SOLUTION:

	1 Liter	1 Gallon	20 Liter Cube
Decalcifying Solution, Formic Acid 5%, Aqueous	Part 1049B	Part 1049C	Part 1049E

 

Additionally Needed:

Decalcification End Point Set

Part 1051

 

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

 

APPLICATION:

Newcomer Supply Decalcifying Solution, Formic Acid 5%, Aqueous, provides a moderate rate of decalcification while maintaining cellular morphology.  This solution is a general purpose decalcifier and suitable for all bone specimen types from sternal or iliac crest bone marrow biopsies (light bone) to femoral head and long bone sections (compact bone).

 

METHOD:

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

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

Technique:  Paraffin sections cut at 4 microns on adhesive slides
Solutions: All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. 1. Fix bone for a length of time sufficient for specimen size and type.
      1. 1. See Procedure Note #2.
   2. Adequate bone fixation is essential before decal solution exposure.
   3. Wash fixed specimen in running tap water for 10 minutes.
   4. Submerge fixed bone segment in Decalcifying Solution, Formic Acid 5%, Aqueous, covering specimen at a 20:1 ratio.
      1. 1. See Procedure Notes #3 and #4.
   5. Check specimen regularly for sufficient solution coverage. Change solution daily and do not add or mix fresh solution with old.
   6. Decalcification time will vary, dependent on bone size and weight.
      1. 1. Check light bone samples every 30 to 60 minutes.
         2. Check compact bone samples every 1 to 2 hours.
         3. Bone marrow or light bone biopsies, on average, will decalcify in 4 to 6 hours.
         4. 3 mm thick section of femoral head, on average, will decalcify in 8 to 24 hours.
   7. Check decal completion at regular intervals with Decalcification End Point Set (Part 1051) to deter over-decalcification.
      1. 1. See Procedure Note #5.
   8. Wash in running tap water when decalcification is complete.
      1. 1. Wash small samples 30-60 minutes.
         2. Wash larger bones 1-4 hours.
         3. Additional trimming of decaled bone can occur at this point to size and thickness suitable for tissue processing.
   9. Proceed with tissue processing procedure for bone specimens.
   10. Trim block and section bone. If trimming or sectioning is impaired due to bone hardness, surface decalcification is recommended.
   11. Perform surface decalcification: Soak exposed bone surface side down in Decalcifying Solution, Formic Acid 5%, Aqueous for 15-60 minutes. Rinse block with distilled water to remove corrosive acids and re-section.
       1. 1. See Procedure Note #6.

 

PROCEDURE NOTES:

1. 1. Other fixatives suitable for bone specimens include: AZF Fixative (Part 1009), B-5 Fixative Modified, Zinc Chloride (Part 1015), Bouin Fluid (Part 1020), Zamboni Fixative (Part 1459) and Zinc Formalin Fixative (Part 1482).
   2. Reduce size of a large bone by bisecting bone into smaller pieces, removing excess soft tissue for faster fixation. Maximum bone thickness of 3-5 mm is recommended.
   3. Decal solution should be in contact with all specimen surfaces. For multiple pieces, ensure pieces are separated or suspended and not in direct contact or stacked on each other.
   4. Enhance decal with low-speed agitation shaker, rotator or stir plate.
   5. Decalcification end-point testing can also be done with specimen radiography. Physical probing of bone is not recommended.
   6. Only a few calcium-free sections will be obtained after surface decalcification. Repeat the process for additional sections.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 338-343.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 6-11.
   3. Urban, Ken. “Routine Decalcification of Bone.” Laboratory Medicine 12.4 (1981): 207-212.
   4. Villanueva, Anthony. “Experimental Studies in Demineralization and Its Effects on Cytology and Staining of Bone Marrow Cells.” The Journal of Histotechnology 9.3 (1986): 155-161.
   5. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	1 Liter	1 Gallon	10 Liter Cube
Decalcifying Solution, EDTA/Sucrose	Part 1048B	Part 1048C	Part 1048D

 

Additionally Needed:

Decalcification End Point Set

Part 1051

 

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

  

APPLICATION:

Newcomer Supply Decalcifying Solution, EDTA/Sucrose procedure uses a chelating agent with an added TRIS buffer for gentle bone decalcification.  Decalcification rate will be slower but preservation of cellular morphology is excellent and viability of staining for enzymes, immunohistochemistry antigenicity and electron microscopy is maintained. This solution is not recommended for use when proteoglycan preservation in articular cartilage is important.

 

METHOD:

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

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

Technique:  Paraffin sections cut at 4 microns on adhesive slides
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

PROCEDURE:

1. 1. Fix bone for a length of time sufficient for specimen size and type.
      1. 1. See Procedure Note #2.
   2. Adequate bone fixation is essential before decal solution exposure.
   3. Wash fixed specimen in running tap water for 10 minutes.
   4. Submerge fixed bone segment in Decalcifying Solution, EDTA/Sucrose, covering specimen at a 20:1 ratio.
      1. 1. See Procedure Notes #3 and #4.
   5. Check specimen daily for sufficient solution coverage. Change solution at least daily to ensure chelating agent is not depleted by its reaction with calcium. Do not add or mix fresh solution with old.
   6. Decalcification with Decalcifying Solution, EDTA/Sucrose can take from 2-14 days, dependent on specimen type, thickness and weight. Larger bones may require longer decal exposure.
   7. Check decal completion at regular intervals with Decalcification End Point Set (Part 1051) to deter over-decalcification.
      1. 1. See Procedure Note #5.
   8. Wash in running tap water when decalcification is complete.
      1. 1. Wash small samples 30-60 minutes.
         2. Wash larger bones 1-4 hours.
         3. Additional trimming of decaled bone can occur at this point to size and thickness suitable for tissue processing.
   9. Proceed with tissue processing procedure for bone specimens.

 

PROCEDURE NOTES:

1. 1. Other fixatives suitable for bone specimens include: AZF Fixative (Part 1009), B-5 Fixative Modified, Zinc Chloride (Part 1015), Bouin Fluid (Part 1020), Zamboni Fixative (Part 1459) and Zinc Formalin Fixative (Part 1482).
   2. Reduce size of a large bone by bisecting bone into smaller pieces, removing excess soft tissue for faster fixation. Maximum bone thickness of 3-5 mm is recommended.
   3. Decal solution should be in contact with all specimen surfaces. For multiple pieces, ensure pieces are separated or suspended and not in direct contact or stacked on each other.
   4. Enhance decal with low-speed agitation shaker, rotator or stir plate.
   5. Decalcification end-point testing can also be done with specimen radiography. Physical probing of bone is not recommended.

