Lung infected with acid fast bacteria (AFB).

The Form Zero product will produce excess sulfite once the reaction has been completed for neutralizing the formalin waste.  These test strips will test for any residual sulfite that is in the treated solution.  Upon confirming a positive response on the test strip means the neutralization on the formalin waste is complete and now can be safely sewer disposed

PRODUCT:

	30 cc Tube
Mount-Quick Mounting Media	Part 6270

 

Additionally Needed for Tissue/Cell Transfer Procedure:

Xylene, ACS	Part 1445
HistoTec™ Pen	Part 5725
Scalpels, Finger        OR Scalpels, Flat Edge	Part 6810     OR Part 5545
Superfrost® Plus Slides               OR NewSilane Adhesive Slides	Part 6203W     OR Part 5070
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842

 

APPLICATION:

Newcomer Supply Mount-Quick Mounting Media is a multi-use product with applications as a standard mounting media, a liquid coverslip (coverglass free) mounting media and for tissue transfer procedures.

The tissue transfer procedure involves the use of Mount-Quick to transfer sections from a prepared microscope slide when specimen availability is limited and further diagnostic workup is needed. Staining quality will not be affected and histochemical, immunohistochemistry (IHC) and cytochemical procedures performed on transferred samples produce reliable results.  Examples when tissue transfer is applicable include:

• • • Paraffin sections of small biopsies/minimal tissue
    • Biopsies with a small foci of neoplasm
    • Paraffin block is unavailable
    • Cytology preparations, fine needle aspirates (FNA)
    • Transfer of tissue from an uncharged slide to charged slide(s)
    • Preparing multiple slides from a single slide preparation
    • Restoring broken slides back to a whole mount

 

This tissue transfer procedure takes time to obtain a well-prepared end product. Practicing the technique prior to performing on limited specimens is suggested. Reduced timings/short cuts shouldn’t be taken.

 

METHOD:

Coverslipping:  Paraffin or frozen sections
Tissue/Cell Transfer: Stained and unstained paraffin sections, cytology preparations and smears.

 

COVERSLIPPING PROCEDURE:

1. 1. Complete staining procedure: dehydrate, clear with xylene, blot excess xylene from slide edges.
   2. Apply Mount-Quick Mounting Media to fully cover tissue section(s).
      1. 1. Standard coverglass; install one edge first, allowing air bubbles to escape as opposite edge is lowered to slide.
         2. Liquid coverglass (coverglass free); allow to dry flat for 30 minutes before handling and viewing microscopically.
   3. Dry slides a minimum of 24-48 hours before filing.

 

TISSUE/CELL TRANSFER PROCEDURE:

1. 1. Mark tissue area(s) that is the focus of tissue transfer on underside of the original slide with a HistoTec Pen (Part 5725).
   2. For previously stained sections:
      1. 1. Soak slide in xylene until coverslip is removed without damage to tissue section.
         2. Run slide through several changes of xylene to ensure removal of residual mounting media.
         3. Do not proceed through alcohol steps.
         4. See Procedure Note #1.
   3. For unstained paraffin sections or smears:
      1. 1. Soak slides in several changes of xylene, a minimum of 10 minutes. Do not proceed through alcohol steps.

1. 4. Spread Mount-Quick Mounting Media on xylene-coated slides with an applicator stick. Use sufficient Mount-Quick to form a meniscus over entire tissue section or cell preparation.
      1. 1. Slide must be xylene coated and not allowed to dry out.
   5. Place slide in a flat horizontal position in a 60°C oven for 90 minutes to 2 hours or in a 37°C oven overnight until meniscus hardens.
      1. 1. See Procedure Note #2.
   6. Mark surface of hardened mounting media meniscus to correspond with areas marked on underside of the slide (Step #1) with HistoTec Pen to maintain tissue orientation in transfer process.
   7. Immerse slide in a Coplin jar of water warmed in a 45°- 60°C water bath; soak for 90 minutes to 3 hours.
   8. Slowly pry meniscus off at edges with a scalpel blade. If meniscus does not peel off easily, return to warm water and continue soaking.
      1. 1. See Procedure Note #3.
   9. Moisten charged slides to receive transferred segments with water.
   10. To transfer one intact segment; place on moistened charged slide, meniscus side up.
   11. To transfer multiple sections; cut removed meniscus with water moistened scalpel blade into segments as marked on medium surface.
       1. 1. Place each cut segment, meniscus side up, onto moistened charged slide in sequential order.
   12. Soak gauze in warm water, place over tissue section. With thumb, press gauze flat transferring water to the section, flattening edges and sealing transferred section to slide.
       1. 1. If a transferred section is placed meniscus down, the specimen will not adhere and will be lost in further steps.
   13. Place slide(s) in a 37°- 60°C oven in a flat horizontal position for 1 hour or longer to securely adhere transferred section to slide.
   14. Soak slide(s) in four changes of xylene for 10 minutes each or longer until all Mount-Quick compound is completely dissolved.
       1. 1. See Procedure Note #4.
   15. Re-hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each. Wash well with distilled water.
   16. Specimen is ready for histochemical, IHC or cytochemical staining.

 

PROCEDURE NOTES:

1. 1. If any residual mounting medium remains on the slide, tissue transfer and staining may be compromised.
   2. The meniscus of Mount-Quick Mounting Media must thoroughly harden (but remain pliable) on the slide before proceeding. To avoid brittleness do not exceed 60°C during the hardening process.
   3. If a hotplate was used to initially dry slides or melt paraffin after original sectioning, it will be difficult to remove an entire intact section from the slide. Extended soaking of mounting media and tissue section does not improve results.
   4. If Mount-Quick is not completely dissolved from the tissue section, subsequent staining may be compromised.
   5. The use of xylene substitutes has not been tested with this product.

 

REFERENCES:

1. 1. Gill, Gary. Cytopreparation: Principles & Practice, Essentials in Cytopathology. New York: Springer Science and Business Media, 2013. 393-395.
   2. Kubier, Patty and Rodney Miller. “Tissue Protection Immunohistochemistry.” American Journal of Clinical Pathology 117 (2002): 194-198.
   3. Miller, Rodney and Patty Kubier. “Immunohistochemistry on Cytologic Specimens and Previously Stained Slides (When No Paraffin Block is Available).” The Journal of Histotechnology 25.4 (2002): 251-257.
   4. Sherman ME, D Jimenez-Joseph, MD Gangi and RR Rojas-Corona. “Immunostaining of Small Cytologic Specimens. Facilitation with Cell Transfer.” Acta Cytologica 38.1 (1994): 18–22.
   5. Zu, Youli, Maryann Gangi and Grace Yang, “Ultrafast Papanicolaou Stain and Cell-Transfer Technique Enhance Cytologic Diagnosis of Hodgkin Lymphoma.” Diagnostic Cytopathology 27.5 (2002): 308-311.
   6. Modifications developed by Newcomer Supply Laboratory.

Out/In Cards mark where a slide has been removed for easy replacement.

