These pads are used to hold biopsies in place and prevent them from being lost during processing. They are made of a polyester urethane foam which is always verified for consistency throughout to achieve optimum solvent flow. This pad will fit the majority of tissue cassettes for this purpose.  Will resist temps. from -40°C to 121°C.

Disposable, deep mold. Inexpensive enough to discard after use or strong enough to be reused. Designed to facilitate easy block release.

CLICK HERE for MOLD RELEASE, in Reagents Chemicals & Buffers Section.

 

MICROSCOPE GLASS COVER SLIPS PACKAGING:

• Each case comes with 10 one ounce boxes
• Each box shrink sealed with desiccant

 

QUALITY GLASS COVER SLIPS WITHOUT THE COST!

Available in the following sizes:

• 18 x 18 mm
• 22 x 22 mm
• 24 x 24 mm
• 24 x 40 mm
• 24 x 50 mm
• 24 x 60 mm

Easy to use timer/stopwatch with 4 independent timing channels with visual & audible alarms. Readout on 6-digit LCD. Spring clip, magnet & flip stand. Includes battery. Measures 61 x 71 x 25mm.

 

The plastic Tissue Block Modular Storage Drawers provide permanent storage and identification of embedding cassettes and rings.

• Each drawer holds up to 250 cassettes or 165 embedding rings
• Stackable with interlocking ridges on top and bottom
• Made of high impact resistant plastic
• Dimensions: 15 7/8″ x 9 1/8″ x 2 1/8″
• Identification labels included

 

A durable plastic slide staining tray that is priced to be disposable, but can be used more than once!

 

FEATURES OF THE DISPOSABLE SLIDE STAINING TRAYS:

• • Stain, rinse, and dry slides on a single working tray
  • A clean and fresh working surface area every time
  • Dark lid protects slides for light-sensitive applications
  • Compact size and recessed handles for easy transport from work area to sink
  • Disposable

 

SPECIFICATIONS:

• • Made from a polypropylene and polyethylene blend for stability
  • Color: Black
  • Up to eight slides fit comfortably onto the base
  • Deep well holds up to 38 ml
  • 2 convenient pour spouts for quick disposal of unwanted liquid waste
  • 12 x 5 x 1.2 in (30.5 x 12.7 x 3 cm)

 

INCLUDES:

• • 4 Black Base/Trays
  • 1 Black Lid

Tissue-Plus^® OCT (Optimal Cutting Temperature) Compound formula has been used extensively in labs since 1999 as an embedding medium for frozen tissue specimens. Compares to Sakura Tissue-Tek OCT.

 

TISSUE-PLUS OCT SPECIFICATIONS:

• Clear, colorless solution free of particles.
• Provides a stable specimen matrix and superior section quality at cryostat temperatures from -10°C and below.
• Leaves no residue on slides, eliminating undesirable background staining.
• Will not dull cryostat blades.
• Does not separate like some other freezing compounds.
• Provides a consistent sectioning substrate from the first use to the last.
• Convenient 4 oz. spout plastic container.

Black, fine-tip ink pen designed for histo and cyto labs. Writes without smearing on both frosted and plain glass slides along with plastic cassettes. Works well with high pH citrate buffer HIER. Dries quickly.

A microscope slide box for slides larger than 3″ x 2″.

• Sturdy ABS material
• Soft cork lining in bottom
• Identification card on inside lid
• Stackable with other boxes

Each envelope contains a stainless steel 4″ forcep, a plastic handled scalpel with a #11 blade & a pointed scissors. Avoid washing & exposure to infectious disease by simply disposing of them after use.

Ultra fine-tip, black ink pen that writes and dries quickly. Works on frosted slides and plastic cassettes. Aqueous based ink. Withstands histo & cyto stains and chemistries. Keeps working and gets the job done. A lot of pen for the buck!

SOLUTION:

	100 ml
Crystal Violet Stain, Lieb, Alcoholic	Part 10421A

 

Additionally Needed:

Amyloid, Animal Control Slides	Part 4031
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 Crystal Violet Stain, Lieb, Alcoholic is used to provide a rapid screening method for amyloid deposits in tissue sections.  This procedure has low sensitivity and should be considered as an amyloid screening technique and not an amyloid specific stain.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 10-12 microns

1. 1. See Procedure Note #1.

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. See Procedure Note #2.
   3. Stain sections in Crystal Violet Stain, Lieb, Alcoholic for 10 minutes.
   4. Rinse well in tap water.
   5. Blot water from slides; allow slides to air-dry in a vertical position.
   6. Coverslip air-dried sections with compatible mounting medium.
      1. 1. See Procedure Note #3.

 

RESULTS:

Amyloid	Purple/violet
Background	Purple/blue

 

PROCEDURE NOTES:

1. 1. For best results cut sections at 10-12 microns to provide better definition and more intense amyloid staining.
   2. Drain slides after each step to prevent solution carry over.
   3. Avoid the use of aqueous based mounting mediums which will cause bleeding/diffusion of the stain from the tissue section.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization step.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 154-155.
   2. Lieb, E. “Permanent Stain for Amyloid.” American Journal of Clinical Pathology 17 (1947). 413-414.
   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. 154.
   4. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 177.
   5. Modifications developed by Newcomer Supply Laboratory.

GRAM, HUCKER-TWORT STAIN KIT INCLUDES:                                                                                   

		Part 9125A
Solution A:	Crystal Violet-Oxalate Stain, Alcoholic	250 ml
Solution B:	Iodine, Lugol’s, Aqueous	250 ml
Solution C:	Neutral Red Stain 1%, Alcoholic	125 ml
Solution D:	Fast Green Stain 1%, Alcoholic	50 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
Acetone, ACS	Part 10014

 

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

 

APPLICATION:

Newcomer Supply Gram, Hucker-Twort Stain Kit is a rapid procedure for staining gram-positive and gram-negative bacteria without picric acid.  The Twort Stain combines Neutral Red and Fast Green, for clear detection of red gram-negative bacteria with a green counterstain.

 

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 A: Crystal Violet-Oxalate Stain, Alcoholic with high quality filter paper.

 

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 Note #1.
   4. Stain in freshly filtered Solution A: Crystal Violet-Oxalate Stain, Alcoholic (Step #2) for 30 seconds.
   5. Rinse quickly in distilled water.
   6. Mordant in Solution B: Iodine, Lugol’s, Aqueous for 20 seconds.
   7. Rinse quickly in distilled water.
   8. Decolorize individually with Acetone, ACS (Part 10014); 2 quick dips.
      1. 1. Or until tissue remains light gray.
   9. Rinse quickly in distilled water.

1. 10. Prepare fresh Twort Stain; combine and mix well.
       1. 1. Solution C: Neutral Red Stain 1%, Alcoholic 9 ml
          2. Solution D: Fast Green Stain 1%, Alcoholic   3 ml
          3. Distilled Water                               30 ml
          4. Use within 30 minutes.
   11. Stain in fresh Twort Stain for 2 minutes.
   12. Rinse quickly in distilled water; carefully blot dry.
   13. Agitate slides quickly in clean Acetone, ACS to remove excess stain and dehydrate; do not use any alcohols.
       1. 1. The use of alcohol will remove Neutral Red.
   14. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Gram-positive bacteria	Dark blue
Gram-negative bacteria	Red
Cytoplasm and red blood cells	Shades of green
Nuclei	Red

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Alan Stevens. Theory and Practice of Histological Techniques. 3rd ed. Edinburgh: Churchill Livingstone, 1990. 290-292.
   2. Culling, C.F.A. Handbook of Histopathological and Histochemical Techniques. 3rd ed. London: Butterworth, 1974. 393-395.
   3. Twort, F.W., “An Improved Neutral Red, Light Green Double Staining for Animal Parasites, Microorganisms and Tissues.” Journal of State Medicine (1924). 351.
   4. Modifications developed by Newcomer Supply Laboratory.

Tech Memo 1: Crystal Violet-Oxalate Stain, Alcoholic for Brown-Brenn Gram Stain

 

SOLUTION:

 

SOLUTION:

	250 ml	500 ml
Crystal Violet-Oxalate Stain, Alcoholic	Part 10422A	Part 10422B

 

Additionally Needed:

Gram, Multi-Tissue, Artificial Control Slides                          OR Gram+ & Gram- Bacteria, Artificial Control Slides	Part 4256      OR Part 4255
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Iodine, Lugol’s, Aqueous	Part 12092
Acetone, ACS	Part 10014
Twort’s Gram Stain Set     Solution A: Neutral Red Stain 1%, Alcoholic     Solution B: Fast Green Stain 1%, Alcoholic	Part 14034

 

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

 

APPLICATION:

Newcomer Supply Crystal Violet-Oxalate Stain, Alcoholic, a component of the Hucker-Twort Gram Stain that stains gram-positive bacteria. The Twort Stain combines Neutral Red and Fast Green for clear detection of red gram-negative bacteria with a green counterstain.

 

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. Filter Crystal Violet-Oxalate Stain, Alcoholic with high quality filter paper.

 

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 Note #1.
   4. Stain in freshly filtered Crystal Violet-Oxalate Stain, Alcoholic for 30 seconds.
   5. Rinse quickly in distilled water.
   6. Mordant in Iodine, Weigert, Lugol’s, Aqueous (Part 12092); 20 seconds.
   7. Rinse quickly in distilled water.
   8. Decolorize individually with Acetone, ACS (Part 10014); 2 quick dips.
   9. Rinse quickly in distilled water.
   10. Prepare fresh Twort Stain (Part 14034); combine and mix well.
       1. 1. Neutral Red Stain 1%, Alcoholic 9 ml
          2. Fast Green Stain 1%, Alcoholic      3 ml
          3. Distilled Water               30 ml
          4. Use within 30 minutes.
   11. Stain in fresh Twort Stain for 2 minutes.
   12. Rinse quickly in distilled water; carefully blot dry.
   13. Agitate slides quickly in clean Acetone, ACS to remove excess stain and dehydrate; do not use any alcohols.
       1. 1. The use of alcohols will remove Neutral Red Stain.