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 338-343.
   2. Callis, Gayle and Diane Sterchi. “Decalcification of Bone: Literature Review and Practical Study of Various Decalcifying Agents, Methods, and Their Effects on Bone Histology.” The Journal of Histotechnology 21.1 (1998): 49-58.
   3. Hao, Zhengling, Vicki Kalscheur and Peter Muir. “Decalcification of Bone for Histochemistry and Immunohistochemistry Procedures.” The Journal of Histotechnology 25.1 (2002): 33-37.
   4. Urban, Ken. “Routine Decalcification of Bone.” Laboratory Medicine 12.4 (1981): 207-212.
   5. Villanueva, Anthony. “Experimental Studies in Demineralization and Its Effects on Cytology and Staining of Bone Marrow Cells.” The Journal of Histotechnology 9.3 (1986): 155-161.
   6. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	500 ml
Crystal Violet Stain 1%, Aqueous, Brown-Hopps	Part 1041A

 

Additionally Needed:

Gram, Multi-Tissue, Artificial Control Slides                            OR Gram+ & Gram- Bacteria, Artificial Control Slides	Part 4256     OR Part 4255
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Iodine, Gram, Aqueous	Part 1140
Acetone, ACS	Part 10014
Basic Fuchsin Stain 0.25%, Aqueous	Part 1011
Gallego Solution	Part 1098
Picric Acid-Acetone 0.05%	Part 13351
Acetone-Xylene 1:1	Part 10015

 

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

 

APPLICATION:

Newcomer Supply Gram Stain, Brown-Hopps, a modification of the original Gram Stain technique, is used for differential staining of gram-positive and gram-negative 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 procedures are designed to be used with Coplin jars filled to 40 ml following the provided staining procedure.

 

STAINING PROCEDURE:     

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Stain slides in Crystal Violet Stain 1%, Aqueous, Brown-Hopps for 2 minutes.
   4. Rinse well in distilled water.
   5. Mordant in Iodine, Gram, Aqueous (Part 1140) for 5 minutes.
   6. Rinse well in distilled water.
   7. Blot excess water from slide; decolorize one slide at a time in Acetone, ACS (Part 10014) until blue stops running; 1-2 dips.
      1. 1. Sections should be very light gray in color.
   8. Quickly rinse in running tap water.
   9. Place in Basic Fuchsin Stain 0.25%, Aqueous (Part 1011) for 5 minutes.
   10. Rinse well in running tap water.
   11. Differentiate sections in Gallego Solution (Part 1098) for 5 minutes.
   12. Rinse in running tap water. Blot water off slides, but not to dryness.
       1. 1. Proceed with Steps #13 to #16 one slide at a time.
   13. Dip quickly in Acetone, ACS; 1-2 quick dips.
   14. Dip directly in Picric Acid-Acetone 0.05% (Part 13351); 3-10 dips.
   15. Dip quickly in Acetone-Xylene 1:1 (Part 10015); 5 dips.
   16. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Gram-positive bacteria	Blue/violet
Gram-negative bacteria	Red
Nuclei	Red
Background tissue	Yellow

               

PROCEDURE NOTES:

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

       

REFERENCES:

1. 1. Brown, Robert C. and Howard C. Hopps. “Staining of Bacteria in Tissue Sections: A Reliable Gram Stain Method.” American Journal of Clinical Pathology 60.2 (1973): 234-240.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 222-224.
   3. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 194-195.
   4. Modifications developed by Newcomer Supply Laboratory.

CI 42555

• Shelf Life is 4 years from date of manufacture.

(use: Mowry Colloidal Iron.)

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.

SOLUTION:

	250 ml
Trichrome Stain, Wheatley Modified	Part 10351A

 

Additionally Needed:

Alcohol, Ethyl Denatured, 70%	Part 10844
Acetic Acid, Glacial, ACS	Part 10010
Alcohol, Ethyl Denatured, 95%	Part 10842
Alcohol, Ethyl Denatured, 100%	Part 10841
Xylene, ACS	Part 1445

 

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

 

APPLICATION:

Newcomer Supply Trichrome Stain, Wheatley Modified provides a ready-to-use solution for rapid staining and permanent slide preparation for detection and identification of intestinal protozoa, flagellates and microsporidia in fecal smears.

 

METHOD:       

Fixation: According to laboratory protocol for fecal/stool samples

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. Prepare a well-made fecal smear from a fresh or fixed specimen with focus on uniform distribution of material.
   2. Fix fresh smears according to specific fixative recommendation.
   3. Place slides in 70% ethyl alcohol (Part 10844); two changes 3 minutes each.
      1. 1. See Procedure Note #2.
   4. Stain in Trichrome Stain, Wheatley Modified for 8-10 minutes.
   5. Prepare Acid-Ethanol Solution; combine and mix well.
      1. 1. Alcohol, Ethyl Denatured, 95% (Part 10842)         100 ml
         2. Acetic Acid, Glacial, ACS (Part 10010)                  0.5 ml     
   6. Differentiate slides in Acid-Ethanol Solution; 3-5 seconds.
   7. Rinse quickly in 100% ethyl alcohol; 2 dips.
   8. Dehydrate in two changes of 100% ethyl alcohol; 3 minutes each.
   9. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

PROCEDURE NOTES:

1. 1. Stool specimens received in modified polyvinyl alcohol (PVA), sodium acetate-acetic acid-formalin fixatives (SAF) or freshly fixed smears in a modified Schaudinn Solution should be prepared and fixed according to manufacturer’s recommendations.
   2. Drain slides after each step to prevent solution carry over.

 

RESULTS:

Nuclear chromatin & chromatoid bodies	Red to purple
Bacteria & ingested RBC’s	Red to purple
Cytoplasm of cysts	Blue/green with purple tinge
Cytoplasm of protozoan trophozoites	Blue/green with purple tinge
Microsporidia spores	Pink/red wall with clear interior
Background	Green

 

REFERENCES:

1. 1. Bauer, John D. Clinical Laboratory Methods. 9th ed. St. Louis: Mosby, 1982. 951-952.
   2. “CDC – DPDx – Diagnostic Procedures – Stool Specimens,” cdc.gov/dpdx/diagnosticprocedures/stool/staining.html
   3. Ryan, Norbert, G. Sutherland, K. Coughlan, M. Globan, J. Doubletree, J. Marshall, R.W. Baird, J. Pedersen and Brian Dwyer. “A New Trichrome-Blue Stain for Detection of Microsporidial Species in Urine, Stool and Nasopharyngeal Specimens.” Journal of Clinical Microbiology 31.2 (1993): 3264-3269.
   4. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 250.
   5. Wheatley, W.B. “A Rapid Staining Procedure for Intestinal Amoeba and Flagellates.” American Journal of Clinical Pathology 21 (1951): 990-991.
   6. Modifications developed by Newcomer Supply Laboratory.

(use: Stock solution for Grocott Solution GMS.)

(use: Working dilution for Grocott Solution GMS.)

SOLUTION:

	250 ml	500 ml	1 Liter
Chrome Alum-Gelatin Adhesive	Part 1033A	Part 1033B	Part 1033C

 

Additionally Needed:

Non-Adhesive Slides:

Part 6215 (Frosted End)

Part 6206 (Colored End)

Part 6210 (Plain)

 

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

 

APPLICATION:

Newcomer Supply Chrome Alum-Gelatin Adhesive provides a blended solution of chrome alum and high-quality gelatin that promotes a strong adhesive bond between tissue sections and non-adhesive microscope slides.  Chrome Alum-Gelatin Adhesive can be used as an additive to water baths or as subbed slide/direct slide coating application to prevent or reduce the loss of tissue sections due to the nature of the tissue,  section thickness or harsh staining treatments, while leaving minimal or no background staining.