Tech Memo 1: May-Grunwald Giemsa Stain for Paraffin Sections

 

SOLUTIONS:

	500 ml	1 Liter
Jenner Stock Stain	Part 1210A	Part 1210B
Giemsa Stock Stain, Wolbach	Part 1121A	Part 1121B

 

Additionally Needed:

Giemsa Control Slides	Part 4240
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Alcohol, Methanol Anhydrous, ACS	Part 12236
Acetic Acid 1%, Aqueous	Part 10012

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

 

APPLICATION:

Newcomer Supply May-Grunwald Giemsa (MGG) Stain procedure for paraffin sections, is used for differential staining of hematopoietic tissue as well as demonstration of some microorganisms.

Jenner Stock Stain combines eosin and methylene blue, while the Giemsa Stock Stain, Wolbach is a mixture of methylene blue, eosin and Azure B. Combining Jenner Stain and Giemsa Stain, Wolbach together into the MGG procedure, provides for intense colorization results.

 

METHOD:

Fixation:  Recommended for hematopoietic tissue

1. 1. Zenker Fixative, Modified, Zinc Chloride (Part 1461)
   2. B-5 Fixative Modified, Zinc Chloride (Part 1015)
   3. Formalin 10%, Phosphate Buffered (Part 1090)

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

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

 

STAINING PROCEDURE:

1. 1. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   2. Rinse in two changes of Methanol (Part 12236); 3 minutes each.
   3. Prepare fresh Working Jenner Stain prior to use; combine, mix well.
      1. 1. Jenner Stock Stain                       20 ml
         2. Distilled Water       20 ml
   4. Place slides in fresh Working Jenner Stain for 6 minutes.
   5. Prepare fresh Working Giemsa Stain; combine, mix well.
      1. 1. Giemsa Stock Stain, Wolbach         3 ml
         2. Distilled Water       47 ml
   6. Place slides, without rinsing, directly from Working Jenner Stain into fresh Working Giemsa Stain for 45 minutes.
   7. Rinse quickly in distilled water.
   8. Differentiate each slide individually in Acetic Acid 1%, Aqueous (Part 10012); 6-10 dips.
      1. 1. Check microscopically for well differentiated nuclei.
   9. Rinse in distilled water.
   10. Dehydrate in two changes each of 95% and 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

Nuclei	Blue/violet
Cytoplasm	Pink/rose to lighter blue shades
Bacteria	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 color range of the stained cells will vary depending upon fixation and degree of differentiation.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 130-131.
   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. 121-122.
   3. Shapiro, Stanley H. and Hilda Laufer. “Observations on Fixation and Staining of Bone Marrow Biopsies.” The Journal of Histotechnology 11.3 (1988): 145-47.
   4. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 157.
   5. Modifications developed by Newcomer Supply Laboratory.

 

Tech Memo 2: May-Grunwald Giemsa Stain for Smears

 

SOLUTIONS:

	500 ml	1 Liter
Jenner Stock Stain	Part 1210A	Part 1210B
Giemsa Stock Stain, Wolbach	Part 1121A	Part 1121B

 

Additionally Needed:

Alcohol, Methanol Anhydrous, ACS

Part 12236

 

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

APPLICATION:

Newcomer Supply May-Grunwald Giemsa (MGG) Stain procedure for smears, is used for differential staining and morphological inspection of peripheral blood smears and bone marrow smears/films.

Jenner Stock Stain combines eosin and methylene blue, while the Giemsa Stock Stain, Wolbach is a mixture of methylene blue, eosin and Azure B. Combining Jenner Stain and Giemsa Stain, Wolbach together into the MGG procedure, provides for intense colorization results.

 

METHOD:

Technique:  Coplin jar or flat staining rack method
Solutions:  All solutions are manufactured by Newcomer Supply, Inc.

 

STAINING PROCEDURE:

1. 1. Prepare a well-made blood smear or bone marrow smear/film with a focus on uniform cell distribution.
   2. Allow smears to thoroughly air-dry prior to staining.
   3. Fix smears by flooding slides with Methanol (Part 12236) for 10-30 seconds.
   4. Prepare fresh Working Jenner Stain prior to use; combine, mix well.
      1. 1. Jenner Stock Stain                       20 ml
         2. Distilled Water       20 ml
         3. Or prepare lesser volumes as needed.
   5. Place slides in fresh Working Jenner Stain for 5-6 minutes.
   6. Prepare fresh Working Giemsa Stain prior; combine, mix well.
      1. 1. Giemsa Stock Stain, Wolbach         5 ml
         2. Distilled Water       45 ml
         3. Or prepare lesser volumes as needed.
   7. Place slides, without rinsing, directly from Working Jenner Stain into fresh Working Giemsa Stain for 30 minutes.
   8. Rinse smears well in distilled water to completely differentiate.
   9. Air-dry slides vertically; examine microscopically.

 

RESULTS:

Erythrocytes	Light pink to light purple
Platelets	Granules – Reddish purple
Lymphocytes/monocytes	Nuclei – Dark purple
	Cytoplasm – Sky blue
Neutrophils	Nuclei – Dark blue
	Granules – Reddish purple
	Cytoplasm – Pale pink
Eosinophils	Nuclei – Blue
	Granules – Red/orange red
	Cytoplasm – Blue
Basophils	Nuclei – Dark blue
	Granules – Purple

 

PROCEDURE NOTES:

1. 1. Timings are suggested ranges. Optimal staining times will depend upon staining intensity preference.
   2. Drain slides after each step to prevent solution carry over.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Hladik Cappellano. Histotechnology: A Self-instructional Text. 4th ed. Chicago: ASCP Press, 2015. 130-131.
   2. McPherson, Richard and Matthew Pincus. Henry’s Clinical Diagnosis and Management by Laboratory Methods. 22nd ed. Philadelphia: Elsevier Saunders, 2011. 522-531.
   3. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 157.
   4. Modifications developed by Newcomer Supply Laboratory.

 

BioSavers™ Biopsy Specimen Wraps are a quality lens paper biopsy wrap that allow optimal infiltration while helping to prevent loss of specimen.

 

SPECIFICATIONS OF BIOSAVERS™ BIOPSY SPECIMEN WRAPS:

• Dimensions: Available in 2″ x 3″ sheets or 2″ x 2″ sheets
• Available in white and pink
• Colors of BioSavers™ Biopsy Wraps sharply contrast to biopsies stained with IPC Blue™ allowing better visualization during embedding & cutting.

 

BIOSAVERS™ BIOPSY SPECIMEN WRAPS MATERIAL BY COLOR:

White & Pink BioSavers™:

• • • Traditional woven lens paper
    • More rigid
    • Able to hold a folded shape that is applied to the biopsy wrap

 

BIOSAVERS™ BIOPSY SPECIMEN WRAPS INSTRUCTIONS FOR USE:

1. Place tissue in the middle of the wrap with plenty of space around all sides.
2. Fold the biopsy specimen wrap in thirds with a slight (about 1/8”) lip of wrap over one edge. (This makes it much easier for the person embedding to open)
3. Fold the remaining ends over to make a closed “envelope” to allow it to fit comfortably in the cassette.