1. 14. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Gram-positive bacteria	Dark blue
Gram-negative bacteria	Red
Cytoplasm and red blood cells	Shades of green
Nuclei	Red

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D. and Alan Stevens. Theory and Practice of Histological Techniques. 3rd ed. Edinburgh: Churchill Livingstone, 1990. 290-292.
   2. Culling, C.F.A. Handbook of Histopathological and Histochemical Techniques (including museum techniques). 3rd ed. London: Butterworth, 1974. 393-395.
   3. Twort, F.W. “An Improved Neutral Red, Light Green Double Staining for Animal Parasites, Microorganisms and Tissues.” Journal of State Medicine 32 (1924). 351.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	100 ml
Cresyl Violet Acetate 2.5%, Aqueous	Part 1039A

 

Additionally Needed:

Luxol Fast Blue (LFB) Control Slides	Part 4407
Luxol Fast Blue (LFB) Stain Set	Part 12218 (for LFB/Cresyl Violet Acetate Stain)
Acetic Acid, Glacial, ACS	Part 10010
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 Cresyl Violet Acetate 2.5%, Aqueous is a metachromatic basic dye stock solution for the demonstration of Nissl substance and nuclei in nerve tissue.  As a working solution, Cresyl Violet Acetate can be utilized in the Luxol Fast Blue (LFB) procedure or used solely as a stand-alone nerve tissue stain.

Cresyl Violet Acetate is also known as Cresyl Violet, Cresyl Fast Violet Acetate and Cresyl Echt Violet Acetate.  Due to characteristics of the dye, the actual percentage of dye concentration in Cresyl Violet powder may vary between lots.  The minimum acceptable dye content for Cresyl Violet certification is noted as 65%.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)

Technique:  Paraffin sections cut at 8-10 microns on adhesive slides

• • • Air-dry a minimum of 30 minutes

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

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare Acetic Acid 10%, Aqueous; combine and mix well.
      1. 1. Acetic Acid, Glacial, ACS (Part 10010) 10 ml
         2. Distilled Water                90 ml
         3. Store at room temperature for up to 1 year.
   3. Prepare Working Cresyl Violet Acetate 0.25%; mix well and filter.
      1. 1. Cresyl Violet Acetate 2.5%, Aqueous 5 ml
         2. Distilled Water              45 ml
         3. Acetic Acid 10%, Aqueous                  7 drops
         4. Heat filtered solution in 57°C oven; hold for Step #7.
         5. See Procedure Notes #3 and #4.

 

STAINING PROCEDURES:

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.
      1. 1. LFB procedure: stop at 95% ethyl alcohol, no distilled water rinse.
         2. Cresyl Violet Acetate stand-alone staining: continue with distilled water rinse. Proceed to Step #6.
         3. See Procedure Notes #1 and #2.
   5. LFB Stain: proceed with LFB Stain Set (Part 12218) protocol through final differentiation step and rinse thoroughly in distilled water.
   6. Continue with LFB Cresyl Violet Acetate counterstain or Cresyl Violet Acetate stand-alone staining.
   7. Stain in filtered, heated Working Cresyl Violet Acetate 0.25% (Step #3) for 6 minutes in 57°C oven.
   8. Rinse in distilled water.
   9. Dehydrate quickly to maintain Cresyl Violet Acetate stain 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:

LFB with Cresyl Violet:	Myelin	Blue
	Nissl substance and nuclei	Violet
	Neurons	Pink to violet
Cresyl Violet alone:	Nissl substance	Purple/dark blue
	Nuclei	Purple to blue
	Neurons	Pale purple-blue
	Background	Colorless

 

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. Due to possible dye percentage variance between lots of Cresyl Violet powder, a solution stronger than 0.25% Working Cresyl Violet Acetate may be necessary for optimal staining results.
   4. For enhanced staining of Working Cresyl Violet Acetate, adjust pH to 4.0 with Acetic Acid 10%, Aqueous.
   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. 366-367.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 191-193, 207-208.
   3. Horobin, Richard and John Kiernan. Conn’s Biological Stains: A Handbook of Dyes, Stains and Fluorochromes for Use in Biology and Medicine. 10th ed. Oxford: BIOS, 2002. 281-282.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	250 ml	500 ml
Carbol Fuchsin Stain, Kinyoun	Part 1031A	Part 1031B

 

Additionally Needed:

Acid Fast Bacteria (AFB) Control Slides	Part 4011
Acid Alcohol 1%	Part 10011
Methylene Blue Stain 0.14%, Alcoholic	Part 12401
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 Carbol Fuchsin Stain, Kinyoun is used in the Kinyoun AFB Stain to demonstrate the presence of acid-fast mycobacteria in tissue sections. Carbol Fuchsin Stain, Kinyoun is a concentrated phenol and basic fuchsin solution that works to permeate the lipoid capsule of acid-fast organisms and renders them resistant to acid alcohol decolorization.

 

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. Filter Carbol Fuchsin Stain, Kinyoun with filter paper before use.

 

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 freshly filtered Carbol Fuchsin Stain, Kinyoun for 15-30 minutes at room temperature. Keep solution covered.
      1. 1. See Procedure Note #3.
   5. Wash in running tap water for 2 to 3 minutes.
   6. Differentiate in Acid Alcohol 1% (Part 10011) until color no longer runs off the slide and sections are pale pink; 3 to 10 rapid dips.
   7. Wash in running tap water 3 to 5 minutes; rinse in distilled water.
   8. Counterstain in Methylene Blue Stain 0.14%, Alcoholic (Part 12401); 3-6 dips.
      1. 1. See Procedure Note #4.
   9. Wash in running tap water for 1 minute; rinse in distilled water.
   10. 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.

 

RESULTS:

Acid-fast bacteria	Bright red
Background	Pale 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. Sections can remain in Carbol Fuchsin Stain, Kinyoun for up to 60 minutes without adverse effect.
      1. 1. Additional differentiation may be required in Step #6.
   4. If preferred, Light Green SF Yellowish Stain 0.1%, Aqueous (Part 12203) can be used as a counterstain in place of Methylene Blue.
      1. 1. Stain for 2-5 dips.
         2. Rinse with one quick dip in distilled water or proceed directly to Step #10 without a distilled water rinse.
   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. 224-226.
   2. Kinyoun. J.J. “A Note on Uhlenhuths Method for Sputum Examination, for Tubercle Bacilli.” American Journal of Public Health 5.9 (1915). 867-870.
   3. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 236-237.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	250 ml	500 ml
Alcian Blue Stain 1%, pH 1.0 Aqueous	Part 1005A	Part 1005B

 

Additionally Needed:

Hydrochloric Acid 0.1N, Aqueous	Part 1207
Nuclear Fast Red Stain, Kernechtrot	Part 1255
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 Alcian Blue Stain 1%, pH 1.0 Aqueous is for the specific staining of sulfated acid epithelial mucins in the Alcian Blue 1%, pH 1.0 procedure.

 

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.

 

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 note #1.
   3. Rinse briefly in Hydrochloric Acid 0.1N, Aqueous (Part 1207).
      1. 1. Save Hydrochloric Acid 0.1N, Aqueous for Step #5.
   4. Stain in Alcian Blue Stain 1%, pH 1.0 Aqueous for 30 minutes.
   5. Rinse briefly in Hydrochloric Acid 0.1N, Aqueous.
      1. 1. Do not wash in water which may alter pH level resulting in nonspecific staining.
   6. Blot slides dry with filter paper.
   7. Counterstain dried slides with Nuclear Fast Red Stain, Kernechtrot (Part 1255) for 5 minutes.
      1. 1. Shake solution well before use; do not filter.
   8. Rinse well in distilled water.
      1. 1. See Procedure Note #2.
   9. 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.

 

RESULTS:

Sulfated acid epithelial mucin	Blue
Cytoplasm and background	Pink
Nuclei	Pink to red

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
   3. Small intestine and colon control slides can be used as positive controls for staining of sulfated acid epithelial mucins.
      1. 1. Alcian Blue pH 2.5, Goblet Cell Control Slides (Part 4051)
         2. Mucin Mucicarmine Control Slides (Part 4455)
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology A Self-Instructional Text. 4th ed. Chicago: ASCP Press, 2015. 148-149.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 173.
   3. Modifications developed by Newcomer Supply Laboratory.

Ultra fine-tip (diameter approximately 0.75mm), dark black ink pen.  Write easily on microscope slides and plastic cassettes.  Will go through the harshest histology chemistries and stay on.  Excellent choice for the toughest applications.

 

 

 

 

 

Molded from acetal, the patented Micromesh Biopsy Tissue Processing & Embedding Cassettes (4 Compartments) 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 without having to use biopsy pads.

 

PACKAGING OF THE MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES (4 Compartments):

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

 

MICROMESH BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES (4 Compartments):

• 1676 square openings (0.38mm) allowing for a greatly improved fluid exchange without having to use biopsy pads.
• Four square compartments each measuring 13 x 13mm.
• Large anterior and posterior slots in both cassette and cover ensure that the cassette will sink rapidly.
• Recessed cover allowing more cassettes to be stacked in automatic labeling machines and tissue processors.
• Anterior writing area is at a 45° angle.

 

This tissue flotation bath is made of a seamless inner-chamber that is easy to fill & clean.  Heats quickly and is simple to operate.  An excellent tissue flotation bath for any histology lab and very easy on the budget!

FEATURES OF THE ECONOMY HISTOLOGY WATER BATH:

• Low Profile 2.3 liter black, finished interior chamber
• Glass lid for easy viewing of interior
• Durable fiberglass insulated body prevents heat loss
• Non-stick coated chamber for easy cleaning
• Thermometer holder
• Temperature range of room temp. to 75ºC

 

DIMENSIONS OF THE HISTOLOGY WATER BATH:

• Overall 10.5″ diameter x 8.25″ H
• Interior 8.25″ diameter x 2.5″ H

 

CERTIFICATION AND APPROVALS:

• CE certified

 

ELECTRICAL SPECIFICATIONS
Voltage	Amps	Hertz	Wattage
120	3.33	60	400

 

WARRANTY DETAILS:

• • The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

 

Manual for Tissue Flotation Bath XH-1001

 

This digital tissue flotation bath has a lighted water basin that produces excellent visibility for the floating tissue section(s).  Excellent value and easy to use!

Features of the Histology Water Bath:

• Low profile design with removable rectangular glass basin
• The water bath is illuminated from the side and provides exceptional viewing of the tissue section(s)
• Easy to read LED display shows both set temperature and actual temperature
• Excellent value!

 

Specifications of the Histology Water Bath:

• Temp Range: Room Temp to 75° C +/- 1° C
• Capacity: 2 Liters (1/2 gallon)
• Size Glass Tray (included): 10″ (W) x 6.5″ (D) x 1.75″ (H)
• Overall Size: 14″ x 14″ x 4″
• Shipping Weight: 22 lbs.
• Power Requirements: 300W (110V, 60Hz)

 

Certification and Approvals:

• CE certified

 

ELECTRICAL SPECIFICATIONS
Voltage	Amps	Hertz	Wattage
120	4.17	60	500

 

WARRANTY DETAILS:

• • The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

 

Manual for Tissue Flotation Bath- Digital XH-1003

 

Digital Paraffin Trimmer at an Incredible Price!

