Re: [Histonet] cloudy cornea after Hartman's fix
The cornea consists of cells (nuclei with plenty of cytoplasmic protein) and parallel bundles (lamellae) of collagen fibres, also protein. It is wonderful that the normal cornea is transparent. Doesn't any fixative make the cornea opaque? Davidson's (=Hartmann's) fixative contains plenty of formaldehyde (to add to and slowly crosslink all protein molecules, including collagen fibrils) but perhaps not quite enough ethanol to rapidly coagulate soluble proteins in cytoplasm. (Davidson's 22.2% v/v EtOH is slightly more than in sherry, madeira or vermouth; about half of that in whisky, gin etc.) The acetic acid (about 10%) can be expected to make nuclear DNA molecules and their associated proteins shrink into the patterns we see as typical of different cell-types. Most of the many formalin-acetic-alcohol fixatives contain enough ethanol to bring about protein coagulation before the more slowly reacting formaldehyde does its stuff. As Tony points out, more research is needed. John Kiernan https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Tony Henwood via Histonet Sent: June 27, 2024 7:17 PM To: histonet@lists.utsouthwestern.edu ; Davoli, Katherine A Subject: Re: [Histonet] cloudy cornea after Hartman's fix Hartman's (also known as Davidson's) fixative is sometimes used to reveal lymph nodes in resections where they appear opaque - white. It is not surprising that the tissues of the eye would react the same. I assume that the alcohol causes the bleaching of the tissue (or is it the acetic acid?) - more research needed. Regards, Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired) Principal Scientist, the Children’s Hospital at Westmead (Retired) Adjunct Fellow, School of Medicine, University of Western Sydney. From: Davoli, Katherine A via Histonet Sent: 28 June 2024 04:12 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] cloudy cornea after Hartman's fix Anyone know why the cornea of my pig eye got white/cloudy on dropping it in to Hartman's fixative? I'm used to working with mice where, if this happens I didn't notice. Katherine Davoli, MDiv, HTL(ASCP)cm(they/them/theirs) Lab Manager, Tissue Culture & Histology Cores, U. Pitt Dept of Ophthalmology 7.373 UPMC Mercy Pavilion1622 Locust St., Pittsburgh PA 15219 (412) 624-8508 this number cannot receive texts ___ ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Causes of false positive Congo Red
Dear Greg, This is the same as on p.126 in my copy of Carson's 2nd edition (1997) and also in the 11th and last (2008) edn of Churukian's "Manual of the Special Stains Laboratory". The only stock solution that can be expected to change with time is the Stock Congo red solution, because solutions of dyes with large anions are unstable in the presence of inorganic salts. Churukian (p.195) said it could be kept for 2 months. The correct staining you got with newly made solutions suggests that your earlier stock Congo red stock solution was too old. Evidently you solved the problem yourself! In their "Troubleshooting Histology Stains" book, Horobin & Bancroft (1998, p.45-47) stressed the need for a freshly made dye solution. They also suggested ignoring "pink background" and checking that it's not dichroic. They also listed various Congo-positive and dichroic materials that aren't amyloid. An unidentified yellow compound is often present in Congo red and it may sometimes cause generalized yellow background staining. John Kiernan https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Greg Dobbin Sent: June 20, 2024 8:44 AM To: John Kiernan ; histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Causes of false positive Congo Red You don't often get email from greg.dob...@gmail.com. Learn why this is important<https://aka.ms/LearnAboutSenderIdentification> Hi John, I must apologize again. We used to use the method from Carson's book. We now make up the reagents as follows: Stock alkaline salt solution Sodium chloride. 2g Distilled Water... 20mL Stir until the salt is dissolved, then with continuous stirring on a magnetic stirrer add 80mL of 100% denatured ethanol. Some salt may precipitate out after the ethanol is added. Working alkaline salt solution Stock alkaline stock solution...50 ml 1% Sodium Hydroxide0.5ml Filter and use within 15 minutes Stock Congo red solution Congo red 0.1g Stock alkaline salt solution 50mL Stir well with the magnetic stirrer and let stand overnight or for a minimum of 3 hours if the slides need to be ready the same day that the order was placed. Working Congo red (Congo red) Stock Congo red.. 50ml Sodium hydroxide 1%... 0.5ml Filter and use within 15 minutes. On Thu, Jun 20, 2024 at 9:31 AM Greg Dobbin mailto:greg.dob...@gmail.com>> wrote: Good day John, Very nice to hear from you again! I have been consulting your textbook in my investigations! Sorry about the brevity of the description of our method. I felt like my post was already too long and it might scare off some would-be contributors! :-) And yes, I incorrectly referred to the dichroic green as "fluorescent"-thank you. Our method follows the Puchtler method described on pages 132-3 in Frieda Carson's "Self-Instructional" textbook (1990) as does the hospital that repeated our false-positive Congo Reds. Note, once we re-made our reagents, our results returned to accurate staining. Greg On Thu, Jun 20, 2024 at 2:49 AM John Kiernan mailto:jkier...@uwo.ca>> wrote: Greg, your method is incompletely described in your Histonet post, but it looks quite different from the "traditional" Highman's procedure (Arch. Path. 41:559-562). What method were they using "at another lab" to get correct red amyloid that is green (dichroic, not fluorescent) with crossed polars? John Kiernan https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Greg Dobbin via Histonet mailto:histonet@lists.utsouthwestern.edu>> Sent: June 19, 2024 8:53 AM To: histonet@lists.utsouthwestern.edu<mailto:histonet@lists.utsouthwestern.edu> mailto:histonet@lists.utsouthwestern.edu>> Subject: [Histonet] Causes of false positive Congo Red Hello experts, *Some background:* I know that Congo Red can bind nonspecifically to non-amyloid components such as collagen and elastin under certain conditions (eg Carnoys fixative, insufficient differentiation, insufficient alkalinity, etc). However, everything I have been able to read on the topic suggests that over-staining is "easily" differentiated from true amyloid staining by using polarizing light microscopy. That is, true amyloid produces apple green fluorescence while non-amyloid components produce silver/grey color. *My question:* I want to know if anyone has encountered false positive staining that *is apple green* in color? We had a few bone marrow core biopsies that stained bright green but were later found to be negative when stained at another lab. We subsequently threw out all of our working solutions and made up everything fresh and repeated the previous (false positive) specimens and they were indee
Re: [Histonet] Causes of false positive Congo Red
Greg, your method is incompletely described in your Histonet post, but it looks quite different from the "traditional" Highman's procedure (Arch. Path. 41:559-562). What method were they using "at another lab" to get correct red amyloid that is green (dichroic, not fluorescent) with crossed polars? John Kiernan https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Greg Dobbin via Histonet Sent: June 19, 2024 8:53 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Causes of false positive Congo Red Hello experts, *Some background:* I know that Congo Red can bind nonspecifically to non-amyloid components such as collagen and elastin under certain conditions (eg Carnoys fixative, insufficient differentiation, insufficient alkalinity, etc). However, everything I have been able to read on the topic suggests that over-staining is "easily" differentiated from true amyloid staining by using polarizing light microscopy. That is, true amyloid produces apple green fluorescence while non-amyloid components produce silver/grey color. *My question:* I want to know if anyone has encountered false positive staining that *is apple green* in color? We had a few bone marrow core biopsies that stained bright green but were later found to be negative when stained at another lab. We subsequently threw out all of our working solutions and made up everything fresh and repeated the previous (false positive) specimens and they were indeed negative in our lab as well. *In order to prevent this from happening again, I need to attempt to understand what may have caused this to happen in the first place. * This is where the vast collective knowledge of this group comes in. :-) Can anyone offer some insight as to possible causes? *Our Congo Red method:* Deparaffinize sections and bring them to water. Stain in Hematoxylin for 1 minute Add 0.5ml of 1% Sodium Hydroxide to 50 ml of stock alkaline salt solution. Wash slides in running water Place in *working* alkaline salt solution from step 2 for 20 minutes Add 0.5 ml of 1% Sodium Hydroxide to stock Congo red solution. Start to filter *working* Congo red solution when 15 mins are left in step 6 Place sections in the *working* Congo red from step #8 for 20 minutes. Dehydrate the slides one at a time in 3 changes of absolute ethanol, 6 dips each. Dip the slide 10 times in a coplin of xylene. Continue dehydrating the other slides. Coverslip the slides. *Greg Dobbin* 1205 Pleasant Grove Rd Route 220 York, PE C0A 1P0 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] What is the best xylene substitute for histology?
I liked terpineol (mixed isomers). Smells nice and doesn't harden tissues, but I see that it has now become pretty expensive. Mineral oil USP (also called liquid paraffin and mineral oil, heavy) is OK and cheaper than xylene, but it's not miscible with ethanol. You have to dehydrate specimens in isopropanol and have the oil warmer than 44oC. Tert-butanol is miscible with water, ethanol and paraffin; it's somewhat more expensive than xylene and must be used warm because it freezes at 26oC. It's often used for plant histology. A recently proposed safe and economical alternative clearing agent for paraffin embedding is coconut oil, but it causes much more shrinkage than xylene and also induces ugly artifacts (in rats' prostates) if the clearing time is less than 4 hours. See OA Bright et al. 2024. J. Histochem. Cytochem. 72(4): 233-243. https://www.researchgate.net/profile/Ebenezer-Senu/publication/379429541_Clearing_Properties_Between_Coconut_Oil_and_Xylene_in_Histological_Tissue_Processing/links/661013eb2034097c54f61dbd/Clearing-Properties-Between-Coconut-Oil-and-Xylene-in-Histological-Tissue-Processing.pdf Just a few thoughts! John Kiernan. London, Canada From: Kate Bummer via Histonet Sent: June 7, 2024 10:52 AM To: histonet@lists.utsouthwestern.edu Cc: Kate Bummer Subject: [Histonet] What is the best xylene substitute for histology? Hello Histonetters I'm hoping I can get some recommendations for the best xylene substitute for histology that could be used in the following: * Tissue processor * Deparaffinization * Coverslipper * Mounting media that works with that particular xylene substitute Thank you for your help! Kate SeqMatic ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Inquiry on Tissue Softening for Microtomy
Jay, you are 100% right about "pseudo-scientific mumbo jumbo" for the mechanism. But have you (or anyone else) compared ammonia-water with ordinary pure water for hydrating/softening? Water (with or without dissolved ammonia) won't go through solid wax. The embedded tissue has to be exposed by cutting into the block. Whiffs of ammonia are unpleasant. If they upset some people in the lab, that's a good reason to try odorless water instead. John Kiernan Emeritus, UWO, London, Canada https://publish.uwo.ca/~jkiernan/ = = = From: Jay Lundgren Sent: May 11, 2024 4:40 PM To: John Kiernan Cc: histonet@lists.utsouthwestern.edu ; IGNACIO GONZÁLEZ MASSONI Subject: Re: [Histonet] Inquiry on Tissue Softening for Microtomy I wouldn't say softening, I would say hydrating. Ammonia water accelerates hydration of FFPE blocks. Nobody knows how it works, it's a mystery. Or at least I've never heard a scientific explanation. Only pseudo-scientific mumbo jumbo like "facilitates the removal of paraffin" which is false. Go soak a solid block of paraffin in pure NH4OH for 24 hours, it won't do anything. I was told as a student at AFIP, "It opens the pores of the tissue so water can get in." In other words, pseudo-scientific mumbo jumbo. It works though. Somebody needs to get a $150,000,000 NIH grant and do a research project on how ammonia water hydrates tissue. I worked with some lovely Hmong people in California that called it "crying water". Using it can cause a big interpersonal problem with certain people in the lab though. Interestingly, hypersensitivity to smells is one of the symptoms of autism. Hypersensitivity to smells is also highly correlated with bipolar disorder and heightened emotional reactivity. Sooo... Jay A. Lundgren, M.S., HTL (ASCP) On Fri, May 10, 2024 at 11:14 PM John Kiernan via Histonet mailto:histonet@lists.utsouthwestern.edu>> wrote: If you apply the ammonia to the cut surface of the paraffin block, I suspect that it softens the tissue in the same way as applying water: by entering interstices of the tissue that are not occupied by paraffin molecules. I never tried ammonia for this purpose but in the 1960s to early '70s I occasionally used a proprietary product called Mollifex, which I see is still sold. In 1972 or '73 an elderly technician told me that water was just as good, and I soon found out that he was right. Indeed, water had the advantage of working in 15-30 minutes rather than taking several hours. John Kiernan Emeritus, UWO, London, Canada https://publish.uwo.ca/~jkiernan/ = = = From: IGNACIO GONZÁLEZ MASSONI via Histonet mailto:histonet@lists.utsouthwestern.edu>> Sent: May 10, 2024 8:53 PM To: histonet@lists.utsouthwestern.edu<mailto:histonet@lists.utsouthwestern.edu> mailto:histonet@lists.utsouthwestern.edu>> Subject: [Histonet] Inquiry on Tissue Softening for Microtomy Dear friends at Histonet, I hope this message finds you well. I am reaching out to seek your expertise on a matter that has piqued my interest in the field of histology. I am currently delving into the process of preparing FFPE (formalin-fixed, paraffin-embedded) tissues for microtomy. Specifically, I am curious about the role of ammonia in softening these tissues before sectioning. From my understanding, ammonia serves as an alkaline agent that helps neutralize formalin's effects and facilitates the removal of paraffin, thus aiding in the softening of the tissue. However, I would greatly appreciate if you could provide a more detailed explanation of the chemical interactions at play here. How exactly does ammonia interact with the tissue components to achieve the desired softening effect? Moreover, are there any best practices or safety precautions that one should be aware of when using ammonia in this context? Your insights on this topic would be invaluable to me and would greatly enhance my understanding of the intricacies involved in histological preparations. Thank you for your time and assistance. Warm regards from Santiago of Chile Ignacio, Medical Technologist ___ Histonet mailing list Histonet@lists.utsouthwestern.edu<mailto:Histonet@lists.utsouthwestern.edu> http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu<mailto:Histonet@lists.utsouthwestern.edu> http://lists.utsouthwestern.edu/mailman/listinfo/histonet ____ From: Jay Lundgren Sent: May 11, 2024 4:40 PM To: John Kiernan Cc: histonet@lists.utsouthwestern.edu ; IGNACIO GONZÁLEZ MASSONI Subject: Re: [Histonet] Inquiry on Tissue Softening for Microtomy I wouldn't say softening, I would say hydrating. Ammonia water accelerates hydration of FFPE blocks. Nobody knows how it works, it's a my
Re: [Histonet] Inquiry on Tissue Softening for Microtomy
If you apply the ammonia to the cut surface of the paraffin block, I suspect that it softens the tissue in the same way as applying water: by entering interstices of the tissue that are not occupied by paraffin molecules. I never tried ammonia for this purpose but in the 1960s to early '70s I occasionally used a proprietary product called Mollifex, which I see is still sold. In 1972 or '73 an elderly technician told me that water was just as good, and I soon found out that he was right. Indeed, water had the advantage of working in 15-30 minutes rather than taking several hours. John Kiernan Emeritus, UWO, London, Canada https://publish.uwo.ca/~jkiernan/ = = = From: IGNACIO GONZÁLEZ MASSONI via Histonet Sent: May 10, 2024 8:53 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Inquiry on Tissue Softening for Microtomy Dear friends at Histonet, I hope this message finds you well. I am reaching out to seek your expertise on a matter that has piqued my interest in the field of histology. I am currently delving into the process of preparing FFPE (formalin-fixed, paraffin-embedded) tissues for microtomy. Specifically, I am curious about the role of ammonia in softening these tissues before sectioning. From my understanding, ammonia serves as an alkaline agent that helps neutralize formalin's effects and facilitates the removal of paraffin, thus aiding in the softening of the tissue. However, I would greatly appreciate if you could provide a more detailed explanation of the chemical interactions at play here. How exactly does ammonia interact with the tissue components to achieve the desired softening effect? Moreover, are there any best practices or safety precautions that one should be aware of when using ammonia in this context? Your insights on this topic would be invaluable to me and would greatly enhance my understanding of the intricacies involved in histological preparations. Thank you for your time and assistance. Warm regards from Santiago of Chile Ignacio, Medical Technologist ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Golgi-cox staining
I suggest asking Dr Sami Zaqout, the corresponding author of the paper you cited. A German email address is given on the left side of the first page of the free PDF file for which you provided a web link: https://www.frontiersin.org/articles/10.3389/fnana.2016.00038/ful<https://www.frontiersin.org/articles/10.3389/fnana.2016.00038/full>. He is easy to find with Google and he is now at Qatar University. It's the most thorough description of Golgi-Cox staining that I've seen, and the pictures of results look superb. Vibratome sectioning evidently is better and quicker than embedding in nitrocellulose and cutting with a sliding microtome! The two authors' use of the word "cryoprotectant" was unfortunate in the context of a technique that does not involve freezing. John Kiernan Emeritus, Anatomy, University of Western Ontario, Canada https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Mariela Chertoff via Histonet Sent: April 10, 2024 10:28 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Golgi-cox staining Hi all We made the Golgi cox staining and due to a problem with the vibratome, we left the tissue several days embedded in agarose and they get dryed, It is possoble to recover the brains? it is better to repeat the agarose embebbing o it is better to put the brains in crioprerervate solution to rehidrated and after that put them in agarosa again? We are following the Zaquot protocol https://www.frontiersin.org/articles/10.3389/fnana.2016.00038/full Thanks in advance for your reply Mariela Chertoff, PhD Laboratorio de Neuroepigenetica - QB75 Departamento de Química Biológica Facultad de Ciencias Exactas y Naturales - UBA Ciudad Universitaria Pabellón II Piso 4 Ciudad Autónoma de Buenos Aires C1428EGA - Argentina Tel: 54 11 5285-8680/1/2 email:marielachert...@gmail.com marielachert...@qb.fcen.uba.ar ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Modified Davidson solution
The Davidson/Hartmann fixative is just another alcoholic formaldehyde mixture acidified with acetic acid. Like all others of that ilk it was intended for making up in the lab soon before using. Storage causes slow deterioration. The ethanol slowly gets esterified by the acetic acid, making ethyl acetate, which has no fixative properties. This change is evident from the fruity smell of the ester, which becomes evident after a week or two. All this has been in the textbooks for more than 50 years. Why doesn't every lab spend perhaps $200 on a couple of books? For obvious reasons I recommend my 5th edition (ISBN 9781907904325; $82.50,from the publisher, which is $30 less than Amazon's price), but there are other histotechnology books that are very good. A book may cost much less than a big bottle of a deteriorating mixture of questionable value, and you and your staff could learn a lot by reading. John Kiernan = = = From: Davoli, Katherine A via Histonet Sent: March 5, 2024 4:40 PM To: Histonet ; Naira Margaryan Subject: Re: [Histonet] Modified Davidson solution Hi Naira, I get Hartmann's from Electron Microscopy Sciences Cat# 64133-10 (for 1L, but they have larger sizes I believe) Katherine Davoli, HTL(ASCP)cm(they/them/theirs) Lab Manager, Tissue Culture and Histology Core Facilities U. Pitt Dept of Ophthalmology 7.373 UPMC Mercy Pavilion 1622 Locust St., Pittsburgh PA 15219 (412) 624-8508 this number cannot receive texts From: Naira Margaryan via Histonet Sent: Tuesday, March 5, 2024 1:27 PM To: Histonet Subject: [Histonet] Modified Davidson solution Hello! What is the best company to buy a ready to use Modified Davidson solution? Thanks in advance, Naira ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Davoli, Katherine A via Histonet Sent: March 5, 2024 4:40 PM To: Histonet ; Naira Margaryan Subject: Re: [Histonet] Modified Davidson solution Hi Naira, I get Hartmann's from Electron Microscopy Sciences Cat# 64133-10 (for 1L, but they have larger sizes I believe) Katherine Davoli, HTL(ASCP)cm(they/them/theirs) Lab Manager, Tissue Culture and Histology Core Facilities U. Pitt Dept of Ophthalmology 7.373 UPMC Mercy Pavilion 1622 Locust St., Pittsburgh PA 15219 (412) 624-8508 this number cannot receive texts From: Naira Margaryan via Histonet Sent: Tuesday, March 5, 2024 1:27 PM To: Histonet Subject: [Histonet] Modified Davidson solution Hello! What is the best company to buy a ready to use Modified Davidson solution? Thanks in advance, Naira ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Modified Davidson
The "Davidson's" name is, as Tony says, applied to various mixtures. In these, formalin is acidified with acetic acid in solutions that also contain some alcohol (typically about 33% v/v) but probably not enough to contribute to fixation. The mixture quoted from Latendresse et al. 2002 has only 15% alcohol - about the same as in a good dry sherry. (My gastric mucosa is not yet fixed.) The Histonet archives must surely still contain Dr Bob Richmond's contributions about "Davidson's fixative" in the 1990s or 2000s. Enough from me for now! John Kiernan London, Canada = = = From: Tony Henwood via Histonet Sent: February 8, 2024 5:31 PM To: Naira Margaryan ; Histonet Subject: Re: [Histonet] Modified Davidson There are several formulations (some are probably typos), but this seems to be one commonly cited. Modified Davidson's Fixative: (Latendresse, J. R., Warbrittion, A. R., Jonassen, H., & Creasy, D. M. (2002). Fixation of testes and eyes using a modified Davidson's fluid: comparison with Bouin's fluid and conventional Davidson's fluid. Toxicologic pathology, 30(4), 524-533.) 37–40% formaldehyde30ml Absolute ethanol 15ml Glacial acetic acid5ml Distilled Water 50ml Regards, Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired) Principal Scientist, the Children’s Hospital at Westmead (Retired) Adjunct Fellow, School of Medicine, University of Western Sydney. From: Naira Margaryan via Histonet Sent: Friday, February 9, 2024 8:25:47 AM To: Histonet Subject: [Histonet] Modified Davidson Hello, Could you please share your best recipe for the Modified Davidson? Thanks in advance, Naira ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] O.C.T. what MW PVA and PEG?