SOLUTIONS:

	500 ml	1 Liter	1 Gallon
Hematoxylin Stain, Mayer Modified	Part 1202A	Part 1202B	Part 1202C
Eosin Y Working Solution	Part 1072A	Part 1072B	Part 1072C

 

Additionally Needed For H&E Staining:

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

 

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

 

APPLICATION:

Newcomer Supply Hematoxylin & Eosin (H&E) Progressive Stain is used for screening specimens in anatomic pathology, as well as for research, smears, touch preps and other applications. In progressive staining, tissue is left in the hematoxylin solution just long enough to reach the proper endpoint. It may be necessary to examine the slides at different timed intervals to determine when staining is optimal.

Hematoxylin Stain, Mayer Modified is a ready-to-use progressive hematoxylin that does not require filtering and does not contain chloral hydrate or alcohol. Due to the progressive staining nature of Mayer hematoxylin, over-staining is less likely and an acid alcohol differentiation step is not required in the staining process.

Eosin Y Working Solution is a ready-to-use counterstain with the ability to distinguish between the cytoplasm of different types of cells by staining cytoplasmic components differing shades and intensities of pink to red.

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

 

METHOD:

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

 

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. Stain with Hematoxylin Stain, Mayer Modified, 10-20 minutes, depending on preference of nuclear stain intensity.
   3. Wash well in tap water for 3 minutes.
   4. Blue slides in Lithium Carbonate, Saturated Aqueous (Part 12215) or Scott Tap Water Substitute (Part 1380) for 10 dips.
      1. 1. See Procedure Note #3.
   5. Wash in three changes of tap water; rinse in distilled water.
   6. Drain water from slides; proceed to 70% ethyl alcohol for 10 dips.
   7. Counterstain in Eosin Y Working Solution for 30 seconds to 3 minutes, depending on preference of intensity.
   8. Dehydrate in two changes of 95% ethyl alcohol for 1 minute each and two changes of 100% ethyl alcohol, 10 dips each.   Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.

 

RESULTS:

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

 

PROCEDURE NOTES:

1. 1. Drain slides after each step to prevent solution carry over.
   2. Do not allow sections to dry out at any point during procedure.
   3. Some laboratory tap water may be alkaline enough to accomplish satisfactory bluing, allowing the elimination of Step #4.
   4. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Carson, Freida L. and Christa Cappellano. Histotechnology: A Self-Instructional Text. 5th ed. Chicago: ASCP Press, 2020. 112-115.
   2. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 36-38.
   3. Modifications developed by Newcomer Supply Laboratory.

Tech Memo 1: Wright-Giemsa Stain, Modified for Tissue Sections

 

SOLUTIONS:

	500 ml	1 Liter	1 Gallon
Wright Stain, Modified	Part 1421A	Part 1421B	Part 1421C
Stock Stain	Part 1120A	Part 1120B

 

Additionally Needed:

Control Slides	Part 4240
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Alcohol, Methanol Anhydrous, ACS	Part 12236

 

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

 

APPLICATION:

Newcomer Supply Wright-Giemsa Stain, Modified for Tissue Sections combines a modified Wright’s formula with a Giemsa Stain Solution for differential staining of hematopoietic tissue and demonstration of bacteria that may be present in the sections. This procedure is applicable for either hand or automated staining processes.

 

METHOD:

Fixation:  Recommended for hematopoietic tissue:

1. 1. 1. Zenker Fixative, Modified, Zinc Chloride (Part 1461)
      2. B-5 Fixative Modified, Zinc Chloride (Part 1015)
      3. 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 PREPARTION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare fresh Working Giemsa Stain:
      1. Distilled Water 40 ml
      2. Giemsa Stock Stain (Part 1120) 5 ml

 

STAINING PROCEDURE: 

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Stop at 95% ethyl alcohol.
      1. 1. See Procedure Notes #1 and #2.
   4. Treat slides in two changes of Methanol (Part 12236); 3 minutes each.
   5. Stain in Wright Stain, Modified for 6 minutes.
   6. Stain in fresh Working Giemsa Stain (Step #2); 60°C oven for 60 minutes.
   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:

Nuclei	Blue
Cytoplasm	Pink to red
Bacteria	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
   3. The color range of stained cells may vary depending upon fixation.
   4. If using a xylene substitute, closely follow the manufacturer’s recommendations for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Shapiro, Stanley H., and Hilda Laufer. “Observations on Fixation and Staining of Bone Marrow Biopsies.” The Journal of Histotechnology3 (1988): 145-47.
   2. Sheehan, Dezna C., and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 155-156.
   3. Modifications developed by Newcomer Supply Laboratory.

 

SOLUTION:

 

SOLUTIONS:                                                                                                                 

	500 ml
Alcian Blue Stain, Alcoholic	Part 1004A
Sodium Sulfate 1%, Aqueous	Part 1388A

 

Additionally Needed:

Acetic Acid, Glacial, ACS	Part 10010
Picric Acid, Saturated Alcoholic	Part 1337
Borax (Sodium Borate) 5%, Saturated Alcoholic	Part 1019
Hematoxylin Stain Set, Weigert Iron	Part 1409
Van Gieson Stain	Part 1404
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Alcohol, Ethyl Denatured, 80%	Part 10843

 

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

 

APPLICATION:

Newcomer Supply Sulfated Alcian Blue (SAB) Stain for Amyloid assists in identifying the extraneous protein deposits of amyloidosis, specifically in myocardial and renal biopsy specimens.

 

METHOD:

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

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

 

PRESTAINING PREPARATION:

1. 1. Prepare the following three fresh solutions:
      1. 1. SAB Staining Solution

 Alcian Blue Stain, Alcoholic                               22 ml

 Sodium Sulfate 1%, Aqueous                           22 ml

 Acetic Acid, Glacial, ACS (Part 10010)                      5 ml

Allow to stand for 30 minutes before use.