The use of adhesive slides with gelatin adhesives is not recommended. Only non-adhesive slides should be used.

 

METHOD:

Technique:  Frozen or paraffin sections

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

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

 

PROCEDURES:

Water Bath Method for Paraffin Sections:

1. 1. Fill water bath with distilled water; maintain temperature at 5°-10°C below melting point of embedding medium.
      1. 1. See Procedure Note #2.
   2. Add 10 ml of Chrome Alum-Gelatin Adhesive for each liter of water bath water; combine and mix well.
   3. Float sections onto non-adhesive glass slides. Drain and dry.
      1. 1. See Procedure Note #3.

 

Subbed Slide Preparation:

1. 4. Use only clean and dry non-adhesive slides.
   5. Smear method: Place a large drop of Chrome Alum-Adhesive on slides, spread evenly over surface to create a thin film.
      1. 1. See Procedure Notes #3 and #4.
   6. To sub multiple or racked slides; Dip slides in sufficient amount of Chrome Alum-Gelatin Adhesive for 1-3 minutes, ensuring all slide surfaces are thoroughly coated.
      1. 1. See Procedure Notes #3, #4 and #5.

 

Subbed Slide Method for Paraffin and Frozen Sections:

1. 7. Paraffin Sections: float tissue sections onto dried subbed slides. Drain and dry.
   8. Frozen Sections: pick up sections on dried subbed slides and dry.

 

PROCEDURE NOTES:

1. 1. The use of Chrome Alum-Gelatin Adhesive is not recommended with silver stains due to potential increased background staining.
   2. Clean interior/exterior of water bath on a daily basis to deter contaminates and residual adhesive build-up.
   3. Drain and dry vertically in a dust-free environment. Or dry in 60°C oven for approximately 1 hour.
   4. Store dried subbed slides indefinitely in a slide box at room temperature and low humidity.
      1. 1. Slides will adhere together if not thoroughly dried before storing.
   5. Chrome Alum-Gelatin Adhesive may be difficult to remove from slide racks and glassware.
      1. 1. Wash as soon as possible after use or set aside dedicated racks and glassware for subbing procedure.

 

REFERENCES:

1. 1. Kiernan, J. A. Histological and Histochemical Methods: Theory and Practice. 3rd ed. London, Ontario: Arnold, 2003. 50-51.
   2. Luna, Lee G. Histopathologic Methods and Color Atlas of Special Stains and Tissue Artifacts. Gaitheresburg, MD: American Histolabs, 1992. 584-585.
   3. Marcos, Ricardo, Eduardo Rocha and Rogerio Monteiro. “Strategies to Maximize Adhesion of Thick Paraffin Sections of the Brown Trout Liver for Stereological Purposes.” The Journal of Histotechnology 24.1 (2001): 37-42.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTION:

	1 Liter	1 Gallon	20 Liter Cube
Bouin Fluid	Part 1020A	Part 1020B	Part 1020C

 

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

 

APPLICATION:

Newcomer Supply Bouin Fluid is a ready-to-use picric acid based tissue fixative combined with acetic acid and formalin.  Bouin Fluid penetrates rapidly, fixes evenly, provides crisp nuclear staining and preserves structures with soft and delicate features.

Bouin Fluid is recommended for a variety of specimens including bone marrow clots and biopsies, gastrointestinal tract biopsies, testicular biopsies and lymph nodes.  It also serves as both fixative and mordant for tissues stained with trichrome procedures.

 

METHOD:

Fixation Recommendations:

• • Bone Marrow Clots/Biopsy: 4 hours to 24 hours.
  • Lymph Nodes: Up to 24 hours.
  • Small nodes (5 mm or less) should be halved.
  • Large nodes dissected with no piece thicker than 2-3 mm.
  • Small Biopsies: 4 hours to 24 hours.
  • Fatty Tissue and Lipomas: 48 to 72 hours.

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

 

FIXATION PROCEDURE:

1. 1. Place fresh tissue directly in Bouin Fluid after surgical excision.
      1. 1. See Procedure Notes #1 and #2.
   2. Hold tissue in Bouin Fluid for processing or a maximum of 72 hours.
      1. 1. See Procedure Note #3.
   3. Rinse Bouin fixed tissue thoroughly in running tap water followed by 15 minute minimum wash in 70% ethyl alcohol (Part 10844) prior to processing.
   4. Place on tissue processor in Formalin 10%, Phosphate Buffered (Part 1090) fixation step.
   5. Blocked and sectioned tissues may retain excess picric acid pigment. The yellow picric acid color will normally be removed from tissue sections in the deparaffinization process. Additional methods of removing picric acid are:
      1. 1. Wash deparaffinized tissue sections in running tap water or 70% ethyl alcohol until yellow pigment is removed.
         2. Rinse deparaffinized tissue sections in 70% ethyl alcohol saturated with lithium carbonate until yellow pigment is removed.
   6. Post-fixation applications of Bouin Fluid include use as a mordant to intensify color reactions in trichrome staining procedures.
      1. 1. Refer to trichrome stain protocols for more information.

 

PROCEDURE NOTES:

1. 1. If received in Formalin 10%, Phosphate Buffered, rinse tissue thoroughly in tap water prior to placing in Bouin Fluid.
   2. Bouin Fluid should not be used for preservation of red blood cells, tissues for electron microscopy or for nuclei acid demonstration.
   3. Extended storage in Bouin Fluid is not recommended
      1. 1. After maximum fixation, rinse well in running tap water and transfer Bouin fixed tissue to 70% ethyl alcohol or Formalin 10%, Phosphate Buffered for long-term storage purposes.
   4. Collect Bouin Fluid waste solutions in an appropriately labeled leak proof container for proper disposal.
      1. 1. Do not use metal containers.
         2. Do not use containers with a metal cap or lid.
         3. Do not dispose Bouin Fluid down the drain.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 19-20.
   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. 43, 47, 50.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Sulfated Alcian Blue for Amyloid.)

SOLUTION:

	250 ml	500 ml
Biebrich Scarlet-Acid Fuchsin Stain, Elastic-Trichrome, Aqueous	Part 1016A	Part 1016B

 

Additionally Needed:

Picric Acid, Saturated Alcoholic         OR Bouin Fluid	Part 1337      OR Part 1020
Ferric Chloride 10%, Aqueous	Part 10856
Hematoxylin 5%, Alcoholic	Part 11623
Iodine, Lugol’s, Aqueous	Part 12092
Phosphomolybdic-Phosphotungstic Acid, Aqueous	Part 1332
Aniline Blue Stain, Aqueous	Part 10072
Acetic Acid 1%, Aqueous	Part 10012
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 Modified Verhoeff Elastic-Masson Trichrome Stain combines elastic and trichrome staining for demonstration and definition of elastic fibers of all sizes, connective tissue and nuclei in a single tissue section. This procedure is useful in identifying normal tissue morphology as well as heart, liver, lung and kidney pathologic conditions.

