From relia1 at earthlink.net Fri Nov 5 11:29:05 2021 From: relia1 at earthlink.net (Pam Barker) Date: Fri, 5 Nov 2021 12:29:05 -0400 Subject: [Histonet] Resume Help - Free of charge for my #histopeeps ! Message-ID: <011401d7d262$407e0e70$c17a2b50$@earthlink.net> Hi Histonetters!! Need some help with a resume? Changing jobs? Going for a promotion? Preparing a presentation? Transitioning temp to perm or vice versa? email me at relia1 at earthlink.net for help. No charge #ilovemyhistopeeps #jobs4myhistopeeps Thanks-Pam Right Time, Right Place, Right Move with RELIA! Providing excellent service exclusively to the Histology Community! Thank You! Pam M. Barker Pam Barker President/Senior Recruiting Specialist-Histology RELIA Solutions Specialists in Allied Healthcare Recruiting 5703 Red Bug Lake Road #330 Winter Springs, FL 32708-4969 Phone: (407)657-2027 Cell: (407)353-5070 FAX: (407)678-2788 E-mail: relia1 at earthlink.net https://www.facebook.com/RELIASolutionsforhistologyprofessionals www.linkedin.com/in/reliasolutions #jobs4myhistopeeps #ilovemyhistopeeps #histopeeps Follow my hashtags and make your day great and your career greater!! From ccorbin at bloomu.edu Mon Nov 8 12:50:53 2021 From: ccorbin at bloomu.edu (Corbin, Clay) Date: Mon, 8 Nov 2021 18:50:53 +0000 Subject: [Histonet] Plasma from cap tube Message-ID: Hey folks, I?ve started a project using owl blood. We need to isolate the plasma. Spinning down the blood is the easy part. I?m sort of confused on the best way to draw off the plasma. I?ve heard rumors about hypo needles that do the job, but the ones I?m finding seem too short. Anyone with some hints on the best way to separate formed elements and plasma from capillary tubes? Thanks! Clay From relia1 at earthlink.net Wed Nov 10 11:15:03 2021 From: relia1 at earthlink.net (Pam Barker) Date: Wed, 10 Nov 2021 12:15:03 -0500 Subject: [Histonet] Hi Histonetters!! Resume Tune-up Free of Charge for my Subscribers! Message-ID: <000001d7d656$8139a850$83acf8f0$@earthlink.net> Hi Histopeeps, Resumes they?re not just for job hunting anymore! Did you know it is a great idea to always keep your resume updated? Resumes and CVs are used for many things besides job hunting: Looking at a raise or promotion? ? Does your resume list all of your accomplishments so that you can present them to your supervisor? Ever thought about giving a presentation or writing an article?? ? A great CV always makes a presentation much more polished. How about just for you? ? It feels great to see how far you have come in your career and might help with direction for what?s next. How about if you are transitioning? ? Say going back to permanent work from travel or vice versa or want to move into another area of histology. And of course good old fashioned job hunting! Histopeeps, Let me help you get your resume tuned up!!? It?s free of charge as a service to my Histopeeps!! Here?s what you need to do: Reply to this email with I?m in and we can get started. Job Hunting? Here are some of our Hottest opportunities. All of these are permanent positions. Most are RELIA Exclusives and all of my clients offer excellent compensation, benefits and most offer relocation/sign on bonuses!! You Can Start Before or After the Holidays. The Choice is YOURS!! RELIA Spotlight Opportunity! Work From Home!! Yes YOU asked for it and here is the opportunity!! My client is looking for strong ihc expertise and in exchange for 50% travel to client sites you can base from home!! Contact me for more info! RELIA HOT HISTOLOGY Opportunities MI Kalamazoo Applications Manager IHC MI Kalamazoo Applications Specialist IHC VA Staunton Full time days in the Blue Ridge Mountains! TX Dallas All shifts, Amazing team to work with! FL Ft. Myers AP Training Specialist Do you like to train techs? FL Ft. Myers HT/HTL all shifts full&part time. ***I can help with your Florida license!*** CA Orange County HT/HTL All shifts FULL& PART TIME AVAILABLE!! CA San Diego Brand New Lab!!! CA Modesto IHC Specialist&Histotech positions state of the art lab!! I also have exciting opportunities in: Gainesville, FL Panama City, FL Aurora, CO Chicago, IL And new opportunities coming in all of the time!! I look forward to assisting you!! Thanks-Pam Right Time, Right Place, Right Move with RELIA! Providing excellent service exclusively to the Histology Community! Thank You! ?Pam M. Barker? Pam Barker President/Senior Recruiting Specialist-Histology RELIA Solutions Specialists in Allied Healthcare Recruiting 5703 Red Bug Lake Road #330 Winter Springs, FL 32708-4969 Phone: (407)657-2027 Cell:???? (407)353-5070 FAX:???? (407)678-2788 E-mail: relia1 at earthlink.net https://www.facebook.com/RELIASolutionsforhistologyprofessionals www.facebook.com/PamBarkerRELIA www.linkedin.com/in/reliasolutions #jobs4myhistopeeps #ilovemyhistopeeps #histopeeps Follow my hashtags and make your day great and your career greater!! To unsubscribe please reply to relia1 at earthlink.net with unsubscribe. From sharon at alliedsearchpartners.com Fri Nov 12 08:51:31 2021 From: sharon at alliedsearchpartners.com (Sharon Strickland) Date: Fri, 12 Nov 2021 14:51:31 +0000 Subject: [Histonet] Histotech Jobs in Virginia and Florida Message-ID: Hi Histonet! Allied Search Partners currently has two full time permanent Histotech openings. Below I have listed the locations. If you have an interest, please reply to this message! -Charlottesville, VA area -North Central Florida (1.5 Hours south of Jacksonville, FL) -Relocation Assistance, Full Benefits Package, Day Shifts, New Grads or Seasoned professionals welcome to apply. If you are open to hearing about more please request a copy of the job description! Sharon Strickland Administrative Assistant Allied Search Partners AN MRINETWORK MEMBER Direct (Call) Line: 386.846.7596 From criley at udel.edu Fri Nov 12 10:03:51 2021 From: criley at udel.edu (Charles Riley) Date: Fri, 12 Nov 2021 11:03:51 -0500 Subject: [Histonet] Paraffin embedding following storage in 70% alcohol Message-ID: I am working with a grad student on a project dealing with equine articular cartilage. The protocol she sent me for embedding the tissue samples goes directly from 70% alcohol to the embedding step in paraffin. Correct me if I am wrong but shouldn't the tissue be dehydrated fully and cleared before embedding the samples? From relia1 at earthlink.net Fri Nov 12 11:25:22 2021 From: relia1 at earthlink.net (Pam Barker) Date: Fri, 12 Nov 2021 12:25:22 -0500 Subject: [Histonet] FW: [Shared Post] Strategies for Coping with Histotech Shortage. Top Tips for Managers and Techs. In-Reply-To: References: Message-ID: <00a501d7d7ea$46475cc0$d2d61640$@earthlink.net> From: Histology Dream Job Diva! [mailto:donotreply at wordpress.com] Sent: Friday, November 12, 2021 12:24 PM To: relia1 at earthlink.net Subject: [Shared Post] Strategies for Coping with Histotech Shortage. Top Tips for Managers and Techs. PamatRELIA #ilovemyhistopeeps #jobs4myhistopeeps posted: "Originally posted in the NSH Fixation on Histology Blog." PamatRELIA #ilovemyhistopeeps #jobs4myhistopeeps (relia1 at earthlink.net) shared a post from Histology Dream Job Diva! Strategies for Coping with Histotech Shortage. Top Tips for Managers and Techs. by PamatRELIA #ilovemyhistopeeps #jobs4myhistopeeps Originally posted on Histology Dream Job Diva!: Has there ever been a time when there wasn?t a shortage of histotechs? It just seems to get more critical every year.? I have to be honest I would LOVE to place techs with all of you but they aren?t always out there.? I wanted to write this? Read more of this post PamatRELIA #ilovemyhistopeeps #jobs4myhistopeeps | November 12, 2021 at 9:56 pm | Categories: Uncategorized | URL: https://wp.me/p2AHiM-2g Comment See all comments Like Change your email settings at Manage Notifications. Trouble clicking? Copy and paste this URL into your browser: https://reliasolutionspambarker.wordpress.com/2021/11/12/strategies-for-coping-with-histotech-shortage-top-tips-for-managers-and-techs-2/ Thanks for flying with WordPress.com Thanks-Pam Right Time, Right Place, Right Move with RELIA! Providing excellent service exclusively to the Histology Community! Thank You! Pam M. Barker Pam Barker President/Senior Recruiting Specialist-Histology RELIA Solutions Specialists in Allied Healthcare Recruiting 5703 Red Bug Lake Road #330 Winter Springs, FL 32708-4969 Phone: (407)657-2027 Cell: (407)353-5070 FAX: (407)678-2788 E-mail: relia1 at earthlink.net https://www.facebook.com/RELIASolutionsforhistologyprofessionals www.linkedin.com/in/reliasolutions #jobs4myhistopeeps #ilovemyhistopeeps #histopeeps Follow my hashtags and make your day great and your career greater!! From jkiernan at uwo.ca Fri Nov 12 16:42:06 2021 From: jkiernan at uwo.ca (John Kiernan) Date: Fri, 12 Nov 2021 22:42:06 +0000 Subject: [Histonet] Paraffin embedding following storage in 70% alcohol In-Reply-To: References: Message-ID: Of course you are right! This is yet another example of an error in a procedure informally handed on from person to person! Always work from a book. Even a very old one will be OK for paraffin embedding. John Kiernan. https://www.schulich.uwo.ca/anatomy/people/faculty/emeriti/kiernan_john.html = = = ________________________________ From: Charles Riley via Histonet Sent: November 12, 2021 11:03 AM To: histonet at lists.utsouthwestern.edu Subject: [Histonet] Paraffin embedding following storage in 70% alcohol I am working with a grad student on a project dealing with equine articular cartilage. The protocol she sent me for embedding the tissue samples goes directly from 70% alcohol to the embedding step in paraffin. Correct me if I am wrong but shouldn't the tissue be dehydrated fully and cleared before embedding the samples? _______________________________________________ Histonet mailing list Histonet at lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From carl.hobbs at kcl.ac.uk Sat Nov 13 12:53:40 2021 From: carl.hobbs at kcl.ac.uk (Hobbs, Carl) Date: Sat, 13 Nov 2021 18:53:40 +0000 Subject: [Histonet] Paraffin embedding following storage in 70% alcohol Message-ID: The Most venerable Histologist ( JK) is correct...as always! No way to go from 70% alcohol directly into Pwax Have to remove ALL water before infiltrating with Pwax Yes....also need to replace alcohol with Xylene/Histoclear before infil. with Pwax. Sure, you can use Isopropyl alc to obviate xylene /Histoclear/equiv. but......Jury's out on that Carl Hobbs FIBMS Histology and Imaging Manager Wolfson CARD Guys Campus, London Bridge? Kings College London London SE1 1UL ? 020 7848 6810 From linlue12363 at gmail.com Sat Nov 13 17:43:53 2021 From: linlue12363 at gmail.com (Linda Hines) Date: Sat, 13 Nov 2021 16:43:53 -0700 Subject: [Histonet] Histonet Digest, Vol 216, Issue 5 In-Reply-To: References: Message-ID: STOP On Sat, Nov 13, 2021, 11:00 AM wrote: > Send Histonet mailing list submissions to > histonet at lists.utsouthwestern.edu > > To subscribe or unsubscribe via the World Wide Web, visit > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > or, via email, send a message with subject or body 'help' to > histonet-request at lists.utsouthwestern.edu > > You can reach the person managing the list at > histonet-owner at lists.utsouthwestern.edu > > When replying, please edit your Subject line so it is more specific > than "Re: Contents of Histonet digest..." > Today's Topics: > > 1. Re: Paraffin embedding following storage in 70% alcohol > (John Kiernan) > > > > ---------- Forwarded message ---------- > From: John Kiernan > To: "histonet at lists.utsouthwestern.edu" , > Charles Riley > Cc: > Bcc: > Date: Fri, 12 Nov 2021 22:42:06 +0000 > Subject: Re: [Histonet] Paraffin embedding following storage in 70% alcohol > Of course you are right! > This is yet another example of an error in a procedure informally handed > on from person to person! Always work from a book. Even a very old one will > be OK for paraffin embedding. > John Kiernan. > > https://www.schulich.uwo.ca/anatomy/people/faculty/emeriti/kiernan_john.html > = = = > ________________________________ > From: Charles Riley via Histonet > Sent: November 12, 2021 11:03 AM > To: histonet at lists.utsouthwestern.edu > Subject: [Histonet] Paraffin embedding following storage in 70% alcohol > > I am working with a grad student on a project dealing with equine articular > cartilage. > > The protocol she sent me for embedding the tissue samples goes directly > from 70% alcohol to the embedding step in paraffin. > > > Correct me if I am wrong but shouldn't the tissue be dehydrated fully and > cleared before embedding the samples? > _______________________________________________ > Histonet mailing list > Histonet at lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > > _______________________________________________ > Histonet mailing list > Histonet at lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet From tkngflght at yahoo.com Mon Nov 15 12:11:14 2021 From: tkngflght at yahoo.com (Cheryl) Date: Mon, 15 Nov 2021 12:11:14 -0600 Subject: [Histonet] Increased background and Dako artisan