1. 1. 1. 2. Acetic Acid/Alcohol Solution

 Alcohol, Ethyl Denatured, 95% (Part 10842)            44 ml

 Distilled Water                                                 44 ml

 Acetic Acid, Glacial, ACS                                 10 ml

1. 1. 1. 3. Picric Acid 2%, Alcoholic Working Solution

 Alcohol, Ethyl Denatured, 80% (Part 10843)           30 ml

Picric Acid, Saturated Alcoholic (Part 1337)             10 ml

 

STAINING PROCEDURE: 

1. 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 Acetic Acid/Alcohol Solution (Step #1b) for 2 minutes.
   4. Place slides in SAB Staining Solution (Step #1a) for 2 hours.
   5. Transfer directly to fresh Acetic Acid/Alcohol Solution (Step #1b) for 2 minutes.
   6. Wash well in tap water; rinse in distilled water.
   7. Alkalinize slides in Borax (Sodium Borate) 5%, Saturated Alcoholic (Part 1019) for 30 minutes.
   8. Wash well in tap water; rinse in distilled water.
   9. Prepare Hematoxylin Stain, Weigert Iron (Part 1409); combine, mix well.
      1. 1. Solution A: Ferric Chloride, Acidified   20 ml
         2. Solution B: Hematoxylin 1%, Alcoholic    20 ml

1. 10. Stain in Hematoxylin Stain, Weigert Iron for 5 minutes.
   11. Wash in tap water for 10 minutes; rinse in distilled water.
   12. Differentiate each slide individually; dip in Picric Acid 2%, Alcoholic Working Solution (Step #1c) for 20 seconds.
   13. Rinse briefly in tap water; 2-3 dips.
   14. Counterstain with Van Gieson Stain (Part 1404) for 3 minutes.
   15. 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:

Amyloid deposits	Shades of green; pale to brilliant jade
Fibrin, muscle, cytoplasm	Yellow
Collagen, stroma	Red
Nuclei	Black

Note: Green staining tissues other than amyloid are morphologically distinguishable.

1. 1. Mast cells stain dense bright green, with typical cytoplasmic granularity.
   2. Red blood cells occasionally appear very pale green.
   3. Calcium deposits stain a dirty blue/green.

 

PROCEDURE NOTES:

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

 

REFERENCES:

1. 1. Lendrum, A. C., W. Slidders and D. S. Fraser. “Renal Hyalin: A Study of Amyloidosis and Diabetic Fibrinous Vasculosis with New Staining Methods.” Journal of Clinical Pathology 25 (1972): 373-96.
   2. Pomerance, Ariela, Gerard Slavin and Josephine McWatt. “Experience with the Sodium Sulphate-Alcian Blue Stain for Amyloid in Cardiac Pathology.” Journal of Clinical Pathology, 29 (1976): 22-26.
   3. Modifications developed by Newcomer Supply Laboratory.

SOLUTIONS:

	500 ml	1 Liter
Giemsa Stock Stain, Wolbach	Part 1121A	Part 1121B
Rosin 10%, Alcoholic Stock	Part 13398A

 

Additionally Needed:

Giemsa Control Slides	Part 4240
Xylene, ACS	Part 1445
Alcohol, Ethyl Denatured, 100%	Part 10841
Alcohol, Ethyl Denatured, 95%	Part 10842
Acid Alcohol 1%	Part 10011
Alcohol, Methanol Anhydrous, ACS	Part 12236

 

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

 

APPLICATION:

Newcomer Supply Wolbach Giemsa Stain is used for differential staining of hematopoietic tissue and demonstration of bacteria and rickettsia that may be present in the sections.

 

METHOD:

Fixation:  Recommended for hematopoietic tissue:

1. 1. 1. Zenker Fixative, Modified, Zinc Chloride (Part 1461)
      2. B-5 Fixative Modified, Zinc Chloride (Part 1015)
      3. Formalin 10%, Phosphate Buffered (Part 1090)

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

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

 

PRESTAINING PREPARATION:

1. 1. If necessary, heat dry tissue sections/slides in oven.
   2. Prepare fresh Working Wolbach Giemsa Stain Solution; combine and mix well.
      1. 1. Distilled water           50 ml
         2. Giemsa Stock Stain, Wolbach             2 ml
         3. Methanol (Part 12236)             2 ml

 

STAINING PROCEDURE: 

1. 3. Deparaffinize sections thoroughly in three changes of xylene, 3 minutes each. Hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each.  Wash well with distilled water.
      1. 1. See Procedure Notes #1 and #2.
   4. Treat slides with Acid Alcohol 1% (Part 10011) Solution for 5 minutes.
      1. 1. See Procedure Note #3.
   5. Wash in running tap water for 5 minutes.
   6. Stain in fresh Working Wolbach Giemsa Stain Solution (Step #2) for 60 minutes at room temperature.
      1. 1. See Procedure Note #4.
   7. Differentiate each slide individually in Rosin 10%, Alcoholic until sections are purplish-pink; 5-10 dips. Check microscopically.
      1. 1. Rinse in 100% ethyl alcohol to stop differentiation.
   8. Dehydrate in two changes of 100% ethyl alcohol. Clear in three changes of xylene, 10 dips each; coverslip with compatible mounting medium.
      1. 1. Do not use a 95% ethyl alcohol dehydration step.

 

RESULTS:

Nuclei	Blue/violet
Cytoplasm	Pink/rose to lighter blue shades
Bacteria	Blue
Rickettsia, inclusions	Reddish-purple

 

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. Acid Alcohol treatment ensures an acid pH and improved staining.
   4. For increased staining, stain in Working Wolbach Giemsa Stain Solution at 60°C for 60 minutes.
      1. 1. Additional differentiation may be required.
   5. The color range of cells may vary depending on fixative and degree of differentiation.
   6. If using a xylene substitute, follow manufacturer’s recommendation for deparaffinization and clearing steps.

 

REFERENCES:

1. 1. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 119-120.
   2. Sheehan, Dezna C. and Barbara B. Hrapchak. Theory and Practice of Histotechnology. 2nd ed. St. Louis: Mosby, 1980. 156-157.
   3. Wolbach, S. Burt and John Todd. The Etiology and Pathology of Typhus. S.l.: Harvard University Press, 1922. 13-14.
   4. Modifications developed by Newcomer Supply Laboratory.

• • • Aminosilane Coated Slides
    • Available in Blue, Green, Pink, White and Yellow
    • Sold by the case
    • Great pricing!

Out/In Cards mark where a cassette has been removed for easy replacement.

 

Eliminates Harmful Vapors in Minutes!

PolyForm-F™ Formalin/Formaldehyde Neutralizer is a free granule spill product for the safe and rapid control of harmful vapors from formaldehyde, formaldehyde-based embalming solutions, glutaraldehyde, 10% buffered formalin and other aldehydes, including: Cidex, Cidex-OPA, Metracide, Wavacide & OmniCide.