 

METHOD:

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

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

 

STAINING PROCEDURE:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Mordant in Picric Acid, Saturated Alcoholic (Part 1337) for 5 minutes or Bouin Fluid (Part 1020) at 56°C for 1 hour.
      1. 1. See Procedure Note #3.
         2. Bouin Fluid mordant; Cool at room temperature for 5-10 minutes before proceeding.
         3. Skip Step #2 if tissue was originally Bouin fixed.
   3. Wash well in running tap water; rinse in distilled water.
   4. Prepare Verhoeff Working Solution:
      1. 1. Hematoxylin 5%, Alcoholic (Part 11623) 20 ml
         2. Ferric Chloride 10%, Aqueous (Part 10856) 12 ml
         3. Iodine, Lugol’s, Aqueous (Part 12092)                   8 ml
   5. Stain in Verhoeff Working Solution for 15 minutes.
   6. Rinse in several changes of tap water.
   7. Prepare fresh Ferric Chloride 2%, Aqueous.
      1. 1. Ferric Chloride 10%, Aqueous 10 ml
         2. Distilled Water 40 ml
   8. Differentiate each slide individually in Ferric Chloride 2%, Aqueous with agitation; 2-10 dips.
      1. 1. Check differentiation: rinse well in tap water, check microscopically for black elastic staining with gray background.
         2. If needed, repeat in Ferric Chloride 2%, Aqueous until desired elastic differentiation is achieved.
   9. Wash well in running tap water.
   10. Stain in Biebrich Scarlet-Acid Fuchsin Stain, Elastic-Trichrome, Aqueous; 3 minutes.

1. 11. Rinse in distilled water for 10 minutes.
   12. Differentiate in Phosphomolybdic-Phosphotungstic Acid, Aqueous (Part 1332) for 15 minutes.
       1. 1. Collagen should be colorless and muscle red.
   13. Transfer directly to Aniline Blue Stain, Aqueous (Part 10072); 3 minutes.
   14. Differentiate in Acetic Acid 1%, Aqueous (Part 10012); 3 minutes.
   15. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Elastin	Blue-black
Muscle, keratin & cytoplasm	Red
Collagen	Blue
Nuclei	Red-brown to blue-black

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. The use of:
      1. 1. Picric Acid, Saturated Alcoholic will reduce staining time.
         2. Bouin Fluid requires longer exposure but enhances Biebrich Scarlet-Acid Fuchsin staining (Step #10).
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. 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.
   2. Garvey, Winsome. “Modified Elastic Tissue-Trichrome Stain.” Stain Technology 59.3 (1984): 213-216.
   3. Landas, Steve, M.T. Maher Strum and Karen Ellison. “Rapid Convenient Elastachrome Stain.” The Journal of Histotechnology 14.3 (1991): 191-192.
   4. Modifications developed by Newcomer Supply Laboratory.

(use: Masson or McLetchie Trichrome Stain.)

(use: Brown-Hopps mod. Gram Stain.)

(use: Brown-Brenn mod. Gram Stain. Can be used for Brown-Hopps)

CI 42500

• Shelf Life is 4 years from date of manufacture.

 

(use: Fluorescent dye.)

SOLUTION:

CI 42780

• Shelf Life is 4 years from date of manufacture.

 

 

Tech Memo 2:  Trichrome Stain, McLetchie, Aniline Blue

 

SOLUTION:                                                                                                   

	250 ml	500 ml
Aniline Blue Stain, Aqueous	Part 10072B	Part 10072C

 

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
Biebrich Scarlet-Acid Fuchsin Stain, Aqueous	Part 10161
Iodine, Weigert & Lugol, Aqueous	Part 12092
Phosphotungstic Acid 2%, Alcoholic	Part 13342

 

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

 

APPLICATION:

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

 

METHOD:

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

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

 

STAINING PROCEDURE:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   3. Place in Biebrich Scarlet-Acid Fuchsin Stain, Aqueous (Part 10161) for 5 minutes.
   4. Rinse slides in several changes of distilled water.
   5. Place in Iodine, Weigert & Lugol, Aqueous (Part 12092) for 2 minutes.
   6. Rinse slides in several changes of distilled water.
   7. Differentiate one slide at a time in Phosphotungstic Acid 2%, Alcoholic (Part 13342) for 15-30 seconds with gentle agitation.
      1. 1. To avoid over-differentiation do not exceed 30 seconds.
         2. If sections are over-differentiated, wash well in distilled water and repeat Steps #3 through #7.
   8. Rinse quickly in several changes of distilled water.
   9. Stain in Aniline Blue Stain, Aqueous for 1-3 minutes.
   10. Rinse in several changes of distilled water.
   11. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

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

 

PROCEDURE NOTES:

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

 

REFERENCES:

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

(use: Reticulum, Gordon & Sweets; Lester King, Bielschowsky.)

(FYI: No ketone and less Isopropyl.)
See also Ethyl Alcohol Denatured.

(FYI: No ketone and less Isopropyl.)
See also Ethyl Alcohol Denatured.

(FYI: No ketone and less Isopropyl.)
See also Ethyl Alcohol Denatured.

(FYI: No ketone and less Isopropyl.)

See also Ethyl Alcohol Denatured.

CI 74240

• Shelf Life is 4 years from date of manufacture.

 

(use: Gram Stain Brown-Brenn)

(use: Brown-Hopps Gram Stains.)

(use: Various, including Brown-Brenn & Hucker-Twort Gram Stains.)

(use: Warthin-Starry Method.)

(use: H&E, AFB, Brown-Brenn, PAS & Fite Stains.)

Why pay for sharpening when they’re so affordable!!!

 

3’x5′, 7/8 inch

 

2’x3′, 7/8 inch

SOLUTION:                           

	30 cc Bottle
Mount-Quick Aqueous	Part 6271A

 

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

 

APPLICATION:                                                                                                      

Newcomer Supply Mount-Quick Aqueous is a ready-to-use liquid mounting medium for histological techniques that require an aqueous based mounting medium and when dehydrating and clearing agents will adversely affect the stain, such as:  lipid/fat stains, crystal violet stains and solvent soluble immunoperoxidase chromogenic procedures.

Mount-Quick Aqueous mounting medium is not recommended for use in immunofluorescence procedures.

 

METHOD:

Technique:  Paraffin or frozen sections

 

PROCEDURE:

1. 1. Initial use of Mount-Quick Aqueous; with a clean blade, slice open the bottle tip at a slight angle in order to allow a controlled flow of mounting medium and minimal development of air bubbles.
      1. 1. Keep Mount-Quick Aqueous tightly capped when not in use to avoid evaporation and maintain solution viscosity.
   2. Complete staining procedure and final rinsing step; blot excess water from slide edges.
   3. Apply Mount-Quick Aqueous mounting medium to flow over and fully cover tissue section.
   4. Install coverglass, one edge first, allowing air bubbles to escape as the opposite edge is lowered to slide.
      1. 1. See Procedure Notes #1 and #2.
   5. Wipe off excess mounting medium from sides and bottom of slide.
   6. Dry slide in a horizontal position. Edges of the coverslip will dry sufficiently to hold the coverslip in place.
   7. Prevent overly dried edges and accumulation of air bubbles that may form due to evaporation and long-term storage by sealing coverglass edges with a thin coat of clear nail polish/lacquer. Allow sealed slide to dry thoroughly before filing.

