[Histonet] fume exposure
To All, I work in a lab where I stain and coverslip everything by hand. I coverslip under a fume hood so there isn't much of an issue there. The main issue is my exposue to fumes while staining. I do not stain under a fume hood. My exposure times to these c hemicals is increasing becuase our volume is steadly increasing as well. I can no longer tolerate these fumes and am expirenc ing difficulty breathing and frequent headaches.What are labs that stain there slides by hand doing (types of masks, hoods) to minimize their exposure. If you can recommed any specifics I would appreciate it. Thank you in advance Jenny ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] hand processing schedule late mouse embryos
Under separate cover I am sending a protocol that will work well with your subjects. Since you are doing manual processing you can easily use the reagents I recommend and eliminate xylene and its terpene and alkane based substitutes. René J. --- On Fri, 2/13/09, Nicole Collette collet...@mail.llnl.gov wrote: From: Nicole Collette collet...@mail.llnl.gov Subject: [Histonet] hand processing schedule late mouse embryos To: histonet@lists.utsouthwestern.edu Date: Friday, February 13, 2009, 1:25 PM HI, All, One project finished, another just beginning. I am about to embark on a journey into the land of immunohistochemistry, with late mouse embryos E14.5, E16.5, P0 to examine bone markers in conjunction with LacZ and/or GFP. We have sadly lost our cryostat (so IHC for the GFP on paraffin sections), and our tissue processor - both belonged to a friendly investigator down the hall who has moved on. So, I am processing by hand. For hand-processing, I have had to do some rigging, and I do the wax steps in a hyb oven to try to keep the wax (TissuePrep, Fisher) at around 63-65C, while trying my best to keep the molds, cassettes, and tools with as few giant globs of solidifying wax as possible. As a result of using the hyb oven, we are forced to use Clearene (D-limonene), --or some other xylene substitute that could be recommended--, instead of xylene for the processing. If anyone has a recommendation for a better alternative there (aside from a tissue processor which will have to wait at least until the next grant gets funded--oooh, unless someone has an old one they want to donate, preferably table-top), I'm all ears. My schedule was given to me by a friend who does cartilage, no older than E14.5, and are basically half hour steps for each ethanol, half hour steps for 3 wax steps at the end. Will this be enough time for infiltration of older samples without vacuum? Should I increase my steps to 1 hour for these older embryos? I am optimizing my fixation at 1hour/mm thickness, with the embryos skinned (4% paraformaldehyde in PBS, I decided to start here since I don't yet know much about the problems I might encounter with a particular antigen). I have tried the 30 minute schedule with adult decalcified bones and have not had fantastic sections. I suspect it could be incomplete washing of the EDTA before infiltration, but it's possible that the processing schedule is just not long enough. Any advice? Thanks in advance! Happy Friday! Sincerely, Nicole Collette Lawrence Livermore National Lab/ UC Berkeley ___ 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] need controls
The same appendix that you can use as an HE+ control, usually contains Gram +and- bacteria. René J. --- On Fri, 2/13/09, Jaime Plata enrri...@yahoo.com wrote: From: Jaime Plata enrri...@yahoo.com Subject: Re: [Histonet] need controls To: Histonet@lists.utsouthwestern.edu, Steven Coakley sjchta...@yahoo.com Date: Friday, February 13, 2009, 5:26 PM GI (small intestine) biopsies are other possibilities associated to skin as control. When you mention Grams, are you referring to Bact's positive and negative.? If this is the case you can make them with blood Agar (microbiology) and contaminate the tissue with them. Or use Large intestine ther will be many you jus need to look for them. --- On Fri, 2/13/09, Steven Coakley sjchta...@yahoo.com wrote: From: Steven Coakley sjchta...@yahoo.com Subject: [Histonet] need controls To: Histonet@lists.utsouthwestern.edu Date: Friday, February 13, 2009, 1:51 PM I work in a DermPath lab. Currently we purchase all our controls. Some are Ok but some, especially the grams are not. We'd like to start sectioning. Our own but all the specimens we get are skins. ___ 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] freezing paraffin blocks
Not at all! René J. --- On Fri, 2/13/09, Jennifer Anderson jander...@halozyme.com wrote: From: Jennifer Anderson jander...@halozyme.com Subject: [Histonet] freezing paraffin blocks To: Histonet@lists.utsouthwestern.edu Date: Friday, February 13, 2009, 5:43 PM Hi All. I've just started sectioning Bouin's fixed paraffin embedded human/mouse/rat/pig skin again, tissues processed in a softer wax than I'm used to, so I've been putting the blocks in the freezer at -20 prior to sectioning. Will such cold deleteriously affect the blocks or tissue? Also, is there any harm in cleaning the water bath with a little xylene to remove any wax build-up? Thanks a lot. Jennifer M. Anderson, Scientist Halozyme Therapeutics, Inc. 11404 Sorrento Valley Road San Diego, CA 92121 858-704-8333 jander...@halozyme.com The information transmitted in this email is confidential and is intended only for the person(s) or entity to which it is addressed. Delivery of this message to any person other than the intended recipient(s) is not intended in any way to waive confidentiality or any applicable privilege. Any review, retransmission, dissemination or other use of, or taking of any action in reliance upon, this information by individuals or entities other than the intended recipient is prohibited by Halozyme and may be in violation of applicable laws. If you received this in error, please contact the sender and delete/destroy 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