Our digital paraffin wax trimmer XH-90D model from Premiere is an efficient appliance for removing excess paraffin from embedding tissue cassettes.  The digitally controlled temperature update to the unit allows for the user to set the temperature of the heating plate from room temperature up to 120°C in one degree increments.

Compact tabletop design of the digital paraffin trimmer fits easily in any lab.  The melted paraffin is easily melted away and drains into the removable tray below. Collection tray stays in place by magnetic contact with unit.  Also available is a convenient disposable tray liner that allows easy disposal of the runoff wax.  No need to scrape out the collection tray!

 

FEATURES OF THE PARAFFIN WAX TRIMMER – DIGITAL, PREMIERE:

• • Digital Control Panel
  • Grooved Heating Surface
  • Adjustable Heat Setting
  • Temperature range:  Room Temp. to 120°C
  • Magnetically secured collection tray holds disposable tray liners for easy disposal
  • Trim multiple cassettes simultaneously
  • LED power light
  • Includes 5 disposable drip tray liners

 

DIMENSIONS:

• • Overall size: 11.25″ x 5″ x 8″
  • Plate size:  9″ x 5″

 

ELECTRICAL SPECIFICATIONS
Voltage	Hertz	Wattage
110	60	150W

 

For User’s Manual Click Here

 

The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

 

PARAFFIN DISPENSER XH-4002 FEATURES:

• • 2.3 gallon basin
  • Durable industrial metal outer cabinet for long life
  • Built in temperature controller
  • Stainless steel interior for easy cleaning
  • Heat tolerant metal faucet
  • Heats quickly

 

Temperature Range:   Room Temp to 72° C (tolerance +/- 2 degrees)

Dimensions:

• Overall size 14″ x 16 ¹/₄” x 19″
• Interior size 9″ diameter x 10″

 

CERTIFICATION & APPROVALS:

• • CE certified
  • Cord is CSA & UL listed

ELECTRICAL SPECIFICATIONS
Voltage	Amps	Hertz	Wattage
120	8.83	60	1060

 

 

Click here for Paraffin Dispenser Manual

The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

 

 

This slide warmer’s unique design accommodates twice as many slides as a flat unit of the same size. Also, it makes retrieving the slides easier since the end of the slide is not on a flat surface. Easy to use digital temperature controller allows user to set temperatures from room temp. to 75°C. Removable tray for easy cleaning.

 

SPECIFICATIONS:

• • Capacity:  Holds 40 slides(slide size 3″ x 1″ or 75mm x 25mm)
  • Temperature Range:  Room temp. to 75ºC (+/- 2° C)
  • Overall Size:  14″ x 14″ x 4″H

 

CERTIFICATION & APPROVALS:

• CE certified
• Cord is CSA & UL listed

 

ELECTRICAL SPECIFICATIONS
Voltage	Amps	Hertz	Wattage
120	2.5	60	300

 

The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

 

These Slide Warmers are ideal for use in histology, cytology, pathology and biology for drying & mounting paraffin tissue sections on slides.

 

FEATURES OF THE SLIDE WARMERS:

• Easy to adjust & use
• Thermal heater ensures even heat transfer
• LED Temperature Display
• Temperature settings from room temp. to 75° C (+/- 2° C)
• Available in 2 sizes to fit most needs

 

DIMENSIONS OF THE XH-2002 (6258) SLIDE WARMER:

• Size 10″ x 7″
• Approx. Capacity – 23 slides
• Weight is 10 lbs.
• Power requirements 100W

 

DIMENSIONS OF THE XH-2001 (6259) SLIDE WARMER:

• Size 25″ x 8″
• Approx. Capacity – 66 slides
• Weight is 13 lbs.
• Power requirements 200W

 

CERTIFICATION & APPROVALS:

• CE certified
• Cord is CSA & UL listed

 

ELECTRICAL SPECIFICATIONS – #6258
Voltage	Amps	Hertz	Wattage
120	0.83	60	100

ELECTRICAL SPECIFICATIONS – #6259
Voltage	Amps	Hertz	Wattage
120	1.7	60	204

 

 

The manufacturer warrants this instrument to be free from defects in material and workmanship under normal use for one year from the date of purchase.  It does not cover damage resulting from abuse or misuse, repairs or alterations performed by other than authorized repair technicians, or damage occurring in transit.

SOLUTION: 

	1 Liter	1 Gallon	20 Liter Cube
AZF Fixative	Part 1009A	Part 1009B	Part 1009C
	30 ml vial, 15 ml fill (100/cs)
AZF Fixative Vial	Part 10091B

 

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

 

APPLICATION:

Newcomer Supply AZF (Acetic Acid-Zinc-Formalin) Fixative is a ready-to-use fixative recommended for bone marrow clots and cores, lymph nodes, endoscopic biopsies and immunohistochemical (IHC) studies.

AZF Fixative Vials are available in 30 ml vials prefilled with 15 ml of AZF Fixative for ease of use, storage and transport.

 

METHOD:

Fixation:

• • Bone Marrow Clots: Minimum of 2 hours.
  • Bone Marrow Biopsy: Minimum of 3 hours.
  • Lymph Nodes and Small Biopsies: Minimum of 4 hours.
  • Small nodes (5 mm or less) should be halved.
  • Larger nodes, dissected with no piece thicker than 3 mm.
  • To facilitate grossing, place in AZF Fixative for 1 hour to firm; trim to 2-3 mm.

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

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue in AZF Fixative after surgical excision.
      1. 1. See Procedure Note #1.
   2. Hold tissue in AZF Fixative for processing or maximum of 72 hours.
      1. 1. See Procedure Note #2.
   3. Wash AZF fixed tissue in tap water a minimum of 10 minutes to remove residual zinc.
   4. Decalcify bone marrow specimen as needed.
      1. 1. See Procedure Notes #3 & #4.
   5. Place on tissue processor 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 AZF Fixative.
   2. Extended storage in AZF Fixative is not recommended.
      1. 1. After maximum fixation, wash tissue in running tap water a minimum of 10 minutes.
         2. Transfer AZF fixed tissue to Formalin 10%, Phosphate Buffered for long-term storage purposes.
   3. Nitric acid is not recommended as a decalcification agent following AZF fixation.
   4. Acetic acid in the AZF Fixative may pre-start decalcification process, decreasing overall decalcification time.
   5. Zinc chloride is corrosive, do not discard AZF Fixative solution down the drain.
   6. Neutralize AZF Fixative with sodium carbonate or sodium bicarbonate to precipitate zinc at pH 7.0-8.0.
      1. 1. Approximately 100 grams of sodium bicarbonate will neutralize/precipitate zinc from 1 liter of AZF Fixative.

 

REFERENCES:

1. 1. Bonds, Lian A., Pat Barnes, Kathryn Foucar and Cordelia E. Sever. “Acetic Acid-Zinc-Formalin: A Safe Alternative to B-5 Fixative.” American Journal of Clinical Pathology, 124 (2005): 205-11.
   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. 148, 279.
   3. Naresh, K N, I. Lampert and R. Hasserjian. “Optimal Processing of Bone Marrow Trephine Biopsy: The Hammersmith Protocol.” Journal of Clinical Pathology 59 (2006): 903-11.
   4. Modifications developed by Newcomer Supply Laboratory.

PRODUCTS:                                                       

	Each or 6/Set
PAP Pen Liquid Blocker	Part 6505
PAP Pen Liquid Blocker, Mini	Part 6506

 

APPLICATION:

Newcomer Supply PAP Pen Liquid Blocker, hydrophobic slide markers creates a visible hydrophobic barrier that provides proper surface tension to hold reagents within a targeted area when applied around tissue sections or smears on a microscopic slide. This barrier ensures the amount of reagent needed is reduced and retained for complete tissue coverage and staining reaction. Multiple specimens can be separated by drawn circles or lines applied to the same slide. The PAP Pen, Mini provides a finer pen tip for drawing a thinner barrier film.

PAP Pen Liquid Blockers contain a unique formulation that is water repellent, insoluble in alcohol and acetone, soluble in xylene and temperature resistant up to 120°C.

 

METHOD:

Technique:  Paraffin, frozen sections and smears

• • • Manual immunohistochemistry (IHC) procedures
    • Manual immunofluorescence (IF) procedures
    • Manual In-Situ hybridization (ISH) procedures
    • Manual enzyme procedures

 

PAP PEN ACTIVATION INSTRUCTIONS:

1. 1. Vigorously shake capped pen for approximately 10 seconds.
   2. Remove cap; hold pen upright and depress tip 2-3 times.
   3. Invert pen, press tip lightly on absorbent paper until tip is visually saturated. Release pressure on tip and blot away any excess liquid.
   4. Practice applying a thin PAP Pen liquid barrier on a test slide using minimal pressure. Too much pressure may result in excess barrier solution creating a wider film that could touch tissue edges.
   5. Store PAP Pen Liquid Blocker at room temperature, tightly capped on a horizontal flat surface.
   6. Pen re-activation not required unless tip dries out during storage.

 

PROCEDURE:

1. 7. Paraffin Section Method:
      1. 1. Deparaffinize sections thoroughly in xylene and hydrate through 100% and 95% ethyl alcohols. Wash well with distilled water.
         2. Remove slide from water, blot excess solution from slide and tissue section. Or place long edge of slide on absorbent material to remove excess moisture.
         3. Encircle tissue section on slide surface with PAP Pen as illustrated. Do not touch the pen to any edges of the tissue.
         4. The liquid barrier can be drawn on a slightly damp slide.
         5. See Procedure Notes #1 and #2.
   8. Frozen Section and Smear Method:
      1. 1. Encircle frozen tissue section or smear on slide at room temperature with the PAP Pen. Do not touch the pen to any edges of the tissue section or smear.
         2. The PAP Pen barrier should be applied before fixation or prior to immersion of slide into water or buffer.
         3. See Procedure Notes #1 and #2.
   9. After liquid barrier application, allow barrier film to dry before proceeding with staining procedure.
      1. 1. See Procedure Note #3.

1. 10. Drain/rinse reagents off between staining steps; blotting slide and tissue as needed to remove excess solution.
       1. 1. See Procedure Note #4.
   11. Complete staining; coverslip with compatible mounting medium.

 

PROCEDURE NOTES:

1. 1. Once a tissue section is touched with PAP Pen liquid barrier it cannot be removed. The section remains usable but colorization will result on the touched portion of tissue.
   2. If the tissue section is not completely encircled by the PAP Pen Liquid Blocker barrier film, reagents will not be fully retained on the section and flood out onto the slide. This may compromise adequate tissue coverage by reagents.
   3. If liquid barrier lines are not completely dry prior to staining, a precipitate from reaction with reagents may occur.
   4. The use of Slide Moisture Chamber (68431, 68432) or StainTray™ (6847, 6848) is recommended for manual staining to maintain slide organization and a moist environment during the procedure.