 

PROCEDURE NOTES:

1. 1. In lipid staining procedures, use minimal pressure when applying coverslip or fat/lipid staining may be displaced.
   2. To remove trapped air bubbles under coverslip; soak slide in warm water until coverslip is easily removed. Blot excess water from slide and remount with new coverslip and Mount-Quick Aqueous.
   3. Mount-Quick Aqueous refractive index (1.41) is close to that of glass (1.52).

 

REFERENCES:

1. 1. Bancroft, John D. and Marilyn Gamble. Theory and Practice of Histological Techniques. 6th ed. Oxford: Churchill Livingstone Elsevier, 2008. 693.
   2. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-Instructional Text. 4th ed. Chicago: ASCP Press, 2015. 131-132, 183-185.
   3. Modifications developed by Newcomer Supply Laboratory.

The stainless steel racks have an improved design that allows for the slides to be placed into them with more ease.

 

We have designed and developed an embedding tissue tamper that techs can now easily hold with their fingers or forceps and have complete control of the tamper!  Other tampers on the market are designed solely for operating it with your fingers or a forceps, but not both.

At Newcomer Supply we understand the dilemma, because we have our own histology lab for control slide production.  When embedding, different techs have different styles of how they want to use the tampers.  Some like to keep the embedding forceps in their hands throughout the process and don’t like the idea of putting them down to tamper the tissue in the base molds.  Other techs like the feel and control of holding the tamper with their fingers and don’t like the tampers that can only be used with forceps.

Our solution was to design a single piece aluminum tamper with a handle that extends 24mm above the tamper base.  This allows for great control for techs that like to use their forceps when tampering the tissues, but also affords those that prefer holding the tamper with their fingers the same luxury!  The extended handle with a vinyl cap also allows the surface, where a tech would hold it, to remain cool to the touch!

For a finishing touch, there is also an anodized colored surface on the tamper that makes it more durable and corrosion-resistant.  The colors signify size and give it an artful design.

These embedding tissue tampers are a must have for any grossing lab!

AVAILABLE IN:

• 10mm x 10mm base size (orange, yellow and pink) for smaller base molds
• 16mm x 19mm base size (green, purple and blue) for larger base molds
• Combo pack including 1 of each size –  10mm x 10mm and 16mm x 19mm

See your embedding in a whole new light! These lighted forceps ease identification and location of otherwise undetectable tissue fragments, while reducing embedding time. Increases visibility of specimen through paraffin during embedding, allowing for more accurate orientation. Made of German stainless steel, are submersible and light longevity approx. 20,000 hours. (Each kit includes: one instrument, power pack & battery charger)

SOLUTION: 

	250 ml	500 ml
Fouchet Reagent	Part 1095A	Part 1095B

 

Additionally Needed:

Bile Control Slides	Part 4060
Van Gieson Stain	Part 1404
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

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

 

APPLICATION:

Newcomer Supply Bile Stain, Hall’s Method is for the demonstration of bile (bilirubin) substances in tissue sections and to distinguish bile pigments from other tissue pigments.  The acidity of Fouchet Reagent works to oxidize bilirubin to biliverdin, resulting in a green color development with the Van Gieson Stain serving as a complimentary counterstain.

 

METHOD:

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

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

 

PRESTAINING PREPARATION:

1. If necessary, heat dry tissue sections/slides in oven.
2. Filter Fouchet Reagent with Grade 1 filter paper prior to use.

 

STAINING PROCEDURE:

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. See Procedure Notes #1 and #2.
4. Place slides in freshly filtered Fouchet Reagent for 5 minutes.
5. Wash in three changes of tap water; rinse in distilled water.
6. Stain sections in Van Gieson Stain (Part 1404) for 5 minutes.
7. Rinse quickly in 95% ethyl alcohol.
8. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Bile/bilirubin	Emerald green to olive drab
Connective tissue	Pink to red
Background	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. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 268-269.
2. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 219.
3. Modifications developed by Newcomer Supply Laboratory.

SOLUTIONS:

	100 ml	250 ml	500 ml
Methenamine 3%, Aqueous		Part 12239A	Part 12239B
Silver Nitrate 5%, Aqueous		Part 13805A	Part 13805B
Sodium Borate 5%, Aqueous			Part 13826B
Gold Chloride 0.25%, Aqueous	Part 11287A	Part 11287B
Sodium Thiosulfate 2.5%, Aqueous		Part 13889A	Part 13889B
Light Green SF Yellowish Stain 0.2%, Aqueous		Part 12202A	Part 12202B

 

Additionally Needed:

Urates Control Slides	Part 4700
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 product labels.

 

APPLICATION:

Newcomer Supply Urate Stain, Gomori Methenamine Silver Method is designed to demonstrate urates in tissue sections.  With abnormal accumulations found around joints and in soft tissues, this disturbance in uric acid metabolism is known as gout, with collections of urate crystals referred to as gouty tophi.

Calcium pyrophosphate crystals or pseudogout may also be demonstrated. When viewed with a polarizing filter and red compensator filter, gout and pseudogout can be distinguished.

 

METHOD:

Fixation: Urate crystals are soluble in aqueous solutions. Fix in 100% ethyl alcohol; a minimum of two changes, 4 hours each.
Processing:  Transfer from 100% ethyl alcohol fixative to xylene for 1 hour; proceed with equal parts xylene/paraffin at 58°C for 2 hours. Infiltrate with paraffin for a minimum of 1 hour; embed.
Technique: Chill paraffin blocks in 100% ethyl alcohol; cut sections at 4 microns with minimal water bath exposure.
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

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

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.
   3. Prepare Methenamine Silver Stock Solution.
      1. 1. Methenamine 3%, Aqueous (Part 12239) 50 ml
         2. Silver Nitrate 5%, Aqueous (Part 13805) 2.5 ml
         3. Slowly add silver nitrate; mix to clear milky precipitate.
         4. Store clear stock solution at 2°-8°C for up to 2 months.
   4. Prepare fresh Methenamine Silver Working Solution; mix well.
      1. 1. Methenamine Silver Stock Solution 25 ml
         2. Distilled Water 25 ml
         3. Sodium Borate 5%, Aqueous (Part 13826)   3 ml
   5. Preheat Methenamine Silver Working Solution to 60°C in a water bath.

 

STAINING PROCEDURE:

1. 6. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Rinse in two changes of 100% ethyl alcohol, 10 dips each.
      1. 1. Do not use 95% ethyl alcohol or distilled water steps.
         2. See Procedure Notes #3 and #4.