[Histonet] Problem with Schiff's reagent
Good Morning! I'm a newcomer to the list, and I'll start by pointing out that I'm not a histologist by any means. I am a freshwater ecologist, and we're trying to study mucus secretion behavior in freshwater snails. Last Spring and last summer, we developed a process whereby we could visualize snail mucus trails on glass slides using a periodic acid-Schiff's reagent staining technique. But now, in following the same protocol as used last year for making our own Schiff's reagent, I cannot get the final solution to filter out clear. Recipe I'm using: 900 ml boiling water 10 grams basic fuchsin 25 ml concentrated HCl acid (12 M) 40 grams sodium metabisulfite (this is essentially Sigma Aldrich's ratios, I think) Let this sit for 24 to 72 hours, take 100 ml aliquot, add 0.75 - 1.0 gram ground activated charcoal, stir for 10 minutes, filter through filter paper then through GF/C glass fiber filter. Last summer I got nice, clear (slightly yellow) and very active Schiff's reagent. But now I cannot seem to get the filtrate to be clear. Even after 10 minutes exposure to ground activated charcoal and filtering, the filtrate remains bright orange to dark red and it does not seem to stain mucus trails very well. All the reagents are the same as those used last summer (i.e., less than 7 months old; although the HCl is a bottle several years old from a different lab). Anyone have any troubleshooting suggestions? I don't know the chemistry very well, but the sodium metabisulfite is used for decoloring the initial solution, right? So is the metabisulfite not working for some reason now?? Any help/suggestions would be greatly appreciated. Cheers - Paul Paul E. Brunkow, PhD Department of Biological Sciences Southern Illinois University Edwardsville Edwardsville, ILUSA - SIUE Web Mail ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Problem with Schiff's reagent
Hi Paul, Some people may think this is taking the easy way out, but...you might consider purchasing some commercially prepared Schiff's. It will (should) be consistent product (consistently produced) and the manufacturer has the responsibility of QC'ing the product before it is shipped and sold. There are any number of reliable companies handling Schiff's (I would not favor one over another). You could type Schiff's into your search engine and I'm certain you'd be able to contact a supplier. This may be an answer for you. Good Luck, Tom Jasper Thomas Jasper HT (ASCP) BAS Histology Supervisor Central Oregon Regional Pathology Services Bend, OR 97701 541/693-2677 tjas...@copc.net -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of pbru...@siue.edu Sent: Saturday, February 14, 2009 9:54 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Problem with Schiff's reagent Good Morning! I'm a newcomer to the list, and I'll start by pointing out that I'm not a histologist by any means. I am a freshwater ecologist, and we're trying to study mucus secretion behavior in freshwater snails. Last Spring and last summer, we developed a process whereby we could visualize snail mucus trails on glass slides using a periodic acid-Schiff's reagent staining technique. But now, in following the same protocol as used last year for making our own Schiff's reagent, I cannot get the final solution to filter out clear. Recipe I'm using: 900 ml boiling water 10 grams basic fuchsin 25 ml concentrated HCl acid (12 M) 40 grams sodium metabisulfite (this is essentially Sigma Aldrich's ratios, I think) Let this sit for 24 to 72 hours, take 100 ml aliquot, add 0.75 - 1.0 gram ground activated charcoal, stir for 10 minutes, filter through filter paper then through GF/C glass fiber filter. Last summer I got nice, clear (slightly yellow) and very active Schiff's reagent. But now I cannot seem to get the filtrate to be clear. Even after 10 minutes exposure to ground activated charcoal and filtering, the filtrate remains bright orange to dark red and it does not seem to stain mucus trails very well. All the reagents are the same as those used last summer (i.e., less than 7 months old; although the HCl is a bottle several years old from a different lab). Anyone have any troubleshooting suggestions? I don't know the chemistry very well, but the sodium metabisulfite is used for decoloring the initial solution, right? So is the metabisulfite not working for some reason now?? Any help/suggestions would be greatly appreciated. Cheers - Paul Paul E. Brunkow, PhD Department of Biological Sciences Southern Illinois University Edwardsville Edwardsville, ILUSA - SIUE Web Mail ___ 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] AUTO: Jacquelyn Grewe/Staff/OhioHealth is out of the office .
I will be out of the office starting 02/11/2009 and will not return until 02/16/2009. I will respond to your message when I return. Note: This is an automated response to your message Histonet Digest, Vol 63, Issue 24 sent on 2/14/2009 12:57:22 PM. You will receive a notification for each message you send to this person while the person is away. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] Problem with Schiff's reagent
Paul: If I remember correctly the last time I prepared my own Schiff reagent was in 1955 and it came beautifully BUT it is a real pain to prepare it, and not really worth the work. Since then I purchase my Schiff and avoided all the trouble. If you have the necessary budget I would advise you to buy it from a reputable manufacturer. Mine always came from Sigma. It will eliminate inconsistencies in your research. René J. --- On Sat, 2/14/09, pbru...@siue.edu pbru...@siue.edu wrote: From: pbru...@siue.edu pbru...@siue.edu Subject: [Histonet] Problem with Schiff's reagent To: histonet@lists.utsouthwestern.edu Date: Saturday, February 14, 2009, 12:53 PM Good Morning! I'm a newcomer to the list, and I'll start by pointing out that I'm not a histologist by any means. I am a freshwater ecologist, and we're trying to study mucus secretion behavior in freshwater snails. Last Spring and last summer, we developed a process whereby we could visualize snail mucus trails on glass slides using a periodic acid-Schiff's reagent staining technique. But now, in following the same protocol as used last year for making our own Schiff's reagent, I cannot get the final solution to filter out clear. Recipe I'm using: 900 ml boiling water 10 grams basic fuchsin 25 ml concentrated HCl acid (12 M) 40 grams sodium metabisulfite (this is essentially Sigma Aldrich's ratios, I think) Let this sit for 24 to 72 hours, take 100 ml aliquot, add 0.75 - 1.0 gram ground activated charcoal, stir for 10 minutes, filter through filter paper then through GF/C glass fiber filter. Last summer I got nice, clear (slightly yellow) and very active Schiff's reagent. But now I cannot seem to get the filtrate to be clear. Even after 10 minutes exposure to ground activated charcoal and filtering, the filtrate remains bright orange to dark red and it does not seem to stain mucus trails very well. All the reagents are the same as those used last summer (i.e., less than 7 months old; although the HCl is a bottle several years old from a different lab). Anyone have any troubleshooting suggestions? I don't know the chemistry very well, but the sodium metabisulfite is used for decoloring the initial solution, right? So is the metabisulfite not working for some reason now?? Any help/suggestions would be greatly appreciated. Cheers - Paul Paul E. Brunkow, PhD Department of Biological Sciences Southern Illinois University Edwardsville Edwardsville, ILUSA - SIUE Web Mail ___ 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] fume exposure
Is it so old-fashioned to use xylene in a fume hood? We can't even use paraformaldhyde outside of the hood--Environmental Health and Safety makes sure of that. But we do research, maybe it's different in other types of labs. Emily -- It's like hearing Billy Joel play Piano Man-- joyless for all involved, but demanded by a higher power. --Kevin Murphy, Indiana Jones and the Crystal Skull rifftrax ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] semen analysis
thanks for the email we are a hospital based diagnostic lab and currently do all the testing manually i am actually asking what EQUIPMENT is being used for AUTOMATED analysis (count, conc, motility etc) regards Anne 2009/2/15 Malcolm McCallum malcolm.mccal...@tamut.edu semen analysis can be done using various methods depending on what you are analyzing. I am not certain if you are asking for techniques, or specific procedures. However, fluorescence microscopy can be used to assess simple morphology using mitotracker blue. Fluorescence can also be used with a fast green (I think this is the other dye) and propidium iodide to assess viability. alternatively, you can assess viability using flow cytometry with the same dyes. Also, morphology is typically done using SEM and TEM. there are several techiques for assessing motility. Jill Jenkins at the USGS National Wetlands Laboratory in Lafayette, LA does the flow cytometry. Anne Grippo at Arkansas State University uses the propidium iodide technique, Stan Trauth does the sperm morphology with EM and FM. Also, the Dallas Zoo has specialized instrumentation specifically for sperm work, but the name of the instrument has slipped my mind! They are doing this on non-human animals, so I'm not sure how applicable this is to your question! Malcolm L. McCallum Associate Professor Department of Biological Sciences Texas AM University Texarkana 2600 Robison Rd. Texarkana, TX 75501 O: 1-903-334-6670 H: 1-903-791-3843 Homepage: https://www.eagle.tamut.edu/faculty/mmccallum/index.html VISIT HERPETOLOGICAL CONSERVATION AND BIOLOGY www.herpconbio.org A New Journal Published in Partnership with Partners in Amphibian and Reptile Conservation and the World Congress of Herpetology. Fall Teaching Schedule Office Hours: Ecology: M,W 1-2:40 pm Cell Biology: M 6-9:40 pm (don't ask!) Forensic Science: T,R 10-11:40am Office Hours: MW 12-1, 5-6, TR 11:40-12:30, We live in a time when lemonade is made with artificial flavoring, and furnisher polish is made with fresh lemons. -Alfred E. Neuman -Original Message- From: histonet-boun...@lists.utsouthwestern.edu on behalf of Anne van Binsbergen Sent: Wed 2/11/2009 6:45 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] semen analysis who out there is using automation for semen analysis and what equipment is recommended? i am considering the SQA-V from MES what say you? cheers -- Anne van Binsbergen (Hope) Abu Dhabi UAE ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet -- Anne van Binsbergen (Hope) Abu Dhabi UAE ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
