Re: Re: [Histonet] perfusion question
Hi, Because wash out solution can be 0.9% saline or 0.01 M PBSnear to physiological osmotic pressure... However the perfusion solution (4% PFA in 0.1M PB) is more concentrated, in the sense of osmotic pressureCan you comment on this? You can note the shrinkage of brain after PFA perfusion.~ 2008-12-12 tf 发件人： John Kiernan 发送时间： 2008-12-12 03:16:32 收件人： Neil Fournier 抄送： histonet@lists.utsouthwestern.edu 主题： Re: [Histonet] perfusion question The wash-out solution should have pH and osmotic pressure close to those of the animal's extracellular fluid, to avoid shrinkage or swelling of cells, collagen fibres etc. This can be achieved with simple saline (0.9% NaCl). A buffer prevents acidification of the extracellular fluid by products released from dying cells. Calcium ions (not compatible with phosphate buffers) enhance the preservation of phospholipids of cell membranes, myelin etc. Potassium ions are included in physiological saline solutions such as Ringer-Locke in which tissues and small organs can be kept alive, sometimes for several hours. I don't know of any study of effects of potassium on fixation, but probably someone has looked into it. The formaldehyde should also be dissolved in an isosmotic buffer because the chemical events of fixation occur slowly (several hours). Brain tissue still responds to changes in ambient osmotic pressure after several hours in neutral buffered formaldehyde. In glutaraldehyde, however, the cells are stabilized in 20 minutes. See: Paljarvi L, Garcia JH, Kalimo H (1979) The efficiency of aldehyde fixation for electron microscopy: stabilization of rat brain tissue to withstand osmotic stress. Histochem. J. 11: 267-276. This paper has also has references to several other studies. Traditional fixative mixtures are mostly acidic and rapidly acting, stabilizing the structure of the tissue (for light microscopy) before the development of adverse effects of low pH or osmotic pressure. The subject was also reviewed by J.R.Baker in his book Principles of Biological Microtechnique (1958), pp.75-86. John Kiernan Anatomy, UWO = = = - Original Message - From: Neil Fournier nfourn...@sasktel.net Date: Wednesday, December 10, 2008 14:42 Subject: [Histonet] perfusion question To: histonet@lists.utsouthwestern.edu Is there a rationale for using normal saline (0.9% (w/v) NaCl dissolved in dH2O) over 0.1 M PBS (pH 7.4) as a rinsing solution during intraventricular perfusion of a rat. Would one yield better results over the other? Also is there a raionale for why some people perfuse using PBS made only from monobasic and dibasic sodium phosphate (with 0.9% NaCl) vs. using PBS that also include KCl, sodium phosphate dibasic, NaCl, and potassium phosphate monobasic in the recipe. Thanks for the help Neil ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet - Original Message - From: Neil Fournier nfourn...@sasktel.net Date: Wednesday, December 10, 2008 14:42 Subject: [Histonet] perfusion question To: histonet@lists.utsouthwestern.edu Is there a rationale for using normal saline (0.9% (w/v) NaCl dissolved in dH2O) over 0.1 M PBS (pH 7.4) as a rinsing solution during intraventricular perfusion of a rat. Would one yield better results over the other? Also is there a raionale for why some people perfuse using PBS made only from monobasic and dibasic sodium phosphate (with 0.9% NaCl) vs. using PBS that also include KCl, sodium phosphate dibasic, NaCl, and potassium phosphate monobasic in the recipe. Thanks for the help Neil ___ 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] (reply) silly questions.---PFA
I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! Tony: Do you think this is because of inproper preparation of PFA in his lab, or the common problem in all researchers using PFA? I do think most biomedical labs currently are using PFA to prepare the fixatives! So, anyone has the idea on a correction preparation procedure of 4% PFA? I noticed some of you dissolve PFA powder in NaOH-conditioned alkaline water, then add concentrated PB solution. We here dissolve PFA in concentrated PB solution directly (heat stir for 2-3 hours), then adjust pH to 7.4. We dont have big problem in tissue quailityexcept when one want to cut the brain in a cryostat rather sliding microtome. Many times the brain sections from the cryostat have cheese like holes/cavities, which almost never appear on sliding microtome-prepared sections. 2008-12-12 tf 发件人： Tony Henwood 发送时间： 2008-12-12 06:18:47 收件人： Pat Flannery; histonet@lists.utsouthwestern.edu 抄送： 主题： RE: [Histonet] Silly Question? Pat, I agree with you. In a routine diagnostic histopathology laboratory, it makes little difference what you use. Around the world for over 100 years most labs use 10% neutral buffered formalin made from concentrated 38%(or there abouts) formalin (or formaldehyde). Researchers, though, are a different kettle of fish. They will tend to hang on to misinformed, mystical methods believing they are being scientific. Funny, you would think that they, as a group, would be the ones pushing the boundaries and critically assessing each step of their research, ensuring that they understand what and why they are doing it. (Disclaimer - not all researchers are like this, thank heavens!!) Using a formaldehyde solution made from polyformaldehyde can cause problems. One researcher used it and wondered why their morphology was sub-optimal and their p53 immunohistochemistry was negative. He assured me that he had fixed small samples of tissue for 6 hours in 4% formaldehyde and then processed them using ethanol, xylene and wax. I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! This also explains the loss of p53 staining. I gave him some of our routine 10% phosphate buffered fomalin, asked him to fix overnight, and try agin. Low and behold problem solved. How's that for a Friday Flamming!!! Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC) Laboratory Manager Senior Scientist Tel: 612 9845 3306 Fax: 612 9845 3318 the children's hospital at westmead Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, Westmead NSW 2145, AUSTRALIA -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: Friday, 12 December 2008 3:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I see all 4 on labels for specimens submitted for histology). Is it mostly a matter of personal preference or where you were trained (i.e. force of habit) or is there a valid reason to use each solution (basically the same chemical once in solution, merely buffered or not)? The only answer I've gotten when I've asked is, That's what we always use. Thanks. -Pat Flannery (not a real histologist - I just play one in the lab) ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet * This email and any files transmitted with it are confidential and intended solely for the use of the individual or entity to whom they are addressed. If you are not the intended recipient, please delete it and notify the sender. Views expressed in this message and any attachments are those of the individual sender, and are not necessarily the views of The Children's Hospital at Westmead This note also confirms that this email message has been virus scanned and although no computer viruses were detected, The Childrens Hospital at Westmead accepts no liability for any