 

POLYFORM-F™ FORMALIN/FORMALDEHYDE NEUTRALIZER:

• Harmful vapors are eliminated in 2 to 3 minutes
• Formaldehyde & other aldehydes are neutralized & irreversibly polymerized
• Reaction leaves a non-hazardous biodegradable polymer
• End product is safe & simple for personnel to clean-up & dispose

 

USES OF POLYFORM-F™ FORMALIN/FORMALDEHYDE NEUTRALIZER:

• Hospital & Medical Labs
• Histology & Pathology Labs
• Operating Rooms
• Morgues & Autopsy Suites
• Medical Schools & Universities
• Specimen Transportation
• Shipping & Receiving Areas
• Wherever aldehydes are used

 

DIRECTIONS OF USE FOR POLYFORM-F™ FORMALIN/FORMALDEHYDE NEUTRALIZER:

1. Consult SDS for the spilled chemical solution, to become familiar with its chemical properties and health & safety requirements.
2. Select and wear proper personnel protective equipment as recommended or noted on the SDS for the spilled chemical solution.
3. Evacuate area as necessary to ensure safety of personnel.
4. Eliminate all sources of ignition and insure there is adequate ventilation in the area of the spill.
5. Add PolyForm-F™ Formalin/Formaldehyde Neutralizer around the perimeter of the spill to dike the liquid and prevent spreading.  From the upwind side, cover the entire area from edge to edge at a ratio of approximately one-to-one, completely covering the spill and taking care to avoid vapors and splashing.
6. Once PolyForm-F™ Formalin/Formaldehyde Neutralizer is applied, DO NOT MIX, allow to stand for approximately 12 to 15 minutes.
7. Cleanup spill residue by using a plastic dust-pan and disposable towels, then place the collected spill residue in adequate waste bag.
8. After spill residue has been removed from spill area, wipe up the spill area with cold tap water, using a towel, sponge or mop.
9. “Post-Cleanup” the spill area with mild detergent solution recommended by your  facility for the final floor and/or counter cleanup.
10. In most cases,  the PolyForm-F™ Formalin/Formaldehyde Neutralizer treated spill residue may be disposed of as a non-hazardous waste.

• If the spilled solutions contain heavy metals, then the material must be handled as a potential hazardous waste.
• If human or animal tissue has been in contact with the spilled formalin, then the spill residue material may be handled as a potential bio-medical waste.
• Always dispose of all spill residue waste in accordance with users’ facility recommendations and follow all Federal, State and Local environmental regulations.

“The first line of control for most accidental releases of corrosive materials.”

This unique flowable powder was developed to facilitate the rapid and immediate control of spilled corrosive materials by:

• Solidifying and neutralizing on contact.
• Immediately stopping the spread of hazardous chemicals.
• Reducing hazardous fumes and vapors.
• Reducing the corrosiveness of spilled materials, which reduces chemical attack on floors and other surfaces, as well as on the environment.
• Producing a controlled chemical reaction, rather than the usual violent reaction associated with the neutralization of strong corrosives.
• Eliminating the disposal problems typically associated with generic sorbents.
• Producing a dry powder which can be cleaned up and disposed of as a nonhazardous waste.

 

Directions for Base Control Spill Powder:

1. Consult SDS of spilled material to become familiar with its chemical properties and safety and health requirements.
2. Select and wear proper personal protective equipment for the spilled material.
3. Evacuate area as necesary to ensure the safety of all personnel.
4. Eliminate all sources of ignition and ensure that there is adequate ventilation before applying product.
5. Apply Base Control™ to the spill area, working from the upwind side and start from the outside of spill and working toward the center. If the spilled liquid is running, then apply product downstream of the spill to form a dam.
6. Carefully mix with a non-reactive paddle or shovel until all liquid is solidified.
7. Determine level of neutralization by using a pH test kit.
8. Let solidified/neutralized material cool prior to clean up.
9. Follow final clean up procedures established by your facility or company.
10. Dispose of neutralized waste in accordance with Federal, State and Local environmental regulations.

 

DO Use on the Following Bases:	DO NOT  Use on the following:
Ammonium Hydroxide = aqua ammonia	Chlorine
Anhydrous Ammonia	Hydrogen Peroxide
Monoethanolamine	Oxidizers
Morpholine	Sodium Amide
Potassium Hydroxide = caustic potash	Sodium Hypochloride
Triethylalamine	Sulfurous Fluoride
Sodium Hydroxide = caustic soda
Sodium Metasilicate Solution
Most Alkali Detergents

* The lists given herein are general and do not necessarily include all the materials Acid Handler™, Polyform-F™, Base Control™, and Solvent Handler™ can or cannot be used on. If you would like an exotic species tested, or have questions as to application of the products, call Newcomer Supply or American Bio-Safety, Inc. We will try applying our products to your substance and determine a suitable procedure for spill situations. 

“The first line of control for most accidental releases of corrosive materials.”

This unique flowable powder was developed to facilitate the rapid and immediate control of spilled corrosive materials by:

• Solidifying and neutralizing on contact.
• Immediately stopping the spread of hazardous chemicals.
• Reducing hazardous fumes and vapors.
• Reducing the corrosiveness of spilled materials, which reduces chemical attack on floors and other surfaces, as well as on the environment.
• Producing a controlled chemical reaction, rather than the usual violent reaction associated with the neutralization of strong corrosives.
• Eliminating the disposal problems typically associated with generic sorbents.
• Producing a dry powder which can be cleaned up and disposed of as a nonhazardous waste.

 

Directions for Acid Handler Spill Powder:

1. Consult SDS of spilled material to become familiar with its chemical properties and safety and health requirements.
2. Select and wear proper personal protective equipment for the spilled material.
3. Evacuate area as necessary to ensure the safety of all personnel.
4. Eliminate all sources of ignition and ensure that there is adequate ventilation before applying product.
5. Apply Acid Handler™ to the spill area, working from the upwind side and start from the outside of spill and working toward the center. If the spilled liquid is running, then apply product downstream of the spill to form a dam.
6. Carefully mix with a non-reactive paddle or shovel until all liquid is solidified.
7. Determine level of neutralization by using a pH test kit.
8. Let solidified/neutralized material cool prior to clean up.
9. Follow final clean up procedures established by your facility or company.
10. Dispose of neutralized waste in accordance with Federal, State and Local environmental regulations.

 

DO Use on the Following Acids:	DO NOT  Use on the following:
Acetic Acid	Chlorine
Acetic Anhydride	Concentrated Hydrofluoric Acid
Acetyl Chloride	Hydrogen Peroxide
Aluminum Chloride	Iodic Acid
Chlorosulfonic Acid	Oxidizers
Chromic Acid Solutions-Chromium Waste Haz	Pentrafluoride (“Super-Acids”)
Citric Acid	Picric Acid
Dodecylbensylsulfonic Acid	Sodium Amide
Formic Acid	Sulfurous Fluoride Antimony
Glacial Acetic Acid
Hydrochloric Acid
Muriatic Acid
Hydrofluosilic Acid
Nitric Acid
Perchloric Acid
Phosphoric Acid
Phosphoric Anhydride
Phophorous Pentoxide
Phophorous Trichloride
Sulfonic Acid
Sulfuric Acid
54% Hydrofluoric Acid Solution

 

CLICK HERE for BORAX, cata #1019 in Reagents, Chemicals & Buffers Section.

Solution for setting tissue marking dye in special situations. Spray onto dyed tissue as needed for better adherence through solutions. Can be helpful on fatty tissues.

Hold glass slides with ease. (You’ll have to hide these to keep them.) LOVE ‘EM

 

Click here for product information.

 

 

 

These quality control slides are intended to be used to verify histological techniques and reagent reactivity.  They are to be used for the qualitative purpose of determining positive or negative results, and are not intended to be used for any quantitative purpose.  The first serial section within the control box is stained and provided for your reference.