References: <4407B63F-AABE-44AB-8BE8-7CAF167DAB20.ref@yahoo.com> Message-ID: <4407B63F-AABE-44AB-8BE8-7CAF167DAB20@yahoo.com> Hi guys- Does anyone have a reference for how long the Dako Artisan refrigerated kits can be at room temp? Also - having inconsistent increase in background in GMS. Not sure if it?s the slides or the kit. We?ve heard about the methenamine shortage - do y?all know anything more about this kinda problem?? Cheryl Kerry Please excuse typos-sent from a phone. From igor.deyneko at gmail.com Mon Nov 15 13:18:41 2021 From: igor.deyneko at gmail.com (Igor) Date: Mon, 15 Nov 2021 14:18:41 -0500 Subject: [Histonet] Unstained Slide Storage Message-ID: Hello fellow Histonetters, I would like to ask for your input and get your storage ideas for the unstained slides. In the hospital days, we used the multislide holding cabinets. However, currently, one of my researchers is very adamant about keeping the slides in 100 slide plastic boxes. If you keep your slides in the same boxes, do you have any special types of storage for those boxes that's easy to display and access???? Thank you in advance for your help! Igor Deyneko Novartis Institutes for BioMedical Research, Inc. From samantha.golden at ymail.com Tue Nov 23 10:24:07 2021 From: samantha.golden at ymail.com (Samantha Golden) Date: Tue, 23 Nov 2021 11:24:07 -0500 Subject: [Histonet] Microwave slide drying References: <910829C0-7145-4C10-8731-FC2CAFD44031.ref@ymail.com> Message-ID: <910829C0-7145-4C10-8731-FC2CAFD44031@ymail.com> What are the Histonets thoughts toward drying your slides in the microwave?.. pros/cons I would specifically like feedback regarding the slides being used for IHC. My new lab does this and it?s new to me. Thanks!! Samantha Sent from my iPhone From Richard.Cartun at hhchealth.org Tue Nov 23 10:35:04 2021 From: Richard.Cartun at hhchealth.org (Cartun, Richard) Date: Tue, 23 Nov 2021 16:35:04 +0000 Subject: [Histonet] Microwave slide drying In-Reply-To: <910829C0-7145-4C10-8731-FC2CAFD44031@ymail.com> References: <910829C0-7145-4C10-8731-FC2CAFD44031.ref@ymail.com> <910829C0-7145-4C10-8731-FC2CAFD44031@ymail.com> Message-ID: <81b4ba2e32b44530810c42a2efd19379@hhchealth.org> I have never used microwave drying for IHC slides. I worry that if there is water trapped underneath the tissue section it might cause destruction of the tissue section if it is rapidly heated and "pops". You could run a "side-by-side" comparison and see if you notice a difference. Also, how would you monitor the temperature of the slide? However, my philosophy has always been if it makes the process more efficient and/or less costly without changing the outcome, "do it". Happy Thanksgiving to all who celebrate it. Richard Richard W. Cartun, MS, PhD Director, Histology & The Martin M. Berman, MD Immunopathology/Morphologic Proteomics Laboratory Assistant Director, Anatomic Pathology Department of Pathology & Laboratory Medicine Hartford Hospital 80 Seymour Street Hartford, CT 06102 (860) 972-1596 Office (860) 545-2204 (Fax) -----Original Message----- From: Samantha Golden via Histonet [mailto:histonet at lists.utsouthwestern.edu] Sent: Tuesday, November 23, 2021 11:24 AM To: histonet at lists.utsouthwestern.edu Subject: [Histonet] Microwave slide drying This email is from outside HHC. BE CAREFUL when opening attachments or links from unknown senders. What are the Histonets thoughts toward drying your slides in the microwave?.. pros/cons I would specifically like feedback regarding the slides being used for IHC. My new lab does this and it?s new to me. Thanks!! Samantha Sent from my iPhone _______________________________________________ Histonet mailing list Histonet at lists.utsouthwestern.edu https://urldefense.com/v3/__http://lists.utsouthwestern.edu/mailman/listinfo/histonet__;!!KCs9X-8!IZrcGKqrTAB7YVkmihOC68-4NZ4y4YNfYD8Llt22Q6FJiPjxa6-C6OWEXWgdsjrijH8$ Reminder: This e-mail and any attachments are subject to the current HHC email retention policies. Please save or store appropriately in accordance with policy. This e-mail message, including any attachments, is for the sole use of the intended recipient(s) and may contain confidential and privileged information. Any unauthorized review, use, disclosure, or distribution is prohibited. If you are not the intended recipient, or an employee or agent responsible for delivering the message to the intended recipient, please contact the sender by reply e-mail and destroy all copies of the original message, including any attachments. From amosbrooks at gmail.com Sun Nov 28 17:22:18 2021 From: amosbrooks at gmail.com (Amos Brooks) Date: Sun, 28 Nov 2021 18:22:18 -0500 Subject: [Histonet] Microwaving Slides In-Reply-To: References: Message-ID: Hi Samantha, Microwaves are terrible! I am really not a fan of them in general, but especially for drying slides, and even moreso for slides intended for IHC. There is no way to really monitor the temperature the slides get to. Sure you can get a fnacy one with a probe, but that probe only monitors the solution it is in, not the small bubbles of water under the section or even the section itself. You end up with areas that get *really* hot and areas that are barely heated at all. At best you get an average heating. You would be much better off using an oven and even better a convection oven. A small fan to move the heat around really drys the slides out well Message: 1 Date: Tue, 23 Nov 2021 11:24:07 -0500 From: Samantha Golden To: histonet at lists.utsouthwestern.edu Subject: [Histonet] Microwave slide drying Message-ID: <910829C0-7145-4C10-8731-FC2CAFD44031 at ymail.com> Content-Type: text/plain; charset=utf-8 What are the Histonets thoughts toward drying your slides in the microwave?.. pros/cons I would specifically like feedback regarding the slides being used for IHC. My new lab does this and it?s new to me. Thanks!! Samantha From jordhood at med.umich.edu Tue Nov 30 15:03:05 2021 From: jordhood at med.umich.edu (Hood, Jordan) Date: Tue, 30 Nov 2021 21:03:05 +0000 Subject: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions In-Reply-To: References: <48bd31f00b074079baca10d963e1bc2f@med.umich.edu> <9e1fe5b8-be5e-ae7f-ca29-e57dac959071@shaw.ca> <5a34030c402f4f4b97dec87e4856d6f6@SVDCMBX-MEX024.nswhealth.net> Message-ID: I would like to thank all of you for your advice -- Using Thiosemicarbazide after the periodic acid did the trick, along with making up the silver solution immediately before use. It only took ~30 minutes in the warm silver solution before the basement membranes reached the desired staining intensity. Excellent results on my first try using the TSC, especially when compared to my first two disastrous tests. I am sincerely grateful to you all for helping me so much!! Best wishes, Jordan From: Colleen Forster Sent: Thursday, September 23, 2021 7:28 PM To: Tony Henwood (SCHN) Cc: Bryan Llewellyn ; Hood, Jordan ; histonet at lists.utsouthwestern.edu Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions External Email - Use Caution Make sure the periodic acid is made fresh EACH time you run the stain. That can also make a big difference in the stain quality. Colleen Forster HT(ASCP)QIHC On Thu, Sep 23, 2021 at 6:14 PM Tony Henwood (SCHN) via Histonet > wrote: I agree with Bryan, The introduction of thiosemicarbazide before the silver step improves the staining immensely. I would also look at the periodic acid. Is it too dilute, though 0.5% should work? I usually cover this by using a 1% solution for 20 minutes. Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) Principal Scientist, the Children?s Hospital at Westmead Adjunct Fellow, School of Medicine, University of Western Sydney Tel: 612 9845 3306 Fax: 612 9845 3318 Pathology Department the children's hospital at westmead Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, Westmead NSW 2145, AUSTRALIA -----Original Message----- From: Bryan Llewellyn via Histonet [mailto:histonet at lists.utsouthwestern.edu] Sent: Friday, 24 September 2021 7:47 AM To: Jordan >; Histonet > Subject: Re: [Histonet] Jones' Methenamine Silver Stain for Basement Membranes of Kidney - Issues and Questions Hi, Try the method given in StainsFile at: http://stainsfile.info/stain/metallic/jones.htm Bryan Llewellyn Hood, Jordan via Histonet wrote: > Hello, > > I'm new to histology (and new to histonet), and I work in a small histology lab specializing in animal tissues that receives requests/submissions from researchers. I tried (and failed) to perform a Jones' Methenamine Silver stain on a client's submission of pig kidneys (formalin-fixed, paraffin-embedded, cut at 2.5 microns), and I need some help troubleshooting this stain since my co-workers are stumped, too. I used the following procedure from Rowley Biochemical: > > > ~~~~~ > "Fixation: 10% Buffered Neutral Formalin (F-113) or Bouin's Solution > (F-40) or Zenker's (F-155) > > Sections: Paraffin, 2 microns > > Procedure: Acid washed glassware must be used!!!! > 1. Deparaffinize and hydrate to distilled water. > 2. Oxidize in Periodic Acid 0.5% (F-396-1) 11 minutes. Wash in chloride-free water. > 3. Prepare Methenamine Silver solution by mixing: 42.5 ml Methenamine 3% (F-396-2), 2.5 ml Silver Nitrate, 5% (F-396-3) and 12.0 ml Borate Buffer, pH 8.2 (F-396-4). > 4. Place slides in the solution and the entire jar in a water bath at 70?C for approx. 60-75 minutes. Check under microscope when slides appear medium brown microscopically. Every 10 minutes, once the medium brown color has been established, rinse a slide in 70?C, chloride free water and check under a microscope. Rinse again in hot water and return to the hot staining solution. As the staining time approaches the end point, check the slides, as above, every 1-2 minutes. The entire procedure must be performed quickly to prevent an uneven staining of the tissues. The slides should exhibit a brownish- yellow background, intense black reticulum fibers, and black basement membranes. If the slides become oversaturated, i.e. too black, destain in a dilute Potassium Ferricyanide Solution (F-396-11) for one or two dips. > 5. Rinse well in distilled water. Tone in Gold Chloride 0.2% (F-396-5), 1 minute. If sections are overtoned place in Sodium Metabisulfite, 3% (F-396-12) for 1-3 minutes. Rinse well in distilled water. > 6. Sodium Thiosulfate 3% (F-396-9), 1-2 miutes. Wash in running tap water, 10 minutes. Rinse well in distilled water. > 7. Stain in Harris' Hematoxylin (F-396-6) containing 2-4ml of Glacial Acetic Acid per 100 ml for 5-15 minutes. Wash in water. > 8. Differentiate in Acid Alcohol 1% (F-396-13) until the sections turn red. > 9. Blue section in Ammonia Water, 0.3% (F-396-14). Wash thoroughly. > 10. Counterstain in Eosin Y, 1%, Alcoholic Solution (F-396-7). > 11. Dehydrate in 95% alcohol, absolute alcohol and clear in xylene 3 changes each. Mount. > > Stain Results: > Basement membranes, reticulum fibers: Black > Nuclei: Blue > Cytoplasm, collagen, connective tissue: Pink-orange > > References: Jones, D.B., Amer.J.Path. 27:99 (1951). AFIP Manual of Histolocical Staining Methods, 3rd ed., Ed. L. Luna: NY: McGraw-Hill Publications, c. 1968, p. 97." > ~~~~~ > > > It became apparent that something went wrong during Step 4 when the slides were in the glass container (not a coplin jar - we have ten slides that we need to stain so we're using a rectangular glass container that holds ten slides on their sides - it does require a metal handle to move, but the handle is flexible and easy to remove after the glass slide rack has been transferred between containers) of silver solution in the water bath because there was lots of precipitate on the slides and floating on the surface of the silver solution. > > In my first test, I used five test slides (extra slides that we cut from the same blocks that were submitted to us). I deparaffinized them in coplin jars (moving them with plastic forceps) and hydrated them to deionized water. I transferred the slides to a glass slide rack that holds ten slides on their sides, added five blank slides that were rinsed in deionized water (so that the displacement of reagents would be equivalent to when we stain our ten "real" slides after testing is complete), and completed Step 2. I don't recall exactly how long the glass container of silver solution and the glass container of deionized water had been heating up in the water bath, but I would estimate ~15-30 minutes. The thermometer said that the water in the bath (not inside the containers) reached ~60-65 degrees Celsius. The silver solution was clear and colorless when I made it up, but by the time I put the slides into the warm silver solution, the solution was beginning to turn a light brown color (though it was still clear and I did not see any precipitate floating around). I removed the metal handle of the glass slide rack after the rack was transferred into the silver solution, but the metal handle did dip into the silver solution briefly. At some point, I noticed precipitate floating around of the surface of the silver solution. After ~80 minutes, I used plastic forceps to remove one test slide from the warm silver solution, dipped it several times into the warm deionized water to rinse it, and wiped off the back of the slide with gauze. The amount of precipitate was so extreme that the gauze did nearly nothing. I showed the slide to one of our pathologists and they could hardly see beyond the precipitate, but said that they couldn't see any staining of the structure that they were looking for (I forget exactly what it was, but I know it's supposed to turn black). > > In my second test (to see if the metal holder was the problem) that I performed immediately after the first test, I used one test slide. I deparaffinized it in the same coplin jars as before (moving it with plastic forceps) and hydrated it to deionized water. I used new glass containers for the periodic acid and deionized water rinse in Step 2, for making the silver solution in Step 3, and for the warm silver solution and warm deionized water in Step 4. I used plastic forceps to move the slide into the periodic acid, and propped it up in the container so that no glass rack or metal handle was used at all. I used plastic forceps to transfer the slide to the deionized water rinse, and dunked it several times and swished the slide around a bit. I used plastic forceps to transfer the slide into the warm(-ish) silver solution and propped it up against the side again. After approximately 20 minutes, I saw precipitate floating around, and I used plastic forceps to remove the slide from the silver solution. I dipped the slide into the warm(-ish) deionized water several times, and saw that the precipitate was again covering the slide and the tissue so I stopped there for the day. > > We purchased all of the reagents listed in the above procedure from Rowley Biochemical (except for the Glacial Acetic Acid mentioned in Step 7, but I didn't even get that far). > > Questions: > > 1. Could this indicate that the acid-washing was not done correctly? I made up a ~1% Hydrochloric Acid solution (with deionized water) and filled a plastic bin with the solution (I rinsed the bin with deionized water first). I then submerged all glassware (in several batches) for at least five minutes, then rinsed well with deionized water (not by filling a bin - I just used the hose of deionized water in our lab sink and poured it over the glassware) and left them to air-dry overnight. > > 2. Are using acid-washed glassware and avoiding metal even necessary precautions after the sodium thiosulphate in Step 6? I read that sodium thiosulphate "stops the reaction," and the procedure stops specifically saying to use deionized water after Step 6 and starts saying to use just "water" or "tap water." My lab refers to our waters as either "tap" or "deionized," so I'm assuming that using my deionized water is fine when the procedure calls for "distilled" or "dechlorinated." > > I don't even know enough to ask more questions, but I'm sure many more will arise after I test the stain again next week, so I welcome any and all advice about silver stains, acid-cleaning glassware, and literally anything else... > > Thank you!!! > > Jordan H. > University of Michigan > Ann Arbor, MI > ********************************************************** > Electronic Mail is not secure, may not be read every day, and should > not be used for urgent or sensitive issues > _______________________________________________ > Histonet mailing list > Histonet at lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > _______________________________________________ Histonet mailing list Histonet at lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet This message is intended for the addressee named and may contain confidential information. If you are not the intended recipient, please delete it and notify the sender. Views expressed in this message are those of the individual sender, and are not necessarily the views of NSW Health or any of its entities. _______________________________________________ Histonet mailing list Histonet at lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet -- Colleen Forster HT(ASCP)QIHC BLS Histology and IHC Laboratory Jackson Hall, Room 2-155 321 Church St. SE Minneapolis, MN 55455 612-626-1930 ********************************************************** Electronic Mail is not secure, may not be read every day, and should not be used for urgent or sensitive issues