[Histonet] SPECIAL STAIN PNUEMOCYSTSIS
We currently use a stain called grocotts silver methenimine solution for staining pnuemocystis. We would like to use a hot plate for the procedure instead of a unvented microwave. Does anybody have any formula's the would like to share with the hot plate? Timothy G. Malloy, HT ( ASCP ) A.A.S. tmallo...@hotmail.com This email communication may contain CONFIDENTIAL INFORMATION WHICH ALSO MAY BE LEGALLY PRIVILEGED and is intended only for the use of the intended recipients identified above. If you are not the intended recipient of this communication, you are hereby notified that any unauthorized review, use, dissemination, distribution, downloading, or copying of this communication is strictly prohibited. If you have received this communication in error, please immediately notify us by reply email, delete the communication and destroy all copies.___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] training materials
Not an immediate help, but there will be a NSH teleconference May 27, 2009 on embedding. PARAFFIN EMBEDDING AND PROCESS IMPROVEMENT Date: May 27, 2009 Time: 1:00PM EST Presented by Joelle Weaver, HTL(ASCP), Blanchard Valley Hospital, Findlay, Ohio; Riverside Methodist Hospital, Columbus, Ohio; Histology Education Coordinator, Columbus State Community College, Columbus, Ohio This presentation is a good review of fundamentals for everyone who routinely embeds a variety of tissue types. The embedding step is often overlooked in process improvement initiatives. At many labs, the embedding step has remained relatively static. What is being done at your facility during embedding to eliminate contamination as well as orientate tissue correctly and optimally for sectioning quality? This teleconference will begin with a review of the fundamentals of tissue embedding and orientation, covering basic principles of tissue sampling and specimen inking that may assist the embedding histologist. The impact of well infiltrated, correctly spaced, properly oriented specimens is strongly correlated to both ease of sectioning as well as section quality. Therefore, it is worth taking the time to consider this topic with fresh eyes and to re-evaluate embedding methodology when other changes are made in the overall process. For info on this and other NSH Teleconferences, go to: http://www.e-guana.net/organizations.php3?orgid=111typeID=1184action=print ContentTypeHomeUser_Session=41327fea4e36d34fede5e76bf55697d2 If having problems with the link, go to: www.nsh.org Click on Continuing Education on the left Click on Earn Continuing Education Click on Partipate in Teleconferences, about half way down the page The 2009 Schedule is available. All are on Wednesdays, from 1-2 pm Eastern time. Cost is $125 per teleconference, or if order all 11 by Jan. 27, 2009, it's only $1100, which is $100 each. You can have as many people listen as you want, and they all earn CE certificates. (If you have 10 people working in your lab, that bring the cost down to $12.50 per person for each to earn 1 hour CE. The Jan 2009 is Working with Difficult People, so invite everyone from all the labs and the pathology office and the pathologists, and everyone can learn to get along with earning 1 hour CE!) About 1 week before the teleconference, your lab will be sent a link to the PowerPoint and any additional handouts. The day of the talk, call up a phone number, listen to the speaker through speaker phone and be able to ask questions, and have everyone sign in, which is then faxed to NSH. Everyone who attended and signed in earns 1 hour CE. About 1-2 months after the teleconference, the lab will receive a CD with the PowerPoint, speaker's talks, any additional handouts, and a 4 question test. So if anyone couldn't attend the original teleconference, they can sit down when there is some free time in the lab, listen to the conference, view the PowerPoint, read the additional handouts, and now take a 4 question test, which they take and fax/mail into NSH, and they can get 1 hour CE, up to 2 years later. Also, your lab now has good training modules for anyone at any time. Peggy A. Wenk, HTL(ASCP)SLS Representing NSH as the NSH Teleconference Coordinator -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Jennifer Johnson Sent: Thursday, December 11, 2008 11:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] training materials Can anyone suggest a really good book, atlas, etc. for embedding? The girl that took my place at my last job is having a really hard time (especially with skin) and I told her I would ask the experts. Thanks, Jennifer Johnson, HTL (ASCP) _ Send e-mail faster without improving your typing skills. http://windowslive.com/Explore/hotmail?ocid=TXT_TAGLM_WL_hotmail_acq_speed_1 22008___ 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 wax
We have used Paraplast Plus for years. For both processing and embedding. Betsy Molinari HT(ASCP) Texas Heart Institute Cardiovascular Pathology 6770 Bertner Ave MC 1-283 Houston, TX 77030 832-355-6524 832-355-6812 (fax) -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Histonet Alias Sent: Thursday, December 11, 2008 9:07 AM To: Charles, Roger Cc: histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Paraffin wax I have had very good luck with Paraplast Plus for infiltration and using the Paraplast Xtra for embedding. On Thu, Dec 11, 2008 at 10:00 AM, Charles, Roger rchar...@state.pa.uswrote: Hello, Could someone share their knowledge of the Paraplast line of paraffin with me? We are being forced to change from the TissuePrep line. Thanks Roger Charles Microbiologist Pennsylvania Veterinary Laboratory 2305 N Cameron St Harrisburg, PA 17110 717-787-8808 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet -- Al Ias HT(ASCP) Histology Manager Pathology Laboratory United States ___ 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] (reply) silly questions.---PFA