 

REFERENCES:

1. 1. Grizzle, William, Cecil Stockard and Paul Billings. “The Effects of Tissue Processing Variables Other Than Fixation on Histochemical Staining and Immunohistochemical Detection of Antigens.” The Journal of Histotechnology 24.3 (2001): 213-219.
   2. Vidwans, Malavika, Srinivas Mandavilli, Wanda Nethers and Richard Cartun. “Fine-Needle Aspiration Diagnosis of a Neck Mass Using Immunocytochemical Stains Performed on Stained Cytology Slides.” The Journal of Histotechnology 25.4 (2002): 275-277.
   3. Modifications developed by Newcomer Supply Laboratory.

PRODUCTS:                                                       

	Each or 6/Set
PAP Pen Liquid Blocker	Part 6505
PAP Pen Liquid Blocker, Mini	Part 6506

 

APPLICATION:

Newcomer Supply PAP Pen Liquid Blocker, hydrophobic slide markers creates a visible hydrophobic barrier that provides proper surface tension to hold reagents within a targeted area when applied around tissue sections or smears on a microscopic slide. This barrier ensures the amount of reagent needed is reduced and retained for complete tissue coverage and staining reaction. Multiple specimens can be separated by drawn circles or lines applied to the same slide. The PAP Pen, Mini provides a finer pen tip for drawing a thinner barrier film.

PAP Pen Liquid Blockers contain a unique formulation that is water repellent, insoluble in alcohol and acetone, soluble in xylene and temperature resistant up to 120°C.

 

METHOD:

Technique:  Paraffin, frozen sections and smears

• • • Manual immunohistochemistry (IHC) procedures
    • Manual immunofluorescence (IF) procedures
    • Manual In-Situ hybridization (ISH) procedures
    • Manual enzyme procedures

 

PAP PEN ACTIVATION INSTRUCTIONS:

1. 1. Vigorously shake capped pen for approximately 10 seconds.
   2. Remove cap; hold pen upright and depress tip 2-3 times.
   3. Invert pen, press tip lightly on absorbent paper until tip is visually saturated. Release pressure on tip and blot away any excess liquid.
   4. Practice applying a thin PAP Pen liquid barrier on a test slide using minimal pressure. Too much pressure may result in excess barrier solution creating a wider film that could touch tissue edges.
   5. Store PAP Pen Liquid Blocker at room temperature, tightly capped on a horizontal flat surface.
   6. Pen re-activation not required unless tip dries out during storage.

 

PROCEDURE:

1. 7. Paraffin Section Method:
      1. 1. Deparaffinize sections thoroughly in xylene and hydrate through 100% and 95% ethyl alcohols. Wash well with distilled water.
         2. Remove slide from water, blot excess solution from slide and tissue section. Or place long edge of slide on absorbent material to remove excess moisture.
         3. Encircle tissue section on slide surface with PAP Pen as illustrated. Do not touch the pen to any edges of the tissue.
         4. The liquid barrier can be drawn on a slightly damp slide.
         5. See Procedure Notes #1 and #2.
   8. Frozen Section and Smear Method:
      1. 1. Encircle frozen tissue section or smear on slide at room temperature with the PAP Pen. Do not touch the pen to any edges of the tissue section or smear.
         2. The PAP Pen barrier should be applied before fixation or prior to immersion of slide into water or buffer.
         3. See Procedure Notes #1 and #2.
   9. After liquid barrier application, allow barrier film to dry before proceeding with staining procedure.
      1. 1. See Procedure Note #3.

1. 10. Drain/rinse reagents off between staining steps; blotting slide and tissue as needed to remove excess solution.
       1. 1. See Procedure Note #4.
   11. Complete staining; coverslip with compatible mounting medium.

 

PROCEDURE NOTES:

1. 1. Once a tissue section is touched with PAP Pen liquid barrier it cannot be removed. The section remains usable but colorization will result on the touched portion of tissue.
   2. If the tissue section is not completely encircled by the PAP Pen Liquid Blocker barrier film, reagents will not be fully retained on the section and flood out onto the slide. This may compromise adequate tissue coverage by reagents.
   3. If liquid barrier lines are not completely dry prior to staining, a precipitate from reaction with reagents may occur.
   4. The use of Slide Moisture Chamber (68431, 68432) or StainTray™ (6847, 6848) is recommended for manual staining to maintain slide organization and a moist environment during the procedure.

 

REFERENCES:

1. 1. Grizzle, William, Cecil Stockard and Paul Billings. “The Effects of Tissue Processing Variables Other Than Fixation on Histochemical Staining and Immunohistochemical Detection of Antigens.” The Journal of Histotechnology 24.3 (2001): 213-219.
   2. Vidwans, Malavika, Srinivas Mandavilli, Wanda Nethers and Richard Cartun. “Fine-Needle Aspiration Diagnosis of a Neck Mass Using Immunocytochemical Stains Performed on Stained Cytology Slides.” The Journal of Histotechnology 25.4 (2002): 275-277.
   3. Modifications developed by Newcomer Supply Laboratory.

• Holds up to 76 cassettes
• Cassettes stack vertically (label side up)
• Snap in place lid

 

 

Dimensions:  1 3/4″ x 5 1/4″ x 7 1/2″

Removable dividers, a hinged lid and handles make this large capacity basket versatile and convenient.  This is a ‘must-have’ item for your lab!  Holds up to 150 cassettes.

Removable dividers and lid with handle.  Holds up to 100 cassettes.

Removable dividers and lid with handle.  Holds up to 50 cassettes.

                                                            

Newcomer Supply Wright Stain, Unbuffered for Smears is 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.

 

STAINING PROCEDURE:

1. Prepare within an accepted time frame, a well-made blood smear or bone marrow smear/film per your laboratories protocol, with a focus on uniform cell distribution.
2. Allow slides to thoroughly air-dry prior to staining.
3. Place slides on a flat staining rack suspended over a sink.
4. Fix smear by flooding slide with methanol for 10-30 seconds.
5. Shake off excess methanol; flood each slide with 1 ml Wright Stain Solution for 3 to 5 minutes.

1. Filter Wright Stain Solution prior to use.
2. See Procedure Notes #1 and #2.

6. Retain Wright Stain Solution on slides, directly add 1 ml of Wright Buffer Solution, pH 6.8 to each slide; gently agitate to mix. Stain for an additional 6 to 10 minutes.
7. Wash well in distilled water; rinse thoroughly in running tap water.
8. Air-dry slides in a vertical position, then examine microscopically.
9. If coverslip is preferred, allow slides to air-dry 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. The timings provided in this procedure 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 may require a longer staining time. Bone marrow smears/films may also require a longer staining time.
3. The color range of the stained cells may vary depending upon the pH of the buffer and the pH of the rinse water used.

1. Alkalinity is indicated by red blood cells being blue-grey and white blood cells only blue.
2. Acidity is indicated by red blood cell 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. 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.

APPLICATION:

Newcomer Supply Wright-Giemsa, Romanowsky Stain for Smears is deemed the classic Wright-Giemsa stain for hematology. It is designed to demonstrate differential staining of cell types in peripheral blood smears and bone marrow smears/films as well as a method for detecting parasites, bacteria, and inclusion bodies.

A Romanowsky-type stain refers to a stain made from water-soluble eosin, methylene blue and methanol.  Wright-Giemsa stains, comprised of polychrome methylene blue, azure B and eosin Y dyes, are classified as Romanowsky stains.

 

METHOD:

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 staining procedure provided below.

 

STAINING PROCEDURE:

1. Prepare within an accepted time frame, a well-made blood smear or bone marrow smear/film per your laboratories protocol, with a focus on uniform cell distribution.
2. Allow smear to thoroughly air-dry prior to staining.
3. Fix smear in Alcohol, Methanol Anhydrous (Part 12236) for 3-5 minutes.
4. Air-dry slides in a vertical position.
5. Prepare Wright-Giemsa, Romanowsky Working Stain Solution; combine, mix well and filter if particulates are present.
   1. For thin smears:

Giemsa Stock Stain, Romanowsky                                20 ml

Wright Stain Buffer, pH 6.8                             20 ml

2. For thick smears:

Giemsa Stock Stain, Romanowsky                                  4 ml

Wright Stain Buffer, pH 6.8                             36 ml

6. Stain in Wright-Giemsa, Romanowsky Working Stain Solution for 30-45 minutes.
   1. See Procedure Notes #1 and #2.
7. Wash in distilled water.
8. Air-dry slides in a vertical position; examine microscopically.
9. If coverslip is preferred, air-dry slides and coverslip with compatible mounting medium.

 

RESULTS:                  

Erythrocytes	Orange-pink to rose
Platelets	Red to purple granules with light blue halo

 

Granulocytes

Neutrophils	Nucleus – Dark blue to violet
	Cytoplasm – Pink
	Granules – Purple to lilac
Eosinophils	Nucleus – Blue
	Granules – Orange to pink
Basophils	Nucleus – Deep blue to violet
	Granules – Deep blue to violet

 

Mononuclear Cells

Lymphocytes	Nuclei – Deep blue to violet
	Cytoplasm – Light blue
Monocytes	Nuclei – Light blue/purple
	Cytoplasm – Pale gray/blue
Mast cells	Nuclei – Deep blue to violet
	Granules – Deep blue-violet
Malarial parasites	Nucleus – Red chromatin dot
	Cytoplasm – Blue
Bacteria	Blue

 

PROCEDURE NOTES:

1. The timings provided in this procedure 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 may require a longer staining time. Bone marrow smears/films may also require a longer staining time.
3. The color range of the stained cells may vary depending upon the pH of the buffer and the pH of the rinse water used.
   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. Bauer, John D. Clinical Laboratory Methods. 9th ed. St. Louis: Mosby, 1982. 111-112.
2. Conn’s Biological Stains. Edited by Richard Horobin and John Kiernan. 10th ed. Oxford, UK: BIOS Scientific Publishers, 2002. 303-312.
3. Lillie, R. D., and Harold Fullmer. Histopathologic Technic and Practical Histochemistry. 4th ed. New York: McGraw-Hill, 1976. 744-748.
4. McPherson, Richard and Matthew Pincus. Henry’s Clinical Diagnosis and Management by Laboratory Methods. 22nd ed. Philadelphia:  Elsevier Saunders, 2011. 522-532.
5. Modifications developed by Newcomer Supply Laboratory.

