Re: [Histonet] dehydration time for a relatively large sample
Thank you for your advice! I was going to try if the current sample that was embedded by a technician at the histological service center. This is the first time that i asked for an expert to prepare my sample. Then I sectioned the sample by myself. However, the muscles in the section always looks very dry or are unable to be penetrated with wax, thus the sections do not look that great I am not an expert at all for preparing histological sections, so if the sample embedded by an expert, I am not sure how to fix the problem. I may try it again following your time schedule, but i am a bit tight schedule to end this project. If i try i will let you know how it turns out. I currently reside in montreal, Canada and am studying as a PhD student. I really appreciate your help. rui From: k...@regionsjaelland.dk To: ru...@hotmail.com Subject: SV: [Histonet] dehydration time for a relatively large sample Date: Thu, 21 May 2015 13:24:56 + Hi Rui Sorry for a little late response. I tried to write to you yesterday but my internet failed. Tissue is so much easier with a processor, but life is not always easy and we have to do things the possible way ;-) First I have to tell you, that I never have tried to process tissue by hand. and I have never tried to process zebra finch. BUT I have been helping lot of other people with protocols, so I think I'm able to help you too. maybe we need to adjust the protocol. ... But lets see how it will work. First make sure your tissue are well fixed before decalcination. 1 day in formalin. Decalcinate the tissue as short time as possible but make sure it's with out calcium before processing. 70% ethanol 3 hours 96 % ethanol 5-7 hours 99 % over night ( about 12 hours ) 99 % so it will fit your work ( change 99% 2 times) Clearens over night (about 12 hours ) Clearens so it will fit your work ( change clearens 2 times ) Paraffin 2-3 days ( change paraffin 3 times ) When you embed the tissue please notise if you see lot of small bobles. The bobles can be a sign of poorly infiltretet paraffin (water in the tissue) Please let my know how things go, because we maybe need some adjustment. I would like to know where you are from. I have helped all over the would and it's fun to know. Kind regards Karen it's___ Fra: Rui TAHARA [ru...@hotmail.com] Sendt: 20. maj 2015 15:44 Til: histonet@lists.utsouthwestern.edu Emne: Re: [Histonet] dehydration time for a relatively large sample Hi, Thanks for prompt response. We unfortunately do not have a processor in our lab at university..The protocol i wrote was working in quail embryonic samples (just before hatching). I process the tissue manually and cannot process the tissue with strict time schedule. Thus, I need to leave a sample overnight at some point. I cut the zebra finch head into anterior (beak) and posterior (brain) region and mid-sagittally in both. so each tissue sample is about 0.5 X 1 X 1cm cube at maximum. I have also tried short time schedule compared the one i wrote in previous email, for similar sized sample (zebra finch beak). However, it never worked. Thus i prolonged the each step for the latest sample. It would be great if you could provide me the proper time schedule. rui From: k...@regionsjaelland.dk To: ru...@hotmail.com Subject: SV: [Histonet] dehydration time for a relatively large sample Date: Wed, 20 May 2015 06:18:24 + Hi Rui Which processor unit do you use to me it seems like a wrong protocol. Maybe I can help you set up a better protocol - if you want - but then I need to know size og your tissue, processor unit and what kind of clearens you use. Kind regards Karen Supervisor tissue processing Denmark -Oprindelig meddelelse- Fra: Rui TAHARA [mailto:ru...@hotmail.com] Sendt: Wednesday, May 20, 2015 7:11 AM Til: histonet@lists.utsouthwestern.edu Emne: [Histonet] dehydration time for a relatively large sample Hi, I am wondering if prolong dehydration time with 95 and 100% ethanol would brittle the sample for paraffin sectioning. I have been trying to section adult zebra finch beak and processed several samples, however, i failed to obtain a good section. It appears that the paraffin did not penetrated the tissue. This may be derived from incomplete dehydration and clearing before paraffin. Because i have read somewhere that if the sample was sitting in 95 and 100% ethanol too long it would be brittle and be teared when sectioned. I have processed only a beak (about 1 cmX0.5 mm). Also when is an appropriate time to use vacuum? I am afraid that if i used it at 100% ethanol, the ethanol would evaporate so far I have tried; fix overnight decalcified few days walk from water to 70% ethanol; overnight 85%, 95% (2times change) and 100 % (2 times change
Re: [Histonet] dehydration time for a relatively large sample