1. 7. Incubate slides in preheated Methenamine Silver Working Solution (Step #5) in a 60°C water bath for 30 minutes.
      1. 1. Remove control slide, rinse in warm distilled water, check microscopically for adequate silver development. Crystals should be dark brown/black.
         2. If not sufficiently dark, return to warm silver solution.
         3. Recheck at 2-3-minute intervals for desired intensity.
   8. Rinse well in distilled water.
   9. Tone in Gold Chloride 0.25%, Aqueous (Part 11287) until brown colorization disappears; 5 to 30 seconds.
   10. Rinse well in distilled water.
   11. Place in Sodium Thiosulfate 2.5%, Aqueous (Part 13889); 2-3 minutes.
   12. Wash well in running tap water for 3 minutes; rinse in distilled water.
   13. Counterstain in Light Green SF Yellowish Stain 0.2%, Aqueous (Part 12202) for 1-2 minutes.
   14. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Light Field Microscopy:
	Gout/urate crystals	Black
	Background	Green
Polarized/Red Compensator Filter: (long axes aligned parallel)
	Gout/urate crystals	Yellow, long & needle shaped
	Pseudogout crystals	Blue, shorter & rhomboidal

 

PROCEDURE NOTES:

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

 

REFERENCES: 

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009.255-256, 267-268.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 187-188.
   3. Sheehan, Dezna C. and Barbara B Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 225-226.
   4. Modifications developed by Newcomer Supply Laboratory.

SOLUTIONS:

	250 ml	500 ml	1 Liter
Silver Nitrate 5%, Aqueous	Part 13805A	Part 13805B
Sodium Thiosulfate 5%, Aqueous		Part 1389A	Part 1389B
Nuclear Fast Red Stain, Kernechtrot	Part 1255A	Part 1255C	Part 1255B

 

Additionally Needed:

Calcium Control Slides	Part 4100
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 product labels.

 

APPLICATION:

Newcomer Supply Von Kossa Calcium Stain is an indirect staining method for the demonstration of calcium or calcium salt in tissue sections. This technique is not specific for calcium. Other reducing substances, such as formalin pigment and melanin, will also give a positive reaction.

 

METHOD:

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

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

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. All glassware/plasticware must be acid cleaned prior to use.
      1. 1. See Procedure Notes #1 and #2.

 

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 #3 and #4.
   4. Place in Silver Nitrate 5%, Aqueous (Part 13805) according to the following timings and conditions.
      1. 1. Direct sunlight or ultraviolet light for 10-30 minutes.
         2. In front of a 60-100 watt light bulb for 1 hour or longer.
         3. See Procedure Note #5.
   5. Check slides periodically and remove from light source when control slide shows black-brown deposits macroscopically.
   6. Rinse well in several changes of distilled water.
   7. Place in Sodium Thiosulfate 5%, Aqueous (Part 1389) for 2 minutes.
   8. Rinse well in several changes of distilled water.
   9. Counterstain in Nuclear Fast Red Stain, Kernechtrot (Part 1255) for 5 minutes.
      1. 1. Shake solution well before use; do not filter.    
   10. Rinse well in distilled water.
       1. 1. See Procedure Note #6.
   11. Dehydrate in two changes each of 95% and 100% ethyl alcohol; 10 dips each. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Calcium salts	Black to brown/black
Nuclei	Red
Cytoplasm	Light pink

 

PROCEDURE NOTES:

1. 1. Acid clean all glassware/plasticware (Part 12086) and rinse thoroughly in several changes of distilled water.
   2. No metals of any kind should come in contact with silver solutions to prevent precipitation of silver salts.  Use plastic forceps (Part 5500) or paraffin coated metal forceps.
   3. Drain slides after each step to prevent solution carry over.
   4. Do not allow sections to dry out at any point during procedure.
   5. Direct sunlight is the preferred method. If procedure is in minimal sunlight increased incubation time will be necessary.
   6. Wash well after Nuclear Fast Red Stain, Kernechtrot to avoid cloudiness in dehydration steps.
   7. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 269-270.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 226-227.
   3. Modifications developed by Newcomer Supply Laboratory.

 

The Infinity Microtome Blade was designed for histotechs that use the Accu-Edge and Feather microtome blades.  It performs in the same manner, but lasts longer and costs less!

It is coated with a special Teflon-like substance that consistently provides a smooth and precise section in microtomes and cryostats. Its unique combination of durability and precision competes well against all blades, and is specifically designed to outperform the Feather S35 & R35 and Thermo mx35 blades.

 

FEATURES OF THE INFINTY MICROTOME BLADES:

• Triple faceted, 35° cutting edge
• 80mm length blade
• Incredible longevity and performance
• Ideal for all general paraffin and cryostat use!

 

INIFINITY DISPOSABLE MICROTOME BLADES AVAILABLE IN:

• Low Profile
• High Profile

 

DIMENSIONS:

• Low Profile – 80mm x 8mm x 0.25mm
• High Profile – 80mm x 14mm x 0.30mm

 

Infinity Disposable Microtome Blades are Made in the USA

 

The C.L. Sturkey Select Disposable Microtome blade edges are coated with a teflon finish to provide smooth, wrinkle-free tissue sections.

 

SELECT DISPOSABLE MICROTOME BLADES ARE IDEAL FOR:

• • General Paraffin Sectioning
  • Cryostat Use

 

SELECT DISPOSABLE MICROTOME BLADES ARE AVAILABLE IN:

• • Low Profile
  • High Profile

 

The Select disposable microtome blade model is available in the standard 50 blade dispenser.  The High Profile Select microtome blade is manufactured to the industry standard 0.012″ thickness (C.L. Sturkey’s thinnest high profile blade).

Select Disposable Microtome Blades are Made in the USA

 

 

MONARCH DISPOSABLE MICROTOME BLADES ARE IDEAL FOR:

• • • All general paraffin cutting including: soft tissue, hard tissue, decalcified bone, small biopsies and large tissue.

 

MONARCH DISPOSABLE MICROTOME BLADES ARE AVAILABLE IN:

• • • Low Profile, 50 blade dispenser

 

DIMENSIONS:

• • • 76mm x 8mm x .25mm (H x W x D)

 

Monarch Disposable Microtome Blades are Made in the USA

 

The C.L. Sturkey Extremus disposable microtome blade edges are coated with the hardest of the nitride line coatings.  Extremus high performance disposable microtome blades offer the longest edge life and you can expect to cut two to three times as many sections as other blades on the market today!

 

EXTREMUS DISPOSABLE MICROTOME BLADES ARE IDEAL FOR:

• General purpose needs for a variety of cutting conditions
• Hard Tissue Sections
• Longevity of a disposable microtome blade

 

EXTREMUS DISPOSABLE MICROTOME BLADES ARE AVAILABLE IN:

• Low Profile
• High Profile

 

Extremus Disposable Microtome Blades are Made in the USA

 

 

The CL Sturkey Silver disposable microtome blade edges are coated with a nitride and copolymer finish to provide smooth, wrinkle-free tissue sections.  They are durable, long lasting and a great choice for a general purpose disposable microtome blade!

 

SILVER DISPOSABLE MICROTOME BLADES ARE IDEAL FOR:

• General paraffin sections
• Cryostat use

 

SILVER DISPOSABLE MICROTOME BLADES ARE AVAILABLE IN:

• Low Profile
• High Profile

 

Silver Disposable Microtome Blades are Made in the USA

Our most popular 15cm ruler is now available in a clear plastice!

Flexible plastic with metric on top & bottom sides and flush cut at zero and 15cm mark. Ideal for making precise measurements and being able to view the tissue that is directly beneath the ruler.