 

SOLUTION:

 

GRAM, BROWN-HOPPS STAIN KIT INCLUDES:

		Part 9124A
Solution A:	Crystal Violet Stain 1%, Aqueous, Brown-Hopps	250 ml
Solution B:	Iodine, Gram, Aqueous	250 ml
Solution C:	Basic Fuchsin Stain 0.25%, Aqueous	250 ml
Solution D:	Gallego Solution	250 ml
Solution E:	Picric Acid-Acetone 0.05%	250 ml
Solution F:	Acetone-Xylene 1:1	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed with this kit are two complimentary unstained positive control slides to be used 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
Acetone, ACS	Part 10014

 

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

 

APPLICATION:

Newcomer Supply Gram, Brown-Hopps Stain Kit procedure, is a consistent and reliable Gram stain, that uses Gallego Solution to differentiate gram-negative and gram-positive bacteria.

 

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. Stain slides in Solution A: Crystal Violet Stain 1%, Aqueous, Brown-Hopps for 2 minutes.
   4. Rinse well in distilled water.
   5. Mordant in Solution B: Iodine, Gram, Aqueous for 5 minutes.
   6. Rinse well in distilled water.
   7. Blot excess water from slide; decolorize one slide at a time in Acetone, ACS (Part 10014) until blue stops running; 1-2 dips.
      1. 1. Sections should be very light gray in color.
   8. Rinse quickly in running tap water.
   9. Place in Solution C: Basic Fuchsin Stain 0.25%, Aqueous for 5 minutes.
   10. Rinse well in running tap water.
   11. Differentiate sections in Solution D: Gallego Solution for 5 minutes.
   12. Rinse in running tap water. Blot water off slide(s) but not to dryness.
       1. 1. Proceed with Steps #13 to #16 one slide at a time.
   13. Dip in Acetone, ACS (Part 10014); 1-2 quick dips.

1. 14. Dip in Solution E: Picric Acid-Acetone 0.05%; 3-10 dips.
   15. Dip in Solution F: Acetone-Xylene 1:1; 5 dips.
   16. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Gram-positive bacteria	Blue/violet
Gram-negative bacteria	Red
Nuclei	Red
Background tissue	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. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

       

REFERENCES:

1. 1. Brown, Robert C. and Howard C. Hopps. “Staining of Bacteria in Tissue Sections: A Reliable Gram Stain Method.” American Journal of Clinical Pathology 60.2 (1973): 234-240.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 222-224.
   3. Modifications developed by Newcomer Supply Laboratory.

 

85Get a HANDLE on your frozens!  Specimen chucks with a built-in handle for easy positioning & orientation.

Designed to work on most cryostats. 

• Leica models: 1510, 1800, 1850
• Sakura Cryo 3
• Microm Non Cooled Head models: HM500, HM505, HM525 & HM550

Get a HANDLE on your frozens!

Designed to work on Thermo Cryostat models: FSE, FE & SME

BIELSCHOWSKY, LESTER-KING MODIFIED STAIN KIT INCLUDES:      

		Part 9154A
Solution A:	Silver Nitrate 20%, Aqueous	250 ml
Solution B:	Ammonium Hydroxide 28-30%, ACS	100 ml
Solution C:	Developer	25 ml
Solution D:	Sodium Thiosulfate 5%, Aqueous	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed with this kit are two complimentary unstained positive control slides to be used 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 Bielschowsky, Lester King Modified Stain Kit procedure demonstrates nerve fibers, neurofibrils/tangles, senile plaques and axons.  This stain is instrumental in the diagnosis of Alzheimer’s disease and other neurological disorders.

 

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.

 

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. Preheat Coplin jar of Solution A: Silver Nitrate 20%, Aqueous in water bath to 37°C.
   4. Preheat two Coplin jars of distilled water in 37°C water bath.
      1. 1. Save for slide rinsing/holding in Steps #7 and #10.

 

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 preheated Solution A: Silver Nitrate 20%, Aqueous (Step #3) for 15 minutes.
      1. 1. See Procedure Note #5.
   7. Remove slides from Solution A: Silver Nitrate 20%, Aqueous and hold in 1^st reserved Coplin jar of preheated distilled water.
      1. 1. Save Silver Nitrate 20%, Aqueous for Step #8.
   8. Add Solution B: Ammonium Hydroxide 28-30%, ACS drop by drop into saved Silver Nitrate 20%, Aqueous, swirling until precipitate disappears. Do not go past this point. 
      1. 1. Approximately 10 ml of Ammonium Hydroxide 28-30%, ACS will be required.
   9. Place slides back in Silver Nitrate Solution with added Ammonium Hydroxide in water bath at 37°C for 10 minutes.
   10. Remove slides and hold in 2^nd reserved Coplin jar of distilled water.
       1. 1. Save Ammoniacal Silver Solution for Step #11.

1. 11. Add 1 drop of Solution C: Developer to the saved Ammoniacal Silver Solution with swirling motion.
   12. Return slides to Ammoniacal Silver Solution with added Developer, in 37°C water bath for 5-15 minutes; average time of 6 minutes.
       1. 1. Check slides microscopically at 3 minutes for development of neurons to dark brown.
          2. Then check at 1 minute intervals to avoid over-development.
   13. Rinse thoroughly in distilled water for 5 minutes.
   14. Place in Solution D: Sodium Thiosulfate 5%, Aqueous; 5 minutes.
   15. Rinse thoroughly in tap water.
   16. 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:

Senile plaques, neurofibrils/tangles	Dark brown to black
Neurons	Dark brown
White and gray matter	Yellowish brown
Nerve fibers, axons	Brown to black

 

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. Eight slides per 40 ml of Solution A: Silver Nitrate 20%, Aqueous is recommended for proper silver development.
   6. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 196-199.
   2. King, Lester. “The Impregnation of Neurofibrils.” Yale Journal of Biology and Medicine 14.1 (1941). 59-68.
   3. Modifications developed by Newcomer Supply Laboratory.

 

The CellSafe+ Biopsy Capsules are specifically designed to keep biopsy specimens safe during processing.  The CellSafe+ is made of two hinged interlocking frames with an integral mesh.  This extra fine mesh ensures a high level of specimen security, no tissue damage and low reagent carry over.

The capsule is closed in one direction to fully encapsulate the tissue.  The CellSafe+ can then be placed in a standard cassette for processing.  The tissue can be removed with greater ease and security when compared with conventional foam pads or paper wrap.

 

BENEFITS OF THE CELLSAFE+ BIPOSY CAPSULES:

• X-Ray transparent for breast work
• Blue mesh aids contrast and visualization of light colored biopsies
• Extra fine mesh provides up to 20 times less carry over of solvents than foam pads
• Zero tissue damage or artifacts associated with foam pads
• Substantial time savings when compared to wrapping biopsy in tissue paper

 

DIMENSIONS OF THE CELLSAFE+ BIOPSY CAPSULES:

• 28mm x 25mm x 5mm (H)

 

The Swingsette Biopsy Cassettes in QuickLoad Taped Stacks are taped cassettes to be used with Leica and Sakura Ink Jet printers.  These cassettes are similar to Swingsette Tissue Cassettes but are specially designed to hold biopsy specimens. Made from acetal, they keep specimens safely submerged and are resistant to the chemical reaction of most solvents used in histology laboratories.

 

PACKAGING OF THE SWINGSETTE BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

SWINGSETTE BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Designed to hold small biopsies securely during the embedding process.
• 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.
• Covers packaged separately.
• Also available without QuickLoad Taped Stacks (Part 51301) and in QuickLoad Sleeves (Part 51291).

 

The Swingsette in QuickLoad Taped Stacks is suited for the Leica and Sakura labelers. These cassettes will also load in those cassette labeling instruments in one simple operation.  Made from acetal they keep specimens safely submerged and are resistant to to the chemical action of most solvents used in histology laboratories.

 

PACKAGING OF THE SWINGSETTE TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

SWINGSETTE TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Efficient flow-through slots maximize fluid exchange and ensure proper drainage.
• Easy to assemble disposable cover with special hinge allowing the cassettes to be opened and closed as often as necessary.
• Covers packaged separately.
• Anterior writing area is at a 45° angle.
• Cassettes also available without QuickLoad Taped Stacks (Part 5130) and in QuickLoad Sleeves (Part 5129 ).

 

The Cassettes, Without Lids in QuickLoad Taped Stacks are made to be used with Leica and Sakura Ink Jet printers. The cassettes are made from acetal and keep specimens safely submerged in liquid and are resistant to the chemical reaction of most solvents used in histology laboratories.

PACKAGING OF THE CASSETTES, WITHOUT LIDS IN QUICKLOAD TAPED STACKS:

• • 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

CASSETTES, WITHOUT LIDS IN QUICKLOAD TAPED STACKS:

• • Efficient flow-through slots maximize fluid exchange and ensure proper reagent drainage.
  • Lids sold separately.
  • Each case contains 50 stacks with 40 cassettes per stack.
  • The anterior writing area is at a 45º angle.

 

Histosette II Biopsy Cassettes are similar to Histosette II Tissue Cassettes but specially designed to hold biopsy specimens.  The Histosette II Biopsy cassettes in QuickLoad Stacks are made to be used with Leica and Sakura Ink Jet printers.  These cassettes are made from acetal and keep specimens safely submerged in liquid and are resistant to the chemical reaction of most solvents used in histology laboratories.

 

PACKAGING OF THE HISTOSETTE II BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

HISTOSETTE II BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Designed for biopsy specimens.
• Efficient flow-through slots maximize fluid exchange and ensure proper reagent drainage with 1 mm square openings.
• Pre-attached lid with front hinge and opens from the back of the cassette. The closed lids can be opened many times, always relocking securely.
• The anterior printing area is at a 45° angle and offers an unobstructed view of the writing surface.

 

The Histosette II cassettes in QuickLoad Taped Stacks are specially made to be used with Leica and Sakura Ink Jet printers. These cassettes are made from acetal and keep specimens safely submerged in liquid and are resistant to the chemical action of most solvents used in histology laboratories.

 

PACKAGING OF THE HISTOSETTE II TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

HISTOSETTE II TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Efficient flow-through slots maximize fluid exchange and ensure proper reagent drainage.
• Back mounted locking device securely holds the lid in place.
• Front hinge design allows single-handed cassette manipulation.
• Anterior writing area is at a 45° angle.

 

The Micromesh Biopsy Cassettes in QuickLoad Taped Stacks are taped cassettes to be used with Leica and Sakura Ink Jet printers.  The cassettes are made from acetal and designed to hold biopsy specimens. These patented cassettes keep specimens safely submerged in liquid and are totally resistant to the chemical action of histological solvents. The MICROMESH™ mesh ensures efficient fluid exchange and drainage.