This may be another silly question, but how does one test the concentration of formaldehyde in solution? Thanks, Kathy 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. -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of tf Sent: Friday, December 12, 2008 2:35 AM To: Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed Subject: [Histonet] (reply) silly questions.---PFA I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! Tony: Do you think this is because of inproper preparation of PFA in his lab, or the common problem in all researchers using PFA? I do think most biomedical labs currently are using PFA to prepare the fixatives! So, anyone has the idea on a correction preparation procedure of 4% PFA? I noticed some of you dissolve PFA powder in NaOH-conditioned alkaline water, then add concentrated PB solution. We here dissolve PFA in concentrated PB solution directly (heat stir for 2-3 hours), then adjust pH to 7.4. We dont have big problem in tissue quailityexcept when one want to cut the brain in a cryostat rather sliding microtome. Many times the brain sections from the cryostat have cheese like holes/cavities, which almost never appear on sliding microtome-prepared sections. 2008-12-12 tf 发件人： Tony Henwood 发送时间： 2008-12-12 06:18:47 收件人： Pat Flannery; histonet@lists.utsouthwestern.edu 抄送： 主题： RE: [Histonet] Silly Question? Pat, I agree with you. In a routine diagnostic histopathology laboratory, it makes little difference what you use. Around the world for over 100 years most labs use 10% neutral buffered formalin made from concentrated 38%(or there abouts) formalin (or formaldehyde). Researchers, though, are a different kettle of fish. They will tend to hang on to misinformed, mystical methods believing they are being scientific. Funny, you would think that they, as a group, would be the ones pushing the boundaries and critically assessing each step of their research, ensuring that they understand what and why they are doing it. (Disclaimer - not all researchers are like this, thank heavens!!) Using a formaldehyde solution made from polyformaldehyde can cause problems. One researcher used it and wondered why their morphology was sub-optimal and their p53 immunohistochemistry was negative. He assured me that he had fixed small samples of tissue for 6 hours in 4% formaldehyde and then processed them using ethanol, xylene and wax. I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! This also explains the loss of p53 staining. I gave him some of our routine 10% phosphate buffered fomalin, asked him to fix overnight, and try agin. Low and behold problem solved. How's that for a Friday Flamming!!! Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC) Laboratory Manager Senior Scientist Tel: 612 9845 3306 Fax: 612 9845 3318 the children's hospital at westmead Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, Westmead NSW 2145, AUSTRALIA -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: Friday, 12 December 2008 3:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I see all 4 on labels for specimens submitted for histology). Is it mostly a matter of personal preference or where you were trained (i.e. force of habit) or is there a valid reason to use each solution (basically the same chemical once in solution, merely buffered or not)? The only answer I've gotten when I've asked is, That's what we always use. Thanks. -Pat Flannery (not a real
[Histonet] re: concentration of formaldehyde in soultion: ASTM D2194 - 02(2007) Standard Test Method for Concentration of Formaldehyde Solution
http://www.astm.org/Standards/D2194.htm ASTM D2194 - 02(2007) ASTM D2194 - 02(2007) Standard Test Method for Concentration of Formaldehyde Solutions 2008-12-12 tf 发件人： Walters, Katherine S 发送时间： 2008-12-12 21:49:26 收件人： ti...@foxmail.com; Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed 抄送： 主题： RE: [Histonet] (reply) silly questions.---PFA This may be another silly question, but how does one test the concentration of formaldehyde in solution? Thanks, Kathy 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. -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of tf Sent: Friday, December 12, 2008 2:35 AM To: Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed Subject: [Histonet] (reply) silly questions.---PFA I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! Tony: Do you think this is because of inproper preparation of PFA in his lab, or the common problem in all researchers using PFA? I do think most biomedical labs currently are using PFA to prepare the fixatives! So, anyone has the idea on a correction preparation procedure of 4% PFA? I noticed some of you dissolve PFA powder in NaOH-conditioned alkaline water, then add concentrated PB solution. We here dissolve PFA in concentrated PB solution directly (heat stir for 2-3 hours), then adjust pH to 7.4. We dont have big problem in tissue quailityexcept when one want to cut the brain in a cryostat rather sliding microtome. Many times the brain sections from the cryostat have cheese like holes/cavities, which almost never appear on sliding microtome-prepared sections. 2008-12-12 tf 发件人： Tony Henwood 发送时间： 2008-12-12 06:18:47 收件人： Pat Flannery; histonet@lists.utsouthwestern.edu 抄送： 主题： RE: [Histonet] Silly Question? Pat, I agree with you. In a routine diagnostic histopathology laboratory, it makes little difference what you use. Around the world for over 100 years most labs use 10% neutral buffered formalin made from concentrated 38%(or there abouts) formalin (or formaldehyde). Researchers, though, are a different kettle of fish. They will tend to hang on to misinformed, mystical methods believing they are being scientific. Funny, you would think that they, as a group, would be the ones pushing the boundaries and critically assessing each step of their research, ensuring that they understand what and why they are doing it. (Disclaimer - not all researchers are like this, thank heavens!!) Using a formaldehyde solution made from polyformaldehyde can cause problems. One researcher used it and wondered why their morphology was sub-optimal and their p53 immunohistochemistry was negative. He assured me that he had fixed small samples of tissue for 6 hours in 4% formaldehyde and then processed them using ethanol, xylene and wax. I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! This also explains the loss of p53 staining. I gave him some of our routine 10% phosphate buffered fomalin, asked him to fix overnight, and try agin. Low and behold problem solved. How's that for a Friday Flamming!!! Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC) Laboratory Manager Senior Scientist Tel: 612 9845 3306 Fax: 612 9845 3318 the children's hospital at westmead Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, Westmead NSW 2145, AUSTRALIA -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: Friday, 12 December 2008 3:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I
RE: [Histonet] (reply) silly questions.---PFA