Newcomer Supply, Inc., takes reasonable measures to deliver quality control slides that are as consistent as possible. However, the characteristics of quality control slides may be dissimilar due to variations in the reagents, stains, techniques, laboratory conditions, and tissue sources used.

CD45, LCA is routinely used to aid the differential diagnosis of undifferentiated neoplasms, whenever malignant lymphoma is suspected by the morphological or clinical data.  CD45, LCA labels the cell membrane of the majority of lymphoid cells.

These CD45, LCA control slides are produced from human surgical or autopsy tissues.  At Newcomer Supply we use a manual method of performing immunohistochemical stains, specifically avoiding automation, in order to provide reactive control slides for even the less aggressive methods of staining that our customers may be using.

Please note: A section of adipose tissue has been included on the slide for use as a negative control.

METHOD:

Fixation:                    10% Neutral Buffered Formalin

Technique:                Paraffin sections cut at approximately 4 microns

 

REAGENTS:

Epitope Retrieval:

Target Retrieval Solution, Dako

Catalog #S1699

Diluent:

Antibody Diluent, Cell Marque

Catalog #936B

Blocking Reagent:

3% Hydrogen Peroxide, Newcomer Supply, (30%)

Catalog #1206A

Primary Antibody:

CD45, LCA Monoclonal Antibody (1:100), Cell Marque

Catalog #145M

Negative Antibody:

Mouse Negative Control Serum, Cell Marque

Catalog #932B-02

Detection Kit: 

HiDef Detection^TMHRP Polymer System, Cell Marque

Catalog #954D-20

Chromogen:

DAB Substrate Kit, Cell Marque

Catalog #957D-10

Buffer: 

Tris Buffered Saline, 0.05M, pH 7.6, 10X, Newcomer Supply

Catalog #140304

Hematoxylin:

Hematoxylin Stain Solution, Mayer Modified, Newcomer Supply

Catalog #1202

 

STAINING PROCEDURE:

1. Deparaffinize sections in 3 changes of xylene, then hydrate through graded ethyl alcohols to distilled water.
2. Place slides in preheated (95°C) Target Retrieval Solution in steamer for 20 minutes.  Remove and allow to cool for 20 minutes at room temperature.
3. Rinse in 3 changes of buffer ending with a distilled water rinse.
4. Tap off excess water and apply enough freshly made 3% Hydrogen Peroxide to cover specimen.  Incubate for 5 minutes.
5. Rinse slides gently in distilled water from a wash bottle and place in buffer bath.
6. Tap off excess buffer and apply enough primary antibody (1:100) or negative control reagent to cover specimen.  Incubate for 30 minutes.
7. Rinse slides gently in buffer from a wash bottle, then place in buffer bath for 5 minutes.
8. Tap off excess buffer and apply enough Amplifier to cover specimen.  Incubate for 10 minutes.
9. Rinse slides gently in buffer from a wash bottle and place in buffer bath.
10. Tap off excess buffer and apply enough HRP Polymer to cover specimen.  Incubate for 10 minutes.
11. Rinse slides gently in buffer from a wash bottle and place in buffer bath.
12. Prepare enough substrate-chromogen DAB for the amount of slides being stained.
13. Tap off excess buffer and apply enough DAB to cover specimen.  Incubate for 5 minutes.
14. Rinse slides gently in a distilled water from a wash bottle, then place in distilled water bath.
15. Counterstain lightly with Hematoxylin Stain Solution, Mayer Modified or other aqueous hematoxylin.
16. Blue sections in warm tap water.
17. Dehydrate sections through graded ethyl alcohols, clear in xylene or substitute and coverslip with a compatible mounting medium.

 

RESULTS:

CD45, LCA (Leucocyte Common Antigen)           Brown

Adipose tissue                                                     Negative

 

REFERENCES:

1. Cell Marque HiDef Detection^TM Polymer datasheet.
2. Cell Marque CD45, LCAantibody datasheet.

The enclosed quality control slides are intended to be used to verify histological techniques and reagent reactivity.  They are to be used for the qualitative purpose of determining positive or negative results, and are not intended to be used for any quantitative purpose.  The first serial section within this control box is stained and provided for your reference.

Newcomer Supply, Inc. takes reasonable measures to deliver quality control slides that are as consistent as possible.  However, the characteristics of quality control slides may be dissimilar due to variations in the reagents, stains, techniques, laboratory conditions, and tissue sources used.

CD20, B-Cell is a marker for detecting normal and neoplastic B-Cells.  B-Cells are generally located inside the follicles in lymph nodes and tonsils.

These control slides are produced from human surgical or autopsy tissues.  At Newcomer Supply we use a manual method of performing immunohistochemical stains, specifically avoiding automation, in order to provide reactive control slides for even the less aggressive methods of staining that our customers may be using.

A section of normal human kidney is included on the slide for use as a negative control.  

 

METHOD:

Fixation:                  10% Neutral Buffered Formalin

Technique:              Paraffin sections cut at approximately 4 microns

 

REAGENTS:

Blocking Reagent:	3% Hydrogen Peroxide, Newcomer Supply, (30%)	Catalog #1206A
Primary Antibody:	CD20, Monoclonal Antibody,Cell Marque (1:250)	Catalog #120M-86
Negative Antibody:	Mouse Negative Control Serum, Cell Marque	Catalog #932B-02
Detection Kit:	HiDef Detection™ HRP Polymer System, Cell Marque	Catalog #954D-20
Chromogen:	DAB Substrate Kit, Cell Marque	Catalog #957D-10
Buffer:	Tris Buffered Saline 0.05M pH 7.6, 10X	Catalog #140304
Hematoxylin:	Hematoxylin Stain Solution, Mayer Modified	Catalog #1202

 

STAINING PROCEDURE 

1. Deparaffinize sections in 3 changes of xylene, then hydrate through graded ethyl alcohols to distilled water.
2. Rinse slides in distilled water.
3. Tap off excess water and apply enough 3% Hydrogen Peroxide to cover specimen.  Incubate 5 minutes.
4. Rinse slides gently in distilled water from a wash bottle and place in Tris buffer bath for 5 minutes.
5. Tap off excess buffer and apply enough primary antibody (1:250) or negative control reagent to cover specimen.  Incubate 30 minutes.
6. Rinse slides gently with buffer solution from a wash bottle and place in buffer bath.
7. Tap off excess buffer and apply enough Amplifier solution to cover specimen.  Incubate 10 minutes.
8. Rinse slides gently with buffer solution from a wash bottle and place in buffer bath.
9. Tap off excess buffer and apply enough HRP to cover specimen.  Incubate 10 minutes.
10. Rinse slides gently with buffer solution from a wash bottle and place in buffer bath.
11. Prepare enough substrate-chromogen DAB for the amount of slides being stained.
12. Tap off excess buffer and apply enough of the prepared substrate-chromogen DAB to cover specimen.  Incubate 5 minutes.
13. Rinse slides gently with distilled water from a wash bottle and place in distilled water bath.
14. Counterstain lightly with Hematoxylin Stain Solution, Mayer Modified or other aqueous hematoxylin and blue sections in ammonia or warm tap water if needed.
15. Dehydrate sections through graded ethyl alcohols, clear in xylene or substitute and coverslip with a coverslip with a compatible mounting medium.

