complimentary counterstain for the Fungus/PAS Stain.

 

METHOD:

Fixation:  Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin section 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.

 

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 Periodic Acid 0.5%, Aqueous (Part 13308) for 5 minutes.
   4. Wash in three changes of tap water; rinse in distilled water.
   5. Drain slides of excess water and stain in Schiff Reagent, McManus (Part 1371) for 20 minutes.
   6. Wash gently in lukewarm tap water for 10 minutes to allow pink color to develop.
   7. Counterstain in 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.

(use: Counterstain for GMS Fungus.)

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

(use: Verhoeff Elastic/Masson Trichrome procedure.)

 

• Acetone, Alcohols, and Xylene do not have expiration dates.
• Shelf life is 2 yrs once opened.

(use: Hucker-Twort Gram Stain, Verhoeff Elastic, McLetchie Trichrome and mercury pigment removal.)

SOLUTION:

	1 Liter	1 Gallon
Hollande Fixative	Part 1208A	Part 1208B

 

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

 

APPLICATION:

Newcomer Supply Hollande Fixative, a ready-to-use picric acid based fixative with a copper acetate component that provides excellent morphology preservation, increased nuclear and sub-cellular detail while stabilizing red blood cell membranes. Hollande Fixative is recommended for gastrointestinal tract biopsies and endocrine tissue.

 

METHOD:

Fixation Recommendations:

• • Small Biopsies: A minimum of 1 hour.
  • Larger Biopsies: 4 hours to 24 hours.
  • To facilitate grossing, place in Hollande Fixative for 1 hour to firm; trim to smaller pieces.

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

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue in Hollande Fixative after surgical excision.
      1. 1. See Procedure Notes #1 and #2.
   2. The acidic nature of Hollande Fixative is generally enough to decalcify small bone without additional decalcification steps.
   3. Rinse Hollande fixed tissue thoroughly in running tap water for a minimum of 15 minutes, followed by a minimum of 15 minutes in 70% ethyl alcohol (10844) prior to processing.
      1. 1. See Procedure Notes #3 and #4.
   4. Place on tissue processor starting in Formalin 10%, Phosphate Buffered (Part 1090) fixation step.

 

PROCEDURE NOTES:

1. 1. If received in Formalin 10%, Phosphate Buffered, rinse tissue thoroughly in tap water prior to placing in Hollande Fixative.
   2. Hollande Fixative may grossly stain tissue green. This is normal.
   3. It is essential to thoroughly wash Hollande fixed tissue prior to tissue processing or an insoluble blue phosphate precipitate will form upon exposure to Formalin 10%, Phosphate Buffered.
   4. Extended storage in Hollande Fixative is not recommended.
      1. 1. After maximum fixation, wash tissue in running tap water for a minimum of 15 minutes, then a minimum of 15 minutes in 70% ethyl alcohol.
         2. Transfer Hollande fixed tissue to Formalin 10%, Phosphate Buffered for long-term storage purposes.
   5. Dispose of Hollande Fixative as picric acid hazardous waste according to local/state regulations.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 70.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 21.
   3. Dapson, Janet Crookham and Richard Dapson. Hazardous Materials in the Histopathology Laboratory: Regulations, Risks, Handling, and Disposal. 4th ed. Battle Creek, MI: Anatech, 2005. 150, 265-266.
   4. Goss, Gwen. “Hollande’s Solution as a Routine Tissue Fixative.” Lecture, Region IV Meeting, National Society for Histotechnology, Chicago, June 1990.
   5. Hollande, A. Ch. “Enrichment of Bouin’s Liquid Picric Acid by the Addition of Neutral Copper Acetate.” Report of the Society of Biology 81 (1918): 17.
   6. Modifications developed by Newcomer Supply Laboratory.

 

• Shelf Life is 4 years from date of manufacture.

(use: Gomori Mod. Iron Stain, Prussian Blue & stock for Colloidal Iron.)