Oh, I'd like to know that, too, please! --On Friday, December 12, 2008 7:48 AM -0600 Walters, Katherine S katherine-walt...@uiowa.edu wrote: This may be another silly question, but how does one test the concentration of formaldehyde in solution? Thanks, Kathy 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. -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of tf Sent: Friday, December 12, 2008 2:35 AM To: Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed Subject: [Histonet] (reply) silly questions.---PFA I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! Tony: Do you think this is because of inproper preparation of PFA in his lab, or the common problem in all researchers using PFA? I do think most biomedical labs currently are using PFA to prepare the fixatives! So, anyone has the idea on a correction preparation procedure of 4% PFA? I noticed some of you dissolve PFA powder in NaOH-conditioned alkaline water, then add concentrated PB solution. We here dissolve PFA in concentrated PB solution directly (heat stir for 2-3 hours), then adjust pH to 7.4. We dont have big problem in tissue quailityexcept when one want to cut the brain in a cryostat rather sliding microtome. Many times the brain sections from the cryostat have cheese like holes/cavities, which almost never appear on sliding microtome-prepared sections. 2008-12-12 tf ·¢¼þÈË£º Tony Henwood ·¢ËÍʱ¼ä£º 2008-12-12 06:18:47 ÊÕ¼þÈË£º Pat Flannery; histonet@lists.utsouthwestern.edu ³ËÍ£º Ö÷Ì⣺ RE: [Histonet] Silly Question? Pat, I agree with you. In a routine diagnostic histopathology laboratory, it makes little difference what you use. Around the world for over 100 years most labs use 10% neutral buffered formalin made from concentrated 38%(or there abouts) formalin (or formaldehyde). Researchers, though, are a different kettle of fish. They will tend to hang on to misinformed, mystical methods believing they are being scientific. Funny, you would think that they, as a group, would be the ones pushing the boundaries and critically assessing each step of their research, ensuring that they understand what and why they are doing it. (Disclaimer - not all researchers are like this, thank heavens!!) Using a formaldehyde solution made from polyformaldehyde can cause problems. One researcher used it and wondered why their morphology was sub-optimal and their p53 immunohistochemistry was negative. He assured me that he had fixed small samples of tissue for 6 hours in 4% formaldehyde and then processed them using ethanol, xylene and wax. I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! This also explains the loss of p53 staining. I gave him some of our routine 10% phosphate buffered fomalin, asked him to fix overnight, and try agin. Low and behold problem solved. How's that for a Friday Flamming!!! Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC) Laboratory Manager Senior Scientist Tel: 612 9845 3306 Fax: 612 9845 3318 the children's hospital at westmead Cnr Hawkesbury Road and Hainsworth Street, Westmead Locked Bag 4001, Westmead NSW 2145, AUSTRALIA -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: Friday, 12 December 2008 3:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I see all 4 on labels for specimens submitted for histology). Is it mostly a matter of personal preference or where you were trained (i.e. force of habit) or is there a valid reason to use each solution (basically the same chemical once in
[Histonet] RE: Histonet Digest, Vol 61, Issue 20
Hi - Here we perform the silver step with a hot plate under the hood. As the slides are going into the sodium metabisulfite step, we turn on a hot plate on high. Then we mix the silver solution according to the procedure, put it in a beaker on the hot plate, rinse our slides with microwaved hot DI water a few times, leaving them in hot DI water. We gently swirl the silver solution as it heats and just when it starts to turn color (right before boiling), we dump the hot water off the slides and pour on the silver. The tissue usually stains within 45-60 seconds, a little longer if your silver is not really hot. Rinse with hot DI water afterwards. I was a little dubious when I first heard about it, but we too, do not have a vented microwave, using it only to heat water. This works great, though no one here could tell us who came up with the idea. You do have to use hot water before and after to keep from breaking coplin jars or slides from drastic temp changes. Terri L. Braud, HT(ASCP) Anatomic Pathology Supervisor Laboratory Holy Redeemer Hospital and Medical Center 1648 Huntingdon Pike Meadowbrook, PA 19046 (215) 938-3676 phone (215) 938-3689 fax -Original Message- Message: 9 Date: Fri, 12 Dec 2008 04:52:19 -0500 From: TIMOTHY MALLOY tmallo...@hotmail.com Subject: [Histonet] SPECIAL STAIN PNUEMOCYSTSIS We currently use a stain called grocotts silver methenimine solution for staining pnuemocystis. We would like to use a hot plate for the procedure instead of a unvented microwave. Does anybody have any formula's the would like to share with the hot plate? Timothy G. Malloy, HT ( ASCP ) A.A.S. - CONFIDENTIALITY NOTICE: This E-Mail is intended only for the use of the individual or entity to which it was sent. It may contain information that is privileged and/or confidential, and the use or disclosure of such information may also be restricted under applicable federal and state law. If you received this communication in error, please do not distribute any part of it or retain any copies, and delete the original E-Mail. Please notify the sender of any error by E-Mail. Thank you for your cooperation. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] regarding problems with decalcification of rat maxilla's
Hi, I am a graduate student at University of Illinois, chicago. I have been having tremendous problems with sectioning of paraffin embedded rat maxilla with molars and I searched the Histonet archives and found that people here have invaluable experience, insight and suggestions for various problems. Heres my problem. : As I section the block the section would crumble and tear and has a dusty dry look to it and if I run my fingers over the block I can clean off the dusty apperance of the block. (does that mean it wasnt decalcified well, or paraffin infiltration wasnt enough? or is it possible there is some EDTA salts or left over paraformaldehyde but I did wash the samples with water after being in paraformaldehyde and EDTA ) After decalcification the samples felt very soft and I could bend the maxilla and were flexible and could be cut with an ordinary blade. Here is the protocol I used for decalcifying the samples and the processing details. I would really appreciate if you guys can give me some suggestions as to what I can do about these samples and what must have went worng. Here's the method that I used for rat maxilla's with teeth in them. 1. Fixed in 4% paraformaldehyde for 4 days and then washed with distilled water. 