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

 

METHOD:

Fixation:

1. Bone Marrow:  Bone marrow clot a minimum of 2 hours, for bone marrow biopsy a minimum of 3 hours.
2. Lymph Nodes and Small Biopsies: A minimum of 4 hours.
   1. Small nodes (5 mm or less) should be halved. Dissect larges pieces to maximum thickness of 3 mm.
   2. To facilitate cutting, place tissue in AZF Fixative for 1 hour to firm; then trim to 2-3 mm.

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

 

FIXATION PROCEDURE:

1. Place fresh tissue specimen in AZF Fixative as soon as possible after surgical excision.
   1. See Procedure Note #1.
2. Hold tissue in AZF Fixative until ready to process or maximum of 72 hours.
   1. See Procedure Note #2.
3. Wash AZF fixed tissue thoroughly in tap water for minimum of 10 minutes to remove residual zinc.
4. Decalcify bone marrow specimen as needed.
   1. See Procedure Notes #3 & #4.
5. Place on tissue processor in Formalin 10%, Phosphate Buffered (Part 1090) fixation step.

 

PROCEDURE NOTES:

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. After maximum fixation, wash tissue in running tap water a minimum of 10 minutes.
   1. Transfer AZF fixed tissue to Formalin 10%, Phosphate Buffered for long-term storage purposes.
3. Nitric acid solutions are not recommended for decalcification following AZF fixation.
4. Acetic acid in AZF Fixative may pre-start the decalcification process, decreasing overall decalcification time.
5. Zinc chloride is corrosive, do not discard un-neutralized AZF Fixative solutions down the drain.
6. Neutralize AZF Fixative with sodium carbonate or sodium bicarbonate to precipitate zinc at pH 7.0-8.0. Separate solids from liquid; dispose of according to local/state regulations.
   1. Approximately 100 grams of sodium bicarbonate will neutralize/precipitate zinc from 1 liter of AZF Fixative.

 

REFERENCES:

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.

Newcomer Supply PAP Pen Liquid Blocker, hydrophobic slide markers for staining procedures, 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 surface tension ensures the amount of reagent needed for sufficient reaction is reduced and retained for complete tissue section coverage. Multiple specimens can be separated by drawn circles or lines applied to the same slide.

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. The PAP Pen Liquid Blocker, Mini provides a finer pen tip for drawing a thinner barrier film.

 

METHOD:

Technique:  Paraffin, frozen sections and smears

• Manual staining for:
  • Immunohistochemistry (IHC) procedures
  • Immunofluorescence (IF) procedures
  • In-Situ hybridization (ISH) procedures
  • Enzyme procedures

 

PAP PEN ACTIVATION INSTRUCTIONS:

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.
4. Release pressure on tip and blot away any excess liquid.
5. Practice applying a thin PAP Pen liquid barrier on a test slide using light, minimal pressure. Applying too much pressure may result in excess release of barrier solution, creating a wider film that could touch tissue section edges.

1. See Procedure Note #1.

6. For optimum results; store PAP Pen Liquid Blocker at room temperature, tightly capped on a horizontal flat surface.
7. Pen re-activation is not required unless tip dries out during storage.

 

PROCEDURE:

8. Paraffin Section Method:

1. Deparaffinize sections thoroughly in xylene and hydrate through 100% and 95% ethyl alcohols. Wash well with distilled water.
2. Remove slide(s) from water or buffer; blot excess solution from slide and tissue section(s) with absorbent materials. Or place long edge of the slide on absorbent material to remove excess moisture.
3. Encircle the tissue section(s) on the slightly damp slide surface with the PAP Pen. Do not touch the pen to any edges of the tissue section(s).
4. See Procedure Note #2.
5. Proceed to Step #10.

9. Frozen Section and Smear Method:
   1. Apply PAP Pen barrier before fixation or immersion of slide(s) into water or buffer.
   2. Encircle the frozen tissue section(s) or smear on the slide with the PAP Pen. Do not touch the pen to any edges of the tissue section(s) or smear.
   3. See Procedure Note #2.

10. After liquid barrier application, allow barrier film to dry before proceeding with staining procedure.
    1. See Procedure Note #3.
11. Drain/rinse reagents off between staining steps; blotting slide and tissue as needed to remove excess solution.
    1. See Procedure Note #4.
12. Complete staining process and coverslip with a compatible mounting medium. The barrier film will not affect the coverslipping procedure.

 

PROCEDURE NOTES:

1. Once a tissue section is touched with PAP Pen liquid barrier it cannot be removed. The tissue section remains usable but  colorization may result on the touched portion of tissue.
2. If the tissue section is not completely encircled or segregated by the PAP Pen Liquid Blocker barrier film, reagents will not be fully retained on the tissue section and flood 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 detection reagents may occur.
4. The use of a Slide Moisture Chamber or StainTray™ (Part 68431, 68432, 6848 or 6847) is recommended for manual staining to maintain slide organization and a moist environment during the staining process.

 

REFERENCES:

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.

 

Newcomer Supply PAP Pen Liquid Blocker, hydrophobic slide markers for staining procedures, 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 surface tension ensures the amount of reagent needed for sufficient reaction is reduced and retained for complete tissue section coverage. Multiple specimens can be separated by drawn circles or lines applied to the same slide.