Hi, Thanks for prompt response. We unfortunately do not have a processor in our lab at university..The protocol i wrote was working in quail embryonic samples (just before hatching). I process the tissue manually and cannot process the tissue with strict time schedule. Thus, I need to leave a sample overnight at some point. I cut the zebra finch head into anterior (beak) and posterior (brain) region and mid-sagittally in both. so each tissue sample is about 0.5 X 1 X 1cm cube at maximum. I have also tried short time schedule compared the one i wrote in previous email, for similar sized sample (zebra finch beak). However, it never worked. Thus i prolonged the each step for the latest sample. It would be great if you could provide me the proper time schedule. rui From: k...@regionsjaelland.dk To: ru...@hotmail.com Subject: SV: [Histonet] dehydration time for a relatively large sample Date: Wed, 20 May 2015 06:18:24 + Hi Rui Which processor unit do you use to me it seems like a wrong protocol. Maybe I can help you set up a better protocol - if you want - but then I need to know size og your tissue, processor unit and what kind of clearens you use. Kind regards Karen Supervisor tissue processing Denmark -Oprindelig meddelelse- Fra: Rui TAHARA [mailto:ru...@hotmail.com] Sendt: Wednesday, May 20, 2015 7:11 AM Til: histonet@lists.utsouthwestern.edu Emne: [Histonet] dehydration time for a relatively large sample Hi, I am wondering if prolong dehydration time with 95 and 100% ethanol would brittle the sample for paraffin sectioning. I have been trying to section adult zebra finch beak and processed several samples, however, i failed to obtain a good section. It appears that the paraffin did not penetrated the tissue. This may be derived from incomplete dehydration and clearing before paraffin. Because i have read somewhere that if the sample was sitting in 95 and 100% ethanol too long it would be brittle and be teared when sectioned. I have processed only a beak (about 1 cmX0.5 mm). Also when is an appropriate time to use vacuum? I am afraid that if i used it at 100% ethanol, the ethanol would evaporate so far I have tried; fix overnight decalcified few days walk from water to 70% ethanol; overnight 85%, 95% (2times change) and 100 % (2 times change) ethanol; overnight clearing; 2 days paraffin (2 times change); overnight Any suggestion would be appreciated. Thank you in advance, rui ___ 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] dehydration time for a relatively large sample
Hi, I am wondering if prolong dehydration time with 95 and 100% ethanol would brittle the sample for paraffin sectioning. I have been trying to section adult zebra finch beak and processed several samples, however, i failed to obtain a good section. It appears that the paraffin did not penetrated the tissue. This may be derived from incomplete dehydration and clearing before paraffin. Because i have read somewhere that if the sample was sitting in 95 and 100% ethanol too long it would be brittle and be teared when sectioned. I have processed only a beak (about 1 cmX0.5 mm). Also when is an appropriate time to use vacuum? I am afraid that if i used it at 100% ethanol, the ethanol would evaporate so far I have tried; fix overnight decalcified few days walk from water to 70% ethanol; overnight 85%, 95% (2times change) and 100 % (2 times change) ethanol; overnight clearing; 2 days paraffin (2 times change); overnight Any suggestion would be appreciated. Thank you in advance, rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] soften beak
Thank you for suggestions for softening beak. Yes, I actually processed the whole head for decalcifying first and then place the sample in 10% KOH for 30 mins. It was my first attempt to process beak (including a whole head) thus, i did not know i should have processed softening beak first. Anyway, after placing the beak in 10% KOH for 30 mins, it did not seem to soften enough. However, I did not overly place it in solution because it may destroy the microscopic structure. How soft the nail becomes after 10 % KOH or Nair treatment, such as you can bent the nail easily etc..? Any suggestion would be appreciated. rui *** From: ru...@hotmail.com To: histonet@lists.utsouthwestern.edu Date: Fri, 10 Apr 2015 05:23:01 +0900 Subject: [Histonet] soften beak Hi, I am just wondering what you would recommend to soften avian beak for paraffin processing. I found a protocol that KOH could be used to soften keratin in nail, however, I have not been able to find that this method is applicable specific to beak. Another concern is if i use KOH for an entire head, does KOH affect (e.g. damage) soft tissues? Thank you in advance, rui ___ 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] soften beak
Hi, I am just wondering what you would recommend to soften avian beak for paraffin processing. I found a protocol that KOH could be used to soften keratin in nail, however, I have not been able to find that this method is applicable specific to beak. Another concern is if i use KOH for an entire head, does KOH affect (e.g. damage) soft tissues? Thank you in advance, rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification
Thank you for all the helpful suggestions about this topic. My sample has been in the fixative until it would be decalcified. Thank you again, rui From: r...@leicester.ac.uk To: ru...@hotmail.com Subject: RE: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification Date: Tue, 10 Mar 2015 09:49:00 + Leave it in formalin for as long as possible..good luck -Original Message- From: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] On Behalf Of Rui TAHARA Sent: 04 March 2015 23:39 To: gayle.cal...@bresnan.net; histonet@lists.utsouthwestern.edu Subject: RE: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification Thank you for helpful suggestions. I have further questions. Yes, I have a bird head (probably 1 cm X 1 cm ) stored in 70 % ethanol. But i have a similar size bird head fixed in 3.7% formalin for over night and am actually processing the head to store in 70% ethanol since my lab is just ordering the decalcifying solution. I need to decalcify this sample later. But i am wondering if it is better to keep the sample in formalin for a week or so till i get the decalcification solution or i should store it in 70 % ethanol and then fix it for a few days again later? I am afraid that longer fixative time would affect the sample somehow (e.g. the sample become too rigid?) Thank you, rui From: gayle.cal...@bresnan.net To: histonet@lists.utsouthwestern.edu Date: Wed, 4 Mar 2015 16:25:28 -0700 Subject: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification You wrote: I have an adult bird skull that fixed with formalin and then has been stored in 70% ethanol. I have seen the post that the sample stored in 70% ethanol can be walking back through to series of ethanol to water and can be decalcified if it needs to be. I am wondering if anybody has done this and there is any side effects from decalcification after going through dehydration and rehydration of a sample compared to a general straight forward protocol from decalcification to dehydration? ** ** ** ** I have, in the past, when a weekend arrive, I interrupted acid bone decalcification by removing it from acid decalcifier, a quick water rinse and immersed into 70% alcohol before returning bone to fresh acid decalcifier the next working day. The bones always decalcified without problems but I am sure the decalcification took longer since partially decalcified bone had to rehydrate. I later learned more about dipolar (hope I said that correctly) alcohol slowing and/or stopping ionization of calcium and ceased using 70% alcohol to interrupt acid decalcification. I now use NBF to interrupt decalcification. Interestingly, I learned the alcohol technique from the AFIP bone pathology lab. Alcohol is put into Perenyi's nitric acid decalcifying solutions to slow down or control very rapid nitric acid decalcification. You did not say how big the bird skull was? I suggest immersing the skull back into NBF to let it totally rehydrate for several days (depending on skull size and if the brain is present). I suggest changing NBF if you rehydrate longer than a day. You don't need to go back through an alcohol gradient since many processing schedules have tissue samples going from NBF directly into 70%. If you leave residual alcohol in the bones, the acid decalcification could be slower and hopefully not retarded in any way. It certainly is worth a try. Good luck. Gayle M. Callis HTL/HT/MT(ASCP) ___ 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] decalcification after dehydration
I have an adult bird skull that fixed with formalin and then has been stored in 70% ethanol. I have seen the post that the sample stored in 70% ethanol can be walking back through to series of ethanol to water and can be decalcified if it needs to be. I am wondering if anybody has done this and there is any side effects from decalcification after going through dehydration and rehydration of a sample compared to a general straight forward protocol from decalcification to dehydration? Thank you, rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification
Thank you for helpful suggestions. I have further questions. Yes, I have a bird head (probably 1 cm X 1 cm ) stored in 70 % ethanol. But i have a similar size bird head fixed in 3.7% formalin for over night and am actually processing the head to store in 70% ethanol since my lab is just ordering the decalcifying solution. I need to decalcify this sample later. But i am wondering if it is better to keep the sample in formalin for a week or so till i get the decalcification solution or i should store it in 70 % ethanol and then fix it for a few days again later? I am afraid that longer fixative time would affect the sample somehow (e.g. the sample become too rigid?) Thank you, rui From: gayle.cal...