2. Decalcified using 8% EDTA with change to fresh solution everyday for 3 weeks using the microwave technique. (eg: the maxilla from one rat were decalcified in 40 mL EDTA) 3. After decalcification the samples were washed in distilled water for couple of hours with fresh changes every 30 mins. 4. The samples were then transferred to 50% ethanol for an hour and then 70% ethanol for an hour and then overnight in 70% ethanol. 5. Then 95% ethanol for 1.5 hours with fresh changes every 30 mins 6. Then 100% ethanol for 1.5 hours with fresh changes every 30 mins 7. Then half 100% ethanol half 100% xylene for 20 mins 8. Then Xylene for 3 hours with fresh change every 45 mins (samples could very clear after this step). 9. Half Xylene half paraffin for 30 mins. 10. 100% paraffin (paraplast plus) for 3 hours with fresh change every 45 mins and then embedded in paraffin. I would really appreciate if you guys would guide me as to what I can do so that I can use these samples to cut sections. Thankyou for your time and help. Sincerely, Smit. Add more friends to your messenger and enjoy! Go to http://messenger.yahoo.com/invite/ ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] 7B11 clone of Ki67
Good afternoon, 'netters, I am in desperate need of Ki67 antibody, clone 7B11 preferably, to use on Ventana instruments. I didn't realize we were running low and our usual vendor has no stock. I'd prefer a predilute but only if it is known to work on our instruments. I don't have the time to search this out myself so I am turning to all of you. Thanks, Linda Linda A. Sebree University of Wisconsin Hospital Clinics IHC/ISH Laboratory DB1-223 VAH 600 Highland Ave. Madison, WI 53792 (608)265-6596 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] 7B11 clone of Ki67
Why not use the (30-9) clone? It is much easier to come by. I sue it on my Benchmark, cat# 790-4286. On Fri, Dec 12, 2008 at 1:45 PM, Sebree Linda A lseb...@uwhealth.orgwrote: Good afternoon, 'netters, I am in desperate need of Ki67 antibody, clone 7B11 preferably, to use on Ventana instruments. I didn't realize we were running low and our usual vendor has no stock. I'd prefer a predilute but only if it is known to work on our instruments. I don't have the time to search this out myself so I am turning to all of you. Thanks, Linda Linda A. Sebree University of Wisconsin Hospital Clinics IHC/ISH Laboratory DB1-223 VAH 600 Highland Ave. Madison, WI 53792 (608)265-6596 ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet -- Al Ias HT(ASCP) Histology Manager Pathology Laboratory United States ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Silly Question?
So I have a question about the cross-linking aspect of PFA...while I agree I need it to keep my epitope in place, is there such a thing as OVER-crosslinking (i.e., tissue spending TOO much time in formalin - weeks? months?) that would make my epitope difficult to near-impossible to retrieve? Loving the formaldehyde soap-boxes histonetters get onto... --On Friday, December 12, 2008 10:36 AM -0700 pru...@ihctech.net wrote: i have had this argument with the researchers at the University for 30 years, somewhere back in the day they were told that commercially made formalin had methanol in it (it does but just a little and does not hurt anything in my experience) and that methanol would damage their tissue for IHC, so they think they must use paraformaldehyde and make it fresh themselves. Since new people make it all the time it often does not get made up correctly and their stress over this issue is miss placed as they should be using something commercial for consistancy and paying more attention to adequate time for fixation in reg formalin. Another anoying myth that is difficult to combat with them is that we should limit the fixation time in aldehyde fixatives because it will cross link the proteins masking them for IHC, there fore i am always getting tissue that has not been fixed long enough (at least 24 hrs. to protect it from paraffin processing, because if the proteins are not cross linked they can be alcohol fixed and/or washed away forever), the people in research know about the cross linking fo aldehydes but do not know that cross linking of proteins is a good thing and they also do not know that we have advanced methods HIER or EIER for unmasking the proteins, but we have no way of getting a protein back that has been lost in processing because the sample was not adequately fixed. there i will get off my Friday soap box.. Happy Holidays to all! Patsy Original Message Subject: RE: [Histonet] Silly Question? From: Merced Leiker lei...@buffalo.edu Date: Fri, December 12, 2008 8:12 am To: Edwards, R.E. r...@leicester.ac.uk, 'Pat Flannery' pjfne...@duke.edu Cc: 'histonet@lists.utsouthwestern.edu' histonet@lists.utsouthwestern.edu In research lab situations particularly, one does not have the time or technique for nailing down the ways of making each of the buffers, reagents, and procedures work the right way or the most optimum way...a lot of times it's students or postdocs just focused on getting their project done and not caring how their fixative is made as long as it works to some degree and, alas, it's up to us already over-booked technicians to figure out the best way to make the PFAand we usually don't have a whole day (week, or year) to spend researching the back-and-forth arguments, either! ;-) Merced --On Friday, December 12, 2008 2:04 PM + Edwards, R.E. r...@leicester.ac.uk wrote: You hit the nail on the head That's what we always use, fear of change is a common human condition. -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: 11 December 2008 16:59 To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I see all 4 on labels for specimens submitted for histology). Is it mostly a matter of personal preference or where you were trained (i.e. force of habit) or is there a valid reason to use each solution (basically the same chemical once in solution, merely buffered or not)? The only answer I've gotten when I've asked is, That's what we always use. Thanks. -Pat Flannery (not a real histologist - I just play one in the lab) ___ 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 Merced M Leiker Research Technician II 354 BRB (pkgs) / 140 Farber Hall (letters) School of Medicine and Biomedical Sciences State University of New York at Buffalo 3435 Main St, Buffalo, NY 14214 Ph: (716) 829-6033 Fx: (716) 829-2725 Without my flaws I'm really very boring. - random internet blog commentator ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet Merced M Leiker Research Technician II 354 BRB (pkgs) / 140 Farber Hall (letters) School of Medicine and Biomedical Sciences State
[Histonet] CD123 and TCL-1
Anyone doing IHC for CD123 and TCL-1 on formalin-fixed, paraffin-embedded tissue? Richard Richard W. Cartun, Ph.D. Director, Histology Immunopathology Assistant Director, Anatomic Pathology Hartford Hospital 80 Seymour Street Hartford, CT 06102 (860) 545-1596 (860) 545-0174 Fax ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Biogenex
Would anyone using the Biogenix Xmatrix be willing to speak with me about your experience? I'd appreciate your feedback! Thanks, j Joyce Weems Pathology Manager Saint Joseph's Hospital 5665 Peachtree Dunwoody Rd NE Atlanta, GA 30342 678-843-7376 - Phone 678-843-7831 - Fax Confidentiality Notice: This email, including any attachments is the property of Catholic Health East 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 reply to the sender that you have received the message in error, then delete this message. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
Re: [Histonet] re: concentration of formaldehyde in soultion: ASTM D2194 - 02(2007) Standard Test Method for Concentration of Formaldehyde Solution
I believe that the swiss cheese holes are due to ice crystal formation during freezing, at least that's the rationale we were always taught for using additional fixation in sucrose-formalin after the initial fixation in formalin, i.e. the sucrose would prevent ice crystal formation. Susan - Original Message - From: tf ti...@foxmail.com To: Walters, Katherine S katherine-walt...@uiowa.edu; Tony Henwood antho...@chw.edu.au; Pat Flannery pjfne...@duke.edu; histo...@lists.utsouthwestern.ed histonet@lists.utsouthwestern.edu Sent: Friday, December 12, 2008 10:12 AM Subject: [Histonet] re: concentration of formaldehyde in soultion: ASTM D2194 - 02(2007) Standard Test Method for Concentration of Formaldehyde Solution http://www.astm.org/Standards/D2194.htm ASTM D2194 - 02(2007) ASTM D2194 - 02(2007) Standard Test Method for Concentration of Formaldehyde Solutions 2008-12-12 tf 发件人： Walters, Katherine S 发送时间： 2008-12-12 21:49:26 收件人： ti...@foxmail.com; Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed 抄送： 主题： RE: [Histonet] (reply) silly questions.---PFA This may be another silly question, but how does one test the concentration of formaldehyde in solution? Thanks, Kathy 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. -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of tf Sent: Friday, December 12, 2008 2:35 AM To: Tony Henwood; Pat Flannery; histo...@lists.utsouthwestern.ed Subject: [Histonet] (reply) silly questions.---PFA I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! Tony: Do you think this is because of inproper preparation of PFA in his lab, or the common problem in all researchers using PFA? I do think most biomedical labs currently are using PFA to prepare the fixatives! So, anyone has the idea on a correction preparation procedure of 4% PFA? I noticed some of you dissolve PFA powder in NaOH-conditioned alkaline water, then add concentrated PB solution. We here dissolve PFA in concentrated PB solution directly (heat stir for 2-3 hours), then adjust pH to 7.4. We dont have big problem in tissue quailityexcept when one want to cut the brain in a cryostat rather sliding microtome. Many times the brain sections from the cryostat have cheese like holes/cavities, which almost never appear on sliding microtome-prepared sections. 2008-12-12 tf 发件人： Tony Henwood 发送时间： 2008-12-12 06:18:47 收件人： Pat Flannery; histonet@lists.utsouthwestern.edu 抄送： 主题： RE: [Histonet] Silly Question? Pat, I agree with you. In a routine diagnostic histopathology laboratory, it makes little difference what you use. Around the world for over 100 years most labs use 10% neutral buffered formalin made from concentrated 38%(or there abouts) formalin (or formaldehyde). Researchers, though, are a different kettle of fish. They will tend to hang on to misinformed, mystical methods believing they are being scientific. Funny, you would think that they, as a group, would be the ones pushing the boundaries and critically assessing each step of their research, ensuring that they understand what and why they are doing it. (Disclaimer - not all researchers are like this, thank heavens!!) Using a formaldehyde solution made from polyformaldehyde can cause problems. One researcher used it and wondered why their morphology was sub-optimal and their p53 immunohistochemistry was negative. He assured me that he had fixed small samples of tissue for 6 hours in 4% formaldehyde and then processed them using ethanol, xylene and wax. I looked at the sections and the cell shrinkage (and prominent spaces between cells and connective tissue) indicated that most of the fixation seemed to have occured in the processing ethanols. I asked him for some of the fixative he used, tested the formaldehyde concentration and found it to be less than 0.5%!! This also explains the loss of p53 staining. I gave him some of our routine 10% phosphate buffered fomalin, asked him to fix overnight, and try agin. Low and behold problem solved. How's that for a Friday Flamming!!! Regards Tony Henwood JP, MSc, BAppSc, GradDipSysAnalys, CT(ASC) Laboratory Manager Senior Scientist Tel: 612 9845 3306 Fax: 612 9845 3318 the children's hospital at westmead Cnr
[Histonet] Re: Reduction of autofluorescence using glycine
To reduce aldehyde induced autofluorescence, you can use 100 - 300 mM glycine in pH 7.4 buffer. TRIS buffer or even Dulbeccos PBS will work. You rehydrate the section and then immerse into the glycine solution for 20 minutes, maybe even longer. Glycine works by getting rid (binding?) of free aldehyde groups. You can either treat the tissue prior to processing (after fixation) by immersing for an hour or so, but we simply did the glycine treatment on individual sections. It worked best for us when we did a short length fixation in NBF. This has been discussed at length on Histonet in the past, so do an archive search. One person put a summary together on various methods and what worked best for him. There are other methods for getting rid of autofluorescence although some are less successful than others and one is made from a chemical that is explosive. Try IHCworld website, fluorescence topics or Google access this discussion written by Wright Cell Imaging Faculty, Toronto Western Research Institute, titled: Autofluorescence, Causes and Cures, a must read on the subject. Another trick is to use fluorophores in the near infrared range, the camera sees the fluorescence but no autofluorescence and you cannot see this red fluorophore with the naked eye. Alexa 750 will work if you have the filters and excitation wavelength available. Good luck Gayle M. Callis HTL(ASCP)HT,MT Bozeman MT ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] mouse dendritic cells
Is anyone staining for mouse dendritic cells either in paraffin or frozen sections? If so, what markers/antibodies are you using? Thanks, ANDREA -- ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Mucin blocking reagent for IHC
I am staining CA19-9 on pancreas and have a good amount of mucin trapping the DAB. Is there a product that either blocks mucin or something else that will limit the mucin effect? Aprill Watanabe, B.S. Research Associate Integrated Cancer Genomics Division Tissue Microarray Center (TMA) Translational Genomics Research Institute (TGen) main: 602-343-8822 Fax: 602-343-8840 awatan...@tgen.org www.tgen.org ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Re: rat maxilla processing
Dear Smit You wrote: I am a graduate student at University of Illinois, chicago. I have been having tremendous problems with sectioning of paraffin embedded rat maxilla with molars and I searched the Histonet archives and found that people here have invaluable experience, insight and suggestions for various problems. Heres my problem. : As I section the block the section would crumble and tear and has a dusty dry look to it and if I run my fingers over the block I can clean off the dusty apperance of the block. (does that mean it wasnt decalcified well, or paraffin infiltration wasnt enough? or is it possible there is some EDTA salts or left over paraformaldehyde but I did wash the samples with water after being in paraformaldehyde and EDTA ) After decalcification the samples felt very soft and I could bend the maxilla and were flexible and could be cut with an ordinary blade. Here is the protocol I used for decalcifying the samples and the processing details. I would really appreciate if you guys can give me some suggestions as to what I can do about these samples and what must have went worng. Here's the method that I used for rat maxilla's with teeth in them. 1. Fixed in 4% paraformaldehyde for 4 days and then washed with distilled water. 2. Decalcified using 8% EDTA with change to fresh solution everyday for 3 weeks using the microwave technique. (eg: the maxilla from one rat were decalcified in 40 mL EDTA) 3. After decalcification the samples were washed in distilled water for couple of hours with fresh changes every 30 mins. 4. The samples were then transferred to 50% ethanol for an hour and then 70% ethanol for an hour and then overnight in 70% ethanol. 5. Then 95% ethanol for 1.5 hours with fresh changes every 30 mins 6. Then 100% ethanol for 1.5 hours with fresh changes every 30 mins 7. Then half 100% ethanol half 100% xylene for 20 mins 8. Then Xylene for 3 hours with fresh change every 45 mins (samples could very clear after this step). 9. Half Xylene half paraffin for 30 mins. 10. 100% paraffin (paraplast plus) for 3 hours with fresh change every 45 mins and then embedded in paraffin. I would really appreciate if you guys would guide me as to what I can do so that I can use these samples to cut sections. Thankyou for your time and help. Sincerely, Smit.Reply: You bone sample is underprocessed, poorly infiltrated with paraffin and may not be totally decalcified. When you do the fixation, fix longer than 4 days, as rat maxilla is not a small sample and complicated by teeth in situ. It would be better to perfuse the animal with your PFA solution, then dissect off the maxilla, and immerse into fresh PFA for several days. If perfused then 4 days may be adequate. Rinse with running tap water, and immerse into your EDTA (what is pH?) Also, use 20 times the volume of EDTA to size of tissue, and change the EDTA after a couple of days to replenish the EDTA. EDTA is safe to use over a weekend so you do not have to change it so often. You need to do endpoint determination to know when the bone is decalcified and I will be happy to send a simple weight loss/weight gain method that works for EDTA or use a FAXITRON xray determination. After EDTA, rinse with running tap water for several hours, not distilled water . Please contact me for the endpoint determination, I will send as a separate attachment. To process, start the bone in 70% alcohol. The time changes in each of the alcohols, clearing agent and paraffin are the same. 2 hours per change (30 minutes per change for a total of 1.5 hours is not doing the job!) If you only have a small portion of the maxilla, you can reduce the time to 1.5 hours per change. I hope you have a tissue processor to do this for you, as you need to use vacuum and pressure for the best results. 70% ethanol80% ethanol95% ethanol X 2 changes100% ethanol X 2 changesXylene X 2 changes and to get rid of brittleness, use one change of Richard Allan Clearite 3, then 1 change of xylene. You can reverse the order of these two clearing agents without problems. Paraffin X 3 changes (minimum) at times suggested at NO more than 60C. Heat is an enemy that contributes to hardness of decalcified bone. To achieve vacuum pressure with hand processing, use a vacuum dessicator, and hopefully a heated vacuum oven for the paraffin steps. Use a harder paraffin - there are several on the marked. Tissue Prep 3 is excellent for bone as you want to try and match the hardness of your embedding media to hardness of decalcified bone as much as possible. What is meant by an ordinary blade? A low profile or high profile disposable blade? or are you using a c profile steel knife? High profile blades give much more stability when sectioning bone, and after a very careful trim of block face, soak the block on ice water for a short time, return to holder, and use a brand new sharp edge to section. Do NOT cut away what you have
Re: [Histonet] mouse dendritic cells
Andrea, We stain for murine dendritic cells very frequently. CD11c (HL3 clone) is excellent from BD Biosciences using fresh snap frozen tissue with spleen as the positive control. Fixation is with our beloved 25% ethanol/75% acetone for 5 min at RT then going directly to buffer from fixative. The sections are air dried overnight before fixation. Dec 205, NLDC 145 from Serotec works but we have superior results with the BD Pharm rat antiMouse CD11c HL3 clone. There are a whole series of new DC antibodies out there, used by our FACS tech, but I have not tried them on frozen sections yet. I would still used the same tissue preparation and fixation if we did try them but also toss in cold acetone just for posteriety sake. The CD11c we work with is biotinylated, which simplifies everything since we come back with Streptavidin Alexa dyes for single and double immunofluorescence work these days. It is an Armenian hamster host IgG1. Happy Holidays to you Gayle Callis HTL(ASCP)HT,MT Bozeman MT - Original Message - From: Andrea Hooper anh2...@med.cornell.edu To: Histonet histonet@lists.utsouthwestern.edu Sent: Friday, December 12, 2008 3:11 PM Subject: [Histonet] mouse dendritic cells Is anyone staining for mouse dendritic cells either in paraffin or frozen sections? If so, what markers/antibodies are you using? Thanks, ANDREA -- ___ 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] Silly Question? - Need help quickly!
I tried earlier to send, for this thread, a wonderful paper by Nauta, chronicaling the history his discovery of his tract tracing method, in which serendipity and degradation of formalin played critical roles, not realizing that the size of the attachment would prevent it from going through, so I am trying again with a URL for this paper: http://www.jneurosci.org/cgi/reprint/13/4/1337 Susan - Original Message - From: Rene J Buesa rjbu...@yahoo.com To: Pat Flannery pjfne...@duke.edu; histonet@lists.utsouthwestern.edu; Weems, Joyce jwe...@sjha.org Sent: Thursday, December 11, 2008 12:58 PM Subject: RE: [Histonet] Silly Question? - Need help quickly! Joyce: Methanal, which is the chemical name of formaldehyde, polymerizes. If it forms a polymer of at least 50 molecules or more, it gets solid = para-formaldehyde. Formalin (a trade name as formol is also another trade name)is the 37-50% aqueous solution of formaldehyde (with some additiveses to prevent polymerization). You can prepare BNF using the formalin solution or dissolving the amount of solid para-formaldehydede to get to the concentrationon you desire. The chemical in both solutions is the same = methanal or formaldehyde.René J. --- On Thu, 12/11/08, Weems, Joyce jwe...@sjha.org wrote: From: Weems, Joyce jwe...@sjha.org Subject: RE: [Histonet] Silly Question? - Need help quickly! To: Pat Flannery pjfne...@duke.edu, histonet@lists.utsouthwestern.edu Date: Thursday, December 11, 2008, 12:12 PM I was just going to post a question regarding paraformaldhyde myself! Just last week I believe I remember someone saying that paraformaldehyde and formalin are the same and they had put the same solution in two different containers for one of their researchers because they were so insistent to have two different solutions. Are they the same? Well, today I have a request to put tissue for a researcher in formalin and paraformaldehyde. So Without percentage required, do I use 10% NBF? Do I call somewhere and get paraformaldehyde and make 4% paraformaldehyde? I have asked the surgeon twice for the number for the lab so I can find out - don't have it yet. I have two fresh adrenals in the fridge. Help!! Thanks in advance... Joyce Joyce Weems Pathology Manager Saint Joseph's Hospital 5665 Peachtree Dunwoody Rd NE Atlanta, GA 30342 678-843-7376 - Phone 678-843-7831 - Fax -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Pat Flannery Sent: Thursday, December 11, 2008 11:59 AM To: histonet@lists.utsouthwestern.edu Subject: [Histonet] Silly Question? Please humor me on this if it's obvious (to everyone but me): why do we use paraformaldehyde (which is so inconvenient to make up) rather than buffered formalin or just diluted formaldehyde itself? It seems that around here, some folks prefer paraformaldehyde (either 2% or 4%) and others use formalin, while some others stick to diluted formaldehyde (I see all 4 on labels for specimens submitted for histology). Is it mostly a matter of personal preference or where you were trained (i.e. force of habit) or is there a valid reason to use each solution (basically the same chemical once in solution, merely buffered or not)? The only answer I've gotten when I've asked is, That's what we always use. Thanks. -Pat Flannery (not a real histologist - I just play one in the lab) ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet Confidentiality Notice: This email, including any attachments is the property of Catholic Health East 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 reply to the sender that you have received the message in error, then delete this message. ___ 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] Re: Reduction of autofluorescence using glycine
No, because the glycine acts by reducing the autofluorescene of the free aldehydes (maybe Dr. Kiernan or another knowledgeable person in the chemistry can tell us precisely how) rather than reducing the binding of other staining components to the aldehydes. -Original Message- From: Bob Nienhuis bob.nienh...@gmail.com Date: Fri, 12 Dec 2008 17:16:27 To: Gayle Callisgayle.cal...@bresnan.net Cc: histo...@pathology.swmed.edu Subject: Re: [Histonet] Re: Reduction of autofluorescence using glycine If this works by binding free aldehyde groups that attach to antibodies/ or fluorochromes, or biotinylated whatever. shouldn't it also work for DAB or ABC immunolabeling and reduce background labeling? Bob UCLA / VA Medical Center On Fri, Dec 12, 2008 at 2:08 PM, Gayle Callis gayle.cal...@bresnan.netwrote: To reduce aldehyde induced autofluorescence, you can use 100 - 300 mM glycine in pH 7.4 buffer. TRIS buffer or even Dulbeccos PBS will work. You rehydrate the section and then immerse into the glycine solution for 20 minutes, maybe even longer. Glycine works by getting rid (binding?) of free aldehyde groups. You can either treat the tissue prior to processing (after fixation) by immersing for an hour or so, but we simply did the glycine treatment on individual sections. It worked best for us when we did a short length fixation in NBF. This has been discussed at length on Histonet in the past, so do an archive search. One person put a summary together on various methods and what worked best for him. There are other methods for getting rid of autofluorescence although some are less successful than others and one is made from a chemical that is explosive. Try IHCworld website, fluorescence topics or Google access this discussion written by Wright Cell Imaging Faculty, Toronto Western Research Institute, titled: Autofluorescence, Causes and Cures, a must read on the subject. Another trick is to use fluorophores in the near infrared range, the camera sees the fluorescence but no autofluorescence and you cannot see this red fluorophore with the naked eye. Alexa 750 will work if you have the filters and excitation wavelength available. Good luck Gayle M. Callis HTL(ASCP)HT,MT Bozeman MT ___ 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