Your method makes no sense! It looks like something informally passed along among students and technicians who have never read a book. Cryoprotection means preventing formation of ice crystals or, as it's usually done, minimizing their size. Sucrose is a cryoprotectant; the higher the concentration the better, but strong solutions penetrate the tissue slowly. For fixed specimens dissolve the sucrose in water. (For unfixed, use an isotonic phosphate buffer, but that's only if a day or so at 4C is OK for your needs. Very fast freezing may be needed, especially for muscle.) OCT is not a cryoprotectant; it's goop that surrounds the specimen and enters cracks and spaces but it does not penetrate. OCT serves to hold each section together during transfer from the cryostat's knife onto a glass slide or coverslip. For some more information (much of it derived from Histonet) have a look at the Biological Stain Commission's FAQ, at https://biologicalstaincommission.org/faqlist.htm#CRYPRO. John Kiernan https://www.schulich.uwo.ca/anatomy//people/faculty/emeriti/kiernan_john.html = = = From: Tyrone Genade via Histonet Sent: November 30, 2023 9:46 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] O.C.T. what MW PVA and PEG? Hello, I'm using a cryoprotection protocol that involves 3-stage cryoprotection of 15% glucose O/N, then 30% + 50% OCT O/N and then finally O/N in OCT. Compared to previous protocols this works very well -- even when cutting through eyes and lenses (which had previously given a lot of grief). My issue is that preparing the 30% + 50% OCT is a schlep. The OCT puts up a lot of resistance against mixing with the 60% sucrose. It would be much simpler if I could prepare 30% sucrose with powdered PVA and PEG. Does anyone know what MW polymers of PVA and PEG to use and what concentrations to approximate commercial (Scigen Tissue-Plus) OCT? Thanks Tyrone Genade Ph.D. Quillen College of Medicine ETSU Johnson City, TN ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Tyrone Genade via Histonet Sent: November 30, 2023 9:46 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] O.C.T. what MW PVA and PEG? Hello, I'm using a cryoprotection protocol that involves 3-stage cryoprotection of 15% glucose O/N, then 30% + 50% OCT O/N and then finally O/N in OCT. Compared to previous protocols this works very well -- even when cutting through eyes and lenses (which had previously given a lot of grief). My issue is that preparing the 30% + 50% OCT is a schlep. The OCT puts up a lot of resistance against mixing with the 60% sucrose. It would be much simpler if I could prepare 30% sucrose with powdered PVA and PEG. Does anyone know what MW polymers of PVA and PEG to use and what concentrations to approximate commercial (Scigen Tissue-Plus) OCT? Thanks Tyrone Genade Ph.D. Quillen College of Medicine ETSU Johnson City, TN ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Faded H tissue section
I found the Histo-Logic archive on one of Sakura's web pages. Lee Luna was the editor for several years. Nothing there before about 1980 periodic acid for re-working H The archive is huge and many contributions look good. See https://www.sakuraus.com/getattachment/103878b9-9854-469d-8203-9d9b1c45d850/761 but bear in mind that this is from a company. It's not a peer-reviewed scientific journal. John Kiernan. = = = From: Tony Henwood via Histonet Sent: November 10, 2023 4:49 AM To: Akemi Allison ; jayalakshmy p.s Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Faded H tissue section I thought he may have reported this in Histologic. This trick certainly works. You can notice the nuclear clarity on GIT and to a lesser extent skin biopsies stained with PAS. Regards, Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired) Principal Scientist, the Children’s Hospital at Westmead (Retired) Adjunct Fellow, School of Medicine, University of Western Sydney. From: Akemi Allison<mailto:akemiat3...@gmail.com> Sent: Friday, 10 November 2023 1:30 PM To: jayalakshmy p.s<mailto:psjayalaks...@gmail.com> Cc: Tony Henwood<mailto:afhenw...@outlook.com>; histonet@lists.utsouthwestern.edu<mailto:histonet@lists.utsouthwestern.edu> Subject: Re: [Histonet] Faded H tissue section Hi Tony: I actually gave that tip to Lee back in 1979 when he came to OHSU to give a seminar to the Oregon Histology Society. I found out that when I was doing GMA and the pathologists didn’t like Gills Hematoxylin. They loved my PAS on GMA so I tried using 1% Periodic Acid before using Harris Hematoxylin for H’s on GMA and it turned out beautifully. Guess he shared my tip with the rest of our society. He and I became closer se friends and we shared several tips, including his Movat’s tips which he didn’t publish, but I shared them in Frieda’s 4th Ed. Best, Akemi Allison-Tacha, BS, HT/HTL (ASCP) Sent from my iPhone > On Nov 9, 2023, at 6:11 PM, jayalakshmy p.s via Histonet > wrote: > > Thanks, I'll check it out. > >> On Fri, Nov 10, 2023, 6:58 AM Tony Henwood wrote: >> >> You could treat the decoverslipped section in 1% periodic acid (same as >> used in the PAS technique) for 30 minutes prior to H staining. This >> might improve the H staining. >> >> >> >> I believe Lee Luna suggested this but for the life of me, I can’t find the >> reference! >> >> >> >> Regards, >> >> >> >> Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired) >> >> Principal Scientist, the Children’s Hospital at Westmead (Retired) >> >> Adjunct Fellow, School of Medicine, University of Western Sydney. >> >> >> >> *From: *jayalakshmy p.s via Histonet >> *Sent: *Friday, 10 November 2023 3:43 AM >> *To: *histonet@lists.utsouthwestern.edu >> *Subject: *[Histonet] Faded H tissue section >> >> >> >> Hello, >> I would like to know how to effectively restain a faded H tissue section. >> The color becomes dull when re stained. Somebody please advise. >> Prof. Jayalakshmy. P S >> ___ >> Histonet mailing list >> Histonet@lists.utsouthwestern.edu >> http://lists.utsouthwestern.edu/mailman/listinfo/histonet >> >> >> > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Fixing and processing mouse eyeballs
The Journal of Histotechnology, Vol. 45 No. 4 (Dec 2022) is a special issue devoted to eyes and ocular tissues. Two of the articles are specifically about fixing and processing eyes of mice: J. Pang et 9 al. pp172-181 and J. Li et 9 al. pp161-171. Every member of the NSH should have received a paper copy of the journal. The cover has a splendid photomicrograph (from the Pang paper) of a whole mouse eye. John Kiernan University of Western Ontario London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html <https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html> = = = From: Charles Riley via Histonet Sent: October 11, 2023 5:00 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Fixing and processing mouse eyeballs Does anyone have any helpful tips or protocols for fixing and processing schedules for adult mouse eyes? ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Long term museum specimen storage
I don't know anything about "Jore's fixative" or the rationale of using a very hypertonic unbuffered 4% formaldehyde with magnesium, sodium, chloride and sulphate ions. If brown stuff is now bleeding out of your museum specimens, Jore Juice evidently isn't a good preservative. According to Chapter 26 in the late Charles Culling's excellent book (3rd edn 1974; ISBN: 0407729011) the fixative/preservative for a museum specimen is optimized to preserve colour, which is the red or reddish-brown of haemoglobin and myoglobin. This usually is achieved with Kaiserling's fluid, which contains formalin, potassium acetate and also potassium nitrate (1.5% w/v) as an oxidant. Another approach involves treating specimens with carbon monoxide to convert all haemoglobin etc to a red carboxy derivative. If your museum specimens have already lost all their meaningful colours, a neutral buffered aqueous formaldehyde may be the best that you can provide to preserve the sizes and shapes. 70% alcohol will cause some shrinkage, and it may not be as easy to seal this solvent into a museum container as a watery diluted formalin. John Kiernan = = = From: Rhonda McCormick via Histonet Sent: September 20, 2023 11:39 AM To: Histonet Subject: [Histonet] Long term museum specimen storage Hi All, I am looking to replace the fixative for veterinary specimens that have been preserved as "museum specimens". They are kept in jars in a glass case outside our lab, however, some of the fixative is starting to turn brown (and we've pulled a few jars that have some slight cracks in them). The specimens are currently in Jore's Fixative: 100 mL Distilled water 10 mL 40% Formaldehyde2 g Magnesium Sulfate2 g Sodium Sulfate1 g Sodium Chloride Preserving specimens is new to me. I've never heard of Jore's fixative before and I'm wondering if I could get some advice, please? Do these specimens need to be replaced with the same solution? Could we rinse the specimen and replace the solution with 70% Alcohol? OR would 10% NBF be better to store the specimens in (or something al together different)? We have a varying display of specimens - anywhere from a small porcine optic nerve to a large equine granulosa cell tumor. Realizing it may be different based on the size of the specimen, approximately how often should the solution be changed ? Thank you so much! Any help or insight is much appreciated. Rhonda McCormickRhonda McCormick BS, HT (ASCP)cm Histology Diagnostic Lab Supervisor College of Veterinary Medicine Texas A University ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Formalin pH
Thank you Tony, for drawing attention to that excellent 1991 paper in the Journal of Histotechnology. It was published in the years when the NSH's journal was not included in Current Contents and was not taken by most libraries. Sadly, Norman Hew-Shue died in Toronto in 2022. Papers in J. Histotechnol. (including all back-issues) are now easily available as PDF downloads from libraries that subscribe to the many journals published by Taylor and Francis. That's probably most academic and biomedical libraries. Members of the NSH still get the paper journal and have access to the archive by way of an obscure members-only link on the society's web site; it is very difficult to find. Why won't the NSH make its journal conspicuous and easily accessible to all by way of a single click from https://www.nsh.org/home? Grrr! John Kiernan. = = = From: Tony Henwood via Histonet Sent: September 13, 2023 7:01 PM To: Bacon, Charles ; e...@pigs.ag Cc: HistoNet Subject: Re: [Histonet] Formalin pH Hew-Shue (1991) has described a useful pH indicator for working neutral buffered formalin solutions. Bromocresol purple, when added to formalin solutions, serves as an indicator of pH as well as, from a safety aspect, labelling the solution as formalin making redundant the dangerous “smell” test. At an acidic pH (5.2) the intense indicator colour is yellow whereas at pH 6.8, the colour is purple. A saturated solution of the dye is prepared, and 2 to 4 drops are added to 10 litres of neutral buffered formalin. Other advantages are that the fixative is more readily distinguished from other colourless solutions such as saline, thus preventing accidental misuse, and formalin spills are more easily recognised. Hew-Shue N (1991) “Bromcresol Purple as a Colored Marker and pH Indicator for Ten Percent Neutral Buffered Forrnalin” J Histotechnol 14(4):257-260. Regards, Tony Henwood MSc, BAppSc, GradDipSysAnalys, CT(ASC), FFSc(RCPA) (Retired) Principal Scientist, the Children’s Hospital at Westmead (Retired) Adjunct Fellow, School of Medicine, University of Western Sydney. From: Bacon, Charles via Histonet<mailto:histonet@lists.utsouthwestern.edu> Sent: Thursday, 14 September 2023 6:42 AM To: e...@pigs.ag<mailto:e...@pigs.ag> Cc: HistoNet<mailto:histonet@lists.utsouthwestern.edu> Subject: Re: [Histonet] Formalin pH We pull the COA from the vendors website. Some are better than others but the are usually listed by lot number. I keep them in a network shared folder so they can be pulled by anyone that is asked for the evidence during inspection. Sent from my iPhone > On Sep 13, 2023, at 8:22 AM, "e...@pigs.ag" wrote: > > Sooner or later the people have got to rise up > and tell those brain-dead paper-shuffling regulators > to just stuff it. Their mindless interference is > just wasting time and interfering with people's > ability to keep their own garden. > > -Original Message- > From: Cooper, Brian via Histonet > Reply-To: Cooper, Brian > To: Paula Sicurello, HistoNet > Subject: Re: [Histonet] Formalin pH > Date: Today 10:19 AM > > Ask your vendor to send a certificate of analysis that includes the pH. > That's what we did and now they come with each lot. We do have to bug them > for them occasionally. > > I'm betting your vendor has at least heard about this from other customers so > it should be on their radar at the very least. > > > Thanks, > > > > Brian D. Cooper, HT (ASCP)CMQIHCCM| Histology Supervisor > > Department of Pathology and Laboratory Medicine > > Children's Hospital Los Angeles > > 4650 Sunset Blvd MS#43- Los Angeles, CA 90027 > > Ph: 323.361.3357 > > bcoo...@chla.usc.edu<mailto:bcoo...@chla.usc.edu> > > > From: Paula Sicurello via Histonet > Sent: Tuesday, September 12, 2023 7:12:05 PM > To: HistoNet > Subject: [Histonet] Formalin pH (EXTERNAL EMAIL) > > CAUTION: BE CAREFUL WITH THIS MESSAGE* > This email came from outside CHLA. Do not open attachments, click on links, > or respond unless you expected this message and recognize the email address: > histonet-boun...@lists.utsouthwestern.edu. > > Hello Histoteckies, > What are y'all doing regarding the CAP requirement to monitor the pH of > formalin? > We buy tons and tons of the 5 gallon cubitainers and we are still debating > over how to check the pH. > > Looking forward to your replies. > Toodles! > Sincerely, > > Paula Sicurello > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > https://urldefense.com/v3/__http://secure-web.cisco.com/1dY8pbLOSpzQKfWx-yvsO5rujytRiLi5FacpaIq-afFO3eWHoj801ibzodIvkn6kWJvH0G2K0mQEZsm-bweaI-0WiwrKEp9XDD-CIrZCerqewH6wfWut7f2a
Re: [Histonet] Formalin pH
You can check the pH with a pH meter. This is the most accurate way, but the meter's electrode must be calibrated against at least two standard (usually bought) buffer solutions, such as 4.0 and 7.0. If you don't have a pH meter or the know-how to use one properly, you can use indicator papers, which are inexpensive but deteriorate with storage. With a few "books" of indicator papers covering the range 3 to 8 you should be able to get a value accurate to the nearest 0.2 pH units. I'm assuming that by "formalin" you mean the concentrated solution that is nominally 37-40% by weight of formaldehyde gas in water. Its pH falls with time. For more information about formalin, see https://www.cambridge.org/core/services/aop-cambridge-core/content/view/911F1870567E2487D56A51140EB8EA17/S1551929500057060a.pdf/div-class-title-formaldehyde-formalin-paraformaldehyde-and-glutaraldehyde-what-they-are-and-what-they-do-div.pdf. <https://www.cambridge.org/core/services/aop-cambridge-core/content/view/911F1870567E2487D56A51140EB8EA17/S1551929500057060a.pdf/div-class-title-formaldehyde-formalin-paraformaldehyde-and-glutaraldehyde-what-they-are-and-what-they-do-div.pdf> It's a free download, with >600 citations according to Google Scholar. John Kiernan Emeritus, UWO, London, Canada = = = From: Paula Sicurello via Histonet Sent: September 12, 2023 10:11 PM To: HistoNet Subject: [Histonet] Formalin pH Hello Histoteckies, What are y'all doing regarding the CAP requirement to monitor the pH of formalin? We buy tons and tons of the 5 gallon cubitainers and we are still debating over how to check the pH. Looking forward to your replies. Toodles! Sincerely, Paula Sicurello ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Gram Stain
I don't know a method to obtain selective red coloration of Gram-negative organisms, with yellow for both Gram-positives and "background", and I cannot find one by looking in various books. It may not be possible because Gram-positivity relies on selective retention of an immobilized dye. Gram-negative bacteria stain only because the red dye is excluded by the organisms already filled with an insolubilized blue-purple dye in the Gram-positive cells. The Brown-Hopps modification of Gram staining for paraffin sections clearly separates blue from red bacteria and gives a quite different yellowy-brown counterstain to the "tissue background". This is explained and well illustrated in Freida Carson's textbook. (I have only the 2nd edition, 1997; there is now a 5th, 2020, ISBN 9780891896760.) John Kiernan (London, Ontario) = = = From: Rhonda McCormick via Histonet Sent: August 30, 2023 2:42 PM To: Histonet Subject: [Histonet] Gram Stain Howdy,Does anyone have a recommendation for a Gram stain (or modification of a gram stain) that stains the background yellow with red gram-negative bacteria (no blue - gram positive staining).Thank you! ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Sudan Black B
The Sudan black B method for lipofuscin is exactly the same as for frozen sections, but it is applied to hydrated paraffin sections. Remember that the dye solution needs to be fairly fresh (less than 4 weeks old) and must be filtered immediately before using. Use Sudan black B (CI 26150) powder that has been certified by the Biological Stain Commission. Various controls and other stains are needed to be certain that the pigment is lipofuscin, not melanin or haemosiderin. John Kiernan Anatomy, UWO, London, Canada = = = From: Betsy Molinari via Histonet Sent: August 10, 2023 10:57 AM To: Histonet Subject: [Histonet] Sudan Black B Hi, I have been asked to do a Sudan stain on a heart biopsy for lipofuscin. The biopsy is in a paraffin block. They are looking to better report and understand the IHC. I am totally unfamiliar with this stain. I did some reading but have been unable to find a protocol for paraffin sections. I found a reference to Sheehan & Hrapchak (1973) but unfortunately I don't have that edition. Any ideas would be greatly appreciated . Betsy Molinari HT (ASCP) Texas Heart Institute Cardiovascular Pathology 1101 Bates St. Houston, Texas 77030 832-355-6524 Betsy Molinari, HT (ASCP) Sr. Histology Research Technician CV Pathology Research The Texas Heart Institute (r) 6770 Bertner Avenue, MC 1-283 Houston, TX 77030 Office: 832-355-6524 | Fax: 832-355-6812 Email: bmolin...@texasheart.org texasheart.org<https://www.texasheart.org/> | texasheartmedical.org<https://www.texasheartmedical.org/> | facebook<https://www.facebook.com/Texas.Heart.Institute> | twitter<https://twitter.com/Texas_Heart> CONFIDENTIALITY NOTICE: This email and attachments contain information that may be confidential or privileged. If you are not the intended recipient, notify the sender at once and delete this message completely from your information system. Further use, disclosure, or copying of information contained in this email is not authorized, and any such action should not be construed as a waiver of privilege or other confidentiality protections. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Betsy Molinari via Histonet Sent: August 10, 2023 10:57 AM To: Histonet Subject: [Histonet] Sudan Black B Hi, I have been asked to do a Sudan stain on a heart biopsy for lipofuscin. The biopsy is in a paraffin block. They are looking to better report and understand the IHC. I am totally unfamiliar with this stain. I did some reading but have been unable to find a protocol for paraffin sections. I found a reference to Sheehan & Hrapchak (1973) but unfortunately I don't have that edition. Any ideas would be greatly appreciated . Betsy Molinari HT (ASCP) Texas Heart Institute Cardiovascular Pathology 1101 Bates St. Houston, Texas 77030 832-355-6524 Betsy Molinari, HT (ASCP) Sr. Histology Research Technician CV Pathology Research The Texas Heart Institute (r) 6770 Bertner Avenue, MC 1-283 Houston, TX 77030 Office: 832-355-6524 | Fax: 832-355-6812 Email: bmolin...@texasheart.org texasheart.org<https://www.texasheart.org/> | texasheartmedical.org<https://www.texasheartmedical.org/> | facebook<https://www.facebook.com/Texas.Heart.Institute> | twitter<https://twitter.com/Texas_Heart> CONFIDENTIALITY NOTICE: This email and attachments contain information that may be confidential or privileged. If you are not the intended recipient, notify the sender at once and delete this message completely from your information system. Further use, disclosure, or copying of information contained in this email is not authorized, and any such action should not be construed as a waiver of privilege or other confidentiality protections. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Von Kossa staining
Charles, A handheld light of any kind isn't really suitable because you would have to hold it over the slides for 15 to 60 minutes, according to which variant of the von Kossa method you plan to use (see Lillie & Fullmer 1976 Histopathologic Technic ... 4th ed. pp 539-541). An anglepoise lamp with an old-fashioned 100W bulb is OK, and so is a sunny windowsill. Silver salts absorb at the blue end of the spectrum, so a fluorescent light should be more efficient than an incandescent bulb. If no bright light source is available, it's possible to chemically reduce the silver phosphate and/or carbonate to black colloidal silver, with a traditional photographic developer. The method of Rungby et al.1993 may be better than other post-reduction methods (https://scholar.google.ca/scholar?hl=en_sdt=0%2C5=rungby+1993+calcium+deposits=rungby+1993). I never tried it, but Rungby's paper has collected 104 citations, which is very good for a paper in our field. The von Kossa technique is simply explained in my Histological and Histochemical Methods textbook, 5th edn (2015). The book costs less than 1ml of any antibody. Enough said! John John A. Kiernan Emeritus, Anatomy & Cell Biology, University of Western Ontario London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org = = = -Original Message- From: Charles Riley via Histonet Sent: Wednesday, July 26, 2023 1:23 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Von Kossa staining *** Externally sourced email message *** Can anyone out there who performs Von Kossa staining provide me with any guidelines or suggestions for the light source to use for the Silver nitrate activation? Is a standard handheld black light strong enough or does it need to be a UV sanitizing strength light if using UV versus incandescent bulb exposure? ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet CAUTION: This email was originated from outside StatLab and contains links. Do not click links or open attachments unless you recognize the sender and you know the content is safe. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Fast Green / Sirius Red - Unknown blue features
Your technique is the one first (I think) published by Lopez-De Leon A & Rojkind M (1985) A simple micromethod for collagen and total protein determination in formalin-fixed paraffin-embedded sections. J. Histochem. Cytochem. 33: 737-743. The photos in that paper show some of the collagen almost black - surely taking up both red and green dyes. More recent papers describe exactly the same method, and there are also some variants. Your technique, with an acid rinse after staining for an hour, then quick transition to rapid dehydration in 100% alcohol, is essential for any valid picro-sirius staining. According to the entry for fast green FCF (CI 42053) in Conn's Biological Stains (10th ed, p.180-182), "chemically distinct blue-green dyes have been supplied under this name". Are you sure your fast green FCF is the real McCoy? Is it from a batch certified by the Biological Stain Commission? The jar of dye powder should have a small bluish label, with features that make forgery difficult. See https://biologicalstaincommission.org/certified-biological-dyesstains/ for pictures and other information. There are companies selling "certified stains" that have not been tested and certified by the Biological Stain Commission. Caveat emptor! The Biological Stain Commission is a not-for-profit corporation that has been providing third-party quality control and other services for vendors and users of stains for 100 years. Just a few thoughts; I could add more, but probably this letter already is too long for the Histonet censors. John Kiernan Professor Emeritus, Anatomy & Cell Biology University of Western Ontario, London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org = = = From: David Burk via Histonet Sent: June 21, 2023 5:48 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Fast Green / Sirius Red - Unknown blue features We are working out an alternative method for collagen staining using Fast Green / Sirius Red (Direct Red 80) rather than the standard picrosirius red method as I think it is prettier and easier to see the collagen on a green background. What we’ve noticed, though, is that we are seeing intermediate blue staining in the tissue in particular regions or structures. I’ve not had any success in finding an explanation for this online and was hoping someone on the board may have an idea of what’s going on and what is being stained blue in our tissue sections. We have examined a variety of murine or rat tissues including liver, heart, kidney, lung, skeletal muscle, spleen, brain, pancreas, and even decellularized human adipose tissue. There are, almost always, some structures/features that exhibit a denim blue to lighter blue-green color (at least to my eye) in addition to the expected red-colored structures that we would assume to be collagen, light green cytoplasm, and yellow-ish features stained with picric acid. An interesting tidbit is that these blue-ish stained features are birefringent under polarized light so you would not know their color (with transmitted imaging) was atypical. I don’t want to use a stain if I can’t let people know what a particular color represents and can also cause problems with the quantification of collagen using a color-based approach. Our protocol is as follows: 1. Dewax 2. H2O rinse 3. Stain in a 0.1% Fast Green FCF (C.I. 42053) and 0.1% Direct Red 80 (C.I. 35780) solution dissolved in saturated picric acid for 1 hour at room temperature 4. Dip 5x and then immerse in 0.5% acetic acid for 5 seconds 5. Repeat step 4 6. Dip 5x and then immerse in 100% Ethanol 30 seconds 7. Dehydrate in 100% Ethanol 1 min 8. Repeat step 7 9. 3 x Xylene for 2 min each 10. Coverslip I’m uploading some images from mouse muscle and tumor tissue to the Histonet Image upload site. If that doesn’t work, here are links: Mouse tumor: https://drive.google.com/file/d/1VjOZFzvsQByQLuDtdGPfdwAwap_CVE58/view?usp=sharing Mouse skeletal muscle: https://drive.google.com/file/d/10vT_FKu3-Ad5uemM5gnZmqDCKb3fs2lV/view?usp=sharing Thanks, David Burk ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Peroxidase stain on peripheral smears
What is the Kaplow method? I can't find it in textbooks. A quick Google search brings up only junk papers indicating that a Kaplov method may use carcinogenic benzidine (with wrong spelling) as the chromogen. There are simple, safe and inexpensive methods for histochemical localization of sites of peroxidase activity.in tissue sections or smears of cells. Buy a textbook for about $50, read it, and save your lab a fortune in buying special kits for very simple routine histochemical jobs. My ten cents-worth. June 2023 John Kiernan J. A. Kiernan MB, ChB, PhD, DSc Professor Emeritus, Anatomy & Cell Biology University of Western Ontario, London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org = = = From: Kendra Beechie ND-Bismarck via Histonet Sent: June 12, 2023 3:22 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Peroxidase stain on peripheral smears Hello, I am seeking some guidance in regards to a peroxidase stain. We have always used the Kaplow Method to stain peripheral blood smears, and up until recently, we have had no issues with getting it to work. However, no matter what we try in recent months, we have not been able to get it to work. Several different techs have tried it and we have ordered new reagents, but there are no granules to be seen Does anyone have any experience with peroxidase kits? I can see that Sigma-Aldrich has several available but I'm not sure what to go with Any and all help would be greatly appreciated! Thank you, Kendra Beechie MLS/HTL (ASCP), Technical Consultant CHI St. Alexius Health Bismarck, ND Caution: This email is both proprietary and confidential, and not intended for transmission to (or receipt by) any unauthorized person(s). If you believe that you have received this email in error, do not read any attachments. Instead, kindly reply to the sender stating that you have received the message in error. Then destroy it and any attachments. Thank you. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Kendra Beechie ND-Bismarck via Histonet Sent: June 12, 2023 3:22 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Peroxidase stain on peripheral smears Hello, I am seeking some guidance in regards to a peroxidase stain. We have always used the Kaplow Method to stain peripheral blood smears, and up until recently, we have had no issues with getting it to work. However, no matter what we try in recent months, we have not been able to get it to work. Several different techs have tried it and we have ordered new reagents, but there are no granules to be seen Does anyone have any experience with peroxidase kits? I can see that Sigma-Aldrich has several available but I'm not sure what to go with Any and all help would be greatly appreciated! Thank you, Kendra Beechie MLS/HTL (ASCP), Technical Consultant CHI St. Alexius Health Bismarck, ND Caution: This email is both proprietary and confidential, and not intended for transmission to (or receipt by) any unauthorized person(s). If you believe that you have received this email in error, do not read any attachments. Instead, kindly reply to the sender stating that you have received the message in error. Then destroy it and any attachments. Thank you. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Coffee at the desk
Bob, Why weren't they routinely buffering (or at least neutralizing) the formalin fixatives at Johns Hopkins as recently as 1970? It had all been in the scholarly books (by Pearse, Lillie, etc) for >10 years, and was also in Lee Luna's 1968 Manual of Histologic Staining Methods, published by the USA's Armed Forces Institute of Pathology. I was brung up on cold neutral buffered formalin for enzyme activity histochemistry when I was a medical student doing an interpolated research year in the early 1960s. Cheers, John John Kiernan (London, Canada) = = = From: Bob Richmond via Histonet Sent: June 3, 2023 8:25 AM To: Histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Coffee at the desk This 84 year old pathologist recalls the histopathology laboratory at Johns Hopkins Hospital around 1970, when I was a pathology resident there. Histotechs, often laboratory clerks, sat in front of rows of 400 mL Stender dishes, smoking cigarettes while they hand-stained slides, often carrying out the entire procedure from xylene and descending alcohols, up to final coverslipping. It wasn't the xylene that worried me, so much as the dish full of a 20% solution of picric acid in acetone, that removed most of the copious formalin pigment (since buffering the formalin wasn't permitted). I spoke to the chief histotechnologist about the issue. He responded by stubbing out a lighted cigarette into one of the Stender dishes of xylene. (I'm told you can also do that with gasoline, but not with acetone.) He was never incinerated, but he died of smoking-related disease soon after his retirement. Bob Richmond Maryville TN ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Get the Short Term or Long Term Lab Staffing Coverage You Need
Hear, Hear! Let's see less advertising on Histonet. John Kiernan (London, Canada). = = = From: Jay Lundgren via Histonet Sent: June 2, 2023 1:15 PM To: Melissa Owens Cc: Tom Walls Subject: Re: [Histonet] Get the Short Term or Long Term Lab Staffing Coverage You Need As far as I know, Histonet doesn't really allow advertising for individuals or agencies *looking* for jobs. There are a very few, selective, agencies that are allowed to post on Histonet tastefully and infrequently, but they are buying, not selling. Or histotechs who let others know about open positions in their organization. That's cool. I'm not a moderator, but this is supposed to be a collegial forum for practical histopathology advice. There are plenty of job search sites out there already. But if Histonet does start allowing this, then ***%%%$$$HEY SCRIPPS INSTITUTE HIRE ME$$$*** ***%%%$$$LONG OR SHORT CONTRACT!!!CHEAP$$$%%%*** **%%%$$$BIG TIME IMUNNOHISTOCHEMISTRY$$$%%%*** ***%%%$$$USAF HTL (ASCP) M.S. 30+ YRS!!!$$$%%%*** Sincerely, Jay A. Lundgren, M.S., HTL (ASCP) On Fri, Jun 2, 2023 at 10:50 AM 'Melissa Owens' via Histonet < histonet@lists.utsouthwestern.edu> wrote: > li {display:list-item;text-indent: -1em;} > ul, ol{margin-left: 40px !important; padding-left: 0px > !important;} > > When workloads spike, unexpected projects hit, employees are out > for PTO, Sick or Parental Leave, we got your Lab Covered! Our > Staffing Services for the lab can get the work done quickly. We > have skilled laboratory talent at the ready and range of > temporary staffing options to meet your unique needs. > > · Short Term/Leave Coverage > > · Last Minute Sick Coverage > > · Long Term Leave Coverage > > · Long Term Laboratory Training Program > > · Contract to Hire (Asses a candidate on the job before > committing to hiring them for a permanent role) > > We place skilled laboratory talent across the Anatomic and > Clinical Pathology specializations. > > · No minimum commitment on number of weeks you need > laboratory staffing (Customized to your needs) > > · For Sick Coverage there is 6 hour/day Minimum hours billed > > · All inclusive bill rates (Travel/Pay Rate/Employment > Taxes, etc.) > > Candidates we have right now who can make an impact in your lab! > > Histotech Supervisor/ Manager (2 Available) > > * Certified Histotech (ASCP) with over 20 years of experience > * Seeking Temporary or Permanent work > * Location: Seeking Anywhere along the East coast from FL to ME, > West Coast > * Available: Now > > Histotech > > * ASCP Certified with 10 years of experience > * Primarily looking for Permanent but would also consider > Temp/Travel work > * Georgia > * Available: Starting in July 2023 > > Histotech (5 Available Candidates) > > * ASCP Certified and 5 years of experience > * Looking for Travel/Temp Position but open for Temp-Permanent > (North Carolina) > * Locations of interest: Anywhere in the US, Des Moines, IA, > South Charleston, SC, Birmingham, AL > * Available: Now > > Cytotechnologist-(1 Candidate Available) > > * Several years of experience with ASCP certification and Imaging > Trained > * Seeking primarily Temporary or Temporary to Permanent > * Locations of interest: Anywhere in US (Not State Licensed) > * Available: Now > > Medical Technologist-(7 Available Candidates) > > * Several years’ experience as a Generalists > * Seeking Temporary/Travel > * Locations of Interest: Anywhere in US > * Availability: Now > > Medical Laboratory Technician-Generalist-(3 Available Candidates) > > * Has 2 years’ experience and MLT Certification > * Seeking Permanent or Temporary Work > * Location seeking: Houston, TX area > * Available: Now > > Molecular Technologist – (3 Candidates Available) > > * ASCP Molecular Specialists with 2+ years of experience > * Seeking Permanent, Temp to Permanent or Permanent > * Location: Anywhere in US > * Available: Now > > Phlebotomist- (2 Available Candidates) > > * Certified Phlebotomist (CPT) and ASCP Certified Phlebotomists > and 5-10 years of experience > * Seeking Temp/Travel Work Only > * Location Seeking: Anywhere in US > * Ability to Train other Phlebotomists: YES > * Available: Now > > Medical Assistant with Phlebotomy Experience- (2 Candidates > Available) > > * Has Medical Assistant Diploma with phlebotomy experience and > over 3 years’ experience > * Seeking Temporary or Permanent work > * Location seeking: Houston, TX > * Available: April > > *If you do not see a professional on this list that fits with > what your talent needs are please inquire directly to discuss o