Designed by PAs for the PAs! Flush cut at zero, bold markings every 1/2 cm for easy reads. Identical printing on both top and bottom edges to accomodate both top and bottom readers AND printed identically on both sides. Each side is a working side!

 

Instructions for Aligning the Chuck and Blade Holder

Always wear gloves when working on a Cryostat          

Click here for pdf

 

 

 

 

 

 

 

 

 

 

Every microtome can be ready to do recuts on those precious biopsies! So simple & quick to use, you will be amazed. Only one Universal Microtome Aligner tool needed for ALL models, and it will last forever.

MICROTOME ALIGNER INSTRUCTION MANUAL

Click here for pdf

1. Remove blade from knife holder.
2. Move microtome specimen head all the way back to the “home” position.
3. Insert the Universal Microtome Aligner tool into the specimen head clamp with the bubble level facing up (see Figure #1).

                                                Figure #1

 

4. Release orientation head locking lever so that the orientation head can be adjusted.
5. Slowly move specimen head forward from “home” position until alignment bar on the Universal Microtome Aligner tool is almost touching the backside of the knife holder (see Figure #2).  You may need to manually lower the specimen head with hand wheel to match up the alignment bar with the back side of the knife holder (see Figure #2).

                                                                                                  Figure #2

6. Adjust the gap between knife holder and alignment bar so the left and the right side of alignment bar has the same amount of gap.  You may need to advance or retract specimen head to get the best position.  This will adjust the “Z” axis alignment (see Figure #3).

                                         Figure #3

7. Center the bubble in circle level by adjusting the “X” and “Y” axis (see Figure #3).
8. Once the bubble in the level is centered, lock orientation head with locking lever.
9. Re-check the alignment bar gap (steps 5 & 6).
10. Your microtome orientation head should now be aligned!

 

NOTE:  To receive optimal performance from this Universal Microtome Aligner tool, your microtome MUST BE level and your specimen head clamp must be free of any debris.

 

If the bench or the microtome is found not to be level, here are some suggestions to try and accommodate for this issue:

1. Purchase different types and/or sizes of rubber or felt feet from a local hardware store for adjusting the level foundation beneath the microtome.
2. Use shims of metal or wood to level the base of the microtome.
3. Have your bench top leveled by a maintenance ace or carpenter.
4. If your microtome had adjustable feet, manipulate them to bring your microtome to level.

 

If you cannot level the bench and/or the microtome with the above suggestions, here is a way to work around a non-level surface;

• Match the offset of your microtome or your bench top by physically marking the actual offset on the aligner level.  The new marking becomes the “zero” target mark.  (See diagrams below)

 

 

 

 

 

Superfrost Plus Adhesive Coated Slides are a general purpose adhesive slide.

 

RECOMMENDED USE:

• Histo & IHC Autostainers
• Roche/Ventana Autostainer
• Demanding IHC with antigen retrieval
• Difficult Tissue
• Frozen Tissues

 

SPECIFICATIONS OF THE SUPERFROST PLUS ADHESIVE COATED SLIDES:

• Size: 75 x 25mm, 1mm thick
• Edge Treatment: 90° Corner
• Packaged Approx. 72 slides/box, 20 boxes/case
• Available color: White

An excellent pad to gross on that will absorb & neutralize the formalin from your specimen.

 

FAN PAD ULTRA BENEFITS:

• Dense enough to provide excellent knife suppression
• Soft enough to reduce wear on dissecting blades
• Destroys harmful formalin vapors while you gross
• Use for vapor control in storage of formalin or formalin fixed tissue containers

 

FAN PAD ULTRA LIST OF USES:

• Specimen transport
• Surgery
• Specimen receiving
• Labor & Delivery
• Specimen Storage Areas
• Autopsy

 

ABSORBING/NEUTRALIZING CAPACITY OF THE FAN PAD ULTRA:

• 8″ x 11″   – will abosrb & neutralize up to 100ml of 10% formalin
• 11″ x 15″ – will absorb & neutralize up to 200ml of 10% formalin
• 15″ x 20″ – will absorb & neutralize up to 300ml of 10% formalin

Just one of those little necessities that has to be replaced. Good quality with an even better price.

 

SOLUTION:                                                                                                      

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

 

Additionally Needed:

Acid Fast Bacteria (AFB) Control Slides	Part 4011
Acid Alcohol 1%	Part 10011
Methylene Blue Stain 0.14%, Alcoholic	Part 12401
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

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

APPLICATION:

Newcomer Supply Carbol Fuchsin Stain, Kinyoun is used in the Kinyoun AFB Stain to demonstrate the presence of acid-fast mycobacteria in tissue sections. Carbol Fuchsin Stain, Kinyoun is a concentrated phenol and basic fuchsin solution that works to permeate the lipoid capsule of acid-fast organisms, rendering them resistant to acid alcohol decolorization.

 

METHOD:

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

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

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Filter Carbol Fuchsin Stain, Kinyoun with filter paper before use.

 

STAINING PROCEDURE:

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Stain in freshly filtered Carbol Fuchsin Stain, Kinyoun for 15 minutes at room temperature. Keep solution covered.
      1. 1. See Procedure Note #3.
   5. Wash in running tap water for 2 to 3 minutes.
   6. Differentiate in Acid Alcohol 1% (Part 10011) until color no longer runs off the slide and sections are pale pink; 3 to 10 rapid dips.
   7. Wash in running tap water 3 to 5 minutes; rinse in distilled water.
   8. Counterstain in Methylene Blue Stain 0.14%, Alcoholic (Part 12401); 3-6 dips.
      1. 1. See Procedure Note #4.
   9. Wash in running tap water for 1 minute; rinse in distilled water.
   10. Dehydrate quickly in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Acid-fast bacteria	Bright red
Background	Pale blue

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Sections can remain in Carbol Fuchsin Stain, Kinyoun for up to 60 minutes without adverse effect. Additional differentiation may be required in Step #6.
   4. If preferred, Light Green SF Yellowish Stain 0.1%, Aqueous (Part 12203) can be used as a counterstain in place of Methylene Blue.
      1. 1. Stain for 3-6 dips.
         2. Rinse with one quick dip in distilled water or proceed directly to Step #10 without a distilled water rinse.
   5. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L., and Christa Hladik. Histotechnology: A Self-Instructional Text. 3rd ed. Chicago, Ill.: American Society of Clinical Pathologists, 2009. 224-226.
   2. J.J. “A Note on Uhlenhuths Method for Sputum Examination, for Tubercle Bacilli.” American Journal of Public Health 5.9 (1915). 867-870.
   3. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 236-237.
   4. Modifications developed by Newcomer Supply Laboratory.

 

APPLICATION:

Newcomer Supply Toluidine Blue Stain for Mohs Technique provides a rapid staining method for Mohs micrographic surgery (MMS), useful when evaluating frozen skin samples for basal cell carcinoma (BCC).  Toluidine Blue imparts an identifiable staining pattern if BCC is present that will highlight islands of blue staining basal cell carcinoma and metachromatically stain surrounding mucopolysaccharides/stroma pink.