@bresnan.net To: histonet@lists.utsouthwestern.edu Date: Wed, 4 Mar 2015 16:25:28 -0700 Subject: [Histonet] Re: Bird head stored in 70% alcohol and possible decalcification You wrote: I have an adult bird skull that fixed with formalin and then has been stored in 70% ethanol. I have seen the post that the sample stored in 70% ethanol can be walking back through to series of ethanol to water and can be decalcified if it needs to be. I am wondering if anybody has done this and there is any side effects from decalcification after going through dehydration and rehydration of a sample compared to a general straight forward protocol from decalcification to dehydration? I have, in the past, when a weekend arrive, I interrupted acid bone decalcification by removing it from acid decalcifier, a quick water rinse and immersed into 70% alcohol before returning bone to fresh acid decalcifier the next working day. The bones always decalcified without problems but I am sure the decalcification took longer since partially decalcified bone had to rehydrate. I later learned more about dipolar (hope I said that correctly) alcohol slowing and/or stopping ionization of calcium and ceased using 70% alcohol to interrupt acid decalcification. I now use NBF to interrupt decalcification. Interestingly, I learned the alcohol technique from the AFIP bone pathology lab. Alcohol is put into Perenyi's nitric acid decalcifying solutions to slow down or control very rapid nitric acid decalcification. You did not say how big the bird skull was? I suggest immersing the skull back into NBF to let it totally rehydrate for several days (depending on skull size and if the brain is present). I suggest changing NBF if you rehydrate longer than a day. You don't need to go back through an alcohol gradient since many processing schedules have tissue samples going from NBF directly into 70%. If you leave residual alcohol in the bones, the acid decalcification could be slower and hopefully not retarded in any way. It certainly is worth a try. Good luck. Gayle M. Callis HTL/HT/MT(ASCP) ___ 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] Help for identifying the blue stained structure
Hi, Thank you so much. I was wondering why the image has not been seen on the site, and thought its been processed. Thank you for uploading the image. Rui Date: Sun, 18 May 2014 17:47:21 -0400 From: mha...@histosearch.com To: ru...@hotmail.com; histonet@lists.utsouthwestern.edu Subject: Re: [Histonet] Help for identifying the blue stained structure Hi Rui, You uploaded a tif image and a number of browsers don't support tif images. Jpeg, gif and png images are the best image formats to use because they are universally supported. I converted your image to a jpeg image and posted it at: http://histosearch.com/imageupload/help-for-identifying-the-blue-stained-structure-jpeg/ Best Regards, Marvin Hanna On 05/17/2014 06:39 PM, Rui TAHARA wrote: Hello, I have an embryonic sample that decalcified, paraffin embedded, and stained with Mallory Trichrome (Aniline Blue, Orange G, Acid Fuchsin). I will upload the image in the Histonet Images. This is a cranial region where the bone is being resorbed, so I expected to see the bone (dark blue in trabecular), and red blood, and adjacent white spaces that is being resorbed. Instead, there is a very uniform, granular structure stained with blue without any nuclei at the resorbed regions. This structure looks like almost crystal or some kind of secretion leakage from the ossifying bone. I need a help to identify this blue stained things. I don’t think this is osteoid, because at later stage embryos, there is no bone at this region. If you have some suggestions for other stain to identify this blue thing, or help identifying this, I really appreciate it. Thank you in advance, Rui ___ 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] Help for identifying the blue stained structure
Hello, I have an embryonic sample that decalcified, paraffin embedded, and stained with Mallory Trichrome (Aniline Blue, Orange G, Acid Fuchsin). I will upload the image in the Histonet Images. This is a cranial region where the bone is being resorbed, so I expected to see the bone (dark blue in trabecular), and red blood, and adjacent white spaces that is being resorbed. Instead, there is a very uniform, granular structure stained with blue without any nuclei at the resorbed regions. This structure looks like almost crystal or some kind of secretion leakage from the ossifying bone. I need a help to identify this blue stained things. I don’t think this is osteoid, because at later stage embryos, there is no bone at this region. If you have some suggestions for other stain to identify this blue thing, or help identifying this, I really appreciate it. Thank you in advance, Rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] variable staining of alcian blue
Hello, I am staining paraffin sections (10 microns) with Alcian blue and Nuclear Fast Red as standard protocol as below. The problem is that I see some sections stained with alicain blue show some variability (hope i am able to attach image); alcian blue stain are washed out in some sections even in the same slide. Those sections show pink color in the cartilage. Clearing dehydrate 1% Alcain blue (PH2.5) 30 mins dip in tap water running water 2mins dip deionized water 0.1% Nuclear Fast red 5 mins dip deionized water 30 sec X 2 95% ethanol 1min 100% ethanol 1 mins X 2 clearing Is this something to do with filtering alcian blue or nuclear fast red? I filtered alcian blue using needles to eliminate a big undisolved particle, but I did not filter the nuclear fast red since it seems all dissolved well. also I rarely had the problem last time i used the same staining. I know i need to filter with filter paper but when i tried, even with non-fine filter paper, it absorbed too much dye and was not successful. I tried to wash nuclear fast red with running tap water 1 min and dip in deionized water intead of twice wash of deionized water as described above. However, variable staining of alcian blue still present. Does anybody have same problem before? I appreciate any suggestion to solve this. Thanks. Rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] microtome
Hello, I have undecalcified biological sample embedded in plastic media (MMA). I am looking for faclities that offers self-service microtome (for plastic embedding sectioining) or short time rental microtome with some training around Quebec, Ontraio, NY. I have been trying to find ones in Montreal, however, its been difficult to find self-service microtome. I appreciate if you would provide some information for this. Thank you in advance, Rui ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: [Histonet] Undecalcified sample in paraffin and plastic media
Thank you for your information. Since in our lab we have never used MMA and also no vacuum I decided to ask histological service to embed the sample.I will section it and stain them with Von kossa/Alcian blue by myself. The sample is being processed now and i will see if the images of sample embeded in MMA would work better than the paraffin one for my project. The technician told me its hard to obtain a good section embedded in MMA compared to paraffin. Would you give me some tips or protocol for MMA sectioning? Do i need to use adhesive to place the MMA section on the slide? In general does the staining time for paraffin sample work for MMA sample? Thank you, Rui Date: Wed, 2 Oct 2013 11:46:22 -0500 Subject: Re: [Histonet] Undecalcified sample in paraffin and plastic media From: ratliffj...@gmail.com To: ru...@hotmail.com Rui, Did you need any additional assistance? Please let me know if there is anything I can do to be of assistance to you. Best Regards, Jack Jack L Ratliff, Owner/HistologistRatliff Histology Consultants, LLC389 Nichol Mill LaneFranklin, TN 37067 (615) 236-4901 (o)(615) 236-4962 (f)(317) 281-1975 (c) ratliffjack@gmail.comjratl...@ratliffhistology.com jratl...@ratliffhistology.com (coming soon) On Mon, Sep 30, 2013 at 7:49 AM, Jack Ratliff ratliffj...@hotmail.com wrote: Rui, You will definitely want to consider using plastic media like methyl methacrylate (MMA). It will cause less shrinkage in the tissue during polymerization, you can still cut at a range of 4-12 microns using a rotary microtome and tungsten-carbide knife, any mineralization present in the tissue will infiltrate and polymerize well allowing for enhanced stabilization of tissue and section morphology throughout microtomy, and you can even deplastify the sections with certain MMA formulations to increase staining options. Please let me know if you do wish to continue with plastic media as I have helped many labs to get started with and/or to refine their current capabilities with MMA. Additionally, I would like to point out that I Chair the Hard Tissue Committee (HTC) for the National Society for Histotechnology (NSH). Membership with the NSH has several benefits that could also help you to move forward with your project at your own pace. For example, as a member you will have access to all archived publications of the Journal of Histotechnology (JOH). With this access to the JOH via Manny Publishing, the HTC has created a reference document that collates all relevant publications (1970's to present) that pertain to bone, biomaterials, medical device implants, resin histology, etc., so that one can easily locate and obtain publication information relevant to their niche specific needs. Rest assured that I will be happy to help you either way you choose to move forward. Best Regards, Jack On Sep 23, 2013, at 9:19 PM, Rui TAHARA ru...@hotmail.com wrote: I have undecalcified paraffin embed samples that were sectioned at 10 micron that I want to stain with Von kossa. Because samples are embryonic quail heads (ossification starts to happen) and still soft enough to section with standard rotary microtome with tungsten knife in paraffin. My intention is to 3D reconstruct anatomies based on histological sections. Because of this, I am wondering if I should actually use plastic media rather than paraffin to keep the section shape as consistent as possible. Does plastic embed material actually preserve the consistent shape among sections better than paraffin embed sample? No winkle etc..? Is there any other advantage that I actually should use the plastic media than paraffin for what I want to do? I know downside of plastic media is that in general plastic embedding process are lengthy and plastic embedding material are expensive than the paraffin ones, and are mainly use for bone to support the hard material for sectioning. When I sectioned some ossified samples, beak start to fall off from section and the section show the lines from the possibly scratched knife. Is this indication of paraffin media that does not provide enough strength for sectioning? I thought it may possibly the poor infiltration. In our lab nobody has processed the plastic embedding and sectioning (we have only standard microtome, no vaccum machine. Can I section plastic embed sample with the standard microtome at 10 micron?) so I would like to have any input before actually making a plastic embed sample. Any suggestions would be appreciated. Rui TAHARA Biology Department McGill University ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet ___ Histonet mailing list Histonet
[Histonet] Undecalcified sample in paraffin and plastic media
I have undecalcified paraffin embed samples that were sectioned at 10 micron that I want to stain with Von kossa. Because samples are embryonic quail heads (ossification starts to happen) and still soft enough to section with standard rotary microtome with tungsten knife in paraffin. My intention is to 3D reconstruct anatomies based on histological sections. Because of this, I am wondering if I should actually use plastic media rather than paraffin to keep the section shape as consistent as possible. Does plastic embed material actually preserve the consistent shape among sections better than paraffin embed sample? No winkle etc..? Is there any other advantage that I actually should use the plastic media than paraffin for what I want to do? I know downside of plastic media is that in general plastic embedding process are lengthy and plastic embedding material are expensive than the paraffin ones, and are mainly use for bone to support the hard material for sectioning. When I sectioned some ossified samples, beak start to fall off from section and the section show the lines from the possibly scratched knife. Is this indication of paraffin media that does not provide enough strength for sectioning? I thought it may possibly the poor infiltration. In our lab nobody has processed the plastic embedding and sectioning (we have only standard microtome, no vaccum machine. Can I section plastic embed sample with the standard microtome at 10 micron?) so I would like to have any input before actually making a plastic embed sample. Any suggestions would be appreciated. Rui TAHARA Biology Department McGill University ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Alizarin red/Alcian blue on sections
Does anybody have protocol for Alizarin red/Alcian blue (also counterstain with something else?) on sections. All I can find is either stain or double staining on whole mounts. Thank you in advance, Rui TAHARA PhD student McGill University ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Bone/cartilage/epithelial tissue stain
Hi, Would anyone suggest me what staining is best to color differentiate between cartilage and bone and epithelial tissues in avian embryos? I have been trying Mallory Trichrome for embryos but recently I was suggested that Mallory Trichrome stains cartilage differently in embryos compared to adult samples since Aniline blue stains fiber that may not develop in early embryos. There is some protocol that modified the Mallory Trichrome that could be applied to embryos. However, the resulting colors of all tissues look all purple-ish and difficult to tell the cartilage from the weak blue stain from aniline blue. Currently I am thinking to try out Alcian blue/Hematoxylin and Eosin stain (Ehrlich’s hematoxylin). The purpose of the staining is to look at interaction between ossification and epithelial development so I think alcian blue for staining cartilage works but I am wondering if there is any other staining combination with alcian blue exist for visualizing bone and epithelial tissue (e,g. alcian blue/alizarine red with other staining?). Any suggestion would be appreciated! Rui TAHARA PhD Candidate Biology Department McGill University ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
RE: AW: [Histonet] post-fixation for Mallory Trichrome
Thank you! I have added extra step of 5 min of 1% phosphomolybdic before aniline blue-orangeG, which increased the blue contrast in late stage embryos but not early stage ones. I did not realize that Mallory trichrome react differently between embryonic and adult samples. I just assumed the formalin fixation is causing the staining problem for early embryos. Does formalin fixation cause the different colors or weak staining? If the Mallory Trichrome works differently with developing stages, is mallory trichrome preferably used for adult samples and late stage samples? I found the following ref about modified Mallory Trichrome for embryonic samples. This method is basically use three dyes separately that can help differentiate staining colors like optimized colors achieved in adult samples. I have tried the protocol yet the cartilage does not show sharp enough and most of other tissues are stained purple-ish. Would you suggest some articles that i can refer to for Mallory trichrome in embryos? Thank you, Rui From: gu.l...@gmx.at To: ru...@hotmail.com CC: histonet@lists.utsouthwestern.edu Subject: AW: [Histonet] post-fixation for Mallory Trichrome Date: Sat, 17 Aug 2013 09:26:38 +0200 Hi, I think staining would be improved, when you use Bouin as postfixative ( 1h hour, 60°C). Try to invite a further step between Fuchsin and Anilinblue. When you incubate the slides in 1% phosphomolybdic acid for 5-10 min, you can see decolorization of cardilage. This improves later on anilin-binding. You may control the differentiation-step under microscope. Anilinblue can be reduced to 5 min afterwards. You may also consider, that there's a real difference in cardilage-architecture of the early and late samples, that takes influence on staining-abilities. Gudrun Lang -Ursprüngliche Nachricht- Von: histonet-boun...@lists.utsouthwestern.edu [mailto:histonet-boun...@lists.utsouthwestern.edu] Im Auftrag von Rui TAHARA Gesendet: Freitag, 16. August 2013 23:33 An: histonet@lists.utsouthwestern.edu Betreff: [Histonet] post-fixation for Mallory Trichrome Hello, There seems to be a lot of suggestions for Mallory Trichrome staining involved in Acid fusin, Aniline blue, and orange G. My sample of avian embryos were fixed with Formalin based fixative (4% paraformaldehyde in phosphate-buffered saline and 1% glutaraldehyde) overnight, in case of late embryos bones were decalcified, and followed standard alcohol series, paraffin embedded, and 10 micron sectioned. The slides were dehydrated, stained with 0.5 % Acid fushin for 10min, then without wash, 0.5% Aniline blue and 2 % orange G in 1% phosphomolybdic acid solution for 20 min, then undergone ethanol series, cleared and mounted. Now I know that formalin fixed sample do not present optimized trichrome colors based on your websites and references. Yet the stained sections of late stage embryo do still show near optimized colors whereas those of early stage embryo show almost no blue or very dark blue, almost gray color for cartilages and most of morphologies as purple-redish colors. Since I tested staining on late embryos first I thought staining would work on early embryos. Does anyone provide me explanation why the staining mostly shows red-ish color and not optimized color for cartilage in early embryos? Is that because of the formalin-fixed embryos although late stage embryos fixed with formalin still show the blue for cartilage? Another question is related to fixative. I prefer not to use bousin fixative due to picric acid and it seems that Citrate buffer or Gram’s iodine solution can be substituted to bousin post-fixative. Have anyone tried these solution for Mallory Trichrome? Do those methods show near optimized color for Mallory trichrome? Any suggestion is appreciated. Thank you, Rui TAHARA PhD candidate McGill University, Biology Department ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] post-fixation for Mallory Trichrome
Hello, There seems to be a lot of suggestions for Mallory Trichrome staining involved in Acid fusin, Aniline blue, and orange G. My sample of avian embryos were fixed with Formalin based fixative (4% paraformaldehyde in phosphate-buffered saline and 1% glutaraldehyde) overnight, in case of late embryos bones were decalcified, and followed standard alcohol series, paraffin embedded, and 10 micron sectioned. The slides were dehydrated, stained with 0.5 % Acid fushin for 10min, then without wash, 0.5% Aniline blue and 2 % orange G in 1% phosphomolybdic acid solution for 20 min, then undergone ethanol series, cleared and mounted. Now I know that formalin fixed sample do not present optimized trichrome colors based on your websites and references. Yet the stained sections of late stage embryo do still show near optimized colors whereas those of early stage embryo show almost no blue or very dark blue, almost gray color for cartilages and most of morphologies as purple-redish colors. Since I tested staining on late embryos first I thought staining would work on early embryos. Does anyone provide me explanation why the staining mostly shows red-ish color and not optimized color for cartilage in early embryos? Is that because of the formalin-fixed embryos although late stage embryos fixed with formalin still show the blue for cartilage? Another question is related to fixative. I prefer not to use bousin fixative due to picric acid and it seems that Citrate buffer or Gram’s iodine solution can be substituted to bousin post-fixative. Have anyone tried these solution for Mallory Trichrome? Do those methods show near optimized color for Mallory trichrome? Any suggestion is appreciated. Thank you, Rui TAHARA PhD candidate McGill University, Biology Department ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