Re: [Histonet] history of H staining
Gudrun, your question got me looking through more than a dozen older books, several more recent ones and various articles, but with no clear answer! H wasn't a routine combination in 1902. Pathology and normal human histology textbooks in the 1950s show pictures that are clearly H but with the stain getting scarcely a mention, and this is also true of the most recent (1999) path text on my shelf. Forty-five alum-haematein mixtures were published between 1868 and 1951. Of these, a majority (26) were in the period 1882-1916 and these include the best-known ones: Delafield, Ehrlich, Mayer, Harris etc, but eosin alone was seldom the recommended counterstain before 1890. H has never been the "routine" stain outside the fields of human and veterinary histology and pathology. Other staining combinations are preferred for invertebrates, protozoa, plants and bacteria. My guess is that H gradually became "routine" for pathology in the period 1910-1930. If someone has access to some non-technical textbooks from those decades they might be able to narrow down the dates. I could go on and on, with references etc, but this reply may already be too long for Histonet. John Kiernan. = = = From: Gudrun Lang via Histonet Sent: May 20, 2023 8:46 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] history of H staining Hi all! Does anybody know, when the H stain became that dominant routine-stain in the pathology labs? It was introduced by Wissowzky 1876, but I am curious when our usual histoprocess became worldwide standard. Regards Gudrun Lang ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Mice brain
Immersion in sucrose is for cryoprotection - to minimize damage from formation of ice crystals. It is needed only if you are cutting frozen sections. Specific demonstration of cholinergic neurons is with immunohistochemistry to detect choline acetyltransferase. Use frozen or paraffin sections, according to instructions that come with the primary antibody. For histochemical demonstration of acetylcholinesterase activity (seen also in some neurons that are not cholinergic), you need frozen sections of tissue fixed for 12h in neutral formaldehyde at 4oC. My favourite method for AChE activity was that of Karnovsky & Roots 1964 J. Histochem. Cytochem. 12:219-221. John Kiernan (London, Canada). = = = From: Renee Fisher via Histonet Sent: April 19, 2023 1:34 PM To: Histonet@lists.utsouthwestern.edu Subject: [Histonet] Mice brain Hi, Does anyone know if immersing in sucrose is essential for optimum visualization of cholinergic neurons in mouse brain. Thank you Sent from my iPhone ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Alcian Green Dye for Attwood stain
Alcian green was a mixture of two cationic "Ingrain" dyes: one of ICI's alcian blues (a copper phthalocyanine) and their alcian yellow (CI 12840, a monoazo dye). ICI stopped making these dyes, which were a commercial flop, in the early 1970s. It's extremely unlikely that anything now sold as "alcian green" will be the genuine article. The original "Alcian blue" dyes were unstable even as powders. The staining procedure published by HD Attwood in J. Path. Bact. 76:211 (1958) has to be seen as an oddity from the days when carbohydrate histochemistry was still in its infancy. Any textbook published since the 1970s will provide very simple and rational staining techniques for showing cartilage matrix (basophilic) and keratin (acidophilic) in contrasting colours. John Kiernan London, Canada = = = To: 'histonet@lists.utsouthwestern.edu' Subject: [Histonet] Alcian Green Dye for Attwood stain Hi All, I could use some assistance. I'm gotten a request to do an Attwood stain for squames and cartilage and I'm having a difficult time finding the Alcian Green 2GX dye powder or even a kit if one exists. Another question is the protocol that I've seen from Histonet archives states that you dilute the phloxine in cellosolve. I would just like to ask if anyone has another version of the protocol with any updates to the protocol? Thanks in advance. Debbie Siena ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Debra Siena via Histonet Sent: April 5, 2023 2:47 PM To: 'histonet@lists.utsouthwestern.edu' Subject: [Histonet] Alcian Green Dye for Attwood stain Hi All, I could use some assistance. I'm gotten a request to do an Attwood stain for squames and cartilage and I'm having a difficult time finding the Alcian Green 2GX dye powder or even a kit if one exists. Another question is the protocol that I've seen from Histonet archives states that you dilute the phloxine in cellosolve. I would just like to ask if anyone has another version of the protocol with any updates to the protocol? Thanks in advance. Debbie Siena ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Sudanblack B on FFPET
It's true that Sudan black B won't stain ordinary lipids (fat, phospholipids etc) that are absent from paraffin sections. The lipid in lipofuscin is bound to protein strongly enough to resist extraction during passage through all the solvents used in preparing paraffin sections. Churukian's adaptation of the Lillie & Ashburn method (link to a free download methods book in my recent Histonet email) is OK. It is one of the Biological Stain Commission's tests for certifying Sudan black B. John Kiernan London, Ontario = = = From: AJ Cabral via Histonet Sent: March 28, 2023 4:12 PM To: Tony Henwood Cc: Histonet@lists.utsouthwestern.edu ; Bob Richmond Subject: Re: [Histonet] Sudanblack B on FFPET Sudan Black staining won’t work on FFPET. The alcohols and xylenes used in the tissue processing dissolves the lipids in the tissue. However, you can used formalin fixed tissue as an alternative if no frozen section is available. Rinse the tissue in distilled water for several minutes, pat dry, freeze the tissue on OCT, cut frozen sections and stain in Sudan black. Have you considered looking into acid phosphatase staining for lipofucshin? It is non specific but it can be demonstrated in muscle biopsy. Cheers, Joanna On Tue, Mar 28, 2023 at 12:22 PM Tony Henwood via Histonet < histonet@lists.utsouthwestern.edu> wrote: > I would also let the saturated solution stand for a few days. Like Oil Red > O, it probably needs time to “mature”. I would also use a frozen section of > skin as a control. > > Regards, > > Tony Henwood > Sydney, Australia > > From: Bob Richmond via Histonet<mailto:histonet@lists.utsouthwestern.edu> > Sent: Wednesday, 29 March 2023 4:51 AM > To: Histonet@lists.utsouthwestern.edu histonet@lists.utsouthwestern.edu> > Subject: Re: [Histonet] Sudanblack B on FFPET > > > > > Gudrun Lang in Austria asks: > > > > >>Has anyone experience with Sudanblack B on paraffin slides for staining > [lipofuscin]? A doctor wants the demonstration of the lipoid content of > foamy cells or granulocytes in lung. I've found protocols that have > incubation-times from 10 minutes to over-night. - I've made a saturated > Sudan black B-solution in 70% ethanol and tried it with10 min on liver > without real success.<< > > The main thing you need to do is demonstrate that it isn't hemosiderin with > an iron stain (Perls prussian blue reaction), and perhaps also that it > isn't acid-fast. Lipofuscin can be identified an H & E staining, except for > these considerations. > > Bob Richmond > Maryville, Tennessee > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > http://lists.utsouthwestern.edu/mailman/listinfo/histonet > ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Sudanblack B on FFPET
You probably would get better results with a supersaturated solution of the dye in 60% isopropanol with added dextrin. For details see https://www.urmc.rochester.edu/urmc-labs/pathology/StainsManual/. Click on Table of Contents and follow the links to fined the method. Be sure to use a batch of Sudan black B certified by the Biological Stain Commission. John Kiernan London, Canada = = = From: Gudrun Lang via Histonet Sent: March 28, 2023 10:30 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Sudanblack B on FFPET Hallo! Has anyone experience with Sudanblack B on paraffin slides for staining lipofuszin? A doctor wants the demonstration of the lipoid content of foamy cells or granulocytes in lung. I've found protocols that have incubation-times from 10 minutes to over-night. I've made a saturated Sudanblack B-solution in 70% ethanol and tried it with 10 min on liver without real success. I would appreciate any input and help. Thanks in advance Gudrun Lang Austria ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] IHC staining of tendons and cartilage
You might like to look at this 1999 article from Microscopy Today, about keeping sections on slides. https://publish.uwo.ca/~jkiernan/adhesivs.htm John Kiernan London, Canada. = = = From: Shirley A. Powell via Histonet Sent: March 21, 2023 2:56 PM To: Charles Riley Cc: Histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] IHC staining of tendons and cartilage Hi Charles, Shirley Powell here in humid Georgia. I ran an IHC reference lab here for many years. I had a problem with using charged slides for a lot of the tissues I processed. I used manual and automation methods. My tissues were washing off a lot. I changed to an adhesive for the water bath called Sta-On and I think Surgipath was the company that made it. Surgipath was bought out by Leica but they still sold it. Sta-On was the best adhesive I had found and that worked for me for many years. Whenever I do IHC that is what I use, especially bone, cartilage, bloody specimens, autopsy tissues, they stay on better. Some other companies may be selling it now, like VWR/Avantor. Just Google it. Shirley Shirley Powell, HTL(ASCP) Technical Director Histology Curricular Support Laboratory Pathology Department Mercer University School of Medicine powell...@mercer.edu Phone: 478-301-2374 https://medicine.mercer.edu/ -Original Message- From: Charles Riley via Histonet Sent: Tuesday, March 21, 2023 2:31 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] IHC staining of tendons and cartilage Hello all, I am in a new position and it will potentially require doing a lot of IHC testing on cartilage and tendon samples. I have done some practice runs on my automated stainer and manually and am running into issues with the tissue sections falling off completely or folding over on itself during each process. If anyone does staining like this routinely and has some pointers/tricks to try to get the samples to adhere to the slides better it would be greatly appreciated. I have tried using charged slides from a variety of vendors and get similar results across the board. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu https://nam04.safelinks.protection.outlook.com/?url=http%3A%2F%2Flists.utsouthwestern.edu%2Fmailman%2Flistinfo%2Fhistonet=05%7C01%7Cpowell_sa%40mercer.edu%7C148071115eac460740a508db2a3a6b5e%7C4fb34d2889b247109bcc30824d17fc30%7C0%7C0%7C638150202637381264%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C3000%7C%7C%7C=zZqlEosD%2BmC9X%2FyAsT1k05BSBojNv2uveLaj7nEf730%3D=0 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Conferences 2023
The 2023 meeting of the Biological Stain Commission, marking the organization's 100th anniversary, will be held in Rochester, NY, USA in June 2023. Probably on Friday June 9th. There will be invited presentations and poster sessions. Details are not yet available but should appear on the Commission's web site at some time in the next few weeks. Keep an eye on https://biostain.com. On the BSC web site you can also see what was presented at about 20 earlier BSC meetings (at https://biologicalstaincommission.org/category/annual-meetings/). The BSC web site also has many other links useful to histotechnologists, notably a Glossary and a variety of What's New? items, with conspicuous links on the Home page. The glossary comes up as the top item if you type stain glossary (or similar) into Google. I can see, Lisa, that you won't be travelling from Australia to the USA for a one-day meeting! Your Histonet query has provided an opportunity to advertise the BSC's forthcoming centennial celebration. The availability of BSC-Certified stains (about 60 dyes, as powders) underlies the validity of much medical and other biological research and of all routine staining done in pathology labs with bought pre-made dye solutions. Beware of fake "certified" or similarly named stains sold perhaps from China and India and marketed in Europe and the Americas. These do not carry the BSC's small and difficult-to-forge bottle label, which subtly indicates the history of every truly certified stain. John Kiernan https://biologicalstaincommission.org/category/news-about-dyes-and-stains/ = = = From: Lisa whitham via Histonet Sent: December 8, 2022 3:06 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Conferences 2023 Hello I am a grossing supervisor from Australia and am wishing to attend an international conference in 2023. Can anyone tell me what conferences are being held in 2023? Kind Regards Lisa Whitham ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet From: Lisa whitham via Histonet Sent: December 8, 2022 3:06 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Conferences 2023 Hello I am a grossing supervisor from Australia and am wishing to attend an international conference in 2023. Can anyone tell me what conferences are being held in 2023? Kind Regards Lisa Whitham ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Immunofluorescense staining
If the antibody's supplier does not provide instructions for IHC you will need to test a wide range of concentrations of the primary antibody on sections known to contain the antigen in easily identifiable cell-types or other sites. Use a secondary antibody and detection system that you have already found reliable for primary antibodies raised in the same species as that in which the antibody for flow cytometry was raised. This would probably be a fluorescently labelled secondary antibody if "immunoflourescense" is specified. The researcher probably needs to read a book about immunohistochemistry before expecting someone else to do it for her/him. An investment of $100 or so for a lab, along with a few hours of reading (by boss and workers), can save many hundreds of dollars that might otherwise be spent on antibodies and other reagents that don't work as expected. A good one to start with is Suvarna, S. K., Layton, C., Bancroft, J. D., eds. 2018. Bancroft's Theory and Practice of Histological Techniques 8th ed. London: Churchill Livingstone Elsevier. ISBN: 978-0-7020-6864-5. (NO, I don't have any vested interest!). I'm sure other histonetters will come up with similar and perhaps better suggestions. Beware of working from "protocols" informally passed around among technicians, grad students and research fellows without published references (papers, books) that you can check before investing time, effort and money in a technique that's new to you. Informal lab notes are often locally treated as if they "have Authority, not as the Scribes". In practical science it's probably best to go first with the Scribes, because their writings have been severely reviewed (if in good journals). Good books have references to peer-reviewed papers. I could go on and on, but that's enough of my grumpy advice for now. John Kiernan London, Canada https://www.schulich.uwo.ca/anatomy/people/faculty/emeriti/kiernan_john.html = = = From: Charles Riley via Histonet Sent: August 24, 2022 2:55 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Immunoflouresence staining I have been given an antibody that is used for flow cytometry yet the researcher wants to use it for IF staining. The protocol they provided is for the flow cytometry staining. Will this produce the immunoflourescense results they are looking for or do i need to use a different method? ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] The Passing Of Dr. James McCormick
Thank you, Tom Pella, for the web links. Did James McCormick really invent the first cryostat? I have always seen the early chapters of the late AGE Pearse's Histochemistry book as a good source for the history of the cryostat. In his 2nd (1960), 3rd (1968) and 4th (last, 1980 ISBN 0443019983) editions, Pearse cited many publications about the development of the cryostat in the late 1950s and early 1960s but he did not mention McCormick. I've not seen the 1st (1953) edition of Pearse's book. It may predate the cryostat. As a student in the UK in 1962 I was taught how to use a "Pearse-Slee" cryostat. It was quite similar to ones made since 2000 and used by exploited graduate students to this day. The temperature controls have steadily improved over the decades, and by the 1980s the microtome within the freezer had become something better than the Cambridge Rocker in the Slee cryostats of the 1960s. The anti-roll plate to allow collecting a flat section was in place 60 years ago, and using it was never easy. It is necessary for doing some enzyme activity histochemistry methods (dehydrogenases, cytochrome oxidase etc) on nominally thin (~10um) sections of unfixed tissue. The later invention of the vibrating microtome (Vibratome) has made it quite easy to collect and handle sections of unfixed specimens that have not been frozen, but the sections have to be quite thick (50um or more). That is often OK in research, especially for sections of brains. It is also comparable to traditional frozen sections for surgical pathology, which were cut from pieces of tissue briefly immersed in boiling hot 4% formaldehyde. The sections were cut with a traditional freezing microtome, collected from knife as they melted, and deftly moved onto slides with a paintbrush. This technology is not extinct. John Kiernan = = = From: Tom Pella via Histonet Sent: July 23, 2022 3:44 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] The Passing Of Dr. James McCormick I've skimmed over the posts to Histonet since late June and I haven't seen any post on Histonet, where the passing of Dr. James McCormick was mentioned. He died on June 26th, 2022. I saw this mentioned on Histology Professionals on FB but not here. Here are a few links to online obituaries, the second with some rolling pictures: https://www.legacy.com/us/obituaries/chicagotribune/name/james-mccormick-obi tuary?id=35647547 https://www.swedishhospitalfoundation.org/news/in-memoriam-dr-james-mccormic k His contributions to the field of Histology can't be overstated. He invented the first Cryostat, the first Histology Automated Tissue Processor, the first Embedding Center, the first Tissue Cassette (and many other subsequent cassettes), the first processes to use this instrumentation. His inventions are in use every single day in most Histology Labs worldwide. I only became acquainted with Dr. McCormick for a brief time later in his life on a product collaboration. I found him to be the consummate gentleman; a person in whom ideas were always bubbling to the surface; gracious and intelligent and witty. His passing is a great loss for this community. His public memorial service was held just today. Tom Pella President Ted Pella, Inc. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Dr Freida Carson
Freida Carson was also well known to the Biological Stain Commission. She came to our meetings, and she will be missed by our members. John Kiernan Secretary, Biological Stain Commission Inc. https://biologicalstaincommission.org = = = From: Tony Henwood (SCHN) via Histonet Sent: January 12, 2022 5:25 PM To: Shirley A. Powell ; Willis, Donna G Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Dr Freida Carson I agree, She will be missed 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: Shirley A. Powell via Histonet [mailto:histonet@lists.utsouthwestern.edu] Sent: Thursday, 13 January 2022 7:59 AM To: Willis, Donna G Cc: Histonet (histonet@lists.utsouthwestern.edu) Subject: Re: [Histonet] Dr Freida Carson Hi Donna, So sorry to hear of this. I am proud to have known her and been a recipient of her wealth of knowledge. She contributed so much to the Histology Community. Shirley Shirley Powell, HTL(ASCP) Technical Director Histology Curricular Support Laboratory Pathology Department Mercer University School of Medicine 1550 College St, Macon, GA 31207 O: 478-301-2374/F:478-301-5489 medicine.mercer.edu -Original Message- From: Willis, Donna G via Histonet Sent: Wednesday, January 12, 2022 3:19 PM To: Histonet (histonet@lists.utsouthwestern.edu) Subject: [Histonet] Dr Freida Carson For those of you that are not on Facebook you may not already know that yesterday Freida Carson, PhD became one of our Histology Angels. I had the privilege of having her as my Histology Educator long before she wrote her first edition of Histotechnology. But more precious to me than having her as my educator was that she was a friend. She will be missed. Thank you Freida for all that you have given to the Histology Profession and to me. Rest In Peace singing in the Angels Choir. Donna Willis Anatomic Pathology Manager Baylor Scott Health Baylor University Medical Center 3500 Gaston Ave|Dallas, Texas 75246 214-820-2465 office|214-725-6184 mobile ** The information contained in this e-mail may be privileged, confidential, and/or protected from disclosure. If you are the intended recipient, further disclosures are prohibited without proper authorization. If you are not the intended recipient (or have received this e-mail in error) please notify the sender immediately and destroy this e-mail. Any unauthorized copying, disclosure or distribution of the material in this e-mail is strictly prohibited and no waiver of any attorney-client, work product, or other privilege is intended. No binding agreement on behalf of Baylor Scott & White Health, or any affiliated entity, is permitted by e-mail without express written confirmation by a duly authorized representative of Baylor Scott & White Health. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu https://nam04.safelinks.protection.outlook.com/?url=http%3A%2F%2Flists.utsouthwestern.edu%2Fmailman%2Flistinfo%2Fhistonetdata=04%7C01%7Cpowell_sa%40mercer.edu%7C005224250892499f0a4b08d9d608c6e1%7C4fb34d2889b247109bcc30824d17fc30%7C0%7C1%7C637776155461137075%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C3000sdata=arzYOO05nZbFX5Ng%2Fsse8WA6b1I2OTkeVisiwergLig%3Dreserved=0 ___ Histonet mailing list Histonet@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@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
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@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@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Reprocessing Protocol
It makes no sense to heat a paraffin-infiltrated specimen in saline. Sodium chloride isn't soluble in hot paraffin or in any of the organic solvents used in tissue processing. Heating in water may melt and float out all the wax and rehydrate a specimen, but does it matter if some wax remains? Reprocessing is an attempt to correct the effects of incomplete dehydration. This can be done by taking the specimen back into the clearing agent (xylene or similar) and then into 2 changes of 100% alcohol (methyl, ethyl or isopropyl). Why rehydrate? For a rehydrated specimen, why go "from formalin, using a schedule that would have been of appropriate length as used initially"? This will surely produce, again, a block that is incompletely dehydrated. John Kiernan London, Canada = = = From: Etheridge, Sandra AFF:EX via Histonet Sent: October 12, 2021 5:19 PM To: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Reprocessing Protocol Hi Curt, We have used Taggart's Method quite successfully in the past for poorly processed tissues. You can find it online. The isotonic saline may help to rehydrate your tissues. 1. Melt down the tissue block in the embedding centre block tray area and gently blot off the excess wax. Place the tissue in a newly labelled cassette. 2. Place the cassette into a beaker of isotonic saline (0.9% sodium chloride) and place it in the 65 C incubator/oven for one hour. This will melt the residual wax which will rise to the surface of the saline. 3. Remove the cassette from the saline, drain briefly and place in your processor from formalin, using a schedule that would have been of appropriate length as used initially. 4. Embed and section as per usual. Good luck! Sandra Etheridge -Original Message- From: histonet-requ...@lists.utsouthwestern.edu Sent: October 12, 2021 10:00 AM To: histonet@lists.utsouthwestern.edu Subject: Histonet Digest, Vol 215, Issue 8 [EXTERNAL] This email came from an external source. Only open attachments or links that you are expecting from a known sender. Send Histonet mailing list submissions to histonet@lists.utsouthwestern.edu To subscribe or unsubscribe via the World Wide Web, visit https://can01.safelinks.protection.outlook.com/?url=http%3A%2F%2Flists.utsouthwestern.edu%2Fmailman%2Flistinfo%2Fhistonetdata=04%7C01%7Csandra.etheridge%40gov.bc.ca%7Cbdb6ae0c78de43df498d08d98da1c4da%7C6fdb52003d0d4a8ab036d3685e359adc%7C0%7C0%7C637696548191633375%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C1000sdata=BrEkw92hcrZCE2E7LfNRrM3Zi%2BIxWyNUSevhTPcXqdQ%3Dreserved=0 or, via email, send a message with subject or body 'help' to histonet-requ...@lists.utsouthwestern.edu You can reach the person managing the list at histonet-ow...@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. reprocessing tissue (Curt Tague) -- Message: 1 Date: Tue, 12 Oct 2021 15:14:54 + From: Curt Tague To: "histonet@lists.utsouthwestern.edu" Subject: [Histonet] reprocessing tissue Message-ID: Content-Type: text/plain; charset="us-ascii" I have a problem... some tissue got processed very poorly, there was water in the system somewhere and a few blocks just look burnt.. the nuclei are faint and cloudy, no detail at all. I've tried the process of rehydrating with the 30% formaldehyde, glycerol and sodium acetate solution but they still process poorly, come out very brittle and just don't look good under the scope. Does anyone have a magic bullet to salvage these specimens? I can send a pic directly if it helps. Thanks, Curt -- Subject: Digest Footer ___ Histonet mailing list Histonet@lists.utsouthwestern.edu https://can01.safelinks.protection.outlook.com/?url=http%3A%2F%2Flists.utsouthwestern.edu%2Fmailman%2Flistinfo%2Fhistonetdata=04%7C01%7Csandra.etheridge%40gov.bc.ca%7Cbdb6ae0c78de43df498d08d98da1c4da%7C6fdb52003d0d4a8ab036d3685e359adc%7C0%7C0%7C637696548191633375%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C1000sdata=BrEkw92hcrZCE2E7LfNRrM3Zi%2BIxWyNUSevhTPcXqdQ%3Dreserved=0 -- End of Histonet Digest, Vol 215, Issue 8 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] frozen section problem
Yes, definitely ice crystal holes! If the tissues are unfixed you will have to freeze much more rapidly (isopentane cooled with liquid nitrogen.) If fixed in formaldehyde, cryoprotect by immersing the pieces in 20% sucrose, until they sink. John Kiernan Anatomy & Cell Biology, UWO London, Canada = = = From: Bonello Dorianne M at Health-MDH via Histonet Sent: July 16, 2021 11:25 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] frozen section problem Dear all, We are experiencing freezing artifacts on our frozen sections. Basically, we are seeing cavity-like structures under the microscope, mostly elongated, especially when it's a frozen section on brain tissue. This is most probably happening due to ice crystal formation. We're not using cryospray, relying only on the cryobar boost function. Does anyone has a solution to this problem please? Regards, Dorianne Bonello Allied Health Practitioner (MLS) Histology Laboratory - Pathology Health-Mater Dei Hospital [cid:image001.jpg@01D67184.63288530] T +356 +356 25456434 E dorianne.m.bone...@gov.mt Mater Dei Hospital, Triq id-Donaturi tad-Demm, l-Imsida, Malta MSD 2090 | Tel +356 2545 | https://deputyprimeminister.gov.mt/en/MDH/Pages/Home.aspx<https://deputyprimeminister.gov.mt/en/MDH/Pages/Home.aspx> | https://www.facebook.com/materdeihospital/ Think before you print. Kindly consider your environmental responsibility. This email and any files transmitted with it are confidential, may be legally privileged and intended solely for the use of the individual or entity to whom they are addressed. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Prussian Blue Reaction
Overstained? Doesn't that mean the tissue contains a lot of iron and you are seeing where it is - which was the reason for doing Prussian blue histochemistry. Gudrun Lang correctly says that mineral acids won't remove it. Oxalic acid is said to dissolve Prussian blue (? by chelation); I've never tried this. If it works, you will no longer see where the iron is. To see features other than the distribution of iron, why not just stain another section from the block with a general-purpose stain like Giemsa or H? John Kiernan London, Canada = = = From: Mac Donald, Jennifer via Histonet Sent: June 7, 2021 12:47 AM To: Gudrun Lang Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Prussian Blue Reaction The tissue was overstained and the blue was interfering with interpretation -Original Message- From: Gudrun Lang Sent: Sunday, June 6, 2021 2:18 AM To: Mac Donald, Jennifer Cc: histonet@lists.utsouthwestern.edu Subject: AW: [Histonet] Prussian Blue Reaction EXTERNAL SENDER- Exercise caution with requests, links, and attachments. Hi Jennifer, Why do you want to reduce the staining? I ask, because the impact of hydrochloric acid on the tissue may influence the following results anyway. I think, that the prussian blue pigment cannot be removed in an easy way. It is resistent to solvents and mineral acids. https://nam12.safelinks.protection.outlook.com/?url=http%3A%2F%2Fwww.epsilonpigments.com%2Finorganic-pigment%2Fprussian-blue%2FPrussian-Bluedata=04%7C01%7Cjmacdonald%40mtsac.edu%7C0fbc82a2b13749a4222608d928cbfe52%7Ccc4d4bf20a9e4240aedea7d1d688f935%7C0%7C0%7C637585679205067185%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C2000sdata=KjvijcfrVPGZKGsAn6qX5rMKtulHpmsAzqHEkwz%2B96Y%3Dreserved=0 -for-Solvent-Based-Inks.html On the other hand, if the blue colour doesn't interfere with your following staining, you can try to simple make a "double stain". Regards Gudrun -Ursprüngliche Nachricht- Von: Mac Donald, Jennifer via Histonet [mailto:histonet@lists.utsouthwestern.edu] Gesendet: Sonntag, 6. Juni 2021 06:34 An: histonet@lists.utsouthwestern.edu Betreff: [Histonet] Prussian Blue Reaction Does anyone know of a way to remove/reduce the Prussian blue reaction? Thanks, Jennifer ___ Histonet mailing list Histonet@lists.utsouthwestern.edu https://nam12.safelinks.protection.outlook.com/?url=http%3A%2F%2Flists.utsouthwestern.edu%2Fmailman%2Flistinfo%2Fhistonetdata=04%7C01%7Cjmacdonald%40mtsac.edu%7C0fbc82a2b13749a4222608d928cbfe52%7Ccc4d4bf20a9e4240aedea7d1d688f935%7C0%7C0%7C637585679205067185%7CUnknown%7CTWFpbGZsb3d8eyJWIjoiMC4wLjAwMDAiLCJQIjoiV2luMzIiLCJBTiI6Ik1haWwiLCJXVCI6Mn0%3D%7C2000sdata=gRrUmRDEU3BfcA0rEgQOgBvPHIQ05IRM6WozVZiiR1g%3Dreserved=0 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Protocol for DAPI staining on paraffin sections
Not everyone knows that the product of the Feulgen reaction is fluorescent. If you do the method on paraffin sections, DNA in nuclear chromatin is red with ordinary illumination. If you look with a fluorescence microscope (blue excitation) it shows as a brownish-red fluorescence in the chromatin. A similar fluorescent colour is seen in conventional PAS-stained sections, in mucus, basement membranes, etc. You need a non-fluorescent mounting medium: DPX or one of the various poly(methyl methacrylate)-based media such as Entellan. John Kiernan Professor Emeritus, Dept of Anatomy & Cell Biology University of Western Ontario, London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org<https://biologicalstaincommission.org/> = = = <https://biologicalstaincommission.org> From: Alida Bailleul via Histonet Sent: June 3, 2021 7:53 AM To: Histonet@lists.utsouthwestern.edu Subject: [Histonet] Protocol for DAPI staining on paraffin sections Dear Histonet list, I have made some paraffin blocks of demineralized bone and cartilage. I would like to stain paraffin slides with DAPI (or another DNA stain). Does DAPI staining (or other fluorescent DNA stain) work well on paraffin sections? Does anyone have a protocol they would like to share? I am assuming that histological stains (like the Feulgen stain would work better?). I could also try both. Thank you in advance All the best Alida -- Dr. Alida M. Bailleul Associate Research Fellow/Associate Professor Institute of Vertebrate Paleontology and Paleoanthropology, Chinese Academy of Sciences www.ivpp-avianevolution.com<http://www.ivpp-avianevolution.com> & Research Associate of Paleontology, Museum of the Rockies, Montana State University Google Scholar <https://scholar.google.co.jp/citations?user=Yn4PuWMJ=en> - ResearchGate <https://www.researchgate.net/profile/Alida_Bailleul> ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] IF with permanent mounting media?