 

METHOD:

Fixation: 70% Ethyl Alcohol (Part 10844)
Technique:  Frozen sections cut at 5-7 microns on adhesive slides
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. Fix tissue sections in 70% Ethyl Alcohol for 30-60 seconds.
   1. See Procedure Note #1.
2. Wash well in distilled water to remove excess fixative.
3. Stain slides in Toluidine Blue Stain 0.1%, Aqueous for 30-60 seconds, depending on preference of stain intensity.
4. Wash gently in distilled water to remove excess stain.
5. Dehydrate quickly through one change of 95% ethyl alcohol; 1 quick dip and then two changes 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.
   1. See Procedure Note #2.

 

RESULTS:

Islands of basal cell carcinoma	Deep blue to purple
Surrounding mucopolysaccharides/stroma	Pink to magenta
Background	Blue
Nuclei	Dark blue

 

PROCEDURE NOTES:

1. Section thickness and fixation timing may affect staining quality.
2. Alcohol will work as a differentiator. Proceed quickly through dehydration steps to maintain Toluidine Blue stain.
3. If using a xylene substitute, closely follow the manufacturer’s recommendation for clearing step.

 

REFERENCES:

1. Arnon, Ofer, Ronald Rapini, Adam Mamelak, and Leonard Goldberg. “Mohs Micrographic Surgery: Current Techniques.” IMAJ 12 (2010): 431-35.
2. Gross, Kenneth G. Mohs Surgery: Fundamentals and Techniques. St. Louis: Mosby, 1999. 125-138.
3. Modifications developed by Newcomer Supply Laboratory.

 

Human lung tissue reactive  Anaplastic Lymphoma Kinase (ALK, CD246). Image stained with clone ALK1

Excellent for difficult tissues, frozen sections and special stains (esp. Silvers). Imprinted “ADHESIVE”.

• Size: 75 x 25, 1mm thick
• Packaged Approx. 72/box

Allows for Stat nuclear counter and special staining in 10 seconds. Specially developed for IHC stains. Mountable with xylene and aqueous based mounting media. Stain does not fade with time, no precipitation,no filtration. Usable to the last drop. Shelf life 4 years. Store at 4°C.

Permanent Aqueous Mounting Media

The H&E Mount mounting media is a newly formulated aqueous based permanent mounting media designed for coverslipping H&E stains from water. The use of H&E aqueous mounting media eliminates the need for alcohol dehydration and xylene clearing pre-treatment steps, slides are simply mounted from water post hematoxylin and eosin staining. The H&E Mount provides permanent preservation of H&E stained tissue sections, cell smears and cyto-spins for indefinite storage.

H&E stained slides mounted with H&E Mount do not display any loss of eosin or hematoxylin staining that is often observed when H&E slides are mounted from alcohol and xylene. Therefore, when mounting with H&E Mount, less number of dips (5-10) in eosin is recommended.

 

H&E MOUNTING MEDIA APPLICATION:

H&E Mounting Media is intended for the mounting and preservation of hematoxylin and eosin (H&E) stained biological specimen. It is intended to be applied with coverslips. 

 

FEATURES OF H&E MOUNTING MEDIA: 

• Mount directly from water or alcohol.
• No dehydration or xylene clearing steps required prior to mounting.
• Alcohol differentiation of eosin may be performed prior to mounting.
• Permanent.
• No fading of eosin or hematoxylin.
• Sets in 10 minutes, ready for microscopic examination.
• Excellent for photography.
• Simple soaking in water removes coverslip.

 

H&E STAINING PROGRESSIVE METHOD:

For paraffin sections:  Deparaffinize slides and rinse slides in tap water.

For frozen sections:  Cut sections, fix in 40% formaldehyde for 10 seconds and rinse well with tap water.

1. 1. Immerse in STAT Aqua Hematoxylin (NB305) for 2-5 minutes.  STAT Aqua Hematoxylin does not require post treatment of slides with Ammonia water, slides stained with STAT Aqua Hematoxylin turn blue with tap water rinse alone.

OR

Immerse in Harris hematoxylin for 1-3 minutes, rinse with Ammonia water until slides turn blue (rinse sections well with tap water, too much ammonia carry over will decrease eosin staining).

1. 2. Rinse in tap water until water runs clear.
   3. Stain in eosin with 5-10 dips.
   4. Rinse in tap water.
   5. Blot as much water as possible from the surface of the slides and proceed as below with coverslipping.

 

H&E MOUNTING & DRYING PROTOCOL:

The following is a recommended technique for coverslipping slide specimens. Other techniques that achieve the same basic result are equally acceptable.

1. Take the slides from the final wash (for best results water is quite suitable).
2. Remove excess water by tapping the slides on a paper towel, for best results, blot most of the water or let slides air dry for 1-2 minutes.
3. Place the slides down, face up on a flat surface and apply 1-2 drops (or as needed) of H&E Mount on the lower edge of the coverslip.
4. Bring the slide up to the edge of the coverslip and invert the slide so that the mounting media touches the slide. Gently complete inversion.
5. Mounted slides with H&E mounting media are ready in 10 minutes for microscopic examination.
6. Mounted slides should be properly stored following the microscopic examination. Slides are best stored in the dark and free of dust.

 

SECOND CHANCE COVERSLIPPING: 

If for any reason you end up with an unsatisfactory mounted slide such as one with some air bubbles, remount slides as follows:

1. Allow the coverslip to slip off the end of the slides by holding the slide at a slight vertical angle.
2. Remove excess media as if it were excess water on a slide.
3. Re-apply mounting media and coverslip.
4. Mounted slides should be properly stored following the microscopic examination.

 

COVERSLIP REMOVAL:

When desired, coverslips mounted with H&E Mount may be removed by soaking the slides in warm water. The length required depends on the age of the mounted slides and may vary from 10 minutes to overnight. After softening the mounting media, slowly and gently pull back on the corner of the coverslip until it releases. Then rinse off the remaining mounting media by agitating the un-slipped slide in the warm water for a few moments.

5X Concentrate

(For amplifying Fast Red, BCIP/NBT, Innovex Brown & other substrate chromogens for alkaline phosphatase enzyme staining.)

 

Store multiple 20 place slide trays for easy retrieval.

• Holds five 20-place slide trays (not included)
• Includes 5 colored clasps for easy identification
• Portable
• Refrigerator safe
• Stackable

 

The Stirring Hot Plate has the ability to heat and stir simultaneously or perform each function separately.

FEATURES OF THE STIRRING HOT PLATE:

• Stainless steel heating surface
• Includes temperature probe
• LED display can show either “Set” or “Actual” Temperature
• Includes one 9 x 38mm magnetic stir bar
• Stirring power: 30W
• Temperature range: Room temp. to 300° C
• Stirring Speed:  Start-up to 2,000 rpm

 

DIMENSIONS OF THE STIRRING HOT PLATE:

• Working surface diameter of 6″
• Overall size 11″ x 6.3″ x 4″
• Shipping Weight of 9lbs.

 

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.

The dishes are made of a heavy glass with snug fit lids. They come in three different sizes to accommodate the 20, 30 and 60 place slide racks.

Polyethylene pipet with large bulb. Pipet graduated from 0.5 to 3.0ml.