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. The PAP Pen Liquid Blocker, Mini provides a finer pen tip for drawing a thinner barrier film.

 

METHOD:

Technique:  Paraffin, frozen sections and smears

• Manual staining for:
  • Immunohistochemistry (IHC) procedures
  • Immunofluorescence (IF) procedures
  • In-Situ hybridization (ISH) procedures
  • Enzyme procedures

 

PAP PEN ACTIVATION INSTRUCTIONS:

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.
4. Release pressure on tip and blot away any excess liquid.
5. Practice applying a thin PAP Pen liquid barrier on a test slide using light, minimal pressure. Applying too much pressure may result in excess release of barrier solution, creating a wider film that could touch tissue section edges.

1. See Procedure Note #1.

6. For optimum results; store PAP Pen Liquid Blocker at room temperature, tightly capped on a horizontal flat surface.
7. Pen re-activation is not required unless tip dries out during storage.

 

PROCEDURE:

8. Paraffin Section Method:

1. Deparaffinize sections thoroughly in xylene and hydrate through 100% and 95% ethyl alcohols. Wash well with distilled water.
2. Remove slide(s) from water or buffer; blot excess solution from slide and tissue section(s) with absorbent materials. Or place long edge of the slide on absorbent material to remove excess moisture.
3. Encircle the tissue section(s) on the slightly damp slide surface with the PAP Pen. Do not touch the pen to any edges of the tissue section(s).
4. See Procedure Note #2.
5. Proceed to Step #10.

9. Frozen Section and Smear Method:
   1. Apply PAP Pen barrier before fixation or immersion of slide(s) into water or buffer.
   2. Encircle the frozen tissue section(s) or smear on the slide with the PAP Pen. Do not touch the pen to any edges of the tissue section(s) or smear.
   3. See Procedure Note #2.

10. After liquid barrier application, allow barrier film to dry before proceeding with staining procedure.
    1. See Procedure Note #3.
11. Drain/rinse reagents off between staining steps; blotting slide and tissue as needed to remove excess solution.
    1. See Procedure Note #4.
12. Complete staining process and coverslip with a compatible mounting medium. The barrier film will not affect the coverslipping procedure.

 

PROCEDURE NOTES:

1. Once a tissue section is touched with PAP Pen liquid barrier it cannot be removed. The tissue section remains usable but  colorization may result on the touched portion of tissue.
2. If the tissue section is not completely encircled or segregated by the PAP Pen Liquid Blocker barrier film, reagents will not be fully retained on the tissue section and flood 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 detection reagents may occur.
4. The use of a Slide Moisture Chamber or StainTray™ (Part 68431, 68432, 6848 or 6847) is recommended for manual staining to maintain slide organization and a moist environment during the staining process.

 

REFERENCES:

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 superior and consistently reliable Gram stain that uses Gallego Solution to differentiate and fix staining of gram-negative and gram-positive bacteria in tissue sections.

 

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.

 

STAINING PROCEDURE:

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. 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. 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. Proceed with Steps #13 to #16 one slide at a time.
13. Dip in Acetone, ACS (Part 10014); 1-2 quick dips.

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. 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, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

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 is used to demonstrate nerve fibers, neurofibrils/tangles, senile plaques and axons.  This stain can be 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 staining procedure provided below.  Some solutions in the kit may contain extra volumes.

 

PRESTAINING PREPARATION:

1. If necessary, heat dry tissue sections/slides in oven.
2. All glassware/plasticware must be acid cleaned prior to use.
   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. Reserve two acid cleaned Coplin jars filled with distilled water.
   1. Save for slide rinsing/holding in Steps #7 and #10.

 

STAINING PROCEDURE:

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. See Procedure Notes #3 and #4.
6. Place in preheated Solution A: Silver Nitrate 20%, Aqueous (Step #3) for 15 minutes.
   1. See Procedure Note #5.
7. Remove slides from Solution A: Silver Nitrate 20%, Aqueous and hold in 1^st reserved Coplin jar of distilled water.
   1. Save Silver Nitrate 20%, Aqueous; pour into acid cleaned Erlenmeyer flask for Step #8.
8. Add Solution B: Ammonium Hydroxide 28-30%, ACS drop by drop into the saved Silver Nitrate 20%, Aqueous, swirling until precipitate disappears.
   1. Solution will initially be brown in color and then clear. Do not add excess ammonium hydroxide.
   2. Pour Ammoniacal Silver Solution back into acid cleaned Coplin jar.    
9. Return held slides to Ammoniacal Silver Solution in 37°C water bath for 10 minutes.

10. Remove slides and hold in 2^nd reserved Coplin jar of distilled water.
    1. Save Ammoniacal Silver Solution for Step #11.
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. Check slides microscopically at 3 minutes for development of neurons to dark brown. 
    2. Follow with checks at 1 minute intervals.
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. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
2. Plastic (Part 5500), plastic-tipped or paraffin coated metal forceps must be used with silver solutions to prevent precipitation of silver salts.  No metals of any kind should come in contact with silver solutions. Only glass thermometers should be used.
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, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

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 at www.newcomersupply.com.

 

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. 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. 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. 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. 1. See Procedure Note #3

1. 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.

1. 1. 1. 1. Place slides in a plastic Coplin jar with Solution A: Uranyl Nitrate 1%, Aqueous. Microwave at 70°C for 1 minute.
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. Place slides in a plastic Coplin jar with Solution B: Silver Nitrate 1%, Aqueous. Microwave at 70°C for 1 minute.
      2. Remove from microwave, cover and let sit for 1 minute.
2. 12. Rinse well in several changes of distilled water.
       1. Excessive rinsing may cause nucleus to pick up silver.
   13. Dip 5 times in two changes each of 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. 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. Place slides in a plastic Coplin jar with Reducing Solution. Microwave at 70°C for 1 minute. 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.

1. 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. Plastic (Part 5500), plastic-tipped or paraffin coated metal forceps must be used with any silver solution to prevent precipitation of silver salts. No metals of any kind should be in contact with any silver solution. Only glass thermometers should be used.
   3. Drain slides after each step to prevent solution carry over.
   4. The suggested microwave procedure has been tested at Newcomer Supply. This procedure is a guideline and techniques should be developed for 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 Histotechnology4 (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 a common procedure 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 slides in Solution A: Luxol Fast Blue Stain 0.1%, Alcoholic for 2 hours at 60°C or overnight at 37°C; seal lids 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 with Solution A: Luxol Fast Blue Stain 0.1%, Alcoholic. Microwave at 70°C for 10 minutes.

1. 5. Rinse slides quickly in 95% ethyl alcohol (Part 10842); 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 and white matter are colorless and contrast with stained tissue.
   8. Further differentiate in 70% ethyl alcohol (Part 10844), 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.

8. 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 suggested microwave procedure has been tested at Newcomer Supply. This procedure is a guideline and techniques should be developed for use in your laboratory.
   4. If using a xylene substitute, closely follow the manufacturer’s recommendations 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 Neurology4 (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.

 

APPLICATION:

Newcomer Supply Reticulum, Gordon & Sweets Stain procedure is a silver staining method for demonstration of reticular fibers; regarded as specialized connective tissue fibers.

 

METHOD:

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

Technique:  Paraffin sections cut at 4 microns

Solutions:  All solutions manufactured by Newcomer Supply, Inc.

 

PRESTAINING PREPARATION:

1. If necessary, heat dry tissue sections/slides in oven.
2. All glassware/plasticware must be acid cleaned prior to use.
   1. See Procedure Notes #1 and #2.
3. Prepare Silver Ammoniacal Working Solution in an acid cleaned Erlenmeyer flask.  Save for Step #11.
   1.  Silver Nitrate 10%, Aqueous; 5 ml
   2. Add Ammonium Hydroxide 28-30%, ACS (Part 1006) drop by drop, mix with swirling motion until precipitate completely dissolves.  Do not add any excess Ammonium Hydroxide. 
   3.  Add 5 ml of Sodium Hydroxide 3%, Aqueous.
   4.   Re-dissolve precipitate with Ammonium Hydroxide 28-30%, ACS drop by drop, mix with swirling motion until a faint silver/gray tinge remains.  It is normal for trace precipitate to remain.
   5.  If proceeded too far and solution is completely clear, add Silver Nitrate 10%, Aqueous drop by drop, until one drop causes solution to reach silver/gray tinge.
   6. Bring solution volume to 50 ml with distilled water; filter.

 

STAINING PROCEDURE:

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.   See Procedure Notes #3 and #4.
5. Oxidize in Potassium Permanganate 1%, Aqueous (Part 13393) for 3 minutes.
6. Wash in running tap water for 1 minute; rinse in distilled water.
7. Bleach in Oxalic Acid 1%, Aqueous for 2 minutes or until sections are colorless.
   1.   Oxalic Acid 5% Aqueous (Part 1293)     10 ml
   2.   Distilled water                                 40 ml
8. Wash in running tap water for 1 minute; rinse in distilled water.
9. Sensitize in Ferric Ammonium Sulfate 2.5%, Aqueous; 10 to 15 minutes.
10. Rinse in several changes of distilled water.
11. Impregnate sections in filtered  Silver Ammoniacal Working Solution (Step #3) for 2 minutes.
12. Rinse well in running distilled water for 1 minute.
13. Reduce in Formalin 10%, Phosphate Buffered (Part 1090) for 1 minute.
14. Rinse in running tap water for 3 minutes.
15. Check control microscopically for black reticular fiber development.
    1. See Procedure Note #5.
16. Tone in Gold Chloride 0.2%, Aqueous (Part 11286) for 1-2 minutes.
17. Rinse well in distilled water.
18. Place in Sodium Thiosulfate 5%, Aqueous (Part 1389) for 1 minute.
19. Wash well in tap water for 1 minute; rinse in distilled water.
20. Counterstain with Nuclear Fast Red Stain, Kernechtrot (Part 1255) for 5 minutes.
    1. Shake solution well before use; do not filter.
21. Rinse well in distilled water.
    1. See Procedure Note #6.
22. Quickly 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:

Reticular fibers	Black
Background	Red

 

PROCEDURE NOTES:

1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
2. Plastic (Part 5500), plastic-tipped or paraffin coated metal forceps must be used with silver solutions to prevent precipitation of silver salts.  No metals of any kind should come in contact with silver solutions.
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. If black reticular fibers are not evident or are lightly/poorly stained, return all slides to Silver Working Solution (Step #11) and repeat Steps 11-14 with the same timings.
6. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
7. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 177-179
2. Gordon, Harold, and Henry Sweets. “A Simple Method for the Silver Impregnation of Reticulum.” American Journal of Pathology 12.4 (1936): 545-552.
3. Modifications developed by Newcomer Supply Laboratory.

 

RETICULUM, GORDON & SWEETS STAIN KIT INCLUDES:

		Part 9168A
Solution A:	Potassium Permanganate 1%, Aqueous	250 ml
Solution B:	Oxalic Acid 1%, Aqueous	250 ml
Solution C:	Ferric Ammonium Sulfate 2.5%, Aqueous	250 ml
Solution D:	Silver Nitrate 10%, Aqueous	50 ml
Solution E:	Ammonium Hydroxide 28-30%, ACS	50 ml
Solution F:	Sodium Hydroxide 3%, Aqueous	50 ml
Solution G:	Formalin 10%, Aqueous	250 ml
Solution H:	Gold Chloride 0.2%, Aqueous	250 ml
Solution I:	Sodium Thiosulfate 5%, Aqueous	250 ml
Solution J:	Nuclear Fast Red Stain, Kernechtrot	250 ml
Disposable Plastic Pipettes (12)

APPLICATION:

Newcomer Supply Reticulum, Gordon & Sweets Stain Kit procedure is a silver staining method for demonstration of reticular fibers; regarded as specialized connective tissue fibers.  This stain is useful in the differential diagnosis of certain types of tumors.

 

METHOD:

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

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

 

PRESTAINING PREPARATION:

1. If necessary, heat dry tissue sections/slides in oven.
2. All glassware/plasticware must be acid cleaned prior to use.
   1.  See Procedure Notes #1 and #2.
3. Prepare Silver Ammoniacal Working Solution in an acid cleaned Erlenmeyer flask.  Save for Step #11.
   1. Solution D: Silver Nitrate 10%, Aqueous; 5 ml
   2. Add Solution E: Ammonium Hydroxide 28-30%, ACS drop by drop, mix with swirling motion until precipitate completely dissolves. Do not add excess Ammonium Hydroxide. 
   3.   Add 5 ml of Solution F: Sodium Hydroxide 3%, Aqueous.
   4.   Re-dissolve precipitate with Solution E: Ammonium Hydroxide 28-30%, ACS  drop by drop, mix with swirling motion until a faint silver/gray tinge remains. It is normal for trace precipitate to remain.  
   5.   If proceeded too far and solution is completely clear, add Solution D: Silver Nitrate 10%, Aqueous drop by drop, until one drop causes solution to reach silver/gray tinge.
   6. Bring solution volume to 50 ml with distilled water; filter.

 

STAINING PROCEDURE:

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.  See Procedure Notes #3 and #4.
5. Oxidize in Solution A: Potassium Permanganate 1%, Aqueous for 3 minutes.
6. Wash in running tap water for 1 minute; rinse in distilled water.
7. Bleach in Solution B: Oxalic Acid 1%, Aqueous for 2 minutes or until sections are colorless.
8. Wash in running tap water for 1 minute; rinse in distilled water.
9. Sensitize in Solution C: Ferric Ammonium Sulfate 2.5%, Aqueous; 10 to 15 minutes.
10. Rinse in several changes of distilled water.
11. Impregnate sections in filtered  Silver Ammoniacal Working Solution (Step #3) for 2 minutes.
12. Rinse well in running distilled water for 1 minute.
    1.  See Procedure Note #5.
13. Reduce in Solution G: Formalin 10%, Aqueous for 1 minute.
14. Rinse in running tap water for 1 minute.
15. Check control microscopically for black reticular fiber development.
    1. See Procedure Note #6.
16. Tone in Solution H: Gold Chloride 0.2%, Aqueous for 1-2 minutes.
17. Rinse well in distilled water.
18. Place in Solution I: Sodium Thiosulfate 5%, Aqueous for 1 minute.
19. Wash well in tap water for 1 minute; rinse in distilled water.
20. Counterstain with Solution J: Nuclear Fast Red Stain, Kernechtrot for 5 minutes.
    1.  Shake solution well before use; do not filter.
21. Rinse well in distilled water.
    1. See Procedure Note #7.
22. Quickly 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:

Reticular fibers	Black
Background	Red

 

PROCEDURE NOTES:

1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
2. Plastic (Part 5500), plastic-tipped or paraffin coated metal forceps must be used with silver solutions to prevent precipitation of silver salts.  No metals of any kind should come in contact with silver solutions.
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. This rinse step is critical for good reticulum demonstration.  If rinsing is insufficient, excessive background staining may occur.
6. If black reticular fibers are not evident or are lightly/poorly stained, return all slides to Silver Working Solution (Step #11) and repeat Steps 11-14 with the same timings.
7. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
8. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 177-179
2. Gordon, Harold, and Henry Sweets. “A Simple Method for the Silver Impregnation of Reticulum.” American Journal of Pathology 12.4 (1936): 545-552.
3. Modifications developed by Newcomer Supply Laboratory.

 

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
• Edge Treatment: 90° Corner
• 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: 45° Corner
• White is also available with 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 is used to demonstrate argentaffin substances such as melanin, argentaffin granules of carcinoid tumors, and some neurosecretory granules. This stain is not specific for melanin and argentaffin granules. 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. If necessary, heat dry tissue sections/slides in oven.
2. All glassware/plasticware must be acid cleaned prior to use.
   1. See Procedure Notes #1 and #2.
3. Prepare Fontana Masson Ammoniacal Silver Working Solution in an acid cleaned Erlenmeyer flask:
   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. Do not add excess Ammonium Hydroxide.
   3. Add more Solution A: Silver Nitrate 10%, Aqueous drop by drop until clear solution becomes slightly turbid or cloudy.  The change is subtle.
   4. Let solution stand for 2-4 hours before use.
   5. For use; after standing, filter silver solution. Combine 20 ml of filtered silver solution with 40 ml of distilled water; 60 ml total.

 

STAINING PROCEDURE:

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

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. 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. Shake solution well before use; do not filter.
13. Rinse well in distilled water.
    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. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
2. Plastic (Part 5500), plastic-tipped or paraffin coated metal forceps must be used with any silver solution to prevent precipitation of silver salts.  No metals of any kind should be in contact with any silver solution. Only glass thermometers should be used.
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, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

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, provides 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 PREPARATION:

1. 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. 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); 10-30 seconds.
   6. Drain off
   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 retained 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, 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. 1. Timings provided are suggested ranges. Optimal 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
  • 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 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 or Citrate Buffer 0.01M, pH 6.0.
   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
   3. If using a xylene substitute, closely follow the manufacturer’s recommendations 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 Histotechnology2 (1997): 145-154.
   3. Tacha, David, and Maria Teixeira. “History and Overview of Antigen Retrieval: Methodologies and Critical Aspects.” The Journal of Histotechnology4 (2002): 237-242.
   4. Modifications developed by Newcomer Supply Laboratory.

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

SOLUTIONS: 

APPLICATION:

Newcomer Supply Michel’s Transport Medium and pre-filled Michel’s Transport Medium Vials, provide a stable medium for transport of fresh unfixed tissues, such as renal, rectal, lymph node, skin and oral mucosa biopsies, which will undergo subsequent frozen sectioning and immunofluorescence studies.

• Michel’s Transport Medium is not a fixative and does not have any fixative properties.
• Michel’s Transport Medium is not suitable for transporting cells for flow cytometry or tissues used for fluorescent in-situ hybridization (FISH) studies.

Newcomer Supply Michel’s Wash Solution is used to rinse Michel’s Transport Medium from tissue after transport or storage and prior to freezing.

 

METHOD:

Fixation: Fresh unfixed tissue
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. Place fresh tissue in an adequate amount of Michel’s Transport Medium as soon as possible after excision.
   1. See Procedure Note #1.
2. Ensure that the specimen is completely covered with Michel’s Transport Medium and is free floating.
3. Transport tissue to processing site in Michel’s Transport Medium up to a maximum of five days.
4. During transport or storage, maintain cool to ambient temperatures of 4°C to 22°C.
5. Upon receipt, wash tissue held in Michel’s Transport Medium with Michel’s Wash Solution; three changes, 10 minutes each.
6. Freeze tissue sample(s) per laboratory protocol.
7. Tissue placed in Michel’s Transport Medium may provide adequate results when processed for light microscopy review.
   1. Wash tissue 2-3 minutes in tap water and place in appropriate fixative prior to processing.
   2. See Procedure Note #2.

 

PROCEDURE NOTES:

1. Previously frozen tissue will not provide optimal testing results and should not be used with Michel’s Transport Medium.
2. Tissues held/transported in Michel’s Transport Medium, may provide satisfactory histological results if conditions as outlined in Procedure Steps #3 and #4 are maintained.  Morphology detail will not be equal to that of expediently fixed and processed tissue.

 

REFERENCES:

1. Beutner, Ernst H., Tadeusz Chorzelski and Samuel Bean. Immunopathology of the Skin. 2nd ed. New York: Wiley, 1979. 65.
2. Carson, Freida L., and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 24-25.
3. Chua, Allison, Gina Chua and David Kelly. “Preservation of Acetylcholinesterase Enzyme Activity in Non-Frozen Rectal Biopsy Specimens for Hirschsprung Disease”. The Journal of Histotechnology 35.2 (2012): 80-88.
4. Michel, Beno, Yoram Milner and Kathy David. “Preservation of Tissue-Fixed Immunoglobulins in Skin Biopsies of Patients with Lupus Erythematosus and Bullous Disease”. The Journal of Investigative Dermatology 59.6 (1973). 449-452.
5. Modifications developed by Newcomer Supply Laboratory.

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

SOLUTIONS: 

 

APPLICATION:

Newcomer Supply Michel’s Transport Medium and pre-filled Michel’s Transport Medium Vials, provide a stable medium for transport of fresh unfixed tissues, such as renal, rectal, lymph node, skin and oral mucosa biopsies, which will undergo subsequent frozen sectioning and immunofluorescence studies.

• Michel’s Transport Medium is not a fixative and does not have any fixative properties.
• Michel’s Transport Medium is not suitable for transporting cells for flow cytometry or tissues used for fluorescent in-situ hybridization (FISH) studies.

Newcomer Supply Michel’s Wash Solution is used to rinse Michel’s Transport Medium from tissue after transport or storage and prior to freezing.

 

METHOD:

Fixation: Fresh unfixed tissue
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. Place fresh tissue in an adequate amount of Michel’s Transport Medium as soon as possible after excision.
   1. See Procedure Note #1.
2. Ensure that the specimen is completely covered with Michel’s Transport Medium and is free floating.
3. Transport tissue to processing site in Michel’s Transport Medium up to a maximum of five days.
4. During transport or storage, maintain cool to ambient temperatures of 4°C to 22°C.
5. Upon receipt, wash tissue held in Michel’s Transport Medium with Michel’s Wash Solution; three changes, 10 minutes each.
6. Freeze tissue sample(s) per laboratory protocol.
7. Tissue placed in Michel’s Transport Medium may provide adequate results when processed for light microscopy review.
   1. Wash tissue 2-3 minutes in tap water and place in appropriate fixative prior to processing.
   2. See Procedure Note #2.

 

PROCEDURE NOTES:

1. Previously frozen tissue will not provide optimal testing results and should not be used with Michel’s Transport Medium.
2. Tissues held/transported in Michel’s Transport Medium, may provide satisfactory histological results if conditions as outlined in Procedure Steps #3 and #4 are maintained.  Morphology detail will not be equal to that of expediently fixed and processed tissue.

 

REFERENCES:

1. Beutner, Ernst H., Tadeusz Chorzelski and Samuel Bean. Immunopathology of the Skin. 2nd ed. New York: Wiley, 1979. 65.
2. Carson, Freida L., and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 24-25.
3. Chua, Allison, Gina Chua and David Kelly. “Preservation of Acetylcholinesterase Enzyme Activity in Non-Frozen Rectal Biopsy Specimens for Hirschsprung Disease”. The Journal of Histotechnology 35.2 (2012): 80-88.
4. Michel, Beno, Yoram Milner and Kathy David. “Preservation of Tissue-Fixed Immunoglobulins in Skin Biopsies of Patients with Lupus Erythematosus and Bullous Disease”. The Journal of Investigative Dermatology 59.6 (1973). 449-452.
5. Modifications developed by Newcomer Supply Laboratory.

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

SOLUTIONS: 

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.

1. 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.
   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. 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, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

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 Hopkins2 (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.)

(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.)

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

SOLUTION:

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

 

Additionally Needed:

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

 

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

 

APPLICATION:

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

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

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Decalcified paraffin sections cut at 4 microns

1. 1. 1. See Procedure Note #1.

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

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

 

STAINING PROCEDURE:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #2 and #3.
   2. Wash well in running tap water; rinse in distilled water.
   3. Prepare fresh Weigert Iron Hematoxylin (Part 1409); mix well.
      1. 1. Solution A: Ferric Chloride, Acidfied 20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic 20 ml
   4. Stain in fresh Weigert Iron Hematoxylin for 10 minutes.
   5. Wash in running tap water for 10 minutes; rinse in distilled water.
      1. 1. See Procedure Note #4.
   6. Prepare 0.25% Fast Green Stain, Aqueous; combine and mix well.
      1. 1. Fast Green Stain 2.5%, Aqueous (Part 10852) 5 ml
         2. Distilled Water 45 ml
   7. Stain in 0.25% Fast Green Stain, Aqueous for 5 minutes.
   8. Rinse directly in Acetic Acid 1%, Aqueous (Part 10012); 10-15 seconds.
   9. Place directly in Safranin O Stain 1%, Aqueous for 5 minutes.
   10. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Cartilage	Red to orange
Mucin and mast cell granules	Red to orange
Bone, connective tissue and cytoplasm	Green
Nuclei	Black

 

PROCEDURE NOTES:

1. 1. Fixatives and decalcifying agents such as EDTA, nitric acid and hydrochloric acid can work to extract proteoglycans and result in weak Safranin O cartilage staining.
   2. Drain slides after each step to prevent solution carry over.
   3. Do not allow sections to dry out at any point during procedure.
   4. If Weigert Iron Hematoxylin is not completely washed from sections, nuclear and cytoplasmic staining will be compromised.
   5. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Bancroft, John D., and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 346-347.
   2. Callis, Gayle and Diane Sterchi. “Decalcification of Bone: Literature Review and Practical Study of Various Decalcifying Agents, Methods, and Their Effects on Bone Histology.” The Journal of Histotechnology 1 (1998): 49-58.
   3. Chevrier, Anik, Evgeny Rossomacha, Michael D. Buschmann, and Caroline D. Hoemann. “Optimization of Histoprocessing Methods to Detect Glycosaminoglycan, Collagen Type II, and Collagen Type I in Decalcified Rabbit Osteochondral Sections.” The Journal of Histotechnology 3 (2005): 165-175.
   4. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 256.
   5. Modifications developed by Newcomer Supply Laboratory.

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

SOLUTION:

SOLUTION:

	1 Liter
Zamboni Fixative	Part 1459A

 

APPLICATION:

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

 

METHOD:

Fixation:

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

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

 

FIXATION PROCEDURE:

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

 

PROCEDURE NOTES:

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

 

REFERENCES:

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

(use: Fite Stain for Leprosy & Nocardia.)

 

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

 

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

CI 50240

• Shelf Life is 4 years from date of manufacture.

 

SOLUTION:

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

 

Additionally Needed:

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

 

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

 

APPLICATION:

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

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

 

METHOD:

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

 

PROCEDURE:

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

 

PROCEDURE NOTES:

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

 

REFERENCES:

1. 1. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 70.
   2. Huang, W.M, S.J. Gibson, P. Facer, J. Gu and J.M. Polak. “Improved Section Adhesion for Immunocytochemistry Using High Molecular Weight Polymers of L-Lysine as a Slide Coating.” Histochemistry2 (1983): 275-279.
   3. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

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

 

Additionally Needed:

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

 

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

 

APPLICATION:

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

 

METHOD:

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

 

PRESTAINING PREPARATION:

1. 1. Prepare 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. Filter Wright Stain Solution prior to use with quality filter paper.
      1. 1. Filter sufficient stain to allow 1 ml of stain per slide.

 

STAINING PROCEDURE:

1. 4. Place slides on flat staining rack suspended over sink.
   5. Fix by flooding slides with Methanol (Part 12236) for 10-30 seconds.
   6. Drain off Methanol.
   7. Flood each slide with 1 ml of filtered Wright Stain for 3-5 minutes.
      1. 1. See Procedure Notes #1 and #2.
   8. Retain Wright Stain on slides.
   9. Directly add 1 ml of Wright Stain Buffer, pH 6.8 (Part 1430) to each slide; agitate gently to mix with retained 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, 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. 1. Timings provided are suggested ranges. Optimal times will depend upon staining intensity preference.
   2. Smears containing primarily normal cell populations require minimum staining time; immature cells and bone marrow smears/films may require longer staining time.
   3. The color range of stained cells may vary depending on buffer pH and pH of rinse water.
      1. 1. Alkalinity is indicated by red blood cells being blue-grey and white blood cells only blue.
         2. Acidity is indicated by red blood cells being bright red or pink and lack of proper staining in white blood cells.
         3. If necessary adjust buffer pH accordingly to 6.8 +/ – 0.2.

 

REFERENCES:

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

(use: Wolbach Giemsa.)

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

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

 

APPLICATION:

Newcomer Supply Victoria Blue Stain, Alcoholic, a uniquely blended stain solution, is used for demonstration of connective tissue, elastic fibers and fibrosis.  Other applications include staining of copper-associated protein in liver sections.

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 5 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 staining procedure provided below.

 

STAINING PROCEDURE:

1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each.  Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
   1. See Procedure Note #1.
2. Prepare fresh Potassium Permanganate-Sulfuric Acid Working Solution; combine and mix well.
   1. Potassium Permanganate 1%, Aqueous (Part 13393)      10 ml
   2. Sulfuric Acid 1%, Aqueous (Part 14012)                          10 ml  
   3. Distilled Water                                                           40 ml  
3. Place slides in Potassium Permanganate-Sulfuric Acid Working Solution for 5 minutes.
4. Treat with Sodium Bisulfite 1%, Aqueous (Part 13821) for 2 minutes or until sections are colorless.
5. Wash slides well in running tap water.
6. Rinse in 70% ethyl alcohol (Part 10844) for 2 minutes.
7. Stain in Victoria Blue Stain, Alcoholic for a minimum of 4 hours.
   1. See Procedure Note #2.
8. Differentiate in 70% ethyl alcohol for 1-3 minutes or until background is completely decolorized.
9. Wash slides well in running tap water.
10. Counterstain in Nuclear Fast Red Stain, Kernechtrot (Part 1255) for 5 minutes.
11. Wash in running tap water for 5 minutes.
12. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Elastic fibers	Blue
Copper-associated protein	Blue (if present in liver sections)
Nuclei and cytoplasm	Red

 

PROCEDURE NOTES:

1. Drain staining rack/slides after each step to prevent solution carry over.
2. For best results, overnight staining at room temperature in Victoria Blue Stain, Alcoholic is recommended.
3. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 296-297.
2. Prophet, Edna B., Bob Mills, Jacquelyn Arrington, and Leslie Sobin. Laboratory Methods in Histotechnology. Washington, D.C.; American Registry of Pathology. 11992. 210-211.
3. Tanaka, Kaoru, Wataru Mori, and Koji Suwa. “Victoria Blue-Nuclear Fast Red Stain for HBs Antigen Detection in Paraffin Section.”  Pathology International 31.1 (1981): 93-98.
4. Tsutsumi, Yutaka, Noboru Onoda, and Yoshiyuki Osamura. “Victoria Blue-Hematoxylin and Eosin Staining: A Useful Routine Stain for Demonstration of Venous Invasion by Cancer Cells.” The Journal of Histotechnology 13.4 (1990): 271-274.
5. Modifications developed by Newcomer Supply Laboratory.

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

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

SOLUTION:

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

 

Additionally Needed:

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

 

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

 

APPLICATION:

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

 

METHOD:

Fixation: Formalin 10%, Phosphate Buffered (Part 1090)
Technique:  Paraffin sections cut at 4 microns

1. 1. 1. 1. See Procedure Note #1.

Solutions:  All solutions manufactured by Newcomer Supply, Inc.

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

 

PRESTAINING PREPARATION:

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

 

STAINING PROCEDURE:

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

       Microwave Modification:  See Procedure Note #4.

1. 1. 1. 2. Place slides in a plastic Coplin jar containing Bouin Fluid and microwave for 5 minutes at 60° Allow slides to sit an additional 10 minutes in solution.

1. 5. Wash well in running tap water; rinse in distilled water.
   6. Prepare fresh Weigert Iron Hematoxylin (Part 1409); combine and mix well.
      1. 1. Solution A: Ferric Chloride, Acidified 20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic          20 ml
   7. Stain slides in fresh Weigert Iron Hematoxylin for 10 minutes.
   8. Wash in running tap water for 10 minutes; rinse in distilled water.

1. 1. 1. 1. See Procedure Note #5.

• 9. Stain in Trichrome Stain, Gomori One-Step, Aniline Blue for 20 minutes.
  10. Differentiate in Acetic Acid 0.5%, Aqueous (Part 100121) for 2 minutes.
  11. Rinse quickly in distilled water.
  12. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

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

 

PROCEDURE NOTES:

1. 1. Using ammonium hydroxide to soak/face tissue blocks will alter the pH of tissue sections and diminish trichrome staining.
   2. Drain slides after each step to prevent solution carry over.
   3. Do not allow sections to dry out at any point during procedure.
   4. The suggested microwave procedure has been tested at Newcomer Supply. This procedure is a guideline and techniques should be developed for your laboratory.
   5. If Weigert Iron Hematoxylin is not completely washed from tissue sections, nuclear and cytoplasmic staining may be compromised.
   6. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

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

 

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

(use: Steiner Silver Impregnation Methods.)

(use: Schmorl Melanin Stain.)

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

 

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

 

SOLUTION:                                                                                                                                               

APPLICATION:

Newcomer Supply Trichrome Stain, Gomori One-Step, Light Green for frozen muscle biopsies uses a one-step solution that combines a plasma stain with a connective tissue stain.  This procedure provides excellent staining results on fresh non-fixed frozen muscle biopsy sections for the demonstration of muscle fiber morphology and surrounding connective tissue elements.

 

METHOD:

Technique: Frozen muscle sections cut at 8 microns on adhesive slides or coverglass

1. 1. See Procedure Note #1.

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

 

STAINING PROCEDURE:

1. Air-dry frozen muscle sections a minimum of 10 minutes prior to staining.

1. See Procedure Note #2.

2. Stain air-dried frozen muscle sections in Hematoxylin Stain, Harris Modified or Hematoxylin Stain, Harris for 5 minutes.
3. Rinse in running tap water for 3 minutes.

1. Do not differentiate or use a bluing agent after hematoxylin staining.

4. Stain in Trichrome Stain, Gomori One-Step, Light Green for 18-20 minutes in a 38°C-40°C oven.

1. Allow Trichrome Stain, Gomori One-Step, Light Green to reach room temperature prior to use.

4. Prepare Acetic Acid 0.25%, Aqueous; combine and mix well.

1. Acetic Acid 0.5%, Aqueous            20 ml
2. Distilled Water                                 20 ml

6. Differentiate sections in Acetic Acid 0.25%, Aqueous; 1-2 quick dips.
7. 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:                                                                                 

Muscle fibers	Green
Interstitial connective tissue	Light green
Mitochondria	Red
Nemaline rods	Red
Myelinated nerve twigs	Red
Nuclei	Blue

 

PROCEDURE NOTES:

1. For optimal results and minimal tissue section artifact, fresh non-fixed muscle biopsies should be expediently snap frozen using an isopentane (2-Methylbutane) – liquid nitrogen freezing method.
2. Do not fix sections or use a Bouin Fluid mordant prior to staining.  Exposure to a fixative or mordant will alter staining results.
3. If using a xylene substitute, closely follow the manufacturer’s recommendations for clearing step.

 

REFERENCES:

1. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 328-329.
2. Dubowitz, Victor, and Caroline A. Sewry. Muscle Biopsy: A Practical Approach. 2nd ed. London: Baillière, 1985.30.
3. Mitchell, Jean and Andrew Waclawik. “Muscle Biopsy in Diagnosis of Neuromuscular Disorders: The Technical Aspects, Clinical Utility, and Recent Advances.” The Journal of Histotechnology 30.4 (2007): 257-269.
4. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 191-192.
5. Modifications developed by Newcomer Supply Laboratory.

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

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

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