Espada J, Juarranz A, Galaz S, Canete M, Villanueva A, Pacheco M, Stockert JC (2005) Non-aqueous permanent mounting for immunofluorescence microscopy. Histochem. Cell Biol. 123: 329-334. https://doi.org/10.1007/s00418-005-0769-2 A PDF can be downloaded from the Google Scholar entry for this article. John Kiernan London, Canada = = = From: Morken, Timothy via Histonet Sent: May 28, 2021 11:31 AM To: Histonet Subject: [Histonet] IF with permanent mounting media? Has anyone tried using xylene/permanent mounting media for immunofluorescence stains? I had a question from a pathologist who wondered if we could do this. I have never heard of anyone doing it. Tim Morken Supervisor, Electron Microscopy/Neuromuscular Special Studies Department of Pathology UC San Francisco Medical Center ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Luxol fast blue for staining myelin. Validity of sources.
This Histonet query is addressed to vendors of stains (powders and/or solutions), and to anyone who makes up luxol fast blue in the lab, avoiding the high cost of buying and transporting the flammable ready-made staining solution. Most diagnostic labs buy ready-made staining solutions and follow a vendor's instructions, whereas research labs are likely to buy dye powders, make up their own solutions, and use them according to published books or papers. This means that the largest amounts of luxol fast blue (LFB) powder are sold to vendors of staining solutions. Am I right? Are vendors having any problems with getting dye powders called luxol fast blue (LFB) that are consistently OK for staining myelin sheaths of axons in sections of brain, spinal cord etc? The LFB name is informally attached to at least three dyes used as stains: LFB G (CI Solvent blue 34) and LFB ARN (CI Solvent blue37) are azo dyes, whereas LFB MBS (CI 74180, Solvent blue 38) is a copper phthalocyanine. All three have been used in similar (but not identical) published staining methods for myelin. LFB MBS is the dye most often used for this purpose, but not much is known about variation in the purity of commercial batches. It has many commercial uses and dozens of trade-names. See http://www.worlddyevariety.com/category/solvent-dyes. (It's free, unlike the Colour Index, but don't believe everything it says.) Luxol fast blue (whichever one) may or may not be the best stain for showing myelinated axons and regions of demyelination, but it probably is the dye most used for doing this job. Is there a need for independent third-party identification, testing and certification of LFB powders? If you have encountered a bad jar of powder labeled luxol fast blue, please reply, including if possible the name on the label and the source. The Biological Stain Commission (BSC) has never offered testing and certification for LFB as a stain for myelin, even though "luxol" dye have been in use for this job since 1953. Should third-party certification be made available for luxol fast blue? Would you submit samples for this service? For more information, see the link below. John Kiernan Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org/for-vendors/ For Vendors and Users of Stains | The Biological Stain Commission<https://biologicalstaincommission.org/for-vendors/> Vendor FAQs: The Biological Stain Commission is a not-for-profit organization which is dedicated to the endorsement of histological materials. It is the mission of the BSC to ensure the quality of biological stains on the market that are sold by many US companies. biologicalstaincommission.org = = = ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] doing Sudan Black B
Hydrated paraffin sections of formaldehyde-fixed tissue are stained the same way as frozen sections of formaldehyde-fixed tissue, but in paraffin sections lipofuscin inclusions are just about the only things that stain. They are, of course, brown or yellow without any staining. If you need to identify lipofuscin inclusions with certainty you'll have to do a few other stains and histochemical tests to distinguish them from ceroid (a similar glycolipoprotein pigment), melanin and haemosiderin. It is not a good idea to use a protocol informally passed on from someone else without checking its origins. Even a method printed in a book can contain errors, and for this reason just about all the methods books published since about 1950 have references to peer-reviewed papers that readers can and sometimes should check. Any book in our field is just a big collection of review articles that have not been reviewed as critically as more specialized reviews in a good journal. The late RD Lillie made important original contributions to pigment histochemistry (two are cited below) and his techniques book (also cited) intelligently reviews the subject and provides technical instructions. So does Pearse's Histochemistry. There are, of course, protocols for staining lipofuscin in all the ordinary histotechniques books. You don't need to have all the latest editions on your lab's bookshelf. The 3rd (1965) edition of Lillie is available for only $8 at https://www.abebooks.co.uk/book-search/title/histopathologic-technic-and-practical-histochemistry/author/r-d-lillie/ Histopathologic Technic and Practical Histochemistry by R D Lillie - AbeBooks<https://www.abebooks.co.uk/book-search/title/histopathologic-technic-and-practical-histochemistry/author/r-d-lillie/> Histopathologic Technic and Practical Histochemistry by Lillie, R D and a great selection of related books, art and collectibles available now at AbeBooks.co.uk. www.abebooks.co.uk Snap it up! The 4th and last (1976) edition of Lillie is now $700 from Amazon, but the 1965 edition is still OK for most of what the book is about, and it's easier to read than his rather rambling 4th edition. Lillie RD, Fullmer HM (1976) Histopathologic Technic and Practical Histochemistry. 4th ed. McGraw-Hill, New York. See comments above - the previous edition is cheaper and for you it may be just as good. Pearse AGE, Stoward PJ (1980,1985,1991) Histochemistry, Theoretical and Applied, 4th edn. Vol. 1. Preparative and Optical Technology. Vol. 2. Analytical Technique. Vol. 3. Enzyme Histochemistry. v. 1,2,3. Churchill-Livingstone, Edinburgh. Pigments are in Vol. 2. Third edn (1968. 2 vols) probably just as good for most labs. Lillie RD (1956) The mechanism of Nile blue staining of lipofuscins. J. Histochem. Cytochem. 4:377-381. I think all the papers in this journal are now free for anyone to download. Lillie RD (1956) A Nile blue staining technic for the differentiation of melanin and lipofuscin. Stain Technol. 31:151-153. Sadly, not free unless you have access to a subscribing library or are a member of the Biological Stain Commission. I hope this answer helps. John Kiernan = = = = From: LEROY H BROWN via Histonet Sent: May 9, 2021 10:29 PM To: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] doing Sudan Black B Hi, I am looking for a protocol for Sudan Black B staining on paraffin embedded tissue. Does anyone have a working stain for this? Thanks LeRoy Brown HT(ASCP) HTL ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Movats
Dear Betsy, Don't say you are sorry for putting a long post on Histonet! To get troubleshooting help you need to say exactly what you did. If you wrote only, "why are my sections brown after Movat staining", nobody would understand your problem. Your procedure starts with an hour in hot Bouin. For many years this has been a routine prior to trichrome stains done on sections of specimens fixed in neutral formaldehyde. It isn't part of Movat's original method (Arch. Path. 60:209-295, 1955), which probably was devised for sections optimally fixed for trichrome staining (in mixtures containing mercuric chloride). Movat's pentachrome is a trichrome method preceded by alcian blue (for no obvious reason) and an iron-haematoxylin for nuclei and elastin. It differs from older trichromes in using a mixture of yellow polyene dyes (saffron) to stain the collagen, instead of the blues or greens as in the Mallory and Masson methods. Your method includes "5% sodium thiosulfate -1 min" after the iron-haematoxylin stain for black nuclei and elastic fibres. This also isn't part of Movat's pentachrome method, and I wonder why. Did you inherit an informal list of instructions passed on within the lab? After a mercuric fixative, hydrated sections are dipped in iodine, followed by thiosulphate, before staining, to remove a black deposit (probably mercurous chloride) introduced by the fixative.I've been seeing similar informal passing of bad staining instructions in research labs for many years. Are you a victim of this trend? The thiosulphate step in your procedure obviously does no harm, because you got the right results with the dog tissues. There may be something different about your human specimens: perhaps inadequate fixation, or excessive acid treatment (if that's what Cal rite is) for decalcification. If the sections of human arteries look OK with a microscope, it might not matter that grossly they are a different colour from the dog small intestine sections. They are, after all, different tissues. A rather long, and not very helpful reply! John Kiernan London, Canada = = = From: Betsy Molinari via Histonet Sent: May 5, 2021 9:46 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Movats Hi Histonetters, I have received several human vessels for paraffin processing and to stain the sections for H and Movats. The H were fine. The human sections turned brownish yellow with the Movats.The control which is canine small intestine was perfect. The protocol is standard Bouins 1hr in 58C waterbath Rinse till yellow disappears Rinse in DH2O 1% Alcian Blue -20 min Rinse in running tap H2O -5min Alkaline alcohol-1hr Rinse 10 min tap H2O Rinse in DH2O Verhoff's Hematoxylin -15 min 3 changes DH2O Differentiate in 2% FeCl Rinse in DH2O 5% sodium Thiosulfate -1min Rinse in running tap-10 min Rinse in DH2O Woodstain scarlet/acid fuchsin-1.5 min Rinse in DH2O Rinse in 0.5% acetic acid water 5% aqueous phosphotungstic acid -2 changes 5 min each Rinse in 5% acetic acid water Rinse in 3 changes absolute ETOH 6% alcoholic Safran solution Absolute alcohol-xylene-coverslip The human slides were fine until the Safran step. When I removed them from the stain into the 100% they were a yellowish brown .Under the scope the colors were there, blue, red, yellow and black. But on the slide the tissue was that brownish yellow. The researcher does not like to strong yellow color. Since my control was fine I question if something was going on with their tissue. I do not know how the tissue was handled before it came into the lab. They were very calcified and were decaled for 1-3 days in Cal Rite. I do know they were not rinsed after decal and were put straight back into 10% NBF before I got them for processing. Should I have used a human control instead of canine? These were very large pieces that were crammed into the cassette. Thanks for the help. Sorry for the long post. Betsy Molinari HT(ASCP) Texas Heart Institute Cardiovascular Pathology 1101 Bates St. Houston,TX 832-355=6524 (lab) 832-355-6812 (fax) Betsy Molinari, HT (ASCP) Sr. Histology Research Technician CV Pathology Research Texas Heart Institute 6770 Bertner Avenue, MC 1-283 Houston, TX 77030 Office: 832-355-6524 | Fax: 832-355-6812 Email: bmolin...@texasheart.org texasheart.org<https://www.texasheart.org/> | texasheartmedical.org<https://www.texasheartmedical.org/> | facebook<https://www.facebook.com/Texas.Heart.Institute> | twitter<https://twitter.com/Texas_Heart> CONFIDENTIALITY NOTICE: This email and attachments contain information that may be confidential or privileged. If you are not the intended recipient, notify the sender at once and delete this message completely from your information system. Further use, disclosure, or copying of information contained in this email is not authorized, and any such action should not be
Re: [Histonet] Animal Histology
Jennifer MacDonald asked, "Can anyone recommend a good book for processing animal tissue?" Here's a list of 14 of the ones published since 1990, with very brief descriptive notes. There are, of course, plenty of good books that are older than these. Lyon,H (1991): Theory and Strategy in Histochemistry. A Guide to the Selection and Understanding of Techniques. Springer-Verlag, Berlin. All on theory; no instructions. All aspects of preparation, staining etc are covered. 32 chapters, by different people; one bibliography. Reprinted 2011. ISBN 9783642737442. Kok,LP; Boon,ME (1992): Microwave Cookbook for Microscopists. Art and Science of Visualization. 3rd ed. Coulomb Press Leyden, Leiden. 432 pages. Plenty of microwave theory. Applications in fixation, infiltr, embedding, decalcif., many staining methods including conn.tiss., cytol., neuro. Enzyme, immuno- & hybridization histochemistry. Sanderson,JB (1994): Biological Microtechnique. (Microscopy Handbooks, 28.) BIOS Scientific Publications & Royal Microscopical Society, Oxford. 224 pages. Detailed paractical instructions, with rationale, & titled bibliography for each of the 7 chapters. Collection, Fixation, Processing, Microtomy, Other prep. methods, Staining, Finishing (mounting etc). Chayen,J; Bitensky,L (1991): Practical Histochemistry. 2nd ed. Wiley, Chichester. 321 pages. Special emphasis on use of unfixed cryostat sections. Presnell,JK; Schreibman,MP (1997): Humason's Animal Tissue Techniques. 5th ed. Johns Hopkins University Press, Baltimore. 572 pages. Humason's Animal Tissue Techniques, 5th edition. Includes chapters on immunohistochem, safety, microwaves, lab hints, suppliers, etc. Hayat,MA (1993): Stains and Cytochemical Methods. Plenum Press, New York. Book of techniques. Many are critically discussed. emphasis on EM, but plenty of LM too. Carson,FL; Hladik,C (2009): Histotechnology. A Self-Instructional Text. 3rd ed. American Society of Clinical Pathologists, Chicago. 400 pages. Freida Carson's textbook. 3rd ed, with Hladik, is 96 pages longer than 2nd (1997); with EM and cytopreparation; glossary. ISBN 9780891895817. (There is a 4th ed, 2015, which I haven't seen.) Kiernan,JA (2015): Histological and Histochemical Methods: Theory and Practice. 5th ed. Scion, Banbury, UK. 606 pages. Kumar,GL; Kiernan,JA (Eds.) (2010): Education Guide: Special Stains and H & E. 2nd ed. Dako North America, Carpinteria, CA. 300 pages. 33 chapters. Photos. PDF file is a free download from https://www.agilent.com/en/dako-pathology-education-guides/. Wick,MR (Ed.) (2008): Diagnostic Histochemistry. Cambridge University Press, New York. 460 pages. Multi-author book. Applications of techniques to path. diagnosis. Chapters by systems, organs, diseases etc. Emphasizes value, economy of pre-1970s (before immuno) histochemical methods. Only Ch1, pp.1-27 describes techniques. Many colour photos. Orchard,G; Nation,B (Eds.) (2012): Histopathology. Oxford University Press, Oxford, UK. 396 pages. Multi-author textbook book. Applications of techniques to path. diagnosis. Chapters mainly by methods, with applications to systems, organs, diseases. One of an Inst. of Biomed. Science series, with web site at www.oxfordtextbooks.co.uk/orc/fbs/. Mulisch,M; Welsch,U (Eds.) (2015): Romeis - Mikroskopische Technik. 19th ed. Springer-Verlag, Berlin. 603 pages. 25 chapters, 3 appendices. All types of microscopy, staining, histochem, etc. (in German) ISBN 9783642551901. Exbrayat,JM (Ed.) (2013): Histochemical and Cytochemical Nethods of Visualization. (Series Ed: Morel,G. Methods in Visualization.) CRC Press, Boca Raton, FL. 335 pages. 13 chapters, 10 contributors. 1-7 are for LM; 8-12 for EM; 13 on image quantification. ISBN 978143987. Suvarna,SK; Layton,C; Bancroft,JD (Eds.) (2018): Bancroft's Theory and Practice of Histological Techniques. 8th ed. Churchill Livingstone Elsevier, London. 672 pages. In this 8th ed., histochem chapters for lipids, proteins, nucleic acids, enzyme activities are now compressed into Appx I (of VIII). Hope this helps. For older books, look in AbeBooks, Amazon etc for ones by GG Brown, HC Cook, CFA Culling, RAB Drury, M Gabe, P Gray, RW Horobin, G Humason, JFA McManus, AGE Pearse. John A. Kiernan Dept of Anatomy & Cell Biology University of Western Ontario London, Canada = = = From: Mac Donald, Jennifer via Histonet Sent: May 3, 2021 1:07 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Animal Histology Can anyone recommend a good book for processing animal tissue? Thanks, Jennifer ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Oil Red O
It's in any textbook in the field of histotechnology published since about 1950. This one from Amazon costs less than $8. Every lab should have a shelf of such books. https://www.amazon.com/Introduction-Histotechnology-Geoffrey-G-Brown/dp/0838543405 [https://images-na.ssl-images-amazon.com/images/I/41xQTLdptPL.jpg1z]m]<https://www.amazon.com/Introduction-Histotechnology-Geoffrey-G-Brown/dp/0838543405> An introduction to histotechnology: A manual for the student, practicing technologist, and resident-in-pathology: Brown, Geoffrey G: 9780838543405: Amazon.com: Books<https://www.amazon.com/Introduction-Histotechnology-Geoffrey-G-Brown/dp/0838543405> An introduction to histotechnology: A manual for the student, practicing technologist, and resident-in-pathology [Brown, Geoffrey G] on Amazon.com. *FREE* shipping on qualifying offers. An introduction to histotechnology: A manual for the student, practicing technologist, and resident-in-pathology www.amazon.com John Kiernan London, Canada = = = From: Niihori, Maki - (mniihori) via Histonet Sent: March 5, 2021 4:08 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Oil Red O We would like to stain Right Ventricle (RV) and Lung (both from rat samples) with Oil Red O. I appreciate if anybody can share a good protocol/kit information with me. Thank you, Maki * Maki Niihori, PhD Life Sciences North Rm# 402, Department of Medicine, The University of Arizona Phone: 520-626-6092 E-mail: mniih...@arizona.edu * ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Other Histonet-like listservers?