 

PACKAGING OF THE MICROMESH BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

MICROMESH BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Designed for biopsy specimens.
• No biopsy pad necessary – 1,676 square openings (0.38 mm) allowing for a greatly improved fluid exchange.
• Large anterior and posterior slots in both cassette and cover ensure that the Micromesh Biopsy Cassette will sink rapidly.
• A large square compartment with a side measuring 27mm is perfect even for needle biopsies.
• One-piece integral lid that does not protrude above the cassette eliminates the need for separate steel lids. They 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 45º angle.
• Cassettes also available without QuickLoad Taped Stacks (Part 5120) and in QuickLoad Sleeves (Part 5123).

 

The Slimsette Biopsy Cassettes in QuickLoad Taped Stacks are taped cassettes to be used with Leica and Sakura Ink Jet printers.  The cassettes are similar to Slimsette Tissue Cassettes but are specially designed to hold biopsy specimens. Made from acetal, they keep specimens safely submerged and are resistant to the chemical reaction of most solvents used in histology laboratories.

 

PACKAGING OF THE SLIMSETTE BIOPSY TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

SLIMSETTE BIOPSY TISSUE PROCESSING & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• Designed for biopsy specimens.
• Efficient fluid exchange and drainage with 392 openings each measuring 1 x 1 mm.
• Recessed cover is pre-attached and easy to remove.
• Lids can be opened and closed as often as necessary and relock securely without danger of specimen loss.
• Anterior writing area is at a 45° angle.
• Also available without QuickLoad Taped Stacks (Part 5127) and 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

 

The Slimsette in QuickLoad Taped Stacks are taped cassettes to be used with Leica and Sakura Ink Jet printers. Made from acetal, they keep specimens safely submerged and are resistant to the chemical action of most solvents used in histology laboratories.

 

PACKAGING OF THE SLIMSETTE TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 40 cassettes/stack; 50 stacks/case; 2,000 cassettes/case.

 

SLIMSETTE TISSUE & EMBEDDING CASSETTES IN QUICKLOAD TAPED STACKS:

• 114 openings each measuring 1 x 5mm maximize fluid exchange and ensure proper reagent drainage.
• The cover does not protrude above the cassette, a great space saving feature.
• The one-piece disposable plastic cover is pre-attached on each cassette and eliminates the need for reusable steel lids.
• The cover can be opened and closed as often as necessary and will relock, reducing the possibility of specimen loss.
• Anterior writing area is at a 45° angle.
• Cassettes also available without QuickLoad Taped Stacks (Part 5126) and in QuickLoad Sleeves (Part 5121).

(use: Trichrome, Gomori One-Step.)

STEINER-STEINER MODIFIED SILVER STAIN KIT INCLUDES:                                                                                   

		Part 9171A
Solution A:	Uranyl Nitrate 1%, Aqueous	250 ml
Solution B:	Silver Nitrate 1%, Aqueous	250 ml
Solution C:	Gum Mastic 2.5%, Alcoholic	175 ml x 2
Ingredient D:	Hydroquinone, Powder	5 grams
Mini Sampling Spoon

 

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 modifications)

 

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

 

APPLICATION:

Newcomer Supply Steiner-Steiner Modified Silver Stain Kit procedure, with included microwave modifications, is a silver technique effective for the demonstration of spirochetes, Helicobacter pylori, Legionella pneumophila, other nonfilamentous bacteria and fungus.

 

METHOD:

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

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/slide in oven.
   2. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.
   3. Preheat Solution A: Uranyl Nitrate 1%, Aqueous to 60°C in a water bath. Save for Step #9.
   4. Preheat Solution B: Silver Nitrate 1%, Aqueous to 60°C in a water bath. Save for Step #11.
   5. Prepare Hydroquinone Solution; combine and mix well.
      1. 1. Ingredient D: Hydroquinone, Powder                5 gm       (or one rounded scoop with reusable mini sampling spoon)
         2. Distilled Water                 25 ml

1. 6. Prepare fresh Reducing Solution by combining in order listed.
      1. 1. Hydroquinone Solution (Step #5)                 25 ml
         2. Solution C: Gum Mastic 2.5%, Alcoholic         15 ml
         3. Solution B: Silver Nitrate 1%, Aqueous   6 ml
         4. Solution will turn milky white after addition of Gum Mastic.
         5. Preheat solution in 45°C water bath. Save for Step #15.
   7. Do not preheat solutions if using Microwave Modifications.

 

STAINING PROCEDURE: 

1. 8. 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 Note #3.
   9. Sensitize in preheated Solution A: Uranyl Nitrate 1%, Aqueous (Step #3) for 10 minutes in a 60°C water bath.

        Microwave Modification: See Procedure Note #4 (page 2).

1. 1. 1. Place slides in a plastic Coplin jar with Solution A: Uranyl Nitrate 1%, Aqueous. Microwave for 1 minute at 70°C.
2. 10. Rinse well in several changes of distilled water.
   11. Place slides in preheated Solution B: Silver Nitrate 1%, Aqueous (Step #4) and incubate in a 60°C water bath for 15 minutes.

        Microwave Modification:

1. 1. 1. 1. Place slides in a plastic Coplin jar with Solution B: Silver Nitrate 1%, Aqueous. Microwave for 1 minute at 70°C.
         2. Remove from microwave, cover and let sit for 1 minute.

1. 12. Rinse well in several changes of distilled water.
       1. 1. Excessive rinsing may cause nuclei to pick up silver.

1. 13. Dip 5 times in two changes of fresh 95% and 100% ethyl alcohols.
   14. Place in Solution C: Gum Mastic 2.5%, Alcoholic for 3 minutes.
   15. Place slides in preheated Reducing Solution (Step #6) in 45°C water bath for 10-30 minutes with frequent agitation. Examine microscopically after 10 minutes of incubation.
       1. 1. Check microscopically by dipping slide in 100% alcohol.
          2. Review for desired staining results.
          3. If necessary, return to warm solution; check every 2-5 minutes until desired results are achieved.

        Microwave Modification: 

1. 1. 1. 1. 1. Place slides in a plastic Coplin jar with Reducing Solution. Microwave for 1 minute at 70°C. Remove from microwave.
            2. Pipette solution twice with plastic pipette to evenly distribute heated solution.
            3. Cover and let sit for 1 minute.
            4. Check microscopically by dipping slide in 100% alcohol.
            5. Review for desired staining results.
            6. If necessary, return to warm solution, check every 1 minute until desired results are achieved.
   16. Directly dehydrate in two changes of 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Spirochetes	Dark brown to black
Helicobacter pylori	Dark brown to black
Legionella pneumophila	Dark brown to black
Nonfilamentous bacteria and fungus	Dark brown to black
Background	Golden brown

 

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. 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.
   5. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Garvey, Winsome. “Some Favorite Silver Stains.” The Journal of Histotechnology 3 (1996): 269-278.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 218-219.
   3. Steiner, Gabriel and Grete Steiner. “New Simple Silver Stain for Demonstration of Bacteria, Spirochetes and Fungi in Sections of Paraffin Embedded Tissue Blocks.” Journal of Laboratory Clinical Medicine 29 (1944). 868-871.
   4. Swisher, Billie. “Modified Steiner Procedure for Microwave Staining of Spirochetes and Nonfilamentous Bacteria.” The Journal of Histotechnology 10.4 (1987): 241-243.
   5. Modifications developed by Newcomer Supply Laboratory.

SET INCLUDES:

		Part 12218A	Part 12218B
Solution A:	Luxol Fast Blue Stain 0.1%, Alcoholic	500 ml	1000 ml
Solution B:	Lithium Carbonate, Saturated Aqueous	500 ml	1000 ml

 

Additionally Needed For LFB/H&E Stain:

Luxol Fast Blue (LFB) Control Slides	Part 4407
Hematoxylin Stain, Harris Modified	Part 1201
Acid Alcohol 1%	Part 10011
Eosin Y Working Solution	Part 1072
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Alcohol, Ethyl Denatured, 70%	Part 10844
Coplin Jar, Plastic	Part 5184 (for microwave modification)

 

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

 

APPLICATION:

Newcomer Supply Luxol Fast Blue (LFB) Stain Set, with included microwave modification, is for the demonstration of myelin in central nervous system and peripheral nerve tissues.

 

The LFB Stain Set is flexible and can be used as a stand-alone without additional stain/counterstain or combined with options such as:

• • LFB/Hematoxylin or LFB/Hematoxylin and Eosin (H&E)
  • LFB/PAS or LFB/PAS/Hematoxylin
  • LFB/Cresyl Violet
  • LFB/Nuclear Fast Red
  • LFB/Silver Nitrate

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090) 
Technique: Paraffin sections cut at 8-10 microns on adhesive slides
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. Prepare Working Lithium Carbonate 0.05%; combine and mix well;
      1. 1. Solution B: Lithium Carbonate, Saturated Aqueous 5 ml
         2. Distilled Water                     95 ml

 

LFB/H&E 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.
      1. 1. Stop at 95% ethyl alcohol; no distilled water rinse.
         2. See Procedure Notes #1 and #2.
   4. Incubate in Solution A: Luxol Fast Blue Stain 0.1%, Alcoholic for 2 hours at 60°C or overnight at 37°C; cover tightly.
      1. 1. To enhance stain, add 0.4 ml of Acetic Acid, Glacial, ACS (Part 10010) to 40 ml of Solution A: Luxol Fast Blue Stain 0.1%, Alcoholic before use.

Microwave Modification: See Procedure Note #3.

1. 1. 1. 1. Place slides in a plastic Coplin jar (Part 5184) with Solution A: Luxol Fast Blue Stain 0.1%, Alcoholic. Microwave for 10 minutes at 70°C.

1. 5. Rinse slides quickly in 95% ethyl alcohol; 2-3 dips.
   6. Rinse in distilled water.
   7. Differentiate slides individually in Working Lithium Carbonate 0.05% (Step #2) for 10-15 seconds with agitation until gray matter and white matter are colorless and contrast with stained tissue.
   8. Further differentiate in 70% ethyl alcohol, until gray and white matter can be distinguished. Do not over differentiate.
   9. Rinse in distilled water.
   10. Check slides microscopically. Continue if additional differentiation is needed. Otherwise proceed directly to Step #12.
       1. 1. One dip in Lithium Carbonate 0.05%, Aqueous (Step #2).
          2. Dip in two changes of 70% ethyl alcohol until green/blue white matter sharply contrasts with colorless gray matter.