 

RESULTS:

B Cells	Brown cytoplasmic staining
Normal Kidney	Negative

 

REFERENCES:

1. Cell Marque HiDef Detection™ HRP Polymer System Kit datasheet.
2. Cell Marque B-Cell, CD20 datasheet.

PRODUCT:                                             

	8 oz Can	12 Can/Case
Permaslip Mounting Medium	Part 6530B	Part 6530B

   

Additionally Needed for Tissue/Cell Transfer Procedure:

Xylene, ACS	Part 1445
HistoTec™ Pen	Part 5725
Scalpels, Finger         OR Scalpels, Flat Edge	Part 6810     OR Part 5545
Superfrost® Plus Slides               OR NewSilane Adhesive Slides	Part 6203W     OR Part 5070
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 Permaslip Mounting Medium is a 3 in 1 mounting medium that can be used as standard mounting medium, liquid coverslip (coverglass free) mounting medium and for tissue transfer procedures.

The tissue transfer procedure involves the use of Permaslip to transfer sections from a prepared microscopic slide when specimen availability is limited and further diagnostic workup is needed.  Staining quality will not be affected and histochemical, immunohistochemistry (IHC) and cytochemical procedures performed on transferred samples produce reliable results. Examples when tissue transfer is applicable include:

• • • Paraffin sections of small biopsies or minimal tissue
    • Biopsies with a small foci of neoplasm
    • Paraffin block is unavailable
    • Cytology preparations, fine needle aspirates (FNA)
    • Transfer of tissue from an uncharged slide to charged slide(s)
    • Preparing multiple slides from a single slide preparation
    • Restoring broken slides back to a whole mount

The tissue transfer procedure takes time to obtain a well-prepared end product. Practicing the technique prior to performing on limited specimens is suggested. Reduced timings/short cuts shouldn’t be taken.

 

METHOD:

Coverslipping: Paraffin or frozen sections
Tissue/cell transfer: Stained and unstained paraffin sections, cytology preparations and smears

 

COVERSLIPPING PROCEDURE:

1. 1. Complete staining procedure; dehydrate, clear with xylene and blot excess xylene from slide edges.
   2. Apply Permaslip Mounting Medium to fully cover tissue section(s).
      1. 1. Standard coverglass; install one edge first, allowing air bubbles to escape as opposite edge is lowered to slide.
         2. Liquid coverglass (coverglass free); allow to dry flat for 30 minutes before handling and viewing microscopically.
   3. Dry slides a minimum of 24-48 hours before filing.

 

TISSUE/CELL TRANSFER PROCEDURE:

1. 1. Mark tissue area(s) that is the focus of tissue transfer on the underside of original slide with a HistoTec Pen (Part 5725).
   2. For previously stained sections:
      1. 1. Soak slide in xylene until coverslip is removed without damage to tissue section.
         2. Run slide through several changes of xylene to ensure removal of residual mounting medium.
         3. Do not proceed through alcohol steps.
         4. See Procedure Note #1.
   3. For unstained paraffin sections or smears:
      1. 1. Soak slides in several changes of xylene, a minimum of 10 minutes. Do not proceed through alcohol steps.
   4. Spread Permaslip Mounting Medium on xylene-coated slides with an applicator stick. Use sufficient Permaslip to form a meniscus over the entire tissue section or cell preparation.
      1. 1. Slide must be xylene coated and not allowed to dry out.
   5. Place slide in a flat horizontal position in a 60°C oven for 90 minutes to 2 hours or in a 37°C oven overnight until meniscus hardens.
      1. 1. See Procedure Note #2.
   6. Mark surface of hardened mounting medium meniscus to correspond with areas marked on underside of the slide (Step #1) with HistoTec Pen to maintain tissue orientation in transfer process.
   7. Immerse slide in a Coplin jar of water warmed in a 45°- 60°C water bath; soak for 90 minutes to 3 hours.
   8. Slowly pry meniscus off at edges with a scalpel blade. If  meniscus does not peel off easily, return to warm water and continue soaking.
      1. 1. See Procedure Note #3.
   9. Moisten charged slides to receive transferred segment with water.
   10. To transfer one intact segment; place on moistened charged slide, meniscus side up.
   11. To transfer multiple sections; cut removed meniscus with water moistened scalpel blade into segments as marked on medium surface.
       1. 1. Place each cut segment, meniscus side up, onto moistened charged slide in sequential order.
   12. Soak gauze in warm water; place over tissue section. With thumb, press gauze flat transferring water to the section, flattening edges and sealing transferred section to slide.
       1. 1. If a transferred section is placed meniscus down, the specimen will not adhere and will be lost in further steps.
   13. Place slide(s) in a 37°- 60°C oven in a flat horizontal position for 1 hour or longer to securely adhere transferred section to slide.
   14. Soak slide(s) in four changes of xylene for 10 minutes each or longer until the Permaslip compound is completely dissolved.
       1. 1. See Procedure Note #4.
   15. Re-hydrate through two changes each of 100% and 95% ethyl alcohols, 10 dips each. Wash well with distilled water.
   16. Specimen is ready for histochemical, IHC or cytochemical staining.

 

PROCEDURE NOTES:

1. 1. If any residual mounting medium remains on the slide, tissue transfer and subsequent staining may be compromised.
   2. The meniscus of Permaslip Mounting Medium must thoroughly harden (but remain pliable) on the slide before proceeding. To avoid brittleness do not exceed 60°C during the hardening process.
   3. If a hotplate was used to initially dry slides or melt paraffin after original sectioning, it will be difficult to remove an entire intact section from the slide. Extended soaking of mounting medium and tissue section does not improve results.
   4. If Permaslip is not completely dissolved from the tissue section, subsequent staining may be compromised.
   5. The use of xylene substitutes has not been tested with this product.

 

REFERENCES:

1. 1. Gill, Gary. Cytopreparation: Principles & Practice, Essentials in Cytopathology. New York: Springer Science and Business Media, 2013. 393-395.
   2. Kubier, Patty and Rodney Miller. “Tissue Protection Immunohistochemistry.” American Journal of Clinical Pathology 117 (2002): 194-198.
   3. Luna, Lee G. Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology. 3rd ed. New York: Blakiston Division, McGraw-Hill, 1968. 67-69.
   4. Miller, Rodney and Patty Kubier. “Immunohistochemistry on Cytologic Specimens and Previously Stained Slides (When No Paraffin Block is Available).” The Journal of Histotechnology 25.4 (2002): 251-257.
   5. Weibel, E. “The Preparation of Serial Microscopic Sections in Form of Plastic Films.” Lecture, Annual meeting of the International Academy of Pathology, Boston, April 1959.
   6. Modifications developed by Newcomer Supply Laboratory.

Available in 7 colors.

Available in 7 colors.

1/8 round.

1/8 round.

Holds 25 slides

• Self locking lid
• Designed for easy stacking
• Cork lined bottom with embossed numerals on bottom of box
• Index paper on inside lid for documented slides

 

Approx 72 slides/box, Sold by the gross.

10 horizontal slots hold 10 to 18 slides (3″ x 1″) effectively. Dimensions: 3″ x 2 1/4″ x 1 1/4″. Includes glass cover. Not for use with slide racks.

Polypropylene, rectangular slide box with hinged flip top lid. 5 slide capacity.

Economical and reusable.  Polyethylene mailers have a sturdy resealable clasp.  Ideal for shipping specimen slides.

Made of sturdy cardboard to protect the slides.  Available to hold one, two or four slides for transport, with or without coverglass.Thumb groove for easy slide removal.

 

Chills to -60F.  Contains no CFC’s or HCFC’s.  Straw attached to can to localize spray onto blocks and fresh tissue.

For pricing and part number,   Click here for product information.

 

For pricing and part number, this product is located in our Laboratory Essentials section.  Click here for product information.

 

An easy way to clean up wax from instruments and counter tops!

 

• Use as an effective mold release
• All natural and non-toxic
• Cucumber melon scent

 

Index Tab Cards allow for easy categorizing and partitioning of slides in the storage file drawers.

 

Dimensions of Card:

3 1/4″ (H)  X  1 1/8″ (W)

Liquid Neutralizer for Formalin Waste.

Click Here for information, part number and pricing.

18 squeezable dispensers. 3 each of black, blue, green, red, yellow & orange in foam reusable base. 4ml dye/dispenser.

Regular size blade with small attached non-slip handle. No need to use a blade without a handle ever again! Ultimate in dexterity.

Cardboard slide trays hold standard (3″ x 1″) slides. The horizontal compartments are recessed protecting the specimens when the cover is closed.

(use: Additional step after fixation to reduce C-J infectivity.)

 

 

See BUFFERS in Reagents, Chemicals & Buffers Section.

 

CLICK HERE for part number and pricing

 

This product is located in our Safety section

 

 

 

CLICK HERE for part number and pricing

 

This product is found in our Grossing & Embedding section

 

CLICK HERE for part number and pricing.

 

This product is located in our Grossing & Embedding section

Unique, permanent mounting media designed for coverslipping H&E stains from water or alcohol. Reduce time in eosin if cover slipping from water. Sets in 10 minutes.

 

CLICK HERE for part number and pricing

Fc Receptor Blocker, Azide Free can be used as a blocking agent to block Fc receptors present on hematopoetic (blood derived), lymphoid cells/tissues and tumor cells and cell lines.  The binding of the Fc region of the primary or the secondary antibody to the Fc receptors present on the surface of leukocytes (white blood cells) or tumor cells gives rise to non-specific binding and is observed as background or non-specific staining in IHC ICC, immunofluorescence and flow cytometry assays.

 

Fc Receptor Blocker, Azide Free does not contain sodium azide or thimerosol and it is therefore suitable for effective blocking of Fc receptors in non-fixed live cells of lymphoid and hematopoietic origin as well as tumor cells and cell lines that carry Fc receptors on their surface.  Fc Receptor Blocker, Azide Free is especially helpful for blocking Fc receptors in tumor cells or cell lines employed in functional assays where sodium azide and other preservatives cause endocytosis and result in cytotoxicity and cell damage.

 

STORAGE CONDITIONS:

Store in refrigerator at 2-8°C through the expiration date noted on the vial label.  DO NOT FREEZE.

 

PRODUCT FORMAT:

Working solution, NO dilution or adjustments necessary.

 

INSTRUCTIONS FOR USE OF THE FC RECEPTOR BLOCKER AZIDE FREE:

For blocking Fc receptors in live cells and for use in functional assays:

Fc Receptor Blocker Azide Free is used for blocking Fc receptors in live cells and cell lines assays that employ live and non-fixed cells; and further for functional assays where presence of sodium azide and other preservatives cause endocytosis, cytotoxicity and cell damage.  Live cell assays and functional assays are varied and are many in numbers.

 

To block Fc receptors, present on live cells and cell lines that express Fc receptors:

Live cells must be exposed to Innovex Fc Blocker for a period of at least 30 minutes.  The optimal incubation time with Fc Blocker must be determined by the lab user for the specific cells, cell lines, species and the assay condition employed.

1. Add 0.2 to 0.5 ml of Fc Receptor Blocker Azide Free for 10⁶ (1 million) cells.
2. Incubate for 30 minutes to 1 hour on ice or at room temperature.
3. Rinse with assay wash buffer or DI water.
4. Proceed with assay.

 

For Flow Cytometry:

1. Use whole blood OR lyse or ficol blood as usual.
2. Add 0.2 ml of Fc Receptor Blocker Azide Free for 10⁶ (1 million) cells.
3. Incubate for 30 minutes on ice or at room temperature.
4. Wash twice in assay wash buffer.
5. Proceed with antibody labeling.

 

(use: Fixative for blood smears.)

Dilute 1 part concentrate to 4 parts DI Water.  One gallon of concentrate makes 5 gallons of 10% NBF.   Thoroughly mixes in 30 seconds.  One case will produce 20 gallons of NBF!

Benefits:

• Reduces shipping costs!
• Saves 80% of valuable storage space!
• Already Buffered!

 

Good news. We eliminated the Chloral Hydrate! Excellent for IHCs.

See HEMATOXYLIN, Mayer in Standard & Special Stains Section.

(use: Fixative for blood smears.)

Ultra-Purified Paraffin wax with additives to enhance infiltration, impart elasticity, and provide cutting ease.  Eliminates “over-hardened” and “under-processed” tissue.  Special closed manufacturing process prevents contaminants and debris from entering the product.  Packaging process ensures zero condensation which means no possibility of water in wax.

Ultra-Purified Paraffin wax with additives to enhance infiltration, impart elasticity, and provide cutting ease.  Eliminates “over-hardened” and “under-processed” tissue.  Special closed manufacturing process prevents contaminants and debris from entering the product.  Packaging process ensures zero condensation which means no possibility of water in wax.

Ultra-Purified Paraffin wax with additives to enhance infiltration, impart elasticity, and provide cutting ease.  Eliminates “over-hardened” and “under-processed” tissue.  Special closed manufacturing process prevents contaminants and debris from entering the product.  Packaging process ensures zero condensation which means no possibility of water in wax.

 

CLICK HERE to see complete kit #9168 in Stain Kits Section

Periodic Acid Methenamine Method

 

CLICK HERE to see GOMORI BASEMENT MEMBRANE Set #1131 in Standard & Special Stains Section

or #9167 in Stain Kits Section