Try this. https://biologicalstaincommission.org/bscglossary.html Glossary of Staining Methods, Reagents, Immunostaining, Terminology and Eponyms. Currently Version 1.2. Version 2.0 with another 300 or so items will go online quite soon. Other items at http://biologicalstaincommission.org include QUESTIONS AND ANSWERS - Staining, histochemistry and histotechnology FAQ (Frequently Asked Questions), Version 2.0 November 2019, at https://biologicalstaincommission.org/faqlist.htm, and information about BSC-certified stains. There's also Bryan Llewellyn's StainsFile site: https://stainsfile.info/xindex.html, Last updated January 2019. John Kiernan J. A. Kiernan MB, ChB, PhD, DSc Professor Emeritus, Dept of Anatomy & Cell Biology University of Western Ontario, London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org = = = From: Morken, Timothy via Histonet Sent: February 11, 2021 11:39 AM To: Histonet Subject: [Histonet] Other Histonet-like listservers? Hi all, I'm going to give a presentation on online histology help sites like Histonet and the NSH Block. Does anyone know of other listservers that can be of help to histotechnologists in all fields? I checked the MSA EM listserver but it is down for some technical reason. Thanks in advance for any help! Tim Morken Supervisor, Electron Microscopy/Neuromuscular Special Studies Department of Pathology UC San Francisco Medical Center ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Alcian Blue staining
Pink with alcian blue? Are you using an alcian blue pH2.5-PAS sequence? If so you should get PAS-positive mucus (not in goblet cells) in the stomach and AB-positive mucus (greenish blue) in the intestine. Intestinal mucus, especially in the duodenum and jejunum, is also PAS +ve, so the cells appear purplish. In the colon the mucus is mostly sulphated rather than sialylated, so it is only AB +ve, even if the stain is done at pH 1. If a PAS stain is done before the AB, PAS +ve mucus also stains quite strongly with AB. For more about this, see Johannes ML & Klessen C (1984) Alcianblue/PAS or PAS/alcianblue. Remarks on a classical technic used in carbohydrate histochemistry. Histochemistry 80: 129-132. (Unfortunately the paper has only black & white photos.) Various dyes are currently sold as "alcian blue" and not all are suitable for all applications. The Biological Stain Commission now certifies alcian blue dyes as either "alcian blue 8G or equivalent" or as "alcian blue variant". See the recently updated (2020) entry for Alcian blue 8G (CI 74240) and other alcian blue dyes at https://biologicalstaincommission.org/new-dyes/. Current issues of interest to vendors and users of dyes and biological stains. | The Biological Stain Commission<https://biologicalstaincommission.org/new-dyes/> Dyes are becoming more expensive! January 2018. BASF, a major dyestuff manufacturing company, recently announced that it has increased its prices for many pigments and dyes by up to 15% worldwide. biologicalstaincommission.org John Kiernan Secretary. Biological Stain Commission = = = From: Charles Riley via Histonet Sent: October 9, 2020 10:14 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Alcian Blue staining We are having an issue where the goblet cells in our control tissue are not staining pink even though the patient tissue is staining beautifully on the same slide. The stain is done manually. What can be some causes for the issues in staining on the control section? ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] human shoulder joint fixation
Thanks for the compliments, Bob, but I've no experience of trying to section anything as big and bony as a human shoulder joint. Car Hobbs and Izak Dimenstein probably will be able to give better advice to Merissa. Incidentally, I wasn't ever a pathologist. I moved from medicine to neuroanatomy (9-5 job!) at an early age. Some of my research could have been called experimental pathology. I also collaborated with some real neuropathologists, especially in the 1990s looking at the cerebral cortex and spinal cord in ALS and other motor neuron diseases. Cheers, John Kiernan. https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html = = = From: Bob Richmond via Histonet Sent: 21 August 2020 17:25 To: Histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] human shoulder joint fixation > > Merissa (where?) asks; >>I am doing some planning for a new project and > wanted to get opinions on fixation of large pieces of tissue. We will have > human shoulders, where we want to preserve the rotator cuff/joint. Cutting > the tissue with a saw will damage the soft tissue, so we were thinking that > post-fixation would be best for cutting slabs. - Does anyone have > experience with fixing such large pieces of tissue? We typically use zinc > buffered formalin for fixation. Would a vacuum work or a vacuum sealer?<< > This is a non-trivial problem, and I hope you're working closely with the pathologist or other investigator who's going to be looking at the slides. Formalin (forget the zinc) penetrates tissue slowly enough that you're not going to get very good fixation if you put the whole specimen in fixative and forget about it for a week or two. Some preliminary dissection is needed to aid fixation, and you're going to need some serious help with that. A valuable resource person on Histonet is John Kiernan, a pathologist turned research anatomist, and an expert on histologic technique. I hope he responds, otherwise try to find him. Bob Richmond Samurai Pathologist Maryville TN ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Apple Green Birefringence in Amyliod slides
For another source of polarizing filters, go to a 3D movie, take home the glasses they provide, and poke out the lenses. They work very nicely as polarizer and analyzer with an ordinary microscope. John Kiernan Anatomy & Cell Biology University of Western Ontario London, Canada = = = From: Morken, Timothy via Histonet Sent: 28 July 2020 13:57 To: Ken M Cc: Histonet Subject: Re: [Histonet] Apple Green Birefringence in Amyliod slides Ken, Yes, polarized light and apple green birefringence is diagnostic for amyloid with congo red and is the best practice. If you have a problem with known control slides there are two possibilities: 1) make up fresh solution. The pH has to be right. Or 2) try other control slides. Maybe you cut through the amyloid area. Because we have hundreds of microscopes in our department most just use polarized film as the polarizer (put over the light source) and another put over the top of the slide as the analyzer. Turn one of the polarizing slides and you will see the birefringence appear. Source: "Polarizing film, 2"" x 2"" , PK/10 (BEST For use as a microscope polarizer)" Cat# S07372 Thermo Fisher Sci Health$36.75 PK/10 "2" x 2" These are polarized film mounted in 2" film holders (like the old Kodachrome slides). Cheap and effective. (and avoids consternation from people losing expensive microscope polarizers) Tim Morken Supervisor, Electron Microscopy/Neuromuscular Special Studies Department of Pathology UC San Francisco Medical Center -Original Message- From: Ken M via Histonet Sent: Tuesday, July 28, 2020 11:43 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Apple Green Birefringence in Amyliod slides Hi everyone. I was wondering if anyone out there has any experience with diagnosing Amyloid tissue using Congo Red stained Kidney using polarized lenses. Is it common to use polarized light to detect Amyloid deposits? Does the absence of the "apple green birefringence" indicate a problem with the control tissue or the control slides? Should this green bifringence always appear to confirm the diagnosis? I know that the tissue should be cut thicker than normal (we usually cut at 5), but in the future maybe we will cut at 7 or 8? ___ Histonet mailing list Histonet@lists.utsouthwestern.edu https://urldefense.proofpoint.com/v2/url?u=http-3A__lists.utsouthwestern.edu_mailman_listinfo_histonet=DwICAg=iORugZls2LlYyCAZRB3XLg=7cy9qXFa73jDX2Iixpjkq1XlWAfHgLLHm33agI_sCKA=2jEHe8Hf3ieiSbv1r-ZtSy-mm4FVj1XtTmUSGcfJmE8=Q_PpmT_KF2fDUvt9ltFVZLp6ctjM3xkK0RsfuUYW73c= ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Fixed frozen non-paraffin mouse brain
And a very good pennyworth it is, Carl! You wrote, "... someone must've originally thought: 'Hang on, if we fix in commercially bought 40% Formalin, it's got 10% methanol added (to slow rate of formaldehyde repolymerisation) ...that will compete with formaldehyde fixation. So, we get coagulative and additive fixation. That is not good, folkslet's get pure and use depolymerised paraformaldehyde: pure methylene glycol polymer'". That's almost how it came about: let's get pure. A fixative made from PFA should have the same composition every time it is freshly prepared. The 10% of methanol (MeOH) in formalin (40% HCHO) isn't enough to coagulate proteins, and neither is the 1% MeOH in 10% formalin (with 4% HCHO). You need 60-70% alcohol to coagulate proteins, viruses etc. Formalin also contains some formic acid; the amount increases with age, from oxidation of the aldehyde by air. Dilution with water always gives an acidic solution. Marble chips can be added bring the pH up to neutrality. Buffering also takes care of the formic acid and can provide a neutral (pH7) or a "physiological" (pH7.4) fixative solution. The usual phosphate buffer also makes the fixative solution approximately iso-osmotic with mammalian extracellular fluid. Before the 1960s, dilution of formalin with with saline (0.9% NaCl) provided "formal saline", which had some advantages over 4% aqueous formaldehyde. See books by J. R. Baker, which are available as free downloads from http://archive.com. Polymerization also increases with age. That's why you see a white precipitate in bottles of formalin stored for a long time. The precipitate is paraformaldehyde (PFA); its presence reduces the amount of formaldehyde that can be easily released by simple dilution of the formalin with water. According to R. Cares (1945: A note on stored formaldehyde and its easy reconditioning. J. Tech. Methods & Bull. Int. Ass. Med. Museums 25, 67-70), milky formalin can be cleared by autoclaving, for 30 m in Kilner jars. I wonder if anyone else has done this? John Kiernan Anatomy & Cell Biology UWO, London, Canada = = = From: Hobbs, Carl via Histonet Sent: 05 July 2020 14:25 To: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Fixed frozen non-paraffin mouse brain Prof. Kiernan, as usual, provides us all with such a depth/breadth of particular information/advice. His Histological and Histochemical methods BIBLE is still my favourite read. Respect Most researchers fix in depolymerised Paraformaldehyde because someone must've originally thought: " Hang on, if we fix in commercially bought 40% Formalin, it's got 10% methanol added ( to slow rate of formaldehyde repolymerisation) ...that will compete with Formaldehyde fixation. So, we get coagulative and additive fixation. That is not good, folkslet's get pure and use depolymerised Paraformaldehyde: pure methylene glycol polymer" I am sure Professor Kiernan can correct my inaccuracies! Anyway..I've never noticed any difference: I've worked in diagnostic labs ( unfixed frozen muscle/renal/rectal bx) and also research labs ( unfixed/ fixed frozen tissues) using both fixing solutions I have not noticed any IHC/IF difference in reactivity. Many primary abs do NOT work even with fixed/unfixed frozensome of them WILL need HIER ( at 90C rather than M/W or pressure cooker AR but, only fixed frozen of course), imho. Part of the problem is whether the antigen is linear or 3D...sorry for simplicity. I can successfully snap-freeze fixed/unfixed rat/ms brain hemispheres without using sucrose ( success measured by lack of holes at the LM level). This is because I was trained in a diagnostic lab to freeze fast but, effectively. It is a technique that requires experience for consistency of successsometimes I fail! The reason most use 20/30% sucrose is to give poor a snap-freezing technique a chance to avoid ice-crystal artefact, as stated by Kiernan). Sucrose is no panacea.technique is everything. My pennyworth-illy Carl Carl Hobbs FIBMS Histology and Imaging Manager Wolfson CARD Guys Campus, London Bridge Kings College London London SE1 1UL 020 7848 6813 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Fixed frozen non-paraffin mouse brain
Dear Ed, Adequate fixation is important. Formaldehyde penetrates quickly but reacts slowly with proteins. 4% formaldehyde made by depolymerizing paraformaldehyde (an insoluble high polymer) is the same as formaldehyde made by 10X dilution of formalin (a mixture of soluble low polymers). For cryoprotection the sucrose must thoroughly penetrate the fixed specimen, which must sink in the concentrated solution. Your Swiss cheese artifact is due to slow freezing. In badly frozen brains the ice crystal holes are as big as large neurons and the tissue architecture is destroyed. Unless specimens are really tiny and frozen super-fast (special methods for electron microscopy), ice crystals always form and show later as holes in the sections. You get bigger holes with bigger specimens and slower freezing. With very fast freezing, a skilled technician can get good cryostat sections even of small unfixed specimens such as muscle biopsies. These are needed for enzyme activity histochemistry methods used in diagnostic pathology and in research. Cryoprotection of fixed specimens slows the growth of ice crystals. With luck, the holes are too small to interfere with light microscope studies of sections. In neuroscience research, quite thick frozen sections of samll animals' brains have been the norm for more than 50 years. About 20 years ago I wrote a chapter that gave some quite detailed instructions and explanations, with references. (Don't do anything important just because a chapter or a review says so; check at least some of the refs!) Here is the reference. Kiernan, J. A. 2002. Freezing and fixation. Chapter 8 in Microscopy and Histology for Molecular Biologists. A User's Guide, ed. Kiernan, J. A. & Mason, I. G. pp. 103-143. London: Portland Press. ISBN 1855781417. Your university's library in Urbana might have the book. It's out-of-print with its publisher. There are used copies on the web for much less than the original price. John Kiernan Emeritus neuroanatomist and histochemist London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html = = = From: Roy, Edward J via Histonet Sent: 04 July 2020 20:08 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Fixed frozen non-paraffin mouse brain As a research lab, we sometimes would like to use paraformaldehyde-fixed but non-paraffin embedded tissues; paraffin embedding alters antigens and necessitates antigen retrieval, but simple fixation does not. We have done the traditional 30% sucrose before OCT and freezing, with cryostat sectioning, but results are inconsistent, sometimes producing Swiss-cheese brains. Does anybody have an alternative to 30% sucrose that is more reliable? I didn’t see anything in the Archives after a search for “30% sucrose”. Thanks very much, Ed Roy Edward J. Roy, PhD Professor Emeritus Department of Molecular and Integrative Physiology University of Illinois at Urbana-Champaign Urbana, IL 61801 217 333-3375 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Brain and spinal cord
10% neutral buffered formaldehyde is good, but a whole human brain needs to be immersed for at least 2 weeks, suspended by a string under the basilar artery to prevent squashing against the bottom of the container. The soft tissue has to be properly hardened before the brain can be sliced and small pieces selected for embedding and sectioning. Fixatives other than formaldehyde can be OK for brains of small animals used in research, but for human neuropathology you need formaldehyde to get the expected results with regularly used stains and with immunohistochemistry. See Adams & Murray 1982 Atlas of post-mortem techniques in neuropathology. ISBN 9780521105682<https://www.abebooks.com/products/isbn/9780521105682?cm_sp=bdp-_-ISBN13-_-PLP>. Secondhand copies cost about US $30. Every lab should have plenty of books. They cost a lot less than making mistakes. John Kiernan Anatomy & Cell Biology University of Western Ontario London, Canada = = = From: Yahoo via Histonet Sent: 15 May 2020 15:42 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Brain and spinal cord Hi All! I’m looking for some suggestions please on fixation for brain tissue and spinal cord submissions from necropsies. We are currently using 10% NBF and ask our pathologists to leave the samples overnight (but that doesn’t always happen!!). Does anyone use alcohol-based fixatives? And if so, how long? Does it affect IHC or any other staining? Do you still process with other routine biopsies (14 hour program)? Thanks! ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] On-line references
Hello, Tom. Some old classics are there for free, most notably JR Baker's "Principles of Biological Microtechnique" (1958), but almost anything more recent has to be bought. There are plenty of cheap older editions of histotechnology books on sites like AbeBooks. Check it out for the last edition of Pearse's Histochemistry! I was amazed. Even the latest editions of books in our field cost only about $100 from the publisher and most are good for several years. Compare this with the price of a few drops of an antibody or (more realistically) a staining machine in which you must only use the liquids sold by its vendor. John Kiernan = = = From: Tom Wells via Histonet Sent: 25 March 2020 14:34 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] On-line references Given that our Institute's library is closed due to the pandemic, is anyone aware of on-line versions of Histotechnology/ Histochemistry textbooks? Thanks. Tom Tom Wells BSc, MEd, MLT, ART Faculty Medical Laboratory Science School of Health Sciences SW03-3088 (604) 412-7594 BCIT ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Glycogen detection; also Spring Forward
Glycogen (MW about 1,000,000) is soluble in water but insoluble in alcohol (Merck Index 12th ed.,1996, p.766). For this reason, non-aqueous coagulant fixatives may have advantages, especially for small specimens or thin layers of cultured cells. Fixation immobilizes cytoplasmic proteins, which then entangle the big long polysacccharide molecules of glycogen, keeping them approximately in their intracellular positions. Formaldehyde penetrates specimens rapidly, but its chemical reactions with proteins, especially the cross-linking that stabilizes structure, are slow (meaning 12-48 hours). During this time, the glycogen in liver cells dissolves and is carried in solution along the direction of the fixative diffusing into the specimen. In each hepatocyte, this intracellular diffusion of glygogen is stopped by each hepatocyte's cell membrane, which has a lipid layers that are unchanged by an aqueous formalin solution. As a result, the stainable glygogen piles up in the side of each hepatocyte furthest from the surface of the specimen. This artifact is often called "polarix=zarion". With processing into paraffin, which removes lipids and coagulates any proteins not yet made insoluble by formaldehyde, glycogen is anchored into place by fixed cytoplasmic proteins, but it can still be attacked and removed by amylase/diastase/spittle. All this has been known for at least 60 years. It's in the textbooks, as Bob Richmond pointed out yesterday. (Or was it the day before?) It's now time for us all to advance our clocks by an hour, go to bed and wake up in time for Church on Sunday! John Kiernan = = = From: Bob Richmond via Histonet Sent: 07 March 2020 13:47 To: Histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Glycogen detection Galina Deyneko (where? asks: >>Does anybody have experience how fix the tissues for successful glycogen ? detection in murine and humane cardiomyocytes. I am wondering maybe the trace of methanol in 10% formalin will dissolve glycogen?? - What would be better process for paraffin embedding or use OCT embedding without fixation? Of course I prefer FFPE blocks, since OCT blocks give bad morphology.<< Ordinary neutral buffered formalin and paraffin embedding should be adequate. R.D. Lillie (3rd ed.) notes good results with Carnoy's fixative, alcoholic formalin, and acetic alcoholic formalin also. The traditional stain for glycogen is periodic acid Schiff (PAS). You verify the presence of glycogen by doing the stain with and without amylase ("diastase") predigestion. (A crude but adequate source of amylase is to just spit on the slide.) Bob Richmond Samurai Pathologist Maryville, Tennessee ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] How to Reduce Tissue Autofluorescence
There's a very brief article (downloadable PDF) from 2002 about suppressing autofluorescence, with a few references, at https://www.researchgate.net/publication/10971457_Suppressing_autofluorescence [https://i1.rgstatic.net/publication/10971457_Suppressing_autofluorescence/links/00463520275ec6a5f100/largepreview.png]<https://www.researchgate.net/publication/10971457_Suppressing_autofluorescence> (PDF) Suppressing autofluorescence - ResearchGate<https://www.researchgate.net/publication/10971457_Suppressing_autofluorescence> A 'read' is counted each time someone views a publication summary (such as the title, abstract, and list of authors), clicks on a figure, or views or downloads the full-text. www.researchgate.net This PDF file also has short peer-reviewed histotechnical tips on 5 other topics. Fun for all there. For something more recent on autofluorescence, try: Davis AS, Richter A, Becker S, Moyer JE, Sandouk A, Skinner J, Taubenberger JK (2014) Characterizing and diminishing autofluorescence in formalin-fixed paraffin-embedded human respiratory tissue. J. Histochem. Cytochem. 62: 405-423. They compared 9 procedures and favoured 3: sodium borohydride, Sudan black B and another dye called eriochrome black T. The last-named dye is CI 14645, Mordant black 11, a monoazo dye very briefly described on page 108 in Conn's 9th edn (1977) with the preferred name chromogen black ETOO; it's not in Conn's 10th edn (2002). Sodium borohydride reacts with aldehydes and probably reduces fixative-induced fluorescence of proteins and the native fluorescence of lipofuscins. The black dyes may work by absorbing more weakly emitted light. Sudan black B can stain lipofuscin black even in in paraffin sections. Using “home brew” reagents is always the best way to go, because you can avoid buying simple products sold at high prices with fancy names. Avoid trying anyone's unexplained "working protocol" because annotated pieces of paper get passed along in labs and can induce well educated people to do things that are obviously wrong. It is necessary to know the reason for each step in a lab procedure. You identify as a research assistant, so you must have a boss. Probably your boss should be online along with you, asking histonetters for advice about reducing autofluorescence. That's quite enough from me, on 9 Feb 2020. John Kiernan (Anatomy, UWO, London, Canada) = = = From: Arun Jyothi S.P via Histonet Sent: 06 February 2020 10:18 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] How to Reduce Tissue Autofluorescence Dear All, Kindly share your working protocol using “home brew” reagents to reduce tissue auto-fluorescence. Thank you Arun Jyothi S.P. Research Assistant Cancer Research RGCB Trivandrum ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Question about gelatin embedding
Gelatin embedding is easy. You infiltrate the specimen and then fix it again in formaldehyde to cross-link the gelatin molecules and make the whole mass isoluble in water. You can than cut frozen sections of any kind: cryostat, or with an old-fashioned freezing microtome collecting thawed sections from the knife with a brush. The formaldehyde-fixed gelatin holds everything together. With a Nissl stain it remains inconspicuous. If you do an H the gelatin will stain red. John Kiernan = = = From: Alonso Martínez Canabal via Histonet Sent: 17 January 2020 16:29 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Question about gelatin embedding Hello, I am here again. I am wondering if someone has good experience embedding in gelatin-albumin for cryostat or vibratome sectioning. Specifically we use brain tissue and is common in free floating techniques non-attached parts of the same section float around and later that generates all sorts of problems. Thank you very much. -- Dr. Alonso Martínez Canabal PhD Profesor Asociado "C" Departamento de Biología Celular, Facultad de Ciencias, UNAM Investigador Nacional "I" 56224833 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Pap stains
As a student in the 1960s I was told by my elders and betters to filter all staining solutions before using. It was very good advice. Filtration does not prolong the life of a stain, but it does remove crud, a material that can be composed of polymerized dyes and bits of previously stained cells or tissues. Fifty years on, I can identify a few staining solutions that never deteriorate into insoluble materials that fall out of solution even after 10+ years. Even for these, filtration before use is good laboratory practice. Not filtering any stain may therefore be bad laboratory practice. Happy New Year, John Kiernan = = = From: Haas, Elizabeth via Histonet Sent: 31 December 2019 11:46 To: S hay Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Pap stains I believe filtering stains daily is a CAP requirement Sent from my iPhone > On Dec 31, 2019, at 9:23 AM, S hay via Histonet > wrote: > > 1. Does everyone filter their pap stains daily? > 2. Are you chaining all other reagents daily? > > Thanks in advance. > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu >http://lists.utsouthwestern.edu/mailman/listinfo/histonet LEGAL DISCLAIMER: This message and all attachments may be confidential or protected by privilege. If you are not the intended recipient you are hereby notified that any disclosure, copying, distribution, or use of the information contained in or attached to this message is strictly prohibited. Please notify the sender of the delivery error by replying to this message and then delete it from your system. Thank you. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Diastase
Why buy? Just think of squeezing a chunk of lemon over your helping of haddock and drool into a small beaker. Remove bubbles by wiping the surface of the collected liquid with the edge of a piece of filter paper. Incubate for 30 minutes at 37C, just the same as for 0.1% malt diastase. Saliva also contains RNase, but this doesn't matter because RNA is not stained by PAS or by other methods for staining glycogen. For a cheap source of RNase you can heat the collected saliva (80C for 10 minutes) to inactivate diastase and any other enzymes. Cool and centrifuge. Use the supernatant at 37C for 1 hour. Some references. Bradbury S (1956) Human saliva as a convenient source of ribonuclease. Quart. J. Microsc. Sci. 97: 323-327. (Free PDF available at https://jcs.biologists.org/content/s3-97/39/323.short). Brown, GG (1978) An Introduction to Histotechnology. New York: Appleton-Century-Crofts. pp. 302 (diastase) & 292 (ribonuclease). Drury RAB, Wallington EA (1967) Carleton's Histological Technique. 4th ed. Oxford University Press, Oxford. pp.163-164 (ribonuclease) & 208 (diastase). John Kiernan Dept of Anatomy & Cell Biology University of Western Ontario, London, Canada = = = From: Paula via Histonet Sent: 26 November 2019 12:16 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Diastase Hello, We've been using STATLabs Diastase for our PAS with diastase digestion but they have a backorder until January. Can anyone recommend an alternative from other vendors? Thank you in advance, Paula ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Metals
Timm's sulphide-silver method is very sensitive, and modifications (mostly by Danscher) even more so. Sulphide-silver methods detect only those metals that have insoluble sulphides (copper and zinc but not aluminium, in Jeanine's list). It is necessary to fix in a special solution containing hydrogen sulphide - stink and also serious safety precautions! Tony's mention of Mallory & Parker's fresh hematoxylin stain prompted me to look it up. The 1939 paper is a free PDF download (Google Scholar: Mallory Parker Hematoxylin Stain Metals). Mallory FB, Parker F (1939) Fixing and staining methods for lead and copper in tissues. Am. J. Pathol. 15: 517-522 and Plates 83-85. The authors noted the importance of fixation (neutral formalin was OK for copper but no good for lead, which needed 95% or 100% alcohol). Like F.B. Mallory's other papers about staining methods, it's rather vague on technical details and has no references. The late Ralph D. Lillie reported more thorough investigations of staining for metals with haematoxylin in his classic book Histopathologic Technic and Practical Histochemistry (4th and last edn 1976, ISBN 0070378622), giving colours of the complexes with 30 metals introduced into tissues. I have tried Lillie's method on paraffin sections of rat tissues containing a few of these, and it works. ISBN 9781907904325 (p.333-334) may be more accessible than Lillie's book, which has become an expensive classic. For the more specific stains mentioned in Tony's message you need to do some critical reading. The best place to start may be Frieda Carson's Histotechnology textbook. ISBN 978-0891896401. Enough about metals for now! John Kiernan London, Canada = = = From: Tony Henwood (SCHN) via Histonet Sent: 30 September 2019 07:22 To: Sanders, Jeanine (CDC/DDID/NCEZID/DHCPP) Cc: Histonet Subject: Re: [Histonet] Metals Two good screening stains are Mallory and Parker’s Fresh Hematoxylin Stain for Metals and Timm’s Silver Sulphide Method for Metals. Malloy's results: Aluminium Blue-black Copper Greenish-blue IronBlue-black LeadBlue ZincBlue For more specific staining: Aluminon Stain for Aluminium Hydroxide Walton’s Stain for Aluminium (Phloxine binds the aluminium) Bedrick et al (1986) method for Zinc Rubeanic Acid Technique for Copper Rhodanine Technique for Copper These methods are quite sensitive but there are some specificity issues. I can provide further details and references if required. Here are some: Ohtsuki, Y., Yamaguchi, T., Sonobe, H., Takahashi, K., Hayashi, K., Takenaka, A., ... & Terao, N. (1989). Stain Technology: A Simplified Aluminum Stain in Paraffin Sections of Bone from Hemodialysis Patients. Stain technology, 64(2), 55-59. Walton, J. R., Diamond, T. H., Kumar, S., & Murrell, G. A. C. (2007). A sensitive stain for aluminum in undecalcified cancellous bone. Journal of inorganic biochemistry, 101(9), 1285-1290. Bedrick, A. E., Ramaswamy, G., & Tchertkoff, V. (1986). Histochemical determination of copper, zinc, and iron in some benign and malignant tissues. American journal of clinical pathology, 86(5), 637-640. 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 From: Sanders, Jeanine (CDC/DDID/NCEZID/DHCPP) via Histonet Sent: Monday, 30 September 2019 20:49 To: 'histonet@lists.utsouthwestern.edu' Subject: [Histonet] Metals Morning all! I need some advice re: protocols to demonstrate metals in FFPE tissues. Metals such as copper, aluminum and zinc. Thanks much! Jeanine Sanders, BS, HT(ASCP), QIHC(ASCP) Centers for Diseases Control and Prevention 1600 Clifton Road NE MS H18-SB Bldg. 18, Rm SB-114 Atlanta, GA 30329 404-639-3590 ___ Histonet mailing list Histonet@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@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Permanent mountant for Oil Red O