1. 11. Rinse thoroughly in distilled water.
   12. Stain with Hematoxylin Stain, Harris Modified (Part 1201) for 1-5 minutes, depending on preference of intensity.
   13. Wash in running tap water for 3 minutes.
   14. Differentiate quickly in Acid Alcohol 1% (Part 10011); 3 dips.
   15. Wash well in running tap water.
   16. Blue in Solution B: Lithium Carbonate, Saturated Aqueous.
   17. Wash well in running tap water.
   18. Counterstain in Eosin Y Working Solution (Part 1072) for 30 seconds to 3 minutes, depending on preference of intensity.
   19. Dehydrate in two changes of 95% 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:

Myelin (white matter)	Blue to blue/green
Gray matter and cytoplasm	Shades of pink to red
Nuclei	Dark 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. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 206-211.
   2. Klüver, Heinrich and Elizabeth Barrera. “A Method for the Combined Staining of Cells and Fibers in the Nervous System.” Journal of Neuropathology and Experimental Neurology 12.4 (1953): 400-403.
   3. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 494-495.
   4. Modifications developed by Newcomer Supply Laboratory.

 

RETICULUM, GORDON & SWEETS STAIN KIT INCLUDES:

 

The MAS-GP adhesive microscope slide surface has an enhanced chemistry coating that is bound on the glass surface and creates a very strong adhesive and hydrophilic surface for tissue sections and cells to adhere to.

The MAS-GP adhesive microscope slide can be widely used for pathology, cytology and immunohistochemistry applications.

The hydrophilic surface of the microscope slide ensures faster draining of the water between the glass surface and the tissue section compared to adhesive slides with a hydrophobic surface.  This can help minimize any water being trapped between the tissue section and the microscope slide when the tissue section is being picked up from the water bath.  Minimizing the trapped water can increase the area of the tissue section being in direct contact with the adhesive coating on the microscope glass and maximizing the adhesive bond between the tissue section and microscope glass.  Thus, creating stronger adhesive properties of the MAS-GP microscope slide.

 

ADHESIVE MECHANISM OF THE MAS-GP ADEHESIVE MICROSCOPE SLIDE:

A coating is applied to the microscope slide surface by a covalent bond of the enhanced aminosilane and OH groups to produce a strong positive charge on the slide.  Tissue sections, having a negative charge, will be attracted to the surface of the glass and adhere to it.  The MAS-GP adhesive microscope slide has a high tissue section & cell adhesive strength.  The microscope slide is highly durable to thermal treatment particularly with microwave and autoclave.

 

RECOMMENDED USE:

• Manual & automated IHC applications
• IF (Immuno Fluorescence)
• HIER (Heat Induced Epitope Retrieval) pH 6 and pH 9 protocols
• Routine histology
• Frozen sectioning

 

SPECIFICATIONS:

• Glass Type:  High Clarity, Low Fluorescence
• Size: 75 x 25mm, 1mm thick
• Surface Wettability:  Hydrophilic
• Edge Treatment:  45° Corner
• Packaged 1,000 slides/case
• Available in White, Blue, Green, Yellow and Pink

 

This slide has remarkably improved adhesion to tissue sections and cells due to the amino groups that bind to the tissue section have a much higher density on the slide than could be achieved by either poly-L-lysine or silane coatings.

 

RECOMMENDED USE:

The MAS adhesive coated slides are excellent for the most challenging tissues and applications:

• Demanding IHC with antigen retrieval
• Veterinary Animal Tissue
• Difficult Tissue (Dry, Fatty or Hard Tissue such as bones, nails, breast, eyes & brain)

 

SPECIFICATIONS:

• Size: 75 x 25, 1mm thick
• Surface Wettability: Hydrophilic
• Packaged 100 slides/box, 10 boxes/case
• Available color: White
• Storage: 15 – 30° C

 

 

The TruBond 380 adhesive microscope slides have an enhanced chemistry coating which creates an incredible slide to work with!

The advanced technology in manufacturing the adhesive surface of the microscope slide produces incredible adhesion to the tissue section even in the most demanding applications.  The surface of the TruBond 380 microscope slide is also hydrophilic producing two very notable features:

• Allows to refloat the tissue in the water bath for repositioning the tissue section
• Allows the microscope slide to be used on all automated IHC platforms

 

The hydrophilic surface of the microscope slide also ensures faster draining of the water between the glass surface and the tissue section compared to adhesive slides with a hydrophobic surface.  This can help minimize any water being trapped between the tissue section and the microscope slide when the tissue section is being picked up from the water bath.  Minimizing the trapped water can increase the area of the tissue section being in direct contact with the adhesive coating on the microscope glass and maximizing the adhesive bond between the tissue section and microscope glass.  Thus, creating stronger adhesive properties of the TruBond 380 microscope slide.

Extensive testing has shown the TruBond 380 Adhesive Microscope Slides outperformed commonly used adhesive microscope slides. They are especially useful in preventing tissue loss when performing high temperature antigen retrieval or enzyme digestion, and are ideal for use in immunostaining of formalin fixed paraffin embedded (FFPE) and frozen tissue where stronger adhesion microscope slides are required. 

 

RECOMMENDED USE:  

• HIER processes when tissue type require extra strong adhesion
• Epitope enhancement and DNA probe procedures
• Autopsy and brain sectioning
• Research applications require extra strong adhesion

 

SPECIFICATIONS:

• Glass Type: High Clarity, Low Fluorescence
• Size: 75 x 25mm, 1mm thick
• Surface Wettability: Hydrophilic
• Edge Treatment: 45° Corner
• Packaged 1,000 slides/case
• Available in White, Blue, Green, Yellow and Pink

 

 

The APS adhesive microscope slide surface is coated with an aminosilane coating that is bound on the glass surface and creates a very strong adhesive surface for tissue sections and cells to adhere to.

The APS adhesive microscope slide can be widely used for pathology, cytology and immunohistochemistry applications.

 

ADHESIVE MECHANISM OF THE APS ADHESIVE MICROSCOPE SLIDES:

A coating is applied to the microscope slide surface by a covalent bond of the aminosilane and OH groups to produce a strong positive charge on the slide.  Tissue sections, having a negative charge, will be attracted to the surface of the glass and adhere to it.  The APS adhesive microscope slide has a high tissue section & cell adhesive strength.  The microscope slide is highly durable to thermal treatment particularly with microwave and autoclave.

 

RECOMMENDED USE:

• Routine histology
• Manual & automated IHC
• IF (Immunofluorescence)
• HIER (Heat Induced Epitope Retrieval)
• Epitope enhancement
• DNA procedures

 

SPECIFICATIONS:

• Glass Type: High Clarity, Low Fluorescence
• Size: 75 x 25mm, 1mm thick
• Surface Wettability: Hydrophobic
• Edge Treatment Options: 45° Corners and 90° Corners
• Packaged 1,000 slides per case
• Available colors: White, Blue, Green, Yellow and Pink

 

 

The NewSilane adhesive coated slides are a very strong adhesive coated slide!  Use for “tricky”, routine or IHC applications.  These adhesive coated slides are an excellent addition to any pathology lab that works with challenging tissue(s) or demanding procedures.

 

RECOMMENDED USE:

The NewSilane Adhesive Coated Slides are recommend for use for procedures with:

• Difficult tissues combined with long incubations
• Extra procedural steps
• Extended antigen retrieval

 

SPECIFICATIONS:

• No word imprint on slide
• Size: 75 x 25mm, 1mm thick
• Edge Treatment: 90° Corner
• Unit: Approx. 72 slides/box, 1,440 slides/case
• Available color: White

• Made of polypropylene
• Can hold 10 slides (1″x3″)
• Interior is grooved to hold slides vertically
• Domed and shallow thread screw cap

 

 

 

FONTANA MASSON STAIN KIT INCLUDES: 

		Part 9105A
Solution A:	Silver Nitrate 10%, Aqueous	250 ml
Solution B:	Ammonium Hydroxide 28-30%, ACS	250 ml
Solution C:	Gold Chloride 0.2%, Aqueous	250 ml
Solution D:	Sodium Thiosulfate 5%, Aqueous	250 ml
Solution E:	Nuclear Fast Red Stain, Kernechtrot	250 ml

 

COMPLIMENTARY POSITIVE CONTROL SLIDES: Enclosed with this kit are two complimentary unstained positive control slides to be used 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 Fontana Masson Stain Kit procedure demonstrates argentaffin substances such as melanin, argentaffin granules of carcinoid tumors, and some neurosecretory granules. This stain is not specific for melanin and argentaffin granules and other reducing substances, such as formalin pigment, will also give a positive reaction.

 

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. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.
   3. Prepare Fontana Working Solution (diamine silver) in an acid cleaned Erlenmeyer flask:
      1. 1. Solution A: Silver Nitrate 10%, Aqueous; 25 ml    
         2. Add Solution B: Ammonium Hydroxide 28-30%, ACS drop by drop, mix with swirling motion until solution clouds, then clears. Use caution to not add excess Ammonium Hydroxide.
         3. Add more Solution A: Silver Nitrate 10%, Aqueous drop by drop until clear solution becomes slightly cloudy.
         4. Let solution stand for 2-4 hours before use.
         5. For use; after standing, filter the solution. Combine 20 ml of this filtered diamine silver solution with 40 ml of distilled water; 60 ml total.

 

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 #3 and #4.
   5. Immerse slides in Fontana Silver Working Solution (Step #3) in a 45°C-60°C water bath for 1 hour.

1. 6. Check slides microscopically; remove control, rinse in warm distilled water. Confirm reaction is complete when granules are dark brown and background is colorless.
      1. 1. Return to heated Fontana Silver Working Solution for longer incubation if indicated.
   7. Rinse well in three changes of distilled water.
   8. Immerse in Solution C: Gold Chloride 0.2%, Aqueous; 10 minutes.
   9. Rinse well in distilled water.
   10. Place in Solution D: Sodium Thiosulfate 5%, Aqueous; 5 minutes.
   11. Rinse well in distilled water.
   12. Counterstain in Solution E: Nuclear Fast Red Stain, Kernechtrot for 5 minutes.
       1. 1. Shake solution well before use; do not filter.
   13. Rinse well in distilled water.
       1. 1. See Procedure Note #5.
   14. 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.

 

RESULTS:

Melanin and argentaffin granules	Black
Nuclei	Pink-red

 

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. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
   6. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 286-287.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 276-277.
   3. Modifications developed by Newcomer Supply Laboratory.

Designed specifically for biopsy processing by using <1mm square openings in the lid and base surfaces for max fluid flow & specimen retention. Covers detached and packaged separately.

We got the PAs involved again in designing this beauty! We took the basic design of the 15cm ruler and stretched it out to 30cm. All metric and the numbers on the top and bottom both start on the left to accommodate both the top and bottom readers. For those of you that flip the ruler for the large tissue section measurements, it’s not only flush cut at zero, but also at 30cm. Not to be out done by the 15cm ruler, this one is also printed on both sides!

Fluoro-Gel is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. The fluorescence is retained during prolonged storage at 4ºC in the dark. This mounting medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC.

 

FLUORO-GEL MOUNTING MEDIUM APPLICATIONS:

• Immunofluorescence, confocal microscopy.

 

FLUORO-GEL MOUNTING MEDIUM PROCEDURE:

1. Bring the vial to room temperature.
2. Rinse slide to be mounted with DISTILLED WATER OR DEIONIZED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light.
3. Turn the vial upside down and open the dropper to remove any air bubbles.
4. Apply 3-4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting slide back and forth or spread evenly with a 0.2 ml plastic pipette tip making sure the tissue is not touched. Excess mounting medium can be removed by touching the edges of slide against paper towel.
5. Let stand at room temperature for about 5 minutes.
6. Apply coverslip carefully avoiding air bubbles.
7. The specimen is ready for visualization under a microscope.
8. One can seal the edges of coverslip with nail polish, any organic mounting medium or our Limonene mounting medium. If a coverslip is not used air bubbles will appear in a few days.
9. For long term storage it is recommended that the slide be stored in the dark at 2-8°C.

 

REMOVAL OF COVERSLIP: 

Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slides several times with warm water to remove all mounting medium. The slide can be remounted again.

 

CLEAR MOUNT MOUNTING MEDIUM APPLICATIONS:

• Mounting of IHC slides that cannot be dehydrated with organic solvents.

 

CLEAR MOUNT INSTRUCTIONS:

1. Bring the vial to room temperature.
2. Rinse slide to be mounted with DISTILLED OR DEIONIZED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface.
3. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 3-4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting the slide back and forth or spread evenly with a 0.2 ml plastic pipette tip making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide against a paper towel.
4. Let stand at room temperature for about 1 to 2 hours. Or the slides can be heated at 37-40°C for 40-60 minutes. The slides are ready for visualization under a microscope.
5. To visualize slide under microscope with 4X, 10X, 20X, or 40X lens coversilp is NOT required.

 

HANDLING AND STORAGE OF CLEAR MOUNT:

This product is stored at room temperature, however, for long term storage 2-8°C is recommended. The pH of this product is more stable at 2-8ºC. DO NOT FREEZE.

 

REMOVAL OF COVERSLIP:

Soak the slide in warm (37°C) distilled or deionized water for 5-10 minutes until the mounting medium is dissolved. Rinse slide with warm water several times to remove all mounting medium; the slide can be remounted again if required.

The Limonene-Mount mounting media is good for preserving tissues and cell smears that can be dehydrated with organic solvents in Immunohistochemistry (IHC). This Organo mounting media is also suitable with alkaline phosphatase chromogens, an organic solvent resistant Super Fast Red. It is an excellent choice for mounting H&E stained slides.

 

LIMONENE-MOUNT MOUNTING MEDIA APPLICATIONS:

Limonene-Mount mounting media is intended for mounting tissues, cell smears in Immunohistochemistry for chromogens that are resistant to organic solvents, and H&E (hematoxylin and eosin) staining.

 

LIMONENE-MOUNT INSTRUCTIONS FOR USE:

1. Dehydrate tissue or cell smear slides with dehydrating agents e.g. 30 to 100% alcohol followed by xylene, toluene or any other dehydrating reagents.
2. Add 2-3 drops of Organo mounting media. Apply coverslip carefully, avoiding air bubbles.
3. Visualize slide under a microscope. The slides can be dried by leaving at room temperature overnight or by heating at 37-40ºC for several hours.

 

HANDLING AND STORAGE OF MOUNTING MEDIA:

Room temperature.

SOLUTION:

	500 ml	1 Liter	1 Gallon
Wright Stain, Modified	Part 1421A	Part 1421B	Part 1421C

 

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, Modified for Smears is a concentrated Wright’s formula for differential staining of cell types in peripheral blood smears and bone marrow smears/films. This procedure is applicable for either hand or automated staining processes.

 

METHOD:

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

 

PRESTAINING PREPARTION:

1. 1. Prepare a well-made blood smear or bone marrow smear/film with a focus on uniform cell distribution.
   2. Allow slides to thoroughly air-dry prior to staining.
   3. Filter Wright Stain, Modified 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, Modified for 3-5 minutes.
      1. 1. See Procedure Notes #1 and #2.
   8. Retain Wright Stain, Modified 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 times.
   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.

SOLUTIONS:

	500 ml	1 Liter	1 Gallon
EDTA Buffer 0.001M, pH 8.0	Part 1056A	Part 1056B	Part 1056C
Citrate Buffer 0.01M, pH 6.0	Part 10355A	Part 10355B	Part 10355C

 

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

 

APPLICATION:

Newcomer Supply Buffer Solutions for Epitope Retrieval procedure provides a choice of two ready-to-use buffers for antigen retrieval. The majority of epitopes/antigens are masked in formalin fixed paraffin embedded (FFPE) tissues.  Antigen retrieval methods improve antibody binding by de-masking the FFPE chemical modification of epitopes through heat induced epitope retrieval (HIER) procedures when performed prior to immunohistochemical (IHC) staining.

No retrieval buffer is optimal for all tissue antigens. The choice of buffer will depend upon the suggested retrieval buffer specific to an individual antibody.  Refer to each antibody datasheet for recommended chemical composition and pH value of retrieval buffer.

• • Part 1056: EDTA Buffer 0.001M, pH 8.0 is an alkaline buffer optimal for use with primary antibodies that require an EDTA buffer at a higher pH for HIER.
  • Part 10355: Citrate Buffer 0.01M, pH 6.0 is an acidic buffer optimal for use with primary antibodies that require a citrate buffer at a lower pH for HIER.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique: Paraffin sections on adhesive slides
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

 

EPITOPE RETRIEVAL PROCEDURE:

1. 1. Choose a HIER procedure that suits the laboratory and anticipated workload.
      1. 1. Instrumentation and methods for HIER include but are not limited to; microwave, pressure cooker and steamer methods.
   2. Validate instrumentation according to manufacturer’s suggested instructions for antigen retrieval methods.
   3. After validation of instrumentation and methodology; 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. Proceed with a validated method of HIER per established protocol implementing either EDTA Buffer 0.001M, pH 8.0 (Part 1056) or Citrate Buffer 0.01M, pH 6.0 (Part 10355).
   5. After completion of HIER, allow sufficient time for slides to cool before proceeding with IHC protocol.

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out 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. 442-445, 458-459.
   2. Shi, Shan-Rong, Richard J. Cote, Lillian L. Young and Clive R. Taylor. “Antigen Retrieval Immunohistochemistry: Practice and Development.” The Journal of Histotechnology 20.2 (1997): 145-154.
   3. Tacha, David and Maria Teixeira. “History and Overview of Antigen Retrieval: Methodologies and Critical Aspects.” The Journal of Histotechnology 25.4 (2002): 237-242.
   4. Modifications developed by Newcomer Supply Laboratory.

PRODUCT IMPROVEMENT NOTICE:  STORAGE RECOMMENDATION IS NOW 15-30°C

SOLUTIONS: 

 

	30 ml vial, 15 ml fill (50/cs)	20 ml vial, 10 ml fill (25/cs)
Michel Transport Medium Vial	Part 12423C	Part 12423E

PRODUCT IMPROVEMENT NOTICE:  STORAGE RECOMMENDATION IS NOW 15-30°C

SOLUTIONS: 

 

	30 ml vial, 15 ml fill (50/cs)	20 ml vial, 10 ml fill (25/cs)
Michel Transport Medium Vial	Part 12423C	Part 12423E

PRODUCT IMPROVEMENT NOTICE:  STORAGE RECOMMENDATION IS NOW 15-30°C

SOLUTIONS: 

 

	30 ml vial, 15 ml fill (50/cs)	20 ml vial, 10 ml fill (25/cs)
Michel Transport Medium Vial	Part 12423C	Part 12423E

GRAM, BROWN-BRENN MODIFIED STAIN KIT INCLUDES:

		Part 9123A
Solution A:	Crystal Violet-Oxalate Stain, Alcoholic	250 ml
Solution B:	Iodine, Gram, Aqueous	250 ml
Solution C:	Acetone-Alcohol 1:1	250 ml
Solution D:	Basic Fuchsin Stain 0.25%, Aqueous	250 ml
Solution E:	Tartrazine Stain 0.25%, Acetic 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 Gram, Brown-Brenn Modified Stain Kit is a simple and rapid procedure for differential staining of gram-positive and gram-negative bacteria with a tartrazine counterstain.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns and smears.
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 may contain extra volumes.

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Filter Solution A: Crystal Violet-Oxalate Stain, Alcoholic.

 

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 freshly filtered Solution A: Crystal Violet-Oxalate Stain, Alcoholic (Step #2) for 1 minute.
   5. Rinse well in distilled water.
   6. Mordant in Solution B: Iodine, Gram, Aqueous for 1 minute.
   7. Rinse well in distilled water, removing excess iodine.
   8. Decolorize in Solution C: Acetone-Alcohol 1:1 until blue stops running; 7-10 dips.
   9. Rinse well in distilled water.
   10. Place in Solution D: Basic Fuchsin Stain 0.25%, Aqueous for 90 seconds.
   11. Rinse well in distilled water.
   12. Dip once in Solution C: Acetone-Alcohol 1:1.
   13. Counterstain in Solution E: Tartrazine Stain 0.25%, Acetic Aqueous for 5-15 seconds.

1. 14. Rinse well in distilled water.
   15. Dehydrate in two changes of 100% ethyl alcohol, 5 dips each. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.
       1. 1. Do not use 95% alcohol in the dehydration step.

 

RESULTS:

Gram-positive bacteria	Blue
Gram-negative bacteria	Red
Nuclei	Red
Background tissue	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. 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. 312-313.
   2. Brown, J.H. and L. Brenn. “A Method for the Differential Staining of Gram Positive and Gram Negative Bacteria in Tissue Sections.”Bulletin of The Johns Hopkins 48.2 (1931): 69-73.
   3. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 188-189.
   4. Modifications developed by Newcomer Supply Laboratory.

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

(use: Masson & Gomori Trichrome & May-Grunwald Giemsa.)

Due to commodity market volatility the price of this product may change.

(use: Bielschowsky Stain)

(use: Von Kossa Calcium Stain, Gomori Basement Membrane & Gomori Methenamine Silver for Urates.)

(use: Original Brown & Brenn Gram Stain, and stock solution for Wolbach Giemsa.)

(use: Grocott Methenamine Silver GMS Stain.)

 

• Shelf Life is 4 years from date of manufacture.

 

(use: Rhodanine Stain for Copper.)

(use: Standard Grocott Methenamine Silver, GMS procedure.)

(use: Gomori Methenamine-Silver for Nitrates.)

CI 75100

• Shelf Life is 4 years from date of manufacture.

 

(use: Working solution for GMS.)

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