Fructose syrup gives a hard set, but it's a bit acidic (I don't know why) and is therefore incompatible with simple basic dyes, but it's OK for Oil red O and the Sudans. Fructose (also called laevulose or levulose) 15 g Distilled water 5 ml Leave at ~60C (in paraffin oven) for 2 or 3 days for all the fructose to dissolve. Don't let the water evaporate! The transparent syrup keeps well for 2-3 months at room temp. With long storage fructose crystallizes under the coverslip; this can be retarded by sealing the adges with a resinous mounting medium such as DPX. (You can buy fructose powder for cooking - about $3 a pound on the the internet. Cf $25-80 from chemical supply houses.) Another good one is polyvinylpyrrolidone: PVP (m.w. 10,000) 25g Water (or a phosphate buffer pH 7.4) 25ml When dissolved (several hours, magnetic stirring), add 1ml glycerol and a small crystal of thymol. Keeps for up to 3 years. Discard if it becomes cloudy. Use the buffered variety if the preparation has been stained or counterstained with a basic dye like toluidine blue or neutral red. This is less viscous than fructose syrup, and also has a lower refractive index, but with evaporation at the edges of the coverslip it gradually (weeks) becomes harder, and its refractive index increases almost to that of a resinous mountant. Probably you won't want to wait before shipping the slides elsewhere. PVP costs more than fructose (about $150 per pound for PVP10). This short article about aqueous mounting media is rather old (1997) but probably still OK: http://publish.uwo.ca/~jkiernan/aqmount.htm John Kiernan London, Canada = = = From: Hagon, Christopher (Health) via Histonet Sent: 15 September 2019 22:43 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Permanent mountant for Oil Red O UNCLASSIFIED Hello Histonetters, We do Oil Red O stains on frozen section post mortem tissue, and need to find a permanent aqueous mounting medium. We used to use the Thermo-Fisher Perm-mount, but can't seem to get it anymore. As they are looking for fat deposits, we can't use any solvent based solutions, and the only aqueous ones we've found aren't permanent. We have to send the slides off site after staining, so the chances of the coverslip moving in transport is fairly high. Does anyone else have this issue and what did you end up using? Thanks in advance, Chris Hagon | Senior Scientist, Anatomical Pathology ACT Pathology | health.act.gov.au<https://health.act.gov.au/> Phone (02) 5124 2874 --- This email, and any attachments, may be confidential and also privileged. If you are not the intended recipient, please notify the sender and delete all copies of this transmission along with any attachments immediately. You should not copy or use it for any purpose, nor disclose its contents to any other person. --- ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Mallory-Azan stain
Mallory's (easy) and AZAN staining (difficult) are different methods! Frank B. Mallory's trichrome stain (Journal of Medical Research 13: 113-136, 1905) is the earliest and one of the simplest of its kind: acid fuchsine followed by a solution containing orange G, aniline blue and phosphotungstic acid (PTA). Martin Heidenhain's trichrome is usually called AZAN (from Azokarmin and Anilinblau, the German names of two of the dyes used. I've not read his original 1916 publiication, but a very thorough account was given by Manfred Gabe in his 1976 Histological Techniques book (ISBN 3540901620), pp. 219-223. I used this quite a bit in the 1990s mostly on paraffin sections of Bouin-fixed decalcified rats' heads. It is a 15-step procedure taking >2 hours and it includes two critical differentiations requiring careful microscopic control. Instructions based on my experiences can be found in Histological and Histochemical Methods (5th ed., 2015, pp.198-200). AZAN gives a wider range of colours than Mallory's or Masson's trichrome or the various one-step trichromes (Cason, Gomori, Gabe). The related Romeis "cresazan" procedure was used to identify at least 6 anterior pituitary cell-types until the 1950s when more rational histochemically based stains were introduced by Adams, Herlant, Pearse and others. Nowadays, immunostainng accurately shows the hormones in pituitary cells, but much more expensively. All trichromes give poor results after simple fixation in neutral formaldehyde. Bouin or (better) a mercuric chloride-containing fixative is needed. Zinc-formalin is probably also OK. (I haven't tried it myself for this purpose). If material fixed in NBF must be used, immerse hydrated paraffin sections in saturated aqueous picric acid either for 2h at 56-60C or overnight at room temperature, then wash well in water before staining. (Bouin's fluid is often used, but its ingredients other than picric acid are unnecessary.) Experiments are needed to learn the mechanism of this "rescue" of staining properties of sections formaldehyde-fixed tissue, which is sometimes wrongly called "mordanting". My guess is that it's comparable to antigen retrieval. It has been claimed that citrate buffer is just as good, though the photos are unconvincing (J. Histotechnol. 26, 133). It should be possible to identify Purkinje fibres with any staining method that shows nuclei and myofibrils, such as H or a trichrome method simpler than AZAN. A glycogen stain such as PAS might show this substance in the otherwise pale areas around the central nuclei of Purkinje fibres. I suggest persuading your researcher to let you try something simpler before attempting Heidenhain's AZAN. Wheater's Functional Histology has a nice photomicrograph of a section stained with H and for endocardial elastin (looks like orcein). Enough rambling! John Kiernan Anatomy & Cell Biology University of Western Ontario London, Canada = = = From: Betsy Molinari via Histonet Sent: 09 September 2019 10:53 To: 'Histonet@lists.utsouthwestern.edu' Subject: [Histonet] Mallory-Azan stain Hi histonetters, I have a researcher that wants to stain Purkinje fibers and has requested a Mallory-Azan stain. I have no experience with this stain. I have looked online for information but am reaching out to you for personal advice. Thanks. Betsy Molinari HT,ASCP Texas Heart Institute 6770 Bertner Ave. Houston, TX 77030 832-355-6524 (lab) 832-355-6812 (fax) Betsy Molinari Sr. Histology Research Technician CV Pathology Research Texas Heart Institute 6770 Bertner Avenue, MC 1-283 Houston, TX 77030 Office: 832-355-6524 | Fax: 832-355-6812 Email: bmolin...@texasheart.org texasheart.org<https://www.texasheart.org/> | facebook<https://www.facebook.com/Texas.Heart.Institute> | twitter<https://twitter.com/Texas_Heart> ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Recent issues with picro sirius red staining (entire liver section become red, no yellow background)
I've never seen the kind of staining you describe, abi jag, with "the complete section become stained as red" but I've never used the method on sections of liver. You should get red collagen and yellow hepatocytes with blue nuclei. The strongly acidic picrosirius stain, applied for an hour, always greatly weakens (differentiates) a prior nuclear stain with Weigert's or Lillie's iron-haematoxylin. It's also quite easy to lose some or all the yellow in the washing and dehydration of the stained sections. Most users of this method are interested only in collagen fibres and do not mind if the nuclear and cytoplasmic colours get lost. The iron-haematoxylin nuclear stain is often omitted. It is necessary to have the right dye. It must be sirius red F3B (= CI 35780 = Direct red 80). This is still used as a textile dye, with several suppliers and trade-names. See http://www.worlddyevariety.com/direct-dyes/direct-red-80.html#respond Direct Red 80 - worlddyevariety.com<http://www.worlddyevariety.com/direct-dyes/direct-red-80.html#respond> www.worlddyevariety.com List of Suppliers: Direct Red F3,Direct Fast Red BA,Direct Fast Red F3B. Tianjin Yadong Group . ACDI Red 8 BLN(Aakash Chemicals & Dyestuffs Inc)Alacodirect Red 2BL(Classic Dyestuffs Inc)AmbidirectRed 3BL( Thai Ambica Chemicals Co Ltd) Anadurm Red D-BA( Albion Colours Ltd) Arid Red 8 BLN ( Aashiana Dyestuffs Inc) Best Direct Supra Red F3B( Oriental Giant Dyes and Chemical Ind Corp) Other dyes with "sirius red" in their trivial names probably are not suitable if they are not the product recognized in the Colour Index as CI 35780, Direct red 80. An example of a different dye is sirius red 4B (= CI 28160 = Direct red 81), which has been prescribed for use in some staining techniques as a dye with properties similar to those of eosin Y and acid fuchsine. The Biological Stain Commission has standards for sirius red F3B as a collagen stain. Dye from a batch that meets their standards will be OK for your method. My Histonet post from which you are using this method must date from the 1990s, when I knew that picro-sirius red solutions were good for 5-6 years. (I would have written 3 years to be cautious.) With more experience with stored and newly made solutions, I feel confident in saying they keep for more than 20 years. It might get contaminated from too much iron-haematoxylin extracted from previously stained slides. I don't know what this would do. The most obvious cause of red cytoplasmic staining by picrosirus is not enough picric acid (yellow powder in the bottom of the bottle) in the staining solution. It's unfortunate that items found with HistoSearch are undated. It doesn't matter in this case, but many Histonet items become outdated after only a year or two; antibodies and automated staining are examples of fields in which you need to know the age. Keep in touch about your sirius red problem. John Kiernan John A. Kiernan MB, ChB, PhD, DSc Professor Emeritus, Anatomy & Cell Biology University of Western Ontario, London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html Also Secretary, Biological Stain Commission, Inc. https://biologicalstaincommission.org = = = From: abi jag via Histonet Sent: 09 August 2019 11:29 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Recent issues with picro sirius red staining (entire liver section become red, no yellow background) Hello Histonetters,I am writing this to seek your help regarding a very recent problem that I am currently facing with Picro Sirius red staining of lab animal (mouse and rat) liver samples. I follow the procedure that was provided by John Kiernan in the histonet archives (please see below), which was working very well. Quite recently, the complete section become stained as red. Usually, collagen in the sections get stained as red with a yellow back ground. Please note that there was no change in the procedure/reagents etc, It will be of great help if you help me in troubleshooting this issue.With my best regards,Abijag Sirius red collagen procedure | | | | Sirius red collagen procedure | | | Solution A. Picro-sirius red Sirius red F3B (C.I. 35782): 0.5 g Saturated aqueous solution of picric acid:500 ml Add a little solid picric acid to ensure saturation (This is important). (Keeps for at least 3 years and can be used many times.) Solution B. Acidified water Add 5 ml acetic acid (glacial) to 1 litre of water (tap or distilled). Procedure Fixation is not critical, The method is most frequently used on paraffin sections of objects fixed adequately (at least 24 hours but ideally 1 or 2 weeks) in a neutral buffered formaldehyde solution. 1. De-wax and hydrate paraffin sections. 2. (Optional, and not usually done) Stain nuclei with Weigert's haematoxylin (as for
Re: [Histonet] Troubleshooting Gomori's Trichrome Stain (Blue Collagen, Richard-Allen) staining
One-step trichrome methods (such as Gomori's) are OK when they work, but there's little you can do when the colours come out wrong. A simple thing to try would be a shorter time in the staining mixture, to reduce diffusion of the more slowly penetrating dye (aniline blue) into cells. Trichrome methods work better after coagulant fixation than after formaldehyde. (The Bouin pre-treament is to offset the undesirable effects of fixation in neutral formaldehyde; picric acid alone works just as well. See also Yu & Chapman 2003 J. histotechnol. 26(2): 131-134.) If you can fix your hearts in Bouin or Carnoy you will get a better result with any trichrome technique. If the one-step method still won't work, use a multi-step trichrome where you can have some control over the actions of the different components. Masson's (which has several variants) is popular; Mallory's has fewer steps. Good luck! John Kiernan Anatomy, University of Western Ontario London, Canada = = = From: abi jag via Histonet Sent: 25 July 2019 08:32 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Troubleshooting Gomori's Trichrome Stain (Blue Collagen, Richard-Allen) staining Dear histo experts,Please provide me with your valuable suggestions for the problem described below.Objective: Staining rat hearts (fibrosis) with Gomori's one step Trichrome Stain (Blue Collagen, Richard-Allen) to demonstrate the collagenProcedure: Paraffin sections of 4 micron thickness; adequately fixed in 10 % NBF, Bouin’s Fluid treatment at 56°C for 1 hour before staining. Follow the procedure exactly recommended by kit insert (please see below)Problem: Non specific diffuse bluish discoloration of cardio myocytes in the normal hearts, which looked completely odd. The staining of same section with picro sirius red came beautiful.Any insights on the potential reasons of this and ways to resolve?Thanks a lot in advance for your time and your vision to make a wealth of knowledge available in histonet. Best regards,Abi Standard Staining Protocol 1. Deparaffinize and hydrate sections to deionized water. 2. Place sections in Bouin’s Fluid at 56°C for 1 hour. 3. Rinse sections in running tap water for 3-5 minutes untilyellow color is removed. 4. Place sections in Working Weigert’s Iron Hematoxylin Stainfor 10 minutes. 5. Rinse sections in running tap water for 5-10 minutes. 6. Stain sections in Trichrome Stain for 15 minutes. 7. Place sections in 1% Acetic Acid Solution for 1 minute. 8. Rinse sections in deionized water for 30 seconds. 9. Dehydrate sections in 95% alcohol for 1 minute. 10. Dehydrate sections in two changes of anhydrous alcohol for 1minute each. 11. Clear sections in three changes of clearing reagent for 1minute each and mount. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet Histonet Info Page - UT Southwestern Medical Center<http://lists.utsouthwestern.edu/mailman/listinfo/histonet> lists.utsouthwestern.edu Histonet -- For the exchange of information pertaining to histotechnology and related fields About Histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] time for penetration of methacarn fixative
Peter Noyce, you did not say what tissue(s) you are planning to fix, or how big the specimens will be. Carnoy (1886) and methacarn (1970) were developed for animal tissues, cleverly balancing the actions of acetic acid and an alcohol in the presence of a hydrophobic solvent (chloroform) that diluted both these ingredients and also enhanced permeation by dissolving lipids. Carnoy's and other acid-alcohol fixatives are also used for plant specimens, which respond differently. If you are working with plant specimens you probably have a book by Steven E. Ruzin (1999) ISBN 0195089561. For the actions of alcohols and acetic acid in fixation of animal tissues, consult any text book of histotechnology or histochemistry published since about 1950. For selfish reasons, I recommend ISBN 9781907904325 (published in 2015) as an item to buy for your lab's bookshelf. Answers to your question are discussed in Chapter 5. With methacarn and other alcohol-acetic fixatives, no slow chemical reactions are involved (an important difference from formaldehyde-containing mixtures). Complete penetration accomplishes the fixation. The volume of fixative and procedure for subsequent processing into paraffin are very important. You need to read the paper by Puchtler et al (1970) and follow the instructions exactly. Probably you should also read Puchtler et al (1968) to use this type of non-aqueous fixative intelligently. PhD students are intelligent. I have added a couple of more recent papers that relate to methacarn. Read Puchtler or a textbook first. Not all modern investigators (especially molecular biologists) understand what the ingredients of fixative mixtures do to the different components of cells and extracellular materials. Current papers with micrographs full of ghastly artifacts abound, even in journals with very high citation indices. There are published mixtures with names like "modified methacarn" that may be OK for extracting RNA but do not have ingredients in correct proportions for minimizing distortion. Here's your list of recommended readings. - - - - - Puchtler H, Waldrop FS, Meloan SN, Terry MS, Connor HM (1970) Methacarn (methanol-Carnoy) fixation. Practical and theoretical considerations. Histochemie 21: 97-116. Puchtler H, Waldrop FS, Conner HM, Terry MS (1968) Carnoy fixation: practical and theoretical considerations. Histochemie 16: 361-371. Uneyama C, Shibutani M, Masutomi N, Takagi H, Hirose M (2002) Methacarn fixation for genomic DNA analysis in microdissected paraffin-embedded tissue specimens. J. Histochem. Cytochem. 50: 1237-1245. Buesa RJ (2008) Histology without formalin? Ann. Diagn. Path. 12: 387-396. - - - - - I wish you well with your research, and hope you will get your PhD while you are still young. John Kiernan London, Canada https://www.schulich.uwo.ca/anatomy/people/bios/emeriti/kiernan_john.html http://biostain.com = = = From: peter noyce via Histonet Sent: 03 May 2019 19:47 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] time for penetration of methacarn fixative Does any one have data for the time it takes for methacarn fixative (60% methanol, 30% chloroform, 10% glacial acetic acid) to penetrate and then fully fix tissue? PW Noyce -PhD student _ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] PAP stains
Charles, What do you mean by "dark nuclei"? Are you asking about the normal colour for the method, or about something you have not seen before that looks wrong? Also, what is "PAP stain"? If "PAP stain" means Papanicoloau, the nuclear stain is Mayer's haemalum. This is a progressive stain; you may need to shorten the time. If "PAP stain" means peroxidase-antiperoxidase (unlabelled antibody-enzyme complex for amplifying detection of HRP-tagged secondary antibodies in immunohistochemistry), nuclei will be dark (usually brown) if they contain the antigen sought by the primary antibody. Nuclear staining might also be a false-positive artifact; if so, it would be present in the simplest of the routine controls (omission of primary antibody). Dark nuclei might also be due to a nuclear counterstain that is too strong. The counterstain should be done in a contrasting colour. Usually it is a progressive haemalum (blue) to contrast with the oxidation product of DAB (brown). Is this a Papanicoloau question or an immunohistochemistry question? Does "PAP stain" have a third meaning? John Kiernan = = = From: Charles Riley via Histonet Sent: 19 March 2019 14:17 To: Histo List Subject: [Histonet] PAP stains What causes dark nuclei in the PAP stain. -- Charles Riley BS HT, HTL(ASCP)CM Histopathology Coordinator/ Mohs ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Words, phrases and names in histotechnology. A free glossary.
Hello histonetters. I'm a HistoNet old-timer, back again after a few years away. It's good to see that a few names are still around from the 1990s. Here is something new that may interest all of us. I send it as a news item; a change from the usual initial question that initiates a Histonet topic. A freely accessible online glossary of words, phrases and eponyms used in histotechnology, histochemistry and immunostaining was published by the Biological Stain Commission (BSC) at the end of December 2018. It includes about 600 entries; it is to be revised and expanded from time to time. (A minor revision was uploaded on 19th January 2019.) Notable features are extensive crosslinks between the entries, and explanations of terminology related to chemical and physical mechanisms involved in the methods. There are also definitions and explanations relating to all the stains (dyes) certified by the BSC. The BSC glossary is readable on screens of all sizes (including mobile phones), and navigation among links is extremely rapid. Check it out directly at https://biologicalstaincommission.org/bscglossary.html. Alternatively, see it in the broader context of the BSC: http://biostain.com. John Kiernan = = = ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] axolotl lymphatics
Instead of an antibody, you might consider enzyme activity histochemistry, which is much less expensive. Demonstration of 5-nucleotidase activity in the presence of levamisole detects lymphatic endothelium. Sections can also be stained for alkaline phosphatase activity in the endothelium of blood vessels. Here are a few references. Kato, S., Yasunaga, A. and Uchida, U. (1991). Enzyme-histochemical method for identification of lymphatic capillaries. Lymphology 24:125-129. Ohkuma, M. (1994). Simultaneous double staining for the blood and lymphatic capillary. Lymphology 27, Suppl:106-107. Okada, E. (1994). An improved enzyme-histochemical method for identification of lymphatic capillaries on paraffin sections. Lymphology 27, Suppl:732-735. Ji, R.C. and Kato, S. (2003). Lymphatic network and lymphangiogenesis in the gastric wall. Journal of Histochemistry and Cytochemistry 51:331-338. Needless to say, none of these relate to amphibian tissues! John Kiernan Anatomy, UWO, London, Canada = = = On 26/07/15, Jason Palmer via Histonet histonet@lists.utsouthwestern.edu wrote: Hi all, I need to find an antibody that will label lymphatic endothelial cells in the axolotl. Does anybody have any experience or ideas? I have tried a couple of our anti-mouse and anti-human Abs for podoplanin and LYVE-1 but no cross-reactivity so far. I have no experience with staining of non-mammalian tissues - maybe an anti-frog Ab would cross react? Does anyone have experience with other amphibians? Thanks, Jason -- Jason Palmer Histology Laboratory Coordinator O'Brien Institute / St Vincent's Institute 42 Fitzroy St, Fitzroy Victoria 3065 Australia tel +61 3 9288 4045 fax +61 3 9416 0926 email: jpal...@svi.edu.au ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Toluidine blue stain for MMA
Yes. Probably hundreds of Histonetters stain plastic sections. Let us all hope they don't all bombard the Histonet listserver with replies to your question. Instructions for staining plastic sections with toluidine blue are in every library that contains books with paper pages, and also (albeit with less authority) in great abundance on the Web. Try typing SEMITHIN STAIN into Google. I just did, and an excellent web site came up on top of the heap. John Kiernan = = = On 27/07/15, Kai Hong via Histonet histonet@lists.utsouthwestern.edu wrote: Hi, is there anyone have an experience with MMA toluidine staining? Im using T7200, T9100, Osteo-bed resin in lab now. Thanks, Kai Research Histotechnologist ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] National Academy of Sciences Confirms That Formaldehyde Can Cause Cancer in a Finding That Has Implications for Anatomic Pathology and Histology Laboratories
Dear Banjo, A reference to the article would be helpful; there must be more to it than one sentence! Formaldehyde has been known for decades to be hazardous, and there are safety regulations in places where it is used. Plenty of old-timers are still alive and well after woking with formaldehyde in the days when there were few or no regulations. I'm one of them. From about 1895 until about 1995 (and perhaps still, in some universities), every medical student spent most of the working day for at least a year with his or her nose and bare hands in a cadaver that had been embalmed in a cocktail containing phenol and formaldehyde. The predominant smell was the phenol, except when dissecting brains, which were fixed and stored in 4% formaldehyde. About 35 years ago, the American Association of Anatomists investigated effects of exposure to embalming chemicals on teachers of anatomy, who are in the dissecting room year after year. The only significant finding was eczema on the hands of some people, long known to be avoidable by wearing rubber gloves. Yes, I too should be able to provide a reference, but this was in the days of paper, which gets thrown out to make room for more paper ... There might be something deep in the archives at http://www.anatomy.org/ Other chemicals used in anatomy, pathology and histology labs also have their dangers; we avoid drinking them, rubbing them into our skin and inhaling their vapours, and we do our best to observe the safety regulations when it comes to getting rid of them. There is no substitute fixative functionally identical to formaldehyde. There are other fixatives, some less hazardous, but they have different effects on staining properties etc. The late Holde Puchtler published papers urging pathologists to use non-aqueous coagulant fixatives for routine fixation of small specimens, with her Carnoy variant methacarn (methanol 60, acetic acid 10, chloroform 30) as the probable best, also good for some modern molecular methods. For this I can provide a few references: Puchtler, H., Waldrop, F.S., Meloan, S.N., Terry, M.S. and Connor, H.M. (1970). Methacarn (methanol-Carnoy) fixation. Practical and theoretical considerations. Histochemie 21:97-116. Cox, M.L., Schray, C.L., Luster, C.N., Stewart, Z.S., Korytko, P.J., Khan, K.N.M., Paulauskis, J.D. and Dunstan, R.W. (2006). Assessment of fixatives, fixation and tissue processing on morphology and RNA integrity. Experimental and Molecular Pathology 80:183-191. Buesa, R.J. (2008). Histology without formalin? Annals of Diagnostic Pathology 12:387-396. Uneyama, C., Shibutani, M., Masutomi, N., Takagi, H. and Hirose, M. (2002). Methacarn fixation for genomic DNA analysis in microdissected paraffin-embedded tissue specimens. Journal of Histochemistry and Cytochemistry 50:1237-1245. Milcheva, R., Janega, P., Celec, P., Russev, R. and Babal, P. (2013). Alcohol based fixatives provide excellent tissue morphology, protein immunoreactivity and RNA integrity in paraffin embedded tissue specimens. Brain Research Protocols 115:279-289. Greer, C.E., Peterson, S.L., Kiviat, N.B. and Manos, M.M. (1991). PCR amplification from paraffin-embedded tissues. American Journal of Clinical Pathology 95:117-124. Tissue processing is extremely simple after non-aqueous coagulant fixation, and most of the stages of a processing machine are not needed. Nuclear chromatin details are much sharper than after formaldehyde. This may not be seen as a blessing by young and middle-aged pathologists. In bygone days the routine fixatives contained mercuric chloride, which gives crisp chromatin and cytoplasmic details. The heterochromatin details probably are artifacts of fixation, but they are useful for identifying cells. John Kiernan Old neuroanatomist and histochemist UWO, London, Canada http://publish.uwo.ca/~jkiernan/ Also Secretary, Biological Stain Commission http://biostain.com = = = On 13/07/15, Adesupo, Adesuyi (Banjo) abades...@nrh-ok.com wrote: Hi, I read this article (National Academy of Sciences Confirms That Formaldehyde Can Cause Cancer in a Finding That Has Implications for Anatomic Pathology and Histology Laboratories) this morning. I wanted to know whether some of you guys out there are using Formaldehyde substitute. Best regards, Banjo Adesuyi, BMLS, HT (ASCP) HTL, QIHC, QLS Histology Supervisor Norman Regional Health System, Norman, OK 73071. Tel: 405- 307- 1145 abades...@nrh-ok.commailto:abades...@nrh-ok.com abades...@nrh-ok.com == CONFIDENTIALITY NOTICE: This e-mail communication and any attachments may contain confidential and privileged information for the use of the designated recipients named above. If you are not the intended recipient, you are hereby notified that you have received this communication in error and that any review, disclosure, dissemination, distribution, or copying
Re: [Histonet] Coverslipping mystery
DPX is a polystyrene mounting medium. In principle you can make your own from published recipes. In practice, everyone buys commercial resinous mounting media. In the 1990s we had trouble similar to what you describe. The commercial DPX was cloudy, and not because of alcohol in our xylene. The Canadian supplier acknowledged the bad DPX and urged us to buy Entellan instead. Entellan is a poly(methacrylate) plastic and is an excellent but expensive mounting medium. Another poly(methacrylate) mountant called CytoSeal was less expensive and also came in a squeeze-easy plastic bottle for delivery onto the slide or coverslip. It's now my routine resious mountant. Good DPX returned to the market in the 2000s, but in old-fashioned bottles and not easy to apply to slides or coverslips. John Kiernan = = = On 09/07/15, Adam Boanas a.boa...@epistem.co.uk wrote: Hello, We are having a problem that is developing into a big issue in our lab and I was wondering if anybody could shed any light on it. Our CV5000 coverslipper has recently started introducing microscopic air bubbles onto the slides during coverslipping. We have been told by our engineer that it is a consequence of the age and use of the motor and that sourcing another for an instrument that old (15yrs) will be v difficult. As such, we have been forced to manually coverslip using DPX and a pipette - manually applying the coverslips to the slide, thus mirroring the action of the coverslipper. This is fine at first and for the next few days the slides look great and very clean. However, after about day 4 -5 days post coverslipping, the slides develop an odd appearance down the microscope which looks like very fine `parched earth / crazy paving` all over the slide - including the section. The excess mountant around the edge of the coverslip also has a very faint, cloudy appearance wh! en this occurs. This of course renders the slide un-useable. Does anyone have a clue what this might be down to / how we can stop it? We are struggling for ideas with this one! - this occurs with fresh DPX also. Many thanks Adam Adam Boanas Senior Research Associate Epistem Ltd 48 Grafton Street Manchester, M13 9XX ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] subbing positively charged slides
Chrome-gelatin cannot be expected to stick well to a glass surface that has been modified by adding aminopropyl (or similar) groups to its silicate ions. See http://publish.uwo.ca/~jkiernan/adhesivs.htm. John Kiernan London, Canada = = = On 09/07/15, Travers, Susan traver...@osu.edu wrote: Does anyone know if it's ok to sub positively charged slides with chrome alum-gelatin. I seem to remember an incident where this did not work- but maybe it's just superstition... it seems as if the chrome alum might be positively charged itself and thus repelled from the surface of the slide... Susan Travers Professor, Biosciences Division of Biosciences, College of Dentistry The Ohio State University 305 West 12th Avenue Columbus, Ohio 43210 traver...@osu.edumailto:traver...@osu.edu traver...@osu.edu (614)361-0800 (cell) (614)292-6366 (work) ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Alizarin Red for rodent Feti
Red bone and blue cartilage: Webb, G.N. and Byrd, R.A. (1994). Simultaneous differential staining of cartilage and bone in rodent fetuses: an alcian blue and alizarin red S procedure without glacial acetic acid. Biotechnic Histochemistry 69:181-185. Redfern, B.G., Wise, L.D. and Spence, S. (2007). An alternative alcian blue dye variant for the evaluation of fetal cartilage. Birth Defects Research (Part B) 80:171-176. Yamazaki, Y., Yuguchi, M., Kubota, S. and Isokawa, K. (2011). Whole-mount bone and cartilage staining of chick embryos with minimal decalcification. Biotechnic Histochemistry 86:351-358. Trueman, D. and Stewart, J. (2014). An automated technique for double staining mouse fetal and neonatal skeletal specimens to differentiate bone and cartilage. Biotechnic Histochemistry 89:315-319. John Kiernan London, Canada P.S. Fetus is a 4th declension Latin noun, so its plural in English is fetuses, not feti! = = = On 06/07/15, Wilson, Carol carol.wil...@ricerca.com wrote: Good Day All, Can someone pass along methodology for processing rodent Feti for Alizarin Red stain and exam? I would appreciate any input/ direction given as this is a foreign topic to me. Thanks, Carol Carol Wilson, HT(ASCP) Associate Scientist III Team Leader/Histopathology Ricerca Biosciences, LLC ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Alcian blue (was Suggestions for staining ground substances ...)
Dear Histonetters, I recently replied to a question about staining extracellular ground substance with alcian blue. Here is some more information. With the alcian blue-PAS sequence it is necessary to have a good alcian blue dye - the same as or functionally equivalent to the original alcian blue 8G. For the PAS-alcian blue sequence, other alcian blues may also be OK. Alcian blue from a batch certified (or rechecked) by the Biological Stain Commission (BSC) 5 or fewer years ago should be good for either procedure. Of the dyes commonly used to make up staining solutions for microscopy, alcian blue is the only only one that sometimes deteriorates, unpredictably and often quite quickly, as a dry powder in the bottle. Some alcian blue powders from the 1950s are still OK, but some very much younger ones are no good 5+ years after passing the BSC's analytical and staining tests. John Kiernan London, Canada http://biostain.com = = = ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Suggestions for staining ground substances in the Heart
Movat's pentachrome is unnecessarily complicated when you want to look only at one component of a tissue. Most ground substance (extracellular material between cells, collagen fibres etc) is weakly stainable with PAS, alcian blue pH2.5 or both. It's usual to stain with alcian blue first (Mowry's method). If you do the PAS first, the PAS-positive materials will also stain with alcian blue and be darker. For the reasons, see: Johannes, M.-L. and Klessen, C. (1984). Alcian blue/PAS or PAS/alcian blue? Remarks on a classical technique used in carbohydrate histochemistry. Histochemistry 80:129-132. Yamabayashi, S. (1987). Periodic acid-Schiff-alcian blue: a method for the differential staining of glycoproteins. Histochemical Journal 19:565-571. Reid, P.E. and Owen, D.A. (1988). Some comments on the mechanism of the periodic acid-Schiff-Alcian blue method. Histochemical Journal 20:651-654. If you can predict the PAS-AB effect without further reading, you already have a good understanding of how the traditional methods of carbohydrate histochemistry work! John Kiernan London, Canada = = = On 30/06/15, Wolfe, Christina christina.wo...@bms.com wrote: Hi all, We are interested in staining ground substances in the heart. Are there other stains that will work beside the Pentachrome? We have tried the Movat's pentachrome (commercial kit) and are able to demonstrate ground substance in the bone with the alcian blue part of this stain. In our hands the goblet cells of the gut and the ground substance in the heart are devoid of staining. We have tried sections cut at 4 and 6 microns. Any thoughts/suggestions? Kristie Christina Wolfe, BSHA, MLT (ASCP), HT, QIHC Drug Safety Evaluation/Bristol-Myers Squibb Pathology Dept. 812-307-2093 This message (including any attachments) may contain confidential, proprietary, privileged and/or private information. The information is intended to be for the use of the individual or entity designated above. If you are not the intended recipient of this message, please notify the sender immediately, and delete the message and any attachments. Any disclosure, reproduction, distribution or other use of this message or any attachments by an individual or entity other than the intended recipient is prohibited. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Removing paraffin sections from glass slides
Why shouldn't the slide surfaces remain positively charged after removing the sections? Water, alcohol and xylene are all routinely used, and don't impair electrostatic forces that help to hold the sections on the slide. The positively charged surface is not a coating that can be washed or rubbed off. Joyce Weems's suggested method did not involve anything that might react chemically with the cationic (amino-hydrocarbon) groups that are covalently bound to the polysilicate structure of the glass. See http://publish.uwo.ca/~jkiernan/adhesivs.htm John Kiernan Anatomy Cell Biology University of Western Ontario London, Canada = = = On 23/06/15, Jamal j.rowa...@alborglaboratories.com wrote: Dear Colleague I just want to remind you.. After all of that procedure of removing the paraffin sections from the positive charged slides. The slides will not be positive charged anymore and not suitable for IHC. Also the cost of the detergents and the chemicals which you used for removing the tissue section is more costly than the slides. Best Regards, Jamal M. Al Rowaihi Anatomic Pathology Supervisor | Al Borg Medical Laboratories | Mobile +966 503629832| j.rowa...@alborglaboratories.com Palestine St, Al Rajhi Building, P.O. Box 52817, Jeddah 21573, KSA | Phone: +966 12 670 0099 | Fax: +966 12 676 4984 | www.alborglaboratories.com -Original Message- From: Weems, Joyce K. [mailto:joyce.we...@emoryhealthcare.org] joyce.we...@emoryhealthcare.org] Sent: Monday, June 22, 2015 5:53 PM To: 'Coffey, Anna (NIH/NCI) [C]'; histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Removing paraffin sections from glass slides This is what I would do... Soak the coverslip off in xylene Either - rehydrate back to water and just wipe the tissue off - then dip the slides in absolute and let dry.. OR - air dry out of xylene and wipe off with a wet gauze - then do the alcohol dip Whichever worked best. Just my 2 cents.. Happy Monday!! Joyce Weems Pathology Manager 678-843-7376 Phone 678-843-7831 Fax joyce.we...@emoryhealthcare.org www.saintjosephsatlanta.org 5665 Peachtree Dunwoody Road Atlanta, GA 30342 This e-mail, including any attachments is the property of Saint Joseph's Hospital and is intended for the sole use of the intended recipient(s). It may contain information that is privileged and confidential. Any unauthorized review, use, disclosure, or distribution is prohibited. If you are not the intended recipient, please delete this message, and reply to the sender regarding the error in a separate email. -Original Message- From: Coffey, Anna (NIH/NCI) [C] [mailto:anna.cof...@nih.gov] anna.cof...@nih.gov] Sent: Monday, June 22, 2015 10:25 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Removing paraffin sections from glass slides Hello, I'm wondering if anyone has experience removing dried unstained paraffin sections from charged glass slides. I don't need to preserve the sections, just want to reuse the slides. Thanks! Anna Anna Coffey, MS, HTL(ASCP)CM Histotechnologist Center for Advanced Preclinical Research Frederick National Laboratory for Cancer Research Leidos Biomedical Research, Inc. Bld 539, 224 Frederick, Maryland 21702 anna.cof...@nih.gov 301-846-1730 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet This e-mail message (including any attachments) is for the sole use of the intended recipient(s) and may contain confidential and privileged information. If the reader of this message is not the intended recipient, you are hereby notified that any dissemination, distribution or copying of this message (including any attachments) is strictly prohibited. If you have received this message in error, please contact the sender by reply e-mail message and destroy all copies of the original message (including attachments). ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Antigen retrieval survey
Your 2 minutes would be better spent looking in an immunohistochemistry textbook. A small but excellent one is Polak, J.M. and Van Noorden, S. (1997). Introduction to Immunocytochemistry, 2nd ed. Royal Microscopical Society Microscopy Handbooks, 37. Oxford: BIOS Scientific Publications. You will find that there is an optimal technique of antigen retrieval for each antigen that has been critically studied. Some conditions (such as pH6, close to 100C for an hour) are OK for many antigens. Some require more alkaline solutions (eg pH9, more section losses!) and a few respond best to heating in a more acid (eg pH2) solution. With lower temperatures (eg 80C) longer times are generally needed. All sorts of chemicals have been included in antigen retrieval solutions, often without obvious reasons or explanations. There are published papers that compare retrieval conditions for antigens of importance in diagnostic pathology. Retrieval can sometimes be achieved without heating, as with proteolytic enzymes or 3M urea. With a survey you may find out which antigen retrieval methods are used by most of those who reply, but you will not learn anything about how to choose and use the methods, or why their discovery about 25 years ago was an important technological advance. Check out this classic paper with Web of Science, Scopus, or Google Scholar: Shi, S.-R., Key, M.E. and Kalra, K.L. (1991). Antigen retrieval in formalin-fixed, paraffin-embedded tissue: an enhancement method for immunohistochemical staining based on microwave oven heating of tissue sections. Journal of Histochemistry and Cytochemistry 39:741-748. The PDF can be downloaded for free. This paper has been cited by thousands of other publications. The titles of recent citing articles may help you find a good retrieval procedure for the antigen that you need to detect immunohistochemically. John Kiernan UWO, London, Canada = = = On 23/06/15, Craig volle...@gmail.com wrote: Hi, I am conducting a short 2 min survey for my science/business class examining current trends for antigen retrieval also known as heat induce epitope retrieval. Response will be greatly appreciated! https://www.surveymonkey.com/s/7989LKR Best, Craig Vollert Graduate Student Department of Pharmacological Pharmaceutical Sciences SR2 521B College of Pharmacy University of Houston ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] ammonium bromide in fixative
Formalin with ammonium bromide is the fixative prescribed for traditional silver and gold techniques for staining neuroglial/x cells in free-floating frozen sections. Adding ammonium bromide lowers the pH of the fixative because of a chemical reaction with formaldehyde. An Argentinian histologist/x called Lascano showed in the 1940s that any sufficiently acidified formaldehyde solution (pH 1.5) was just as good as FAB for Cajal's gold-sublimate (astrocytes) and for silver carbonate methods (oligodendrocytes, microglia). The reaction that lowers the pH is 4NH4+ + 6HCHO --- C6H12N4 + 6H2O + 4H+ (Subscripts and superscripts in this equation will not be honoured in this email!) The 4H+ lowers the pH. The other product, C6H12N4 is hexamethylenetetramine, also known as hexamine in Britain and methenamine in the USA, and used in various histochemical staining methods, notably Grocott's method for fungal cell-walls in sections of animal (including human) tissues. The bromide (Br-) ion of NH4Br is irrelevant, in the FAB fixative and also in Globus's pretreatments (dilute ammonia followed by dilute hydrobromic acid). Globus's bromuration treatment was applied to frozen sections of pieces of CNS that had been fixed in non-acidified formalin. The references for Lascano's work are: Lascano, E.F. (1946a). Influencia del pH en la impregnacion argentica del tejido nervioso. Archivos de la Sociedad Argentina de Anatomia Normal y Patologica 8:105-114. Lascano, E.F. (1946b). Importancia del pH en la fijacion del tejido nervioso. Creacion artificial de fijadores/x tipo formol-bromuro y formol-nitrato de urano de Cajal. Archivos de la Sociedad Argentina de Anatomia Normal y Patologica 8:185-194. Lascano, E.F. (1946c). Influencia del pH del fijador en la coloracion argentica del tejido nervioso. Archivos de la Sociedad Argentina de Anatomia Normal y Patologica 8:272-276. Not everyone agreed with Lascano! Polak, M. (1948). Sobre la importancia del bromuro de amonio de la solucion fijadora de Cajal en la impregnacion argentica del tejido nervioso. Archivos de la Sociedad Argentina de Anatomia Normal y Patologica 10:224-234. Do you really need this information to pass your HTL qualifying exam? It isn't easily found in books or with Google Scholar. I came across these papers quite by chance when looking over some old journals that UWO's library had set aside for throwing out (Gasp!), about 1980. Yes, they had hit a new low! Their services have, however, been exceptionally good for the last 10-15 years. Does anyone still use either Cajal's method for astrocytes or traditional silver carbonate methods to stain oligodendrocytes and microglia? They are difficult for several reasons, take up time, and require an old-fashioned freezing microtome to cut and collect the rather thick sections that are needed. Reliable antibodies for immunostaining glial cell-types have been available for many years, and they work on any kind of section. You need some thickness to appreciate the 3D shapes of astrocytes and oligodendrocytes, however they are stained. There's plenty of histo-history in the traditional neuroglia stains, which defined the cell-types for identification by electron microscopy and immunohistochemistry. John Kiernan Anatomy Cell Biology University of Western Ontario London, Canada = = = On 11/06/15, Walters, Katherine S katherine-walt...@uiowa.edu wrote: Hi all, I am studying to take the HTL certification test and ran across a reference to Formalin Ammonium Bromide. I see that it is very good for central nervous tissue fixation, it must be made fresh and that its pH is 1.5. Does anyone happen to know the reason for ammonium bromide in this fixative? I have been looking online and this has not been explained. Also, has anyone taken this test lately? I am curious as to how much old techniques, such as mercuric fixatives will be included? Thank you, Katherine S Walters Histology Director Central Microscopy Research Facility University of Iowa 76 Eckstein Medical Research Building 431 Newton Road Iowa City, Iowa 52242 319-335-8142 Facility Website: http://cmrf.research.uiowa.edu/ Notice: This UI Health Care e-mail (including attachments) is covered by the Electronic Communications Privacy Act, 18 U.S.C. 2510-2521, is confidential and may be legally privileged. If you are not the intended recipient, you are hereby notified that any retention, dissemination, distribution, or copying of this communication is strictly prohibited. Please reply to the sender that you have received the message in error, then delete it. Thank you. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list
Re: [Histonet] General Genetics (lecture and lab textbooks)
Dear Jorge A. Santiago-Blay, How did you learn enough about General Genetics for science majors to be hired to teach the subject? Do you not own at least one textbook in the field? How did you qualify to teach the subject? With the internet (Google) it will be easy to find the most recent edition of the text from which you studied General Genetics. You could buy the book or ask the publisher for a free copy. Publishers happily provide free textbooks to instructors, hoping that every student will buy a copy of the adopted textbook. Why ask me? My fields of interest, before I retired, were Neuroanatomy and Histochemistry, not Genetics. Does your institution have a way to warn students that an instructor does not know a suitable textbook for his course? Your students may need such a warning! J. A. Kiernan Anatomy Cell Biology University of Western Ontario London, Canada = = = On 08/06/15, Jorge A. Santiago-Blay blayjo...@gmail.com wrote: General Genetics (lecture and lab textbooks) Dear Histonet-Listers: I wish to know if you have recommendations of *excellent* textbooks (lecture and labs) for General Genetics for science majors. Although I just finished teaching an upper division Genetic Analysis course and I am well familiarized with what is available, I am not as familiarized with la creme de la creme for General Genetics. Ideally, I want a textbook that is (list is not comprehensive): 1. As inexpensive as possible (available online, softbound, etc.). 2. Easy to read, understand. and profusely illustrated. 3. Does not assume more than intro. biology (for majors) knowledge. 4. Emphasizes principles of genetics, model organisms, critical thinking, experimental design, ecological and conservation genetics, etc. 5. Has a generous companion packages (incl. PowerPoints, online quiz cartridges - not the ones that reside with the publishers but the ones than reside on campus only, etc.). For anything online, technology is as close to 100% reliable and intuitive as possible. 6. Does not shy away from math and stats (because I do not shy away from those either). 7. Is not divorced of the historical context of genetics. 8. Highlights the importance of genetics in modern life. 9. For labs, all of the above and not just the usual genetics exercises. I want a book that invites exploration and engages all of us in the field. Ideally, with a manual for s/he that may be helping with setting up the lab. If you have any feedback, please kindly send it directly to me, blayjo...@gmail.com With gratefulness, Jorge P.S Apologies for potential duplicate emails. Jorge A. Santiago-Blay, PhD blaypublishers.comblay.cfm http://paleobiology.si.edu/staff/individuals/santiagoblay.cfm ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] tissue fixation-formaldehyde concentrations which is best.
Dear Peter, Some of the information you mention as anecdotal is wrong. Formaldehyde and paraformaldehyde are well documented in original peer-reviewed papers and in all textbooks in the fields of histotechnology and histochemistry. Your anecdote about high concentrations of formaldehyde quickly form a 'shell' in the tissue and will stop good penetration and fixation to the deeper tissues has no basis in published work. Paraformaldehyde is an insoluble polymer, not non polymerized formaldehyde. There is no such thing as 4% paraformaldehyde! It is a sad fact that many labs do not contain even one book about histotechnology. Nearly all books in the field (and there are many) have plenty of references to review articles and original literature about the techniques. There are also several websites that provide links to useful papers. Check out some of the useful links on the Biological Stain Commission's site: http://biologicalstaincommission.org/useful-links/ As a graduate student, you need to work from primary sources or reliable secondary sources. When you defend your thesis, you won't want to justify your fixation or staining method by saying I got the method by asking on an internet listserver. John Kiernan Professor Emeritus Anatomy Cell Biology University of Western Ontario London,Canada = = = On 05/06/15, Peter Noyce pwno...@gmail.com wrote: Formaldehyde 37% (commonly called 100% formalin) compared to 4% ( commonly known as 10% neutral buffered formalin)-in theory the 37% should fix quicker and better BUT anecdotally it is said that high concentrations of formaldehyde quickly form a shell in the tissue and will stop good penetration and fixation to the deeper tissues AND over the years it has been said anecdotally that 4% concentration is the quickest and most complete for all sample (mammal and plant) fixation and preservation-are these true. Please do discuss the methanol or buffers that is in the formaldehyde, or discuss paraformaldehyde (which is non polymerized formaldehyde with no methanol, in water). Regards Peter Noyce PhD student. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Crystallized formalin
The formaldehyde in formalin is present as low polymers. The depolymerize when the liquid is diluted. With storage, especially at lower temperatures, the small polymers join together anf form large insoluble polymers, known as paraformaldehyde. This reduces the effective concentration of formaldehyde in the fixative solution. (At a guess, it might be down from 4% to 3%. ). This probably doesn't matter. Read about formalin and formaldehyde at Microscopy Today 08-01: 8-12 (2000). This is a free download from Microscopy Today's web site, and it includes references to more significant publications. Paraformaldehyde is easily depolymerized. Increase the pH to about 8 with a bead or two of sodium hydroxide, and heat to 60-65C. The paraformaldehyde depolymerizes (dissolves) and you go ahead with dilution and buffering to make a 4% NBF. This information is in every histotechnical textbook published since about 1965. Your description of crystals in old formalin puzzles me. The paraformaldehyde usually settles out as as a finely granular white powder. John Kiernan London, Canada = = = On 26/05/15, Julio Benavides j.benavi...@eae.csic.es wrote: Hi there, we have a bottle of formaline 40% with crystals at the bottom. Can we used it? is there any way to re-suspend the crystals? Better dispose of it? and, BTW, do you know why it crystallized? by its side there is another bottle, same batch, with no crystals. thanks a lot your comments Regards Julio ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Donating Old Microtome
Why get rid of microtomes that still work well? Please explain. John Kiernan London, Canada = = = On 21/05/15, Liette Tougas ltou...@dawsoncollege.qc.ca wrote: Hi Heather, I do not know where you are located but we have 5-6 good old AO microtomes that still work very well that we are willing to donate. They are in excellent condition as they were extremely well maintained over the years and, as a Biomed Lab technology teaching department, they have been used only twice a week for 3 weeks each fall semester since they were purchased, some 30 years ago! They each have a disposable blade holder (high profile) and block clamp (vs original screwing block holder). We easily cut at 4 microns on them. The offer is opened to anyone who can pick them up or pay for shipment. Liette Tougas, RT, B.Sc., M.Sc. Biomedical Laboratory Technology Department Dawson College, Montréal, Qc, Canada 514-931-8731, ext 1519 From: Heather Marlatt [hmarlat...@gmail.com] Sent: May 21, 2015 3:49 PM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Donating Old Microtome I'm posting on behalf of a local researcher. They have been using an older microtome that was previously donated by a hospital. It broke down recently and they cant afford to replace it right now. It is used by the students to cut their own sections so they just need something that can section at 4 microns. They are happy to pay the freight if anyone out there has an old working microtome that they are willing to donate. Thank you!!! Heather ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] optic chiasmata
Slow freezing results in formation of ice crystals. In brain tissue (fresh or formaldehyde-fixed) these destroy the architecture, which is replaced by a sponge-like texture of approximately cell-sized holes. Cryoprotection of fixed tissue with 30% sucrose ameliorates this, but freezing at -4C or -20C is asking too much of the cryoprotective action. You need a surface at or below -80C. A cryostat chuck standing in a slush of solid CO2 and acetone is OK. Isopentane in a small metal can standing in liquid nitrogen is better. Test your technique first on an unimportant piece of white matter about the same thickness as the optic chiasma, and cut test sections through different levels in the specimen, because some parts will freeze more slowly than others. John A. Kiernan Anatomy Cell Biology University of Western Ontario London, Canada = = = On 10/05/15, Salomao Segal sseg...@slu.edu wrote: I intend to use a cryostat to cut 70 - 100 micron thick sections of human optic chiasmata. Tissue is cryoprotected with 30% sucrose solution. My question relates to the freezing process per se. Would it be enough to place the tissue in a -4 freezer to harden and then transfer to the cryostat chamber at say -20 wait a while and cut? Or is it necessary to introduce an intermediary step for freezing? Thanks SS *Solomon Segal, M.D.* *Associate Professor of Anatomy in SurgeryCenter for Anatomical Science and Education (CASE)Department of SurgerySchool of MedicineSaint Louis University* *1402 South Grand Blvd.* *Schwitalla Hall - 3rd Floor - M310* *Saint Louis, MO, 63104office: 314 977 8023laboratory: 314 977 8080* *CASE: 314 977 8027FAX: 314 977 5127e-mail*: sseg...@slu.edu http://medschool.slu.edu/anatomy/ http://slu.academia.edu/SolomonSegal https://sites.google.com/a/slu.edu/segal-laboratory/ https://sites.google.com/a/slu.edu/dr-segal-s-clinical-anatomy-website/ ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Bleach as Cleaning Agent or Decontamination-
You need to be clear about what the concentration means. Household bleach (such as Javex or Clorox) is a 5% aqueous solution of sodium hypochlorite (NaClO). Sometimes it also contains a polymer (not named on the label) to increase the viscosity. Solutions for adding to swimming pools are 10% aqueous sodium hypochlorite and are cheaper (per gm of NaClO) than household bleach. These pool disinfectants are often labelled liquid chlorine, a deceptive name that fosters ignorance among those who didn't pay attention at school. Chlorine boils at -34C at atmospheric pressure and exists as a liquid at ambient temperatures only when compressed in railroad tanks or in the (much smaller) gas shells used as weapons by both sides in the First World War. Dakin's solution, used for cleaning dirty wounds, is 0.45-0.50% sodium hypochlorite, made by dilution with a carbonate-bicarbonate buffer to reduce alkalinity. A stronger solution could be used for non-living surfaces. Aqueous solutions of NaClO are remarkably stable but the solid compound is extremely unstable. The crystaline pentahydrate melts at 18C and is decomposed by reaction with carbon dioxide from the air;. Anhydrous NaClO is obtainable only by freeze-drying and is very explosive. Nobody uses the solid. The solutions are made by reaction of chlorine (gas) with aqueous sodium hydroxide solutions. This can also be done by electrolysis of a sodium chloride solution between inert electrodes. My sources of information are bleach bottle labels (small print) and the Merck Index 12th edition, 1996, checked today. Every lab should have a Merck Index on the shelf of reference books! The phrase 99.9% commercial bleach could not possibly mean 99.9% of either NaOCl or its pentahydrate (solubility 29%). 5.25% (absurd precision!) and 10% bleach probably mean volume dilutions (1:20 or 1:10) of a household hypochlorite bleach without added thickener. A swimming pool 10% sodium hypochlorite solution probably is the cheapest source of chlorine bleach disinfectant without unwanted additives. Dilute it 10-20X with water to swab your possibly infected surfaces. Beware of lab supply companies selling household products at greatly inflated prices. Anyone purporting to sell hospital grade premade bleach needs to be viewed with much suspicion. Think before you buy. John Kiernan London, Canada = = = On 08/05/15, ian bernard ian.bern...@comcast.net wrote: Our facility is moving towards standardization of decontaminants or disinfectants. They prefer all areas use a Sani wipes that kills most pathogens. However, we contend for Anatomic pathology we need our liquid bleach not only as disinfectant or decontaminant but as a cleaning agent for stained lab-ware. You thoughts? Also, what concentration of Bleach (5.25 or 10%) is acceptable for use as both a disinfectant and cleaning agent or should we keep them separate? We used to buy the hospital grade premade bleach at a 5.25% concentration but now they want us instead to buy the 99.9% commercial Bleach and dilute from there. Any suggestions on opaque containers for us to purchase since bleach break down after a time period, at least for disinfecting? V/r IB ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Can't log into Histonet to do anything
You posted this one to Histonet! = = = On 05/05/15, sarah.dys...@stdavids.com wrote: At least you can post...I've been trying to be able to post for over a month, even emailed the list serv. With no response...frustrating because I love this resource! Sarah Dysart Pathology Supervisor -Original Message- From: Kathleen Roberts [mailto:kgrob...@rci.rutgers.edu] kgrob...@rci.rutgers.edu] Sent: Tuesday, May 05, 2015 9:07 AM To: callis, gayle; Histonet Subject: [EXTERNAL] Re: [Histonet] Can't log into Histonet to do anything I tried just now via Safari, and got this error message: Gateway Timeout: can't connect to remote host. Hopefully the IT people can fix it. Kathleen I got Gayle's email as well, but when I went to Histonet's page though Firefox and Google, got the Unable to Connect message when I clicked on a bunch of the Monthly hyperlinks. Same message for everything back until the Histonet, November, 2008 hyperlink. They seem to work prior to this date. Tried IE as well, same result. Brian -Original Message- From: koelli...@comcast.net [mailto:koelli...@comcast.net] koelli...@comcast.net] Sent: Monday, May 04, 2015 3:55 PM To: callis, gayle Cc: Histonet Subject: Re: [Histonet] Can't log into Histonet to do anything I got your message Gayle (through Histonet) although haven't heard much at all weekend otherwise. Use IE. Ray in Lake Forest Park, WA - Original Message - From: Gayle Callis gayle.cal...@bresnan.net To: Histonet histonet@lists.utsouthwestern.edu Sent: Monday, May 4, 2015 3:21:01 PM Subject: [Histonet] Can't log into Histonet to do anything Dear Histonettters,  At the risk of being pesky, is Histonet having problems.  I generally go to Histonet via Firefox/Google and haven't been able to get to the website for two days.  I only hope someone out there can even get this message.  I have tried finding Marvin Hanna's email address.   Stymied and dead in the water.  I would love to unsubscribe, but not sure anyone gets this message.  Gayle Callis ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet - CONFIDENTIALITY NOTICE: This e-mail message, including any attachments, is for the sole use of the intended recipient(s) and may contain confidential or legally privileged information. Any unauthorized review, use, disclosure or distribution is prohibited. If you are not the intended recipient, please contact the sender by reply e-mail and destroy all copies of this original message. - ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet Principal Lab Technician Neurotoxicology Labs Dept of Pharmacology Toxicology Rutgers, the State University of NJ 41 B Gordon Road Piscataway, NJ 08854 (848) 445-1443 FAX (732) 445-6905 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: FW: [Histonet] IHC and oven temperature
The statement quoted by Tony from the Dako manual cannot be true because many antigens have to be exposed to water at 100C in order to be immunostained - antigen retrieval. Denaturation of a macromolecule by heat increases the number of exposed epitopes, which typically are short amino acid sequences that bind specifically to the Fab segments of antibody molecules. On the other hand, it is easy to believe that 60C would denature antibody molecules enough to damage their binding sites and impair or prevent immunostaining. According to AWP Vermeer and W Norde (2000), the Fab segments of IgG were denatured when the temperature of a solution slightly exceeded 60C. (The Thermal Stability of Immunoglobulin: Unfolding and Aggregation of a Multi-Domain Protein Biophysical Journal 78: 394–404.) They found that further heating denatured the Fc segment, but the changed molecules became entangled and aggregated before denaturation was complete. Microwave heating is sometimes used to accelerate immunostaining, but control of the temperature is critical. For example: ME Boon E Marani (1991) The major importance of temperature data in publications concerning microwave techniques European Journal of Morphology 29: 181–183. John Kiernan London, Canada = = = On 30/04/15, Tony Henwood (SCHN) tony.henw...@health.nsw.gov.au wrote: Yes, I read the Dako IPX educational guides (5th ed) and on page 32: No processes should raise tissue temperature to higher than 60oC as this will cause severe loss of antigenicity that may not be recoverable Unfortunately there is no evidence given or cited that validates this statement. Even though this could be right (and there are several papers that have looked at this), this statement is scientifically weak and we should not cite this as truth. Now I do recommend the Dako reference series to my students, and I have contributed to one of these texts myself (Microscopic control of routine HE - know your histology) but I request my students to continue to question what they read and confirm the scientific validity of the information. Regards, Tony From: Joelle Weaver [joellewea...@hotmail.com] Sent: Saturday, 25 April 2015 5:51 AM To: Tony Henwood (SCHN); WILLIAM DESALVO; Preiszner, Johanna Cc: histonet@lists.utsouthwestern.edu Subject: RE: [Histonet] IHC and oven temperature I remember reading that the preferred temperature was about 60 degrees Celsius. I think that this was in the Dako education guides if I'm not mistaken. If that is the case, the citation for the source is probably in that resource available as pdf from their website. Joelle Weaver MAOM, HTL (ASCP) QIHC From: tony.henw...@health.nsw.gov.au To: wdesalvo@outlook.com; preis...@mail.etsu.edu Date: Fri, 24 Apr 2015 09:43:59 + Subject: RE: [Histonet] IHC and oven temperature CC: histonet@lists.utsouthwestern.edu Hi temp drying shown to be a bad idea: Henwood, A., (2005) “Effect of Slide Drying at 80°C on Immunohistochemistry” J Histotechnol 28(1):45-46. Abstract Prolonged high temperature dry heating has been found to be deleterious to the immunohistochemical demonstration of several antigens in formalin-fixed, paraffin- embedded sections. Paraffin sections were dried at 80°C for 7 h and their immunoreactivity was compared with mirror sections dried for 1 h at 60°C. NCL-5D3, CMV, S100, HMB45, and CEA were quite labile to dry overheating whereas AElAE3, HBsAg, HBcAg, HSVII, EMA, chromogranin, and NSE were found to be quite resistant. It is recommended that coated slides (poly-L-lysine or aminopropyltriethoxysilane) and low-temperature drying (60°C) be routinely used for irnmunohistochemistry. From: histonet-boun...@lists.utsouthwestern.edu [histonet-boun...@lists.utsouthwestern.edu] on behalf of WILLIAM DESALVO [wdesalvo@outlook.com] Sent: Tuesday, 21 April 2015 1:56 AM To: Preiszner, Johanna Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] IHC and oven temperature Dry heat compared to wet heat. Do not dry your slides at high heat. You are removing water trapped between slide and paraffin section. Antigen retrieval is an entirely different process. So not try to combine the two processes Sent from my iPhone On Apr 20, 2015, at 8:48 AM, Preiszner, Johanna preis...@mail.etsu.edu wrote: Hi Netters, is there something wrong with this logic: If the tissue needs 95C for HIER, it's ok to dry the slides in an 82C oven. Of course I'll test it before I try it on real specimens, but maybe someone else already knows the answer... Thanks! Hanna Preiszner ETSU/QCOM ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu
Re: [Histonet] Eosin pH
Dissolve it in a buffer (eg acetic acid-sodium acetate) at or near pH 4.5. John Kiernan London, Canada = = = On 23/04/15, Pablo Sanchez-Quinteiro pablo.sanc...@usc.es wrote: Listers, Could you tell me the best way to adjut the eosin pH to 4-5? Thanks in advance Pablo ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Schiffs
Schiff's reagent does not need to be refrigerated. It just needs to be screw-capped so that it doesn't decompose by loss of sulphur dioxide. For decades, Schiff has been stored in many labs at 4C for no good reason. It is used at room temperature in both of its histochemical applications: the Feulgen and PAS reactions. John Kiernan Anatomy, UWO London, Canada = = = On 17/04/15, Bernice Frederick b-freder...@northwestern.edu wrote: Am I missing something? I ordered Schiffs and sigma tells me it was shipped yesterday. Hello, today is Friday and last I recall Schiffs need to be refrigerated! You'd.. think they would realize this. Sorry all, have to vent. Bernice Bernice Frederick HTL (ASCP) Senior Research Tech Pathology Core Facility Robert. H. Lurie Cancer Center Northwestern University 710 N Fairbanks Court Olson 8-421 Chicago,IL 60611 312-503-3723 b-freder...@northwestern.edumailto:b-freder...@northwestern.edu b-freder...@northwestern.edu ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Astrocyte immunohistochemical marker
Immunostaining for GFAP has been the definitive method for astrocyte cytoplasm for 30 years, and good primary antibodies have been around for at least 20 years. The distinction between different kinds of astrocyte is made by staining with serial dilutions of the primary antiserum. Reactive astrocytes have the highest concentrations of glial fibrillary acidic protein (GFAP) in their cytoplasms, and are immunostained by greatly diluted antisera. Normal fibrous astrocytes (in normal white matter and also with processes abutting ependyma, pia and small blood vessels) also contain plenty of GFAP, but are unlikely to be confused with astrocytes around a site of injury or in a tumour. Normal protoplasmic (or velate) astrocytes extend their processes into normal white and grey matter; their cytoplasm contains less GFAP than the cytoplasm of astrocyte in normal white matter. To detect reactive astrocytes you need to do serial dilutions of the primary antibody on sections of normal CNS. For detecting gliosis, use the dilution that fails to stain GFAP in grey matter. This should show astrogliosis strongly, and also the fibrous astrocytes normally present in white matter. A high concentration of anti-GFAP will stain everything in the CNS, because astrocyte processes are everywhere there. The traditional astrocyte stain is Cajal's gold-sublimate. This shows normal and gliotic fibrous astrocytes nicely, but it takes time. For protoplasmic astrocytes you need electron microscopy, wich takes even more time. John Kiernan Anatomy, UWO, London, Canada = = = On 15/04/15, Stephen KumJew stephen@sydney.edu.au wrote: I was wondering if anyone could recommend an antibody (and best dilution) which is more effective than GFAP in staining both resting and active astrocytes in human brain please. Thanks STEPHEN KUM JEW | Senior Technical Officer Discipline of Pathology | School of Medical Sciences THE UNIVERSITY OF SYDNEY Charles Perkins Centre Hub | Building D17 | Camperdown | NSW | 2050 | Australia. Delivery address: The CPC Hub Dock, Orphans School Creek Lane, Camperdown NSW 2050. T +61 2 9036 9027| F +61 2 8627 1606 E stephen@sydney.edu.au | W http://sydney.edu.au/medicine/pathology/ This email plus any attachments to it are confidential. Any unauthorised use is strictly prohibited. If you receive this email in error, please delete it and any attachments. Please consider the environment before printing this email. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Bouin's, formic acid and sirius red staining
Fixation must be adequate before decalcifying. 12-24h in Bouin is OK. My favourite formic decalcifier is that of Clark (1954) Am. J. Clin. Path. 24: 1113-1116. It's a buffer (pH2.0) made by mixing 90% formic acid 250ml, water 750 ml and sodium formate (anhydrous) 34g. Keeps for ever. Check the reference. Never trust a recipe you read on the internet! The picric acid in picro-sirius red is a necessary component of the staining solution. See Junqueira et al 1979 Histochem. J. 11: 447-455. It's important to have the correct red dye; there is more than one sirius red. Use C.I. 35780, Direct red 80. John Kiernan London, Canada = = = On 15/04/15, Tyrone Genade tgen...@gmail.com wrote: Hello, Some questions regarding Bouin's solution. I was told, back when I was doing my PhD and new very little, that I should fix my fish in Bouin's as it will decalsify the bones. Well, Bouin's fixed fish were easier to cut than PFA fixed fish... but I read today that by adding formic acid the decalsification is better. (I must confess, that after Buoin's fixation I still had to soak the tissue face in some dilute nitric acid now and then...) Another reference said that the formaldehyde should be replaced with formic acid. So which is it: add formic acid or replace formaldehyde? And if the former, how much do you add? Second question: a colleague and I want to stain for collagen in diseased kidneys. The fixative of choice for soft tissue is Bouin's... But the staining protocol called for a solution of picric acid and sirius red. Is the picric acid needed if I haven't washed the picric acid from the Bouin's fixation out of the tissue? I was told once that the picric acid was for contrast... Is this BS? Does the picric acid play an important chemical role in the staining? I would like to avoid the need for a bottle of saturated picric acid on the lab shelf here in Iowa where the winter low humidity desiccates everything... I'm hoping this protocol, http://www.ihcworld.com/_protocols/special_stains/sirius_red.htm , can be modified to omit the picric acid. Thanks -- Tyrone Genade Orange City, Iowa tel: (+1) 712 230 4101 http://tgenade.freeshell.org Romans 6:23: The gift of God is eternal life through Christ Jesus our Lord. To find out how to receive this FREE gift visit http://www.alpha.org. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Reference needed
Do you need a source to satisfy ignorant officials? Try a good textbook that shows up in second-hand bookshops. Brown,GC 1978 An Introduction to Histotechnology. New York: Appleton-Century-Crofts. It doesn't seem to have an ISBN. John Kiernan = = = On 13/04/15, Jennifer MacDonald jmacdon...@mtsac.edu wrote: Does anyone have a reference for the theory of: 1. The use of soapy water to prevent fatty tissue from blowing apart in the flotation bath? 2. The use of ammonia water to rehydrate tissues? Many of us use these tricks, but is there a source for the theory? Thanks, Jennifer MacDonald ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Sihler staining
No first-hand knowledge, unfortunately, but in addition to the five references in the 2005 Histonet post that you mentioned - http://lists.utsouthwestern.edu/mailman/htdig/histonet/2005-November/018700.html - there is a more recent review with plenty of technical details and photos: Mu, L. and Sanders, I. (2010). Sihler's whole mount nerve staining technique: a review. Biotechnic Histochemistry 85:19-42. The online version has impressive colour photos; the print version has only BW. The corresponding author is l...@humed.com (Hackensack Univ., NJ). I don't know if he's still there. Hope this helps. John Kiernan London, Canada = = = On 01/04/15, Caroline Miller mi...@3scan.com wrote: Hi Histonet, I just started working for a small start up in San Francisco called 3scan, after 18 years in clinical (human) and academia (animal tissues). We have a microscope that both sections and images at the same time. This means that my job is to get stain into whole mount tissues prior to sectioning. We found the Sihler staining technique for peripheral nerves and have been getting it to work well in animal organs that were taken directly from the animal and fixed. We are now trying embalmed human tissue, which is our final target. We had two samples, the first one came out very ragged and I don't think it had any nerve tissue left in it after the maceration. It was really hard to know where the end point is, it was so subjective, especially when looking at unfamiliar tissues. I am thinking the first sample came out so ragged because we did not put the sample into formaldehyde prior to maceration because it was embalmed and we thought that was enough. We have one more sample that is now dissected and sat in unbuffered 4% formaldehyde. *I wanted to see if anyone familiar with the technique was prepared to have a conversation with me about the end points of each stage, *We have totally read and digested all of John's suggestions from this page: http://lists.utsouthwestern.edu/mailman/htdig/histonet//2005-November/018700.html plus many others, but it would be great to talk to someone first hand. I reached out to Mu and Sanders, but they are no longer at that institution and I can't find them. Or the other person who posted on Sihler to histonet in 2005 - *Maria Mejia* maria @t ski.org. Their email address bounces now histonet%40lists.utsouthwestern.edu?Subject=%5BHistonet%5D%20Sihler%27s%20stainIn-Reply-To= Is there anyone out there with first hand knowledge I can have a chat with? Thanks all, mills aka Caroline :) -- Caroline Miller Director of Histology 3Scan.com 415 2187297 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Tol Blue
Mast cells are stained red-purple by toluidine blue. They are defined as cells with granules that stain metachromatically with cationic dyes. You should seek to enhance, not avoid this effect. Metachromasia means staining in a colour different from that of the dye. Toluidine blue is a blue cationic dye, and it imparts its blue (orthochromatic) colour to anionic materials such as nucleic acids. Some sites in tissues have higher concentrations of anions, and this makes the dye cations pile up in layers. Stacking of dye cations changes their absorption of light, and toluidine blue looks red instead of blue. Mast cells contain heparin, which attracts stacks of small dye cations like those of toluidine blue. John Kiernan London, Canada = = = On 24/03/15, Kimberly Marshall kimbe...@animalreferencepathology.com wrote: ?Hello my fellow Histo Techs. Have a question I just know someone out there can answer for me. In canine tissue, we are having problems with the Tol Blue for mast cell. Am experiencing metachromasia, or the mast cells turning purple not blue. I have attempted to decrease time, or add time, but its not helping. My pathologist says he has had this issue before. So question is. Could it be the mast cell in a dog does not stain the same? Is there another stain that may work? Any help will be much appreciated. Thanks in Advance. Kimberly Marshall H.T.(ASCP) Kimberly Marshall H.T.(ASCP) Histology/Lab Supervisor Toll Free 1-800-426-2099 Fax 801-584-5104 PO Box 17580 Salt Lake City, Utah 84107 www.animalreferencepathology.comhttp://www.animalreferencepathology.com/ Advancing the art and science of veterinary medicine [cid:image001.jpg@01CF8F87.A0BD4830] ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: Fwd: [Histonet] Phosphotungstic acid
Dear Cecilia, I hadn't seen the 2009 paper you mention. Please let me know the reference. I've just sent you another email with an attached LM photo, which might be of interest. John Kiernan = = = On 25/03/15, Cecilia A. cecysta...@gmail.com wrote: Hi John, Thank you for your reply. In a Developmental Dynamics paper in 2009, PTA was used as a simple staining method that produced overall contrast for X-ray imaging soft tissues similar to how Osmium Tetroxide was also used as a soak for this purpose. We were interested in using PTA as it doesn't render the soft tissue black like Osmium Tetroxide does, so that there's still a chance we would be able to do further histology work on the tissue. However, we wonder what PTA might do to the fixed yet unprocessed tissue when it is soaked for this imaging purpose, and if it might affect its viability for staining and IHC work. Thank you, C. On Wed, Mar 25, 2015 at 9:11 AM, John Kiernan jkier...@uwo.ca wrote: PTA can be used on tiny specimens to enhance electron-opacity. For light microscopy, PTA is used in the trichrome methods to enhance differential staining of collagen and cytoplasm, especially in paraffin sections. These uses of PTA are not the same as soak formalin fixed soft tissue for a few hours before it gets processed. What are you hoping to achieve? John Kiernan London, Canada = = = On 25/03/15, Cecilia A. cecysta...@gmail.com wrote: Hello Histonetters, I was wondering if anyone might be able to give some input regarding the use of Phosphotungstic acid in which we want to soak formalin fixed soft tissue for a few hours before it gets processed. Will subjecting it to PTA before processing affect its viability for further histology work (ie, HE staining and IHC)? Thank you very much! C. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Phosphotungstic acid
PTA can be used on tiny specimens to enhance electron-opacity. For light microscopy, PTA is used in the trichrome methods to enhance differential staining of collagen and cytoplasm, especially in paraffin sections. These uses of PTA are not the same as soak formalin fixed soft tissue for a few hours before it gets processed. What are you hoping to achieve? John Kiernan London, Canada = = = On 25/03/15, Cecilia A. cecysta...@gmail.com wrote: Hello Histonetters, I was wondering if anyone might be able to give some input regarding the use of Phosphotungstic acid in which we want to soak formalin fixed soft tissue for a few hours before it gets processed. Will subjecting it to PTA before processing affect its viability for further histology work (ie, HE staining and IHC)? Thank you very much! C. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Block staining with HE
You need to read the paper. It says All routinely used hematoxylins, such as Ehrlich’s, Mayer’s, Harris’ and Delafield’s, proved unsatisfactory STAIN TECHNOLOGY Vol. 56, No. 2, pp. 119-123 (1981) BLOCK STAINING OF MAMMALIAN TISSUES WITH HEMATOXYLIN AND EOSIN IAN F. HINE, Department of Anatomy, Monash University, Clayton, Victoria 3168, Australia. ABSTRACT. Various mammalian tissues were stained en bloc with hematoxylin and eosin after fixation and prior to embedding in paraffin wax and sectioning. The choice of fixative is important and best results are obtained using Worcester’s Fluid, a combination of saturated aqueous mercuric chloride, formaldehyde, and glacial acetic acid. After fixation, blocks of tissue up to 1.5 em thick are stained for seven days in hematoxylin. Excess stain is removed by washing tissues in running water overnight. Tissue blocks then are dehydrated with graded concentrations of ethyl alcohols to 80% and counterstained, with further dehydration, in 0.5% spirit soluble eosin in 90% ethyl alcohol for five days. The tissue is subsequently transferred to 90% ethyl alcohol overnight to differentiate win staining; dehydration is completed in absolute ethyl alcohol. The blocks are cleared in cedarwood oil and briefly in xylene prior to embedding, sectioning, and mounting. Following removal of wax by xylene, coverslips are applied. General morphological and histological features were particularly well differentiated and very selectively and reliably stained by this method. Hine's Worcester's fixative is similar to Stieve's. His haemalum formulation is a little unusual. The counterstain was ethyl eosin (CI 45385), not eosin Y (CI 45380). There is a page of colour photos in which the red/blue colour separation is satisfactory in only two (my opinion). One photo shows a block-stained HE section subsequently stained with Masson's and Verhoeff's, with bluish green collagen and black elastic laminae. John Kiernan London, Canada = = = On 20/03/15, Caroline Miller mi...@3scan.com wrote: Hi Histonet, Happy Friday! I was wondering if anyone had access to this paper: Stain Technol. http://www-ncbi-nlm-nih-gov.ucsf.idm.oclc.org/pubmed/6166093?dopt=AbstractPlustool=cdlotool=cdlotool# 1981 Mar;56(2):119-23. Block staining of mammalian tissues with hematoxylin and eosin. or if anyone had some tips on whole-mount HE staining I would gladly take it! I am just starting on this quest, but so far I have tried vacuum for the haematoxylin overnight with Erlichs and Harris's, and I have some Mayers on order. The Erlichs and harris's really overstained the tissue and I have been destaining for a while. I am thinking Mayer's may be better for its progressive nature, but any advice will be gratefully taken! Thank you in advance for your fantastic advice! yours, Caroline -- Caroline Miller Director of Histology 3Scan.com 415 2187297 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] FW: Masson's trichrome stain
The notion of plastic forceps is new to me. Where did Justine find it? Nothing in any variant of the Masson procedure should be adversely affected by moving slides with stainless steel forceps. Is there a commercial campaign to sell plastic tweezers to Histonetters? John Kiernan = = = On 08/03/15, Linda Margraf lindamarg...@gmail.com wrote: Here is a message from Justine... From: Justine Lanzon [mailto:justinelan...@hotmail.com] justinelan...@hotmail.com] Sent: Thursday, March 05, 2015 5:36 AM To: lindamarg...@gmail.com Subject: Masson's trichrome stain Hi, I am doing a write up on Masson's trichrome stain however I cannot answer these two questions: - Why are plastic forceps used instead of metal ones to hold the stained slide? - Why do we not rinse before Alinine blue? Can you please help me? Many Thanks, Justine Lanzon ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Old slides.
Have you done this? Acetone does not dissolve resinous mounting media and allow removal of coverslips. It's all in the books; buy one. John Kiernan Anatomy Cell Biology, UWO London, Canada = = = On 09/03/15, Jason McGough jmcgo...@clinlab.com wrote: Remove the film coverslip by placing the slide in acetone for a few minutes. Then recoverslip the slide with your current method. Jason McGough, HT(ASCP) Operations Manager Clinical Laboratory of the Black Hills 605-343-2267 jmcgo...@clinlab.com mailto:jmcgo...@clinlab.com jmcgo...@clinlab.com www.clinlab.com http://www.clinlab.com -Original message- From:Bernice Frederick b-freder...@northwestern.edu mailto:b-freder...@northwestern.edu b-freder...@northwestern.edu Sent: Monday, March 9, 2015 1:51 PM To: histonet@lists.utsouthwestern.edu mailto:histonet@lists.utsouthwestern.edu histonet@lists.utsouthwestern.edu Subject: [Histonet] Old slides. Hi all, We received some old slides (1997-1998) that were coverslipped with film. Sakura I would imagine. The issue here is that the coverslips have come up from the slide and the tissue is adhered to the back of the coverslip. They need to be recovered so they can be evaluated. What do you all recommend? We use the CV5030 for coverslipping. I tried one with xylene and mounting media but there were still a couple of air bubbles in there. Thanks, Bernice Bernice Frederick HTL (ASCP) Senior Research Tech Pathology Core Facility Robert. H. Lurie Cancer Center Northwestern University 710 N Fairbanks Court Olson 8-421 Chicago,IL 60611 312-503-3723 b-freder...@northwestern.edu mailto:b-freder...@northwestern.edu b-freder...@northwestern.edu mailto:b-freder...@northwestern.edu b-freder...@northwestern.edu mailto:b-freder...@northwestern.edu b-freder...@northwestern.edu ___ Histonet mailing list Histonet@lists.utsouthwestern.edu mailto:Histonet@lists.utsouthwestern.edu Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: AW: [Histonet] Trichrome Fixation
Quite right! Treating sections of formaldehyde-fixed tissue with Bouin's fluid is not mordanting; binding site retrieval is probably a more accurate term. Picric acid induces a change in the section that improves the selective uptake of the heteropolyacid and the larger dye anions by collagen and of the smaller dye anions by cytoplasm. Picric acid alone is just as good as Bouin for this purpose. The picric acid treatment has the effect of making a section of formaldehyde-fixed tissue more like a section of tissue fixed in a picric acid-containing fixative such as Bouin or Gendre. For spectacular trichrome staining you need a fixative that contains mercuric chloride, like Zenker or SuSa. An example of a mordant in biological staining is iron alum in Heidenhain's method. The subsequently applied haematein solution forms a coloured complex with the iron bound by the sections. The differentiation in iron alum solution then removes bound haematein by forming a water-soluble complex. The word mordant is probably best avoided for staining with pre-formed dye-metal complexes such as carmine, aluminium-haematein (haemalum), Weigert's iron-haematein and gallocyanine-chrome alum. See R.W.Horobin (1982) Histochemistry Stuttgart: Gustav Fischer (ISBN 3437107003 or 0407002480), page 77. John Kiernan = = = On 06/03/15, Gudrun Lang gu.l...@gmx.at wrote: I disagree. Usually a dye binds covalently to the mordant, and a mordant should be a multi-valent metal (like aluminium, ferrum, molybden,..). The dye-mordant-complex binds via the mordant to the substrate. Picric acid of Bouin's is washed out before staining. It may be a matter of definition of the term mordant. Gudrun -Ursprüngliche Nachricht- Von: Johnson, Carole [mailto:cjohn...@nmda.nmsu.edu] cjohn...@nmda.nmsu.edu] Gesendet: Freitag, 06. März 2015 15:10 An: gu.l...@gmx.at Betreff: RE: [Histonet] Trichrome Fixation Boiuns solution acts as a mordant in trichrome stains -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Gudrun Lang Sent: Friday, March 06, 2015 1:13 AM To: 'Amos Brooks' Cc: histonet@lists.utsouthwestern.edu Subject: AW: [Histonet] Trichrome Fixation Hi, years ago we did a stain called CAB (=one step trichrome) regularly on liver-tissue. I don't know if it was because of ignorance or with aim, but it was done without Bouin. The result was blue-grey hepatocytes and darker blue collagen. - also totally different to the result with Bouin (red hepatocytes). I think the Bouin is less a re-fixation than more an binding-site retrival in this context. Gudrun -Ursprüngliche Nachricht- Von: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] histonet-boun...@lists.utsouthwestern.edu] Im Auftrag von Amos Brooks Gesendet: Donnerstag, 05. März 2015 22:01 An: histonet@lists.utsouthwestern.edu Betreff: [Histonet] Trichrome Fixation Hi, It is interesting that you should mention the importance of fixation on the Trichrome stain. I have an image of two murine hearts processed, cut and stained side by side. The only difference between the two is that they were harvested at different times, so one sat in formalin long enough to be properly fixed the other one was placed in the fixative then immediately brought in to be processed from 70% ethanol on. They are *totally* different looking. The red muscle tissue looks more purple therefore less distinct from the blue blood vessels. You get a similar effect with lung bronchial epithelium. Cheers, Amos On Tue, Mar 3, 2015 at 1:00 PM, histonet-requ...@lists.utsouthwestern.edu wrote: Message: 15 Date: Tue, 03 Mar 2015 12:34:33 -0500 From: John Kiernan jkier...@uwo.ca Subject: Re: [Histonet] Masson trichrome and H and E To: Emily Brown talulahg...@gmail.com, histonet@lists.utsouthwestern.edu histonet@lists.utsouthwestern.edu Message-ID: 7390afa3112a.54f5a...@uwo.ca Content-Type: text/plain; CHARSET=US-ASCII If you can't get two colours with HE, don't expect to get the colour scheme right with Masson's trichrome, which needs more skill. If you are hoping to show basement membranes in the kidney, you would do better to use a technically simpler staining method such as picro-sirius red or periodic acid-Schiff. If for some reason you really need three colours, a one-step trichrome such as Gomori's, Cason's or Gabe's might be the way to go rather than Masson's or one of the other multi-step trichromes. Remember that all trichrome methods are greatly influenced by the fixative. A post-fixation treatment of the sections, usually with picric acid, is needed for formaldehyde-fixed tissues. Some alternative post-fixation treatments were proposed by Yu Chapman (2003) J. Histotechnol
Re: [Histonet] Masson trichrome and H and E
If you can't get two colours with HE, don't expect to get the colour scheme right with Masson's trichrome, which needs more skill. If you are hoping to show basement membranes in the kidney, you would do better to use a technically simpler staining method such as picro-sirius red or periodic acid-Schiff. If for some reason you really need three colours, a one-step trichrome such as Gomori's, Cason's or Gabe's might be the way to go rather than Masson's or one of the other multi-step trichromes. Remember that all trichrome methods are greatly influenced by the fixative. A post-fixation treatment of the sections, usually with picric acid, is needed for formaldehyde-fixed tissues. Some alternative post-fixation treatments were proposed by Yu Chapman (2003) J. Histotechnol. 26(2): 131-134, but their coloured photos were not very convincing. Making up staining solutions in-house is always cheaper than buying pre-made solutions. John Kiernan London, Canada = = = On 03/03/15, Emily Brown talulahg...@gmail.com wrote: Hello! I am doing Masson trichrome manually (not with a machine) and I just found out the kit is $800!! I was thinking of buying the ingredients separately, but why the hell is it so expensive??? Also, the people I was going to borrow their reagents from said their aniline blue is not very good and I wanted to replace it. I only need about 250ml, what brand do you guys prefer? l know I can google this, but I want to know what you guys like and what works best. This is for mouse kidney paraffin sections, 4 to 5 microns. Another question, I did H and E and there is no eosin staining. I think the reagents are pretty old, so I thought that might be a problem. Also because my lab is cheap, they were reusing the xylenes and EtOH for both rehydrating and rehydrating. I told my boss this is probably not a good idea as the end steps will have stain in them. And I also think this is why it didn't work! The EtOH is also really old so who know is the 100% is actually even close to 100% any more. I'm buying new reagents, but if you guys think anything else would help, let me know. Also, shoutout to Ann, I know you're reading these!! Join the list!! Emily By bitching and bitching and bitching, they could exhaust the drama of their own horror stories. Grow bored. Only then could they accept a new story for their lives. Move forward. -Chuck Palahniuk, Haunted ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Re: CBG recyler and recycled Formalin
Bob Richmond makes an important point that should also be made clear by any company selling apparatus to recover formaldehyde by distillation of diluted formalin. The recovered product recovered from an aqueous fixative will be an aqueous solution of formaldehyde (and its low polymers), of uncertain concentration. Distillation of full-strength formalin (37%w/w = 40%w/v formaldehyde), done at atmospheric pressure, yields 20-30% of formaldehyde in the distillate, and leaves a higher concentration of formaldehyde and polymerization products in the still. Vacuum distillation or pressure distillation can change the yields to favour the collection of hydrated formaldehyde (methylene glycol) in the distillate. I'm summarizing Chapter 8 in Walker, JF (1964) Formaldehyde, 3rd edition. ISBN 0882752189. Walker's Chapter 8 tabulates conditions for distilling formaldehyde from various concentrations, and takes into account the methanol (about 5%, included in formalin to retard polymerization). The notion of an expensive machine for recycling formalin in a histology lab makes little sense. Why not simply filter (if necessary) and re-use the fixative after removing the specimen? It is still neutral, buffered formaldehyde. John Kiernan London, Canada = = = On 28/02/15, Bob Richmond rsrichm...@gmail.com wrote: Ryan Roy HTL (ASCP) in Manchester NH asks: We are getting a new CBG that recycles xylene , alcohol, and formalin. We purchase buffered formalin. Does anyone know if after recycling the recycled formalin would or would not need be re-buffered? If you distill buffered formalin, the formaldehyde is going to distill over, but not the buffer phosphate, which will remain in the still pot. I suppose you can buy phosphate mixtures to make Lillie's neutral buffered formalin anew, using your recycled formaldehyde. I think you also have to measure the concentration of formalehyde in the distillate, and dilute accordingly. Bob Richmond Samurai Pathologist Maryville TN ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] gelatin
Dear ynw...@u.washington.edu. Boiling makes steam, with bubbles that greatly change tissue structure. Slow freezing is just as bad; the ice crystals make holes that deform and replace the tissue architecture. What are you tryng to find out? It has been known for 100+ years that boiling collagen makes gelatin, and further concentration makes traditional glue. You should involve your boss in future email exchanges. John Kiernan London, Canada = = = On 24/02/15, Yak-Nam Wang ynw...@u.washington.edu wrote: Thank you for your e-mail. Apologies for not explaining treated tissue. We treat the tissue with high intensity focused ultrasound. It can raise the temperature of tissue to boiling in a localized area (millimeter areas). I could use a biochemical assay for collagen and gelatin if we treat a large area, but with single lesions I was hoping I could visualize this. In some treated areas we are almost resulting in liquefaction of the tissue. I am interested to see if we are turning the collagen to gelatin in these areas and what part of the lesion this is happening. You should involve your boss in future email exchanges. Thank you for your thoughts Yak-Nam On Mon, Feb 23, 2015 at 9:47 PM, John Kiernan jkier...@uwo.ca wrote: You need to explain treated tissue. Gelatin is collagen that has been boiled until the protein has lost all its fibrous nature and changed into a water-soluble protein. Gelatin is made permanently insoluble by adequate formaldehyde fixation. It is stained by anionic dyes (such as eosin in the HE method), but it does not show as fibres when you look at the section or smear through a microscope. If this doesn't answer your question, please explain your problem and involve your boss in future email exchanges. John Kiernan London, Canada = = = On 23/02/15, Yak-Nam Wang ynw...@u.washington.edu wrote: Hello, Does anyone know of a stain specific for gelatin? I would like to distinguish between firbous collagen and gelatin in treated tissue. thank you Yak-Nam University of Washington Seattle, WA ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] gelatin
You need to explain treated tissue. Gelatin is collagen that has been boiled until the protein has lost all its fibrous nature and changed into a water-soluble protein. Gelatin is made permanently insoluble by adequate formaldehyde fixation. It is stained by anionic dyes (such as eosin in the HE method), but it does not show as fibres when you look at the section or smear through a microscope. If this doesn't answer your question, please explain your problem and involve your boss in future email exchanges. John Kiernan London, Canada = = = On 23/02/15, Yak-Nam Wang ynw...@u.washington.edu wrote: Hello, Does anyone know of a stain specific for gelatin? I would like to distinguish between firbous collagen and gelatin in treated tissue. thank you Yak-Nam University of Washington Seattle, WA ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet