Re: [Histonet] p2y12 cryo rat problem high background
Hi Esther, I worked extensively with brain and spinal cord sections in the past, in multiple species. You need endogenous quenching step. I would always make my own (commerically available components yielded different results). I used a 0.09% H2O2 in 6% Triton-X 100: (formula: 200mL 1x PBS + 6mL H2O2 + 0.33mL 6% Triton-X 100 solution). Are you samples fixed frozen or fresh frozen? If fixed-frozen, there is no need for the acetone step. In fact, acetone in brain and spinal cord sections makes the tissue brittle. I would recommend this is skipped. If your samples are fresh-frozen, I would recommend a 95% EtOH fix. This is significantly gentler on the tissue. I have no experience with the Envision system, but the polymer system I would always use was from Cell Signaling Technologies. What antibody are you using? On Thu, Nov 15, 2018 at 7:48 AM Kooijman, E.J.M. (Esther) via Histonet < histonet@lists.utsouthwestern.edu> wrote: > Hello Bobbie, > > But should I elimination endogenous peroxidase activity in brain/spinal > cord tissues? > Brains and spinal cord was harvested after cervical dislocation, then the > tissue was snap frozen (isopentane..). > > thanks for your help, > Esther > > -Oorspronkelijk bericht- > Van: Boyce, Bobbie [mailto:bobbie.bo...@nemours.org] > Verzonden: donderdag 15 november 2018 12:21 > Aan: Kooijman, E.J.M. (Esther) > Onderwerp: RE: p2y12 cryo rat problem high background > > Hi Ester, > Try Peroxoblock (Zymed) before your BSA block, but you have to be careful > not to leave it on too long or it will eat your tissue. It's been a while > since I've had to used it. > > > Bobbie Boyce > Histology Specialist III > DuPont Experimental Station > Nemours- Biomedical Research Department > Histochemistry and Tissue Processing Core > 200 Powder Mill Road, Bldg.400 Rm.5240 > Wilmington, DE 19803 > > (lab) 302-651-6771 > (fax) 302-651-5010 > > > > -Original Message- > From: Kooijman, E.J.M. (Esther) via Histonet < > histonet@lists.utsouthwestern.edu> > Sent: Thursday, November 15, 2018 5:53 AM > To: histonet@lists.utsouthwestern.edu > Subject: [Histonet] p2y12 cryo rat problem high background > > **This is an External Email - Please DO NOT open attachments or click > links from unknown senders or unexpected email. ** > > Hello all, > > > > I am trying to stain cryo brain sections from the rat 7um but having a lot > of background. What am I doing wrong. Below the protocol is used and tried > to adjust… > > > > 1- Fix the tissue with cold acetone (-20 ⁰C) for 10 minutes > > > > 2- Let the slide dry for 30 minutes at room temperature. Isolate the > sections with DAKO pen > > > > 3- Block sections with BSA 2% in PBS for 1 hour at room temperature > > > > 4- Add primary antibody in PBS/BSA 1%, overnight at 4⁰C or 1h at room > temperature > > > > 5- Wash 3x5 minutes with PBS/tween-20 0.05% > > > > 6- Add 100 µL Envision solution (goat anti-mouse/rabbit) and incubate > for 1hr at RT > > > > 7- Wash 3x 5 minutes with PBS/tween-20 0.05% > > > > 8- Incubate with DAB (1:50) for 10 min (between 5-10 min; check color > development) > > (wear gloves, carcinogenic!) > > > > 9- Rinse thoroughly with miliQ water > > > > 10- Stain with haematoxylin for 1 min > > > > 11- Wash with running tap water for 5 min > > > > 12- Start the alcohol/xylene series (70% EtOH -> 80% EtOH -> 96% EtOH -> > 100% EtOH -> 100% EtOH/xylene -> xylene -> xylene) > > > > 13- Mount slides with entallan > > Kind regards, > > > > > > Esther Kooijman | Research Technician | Department of Radiology and > Nuclear medicine > > The Netherlands > > > > ___ > Histonet mailing list > Histonet@lists.utsouthwestern.edu > > https://urldefense.proofpoint.com/v2/url?u=http-3A__lists.utsouthwestern.edu_mailman_listinfo_histonet&d=DwIGaQ&c=X2IGR6v8ax_mLhSmU1r3Aw&r=NAc-sZTcJtcdymsLif_fp7ngB_dNBpF-UI64u9fuosc&m=N05bQ_qJ2Li62f83kZl-FK7rp5Ad-spj9JTvwovnnKQ&s=6MYtZ-ICVPPxaohoUjidJjm_mNhwB7nDA1Np5SRxT4w&e= > ___ > 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 with frozen spleen and liver tissue!
Hi all, Can someone please provide me with more details regarding cryosectioning of mouse spleen and liver tissue? Currently, I've been fixing my samples in 4% PFA (they are well fixed - I know that will be the first question asked!), and then cryoprotect the tissue in a series of graded sucrose solutions (15% to 30%) until they sink. However, when I go to place the sections on the slide from the cryostat, they look great initially under the microscope, but once they dry they have poor morphology, especially seen in the liver. I've never run into this issue before, with either brain or spinal cord which are significantly more delicate. I've tried cutting the tissue free-floating as well to see if it was how I was placing the sections on the slide, but nothing seems to work. The morphology in the liver is so terribly compromised that I cannot visualize the sinusoids properly. It is baffling that once they go onto the slide they look okay, but 5 minutes later the tissue appears to "separate" from itself. Does anyone work specifically in frozen tissue sections, liver and spleen in particular? If so, would you be able to help me figure out the best way to generate quality specimens? Thank you! Regards, Allyse Mazzarelli ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Help with liver/heart tissues
Good afternoon, I am having a few issues getting nice morphology from both liver and heart tissues. I currently work only with CNS tissues (brain, spinal cord, DRG, etc.) but recently, our research team have become interested in immunostaining some peripheral tissues, including the heart and liver. All tissue [mouse] is perfusion fixed with 4% PFA and then post-fixed in fresh 4% PFA for at least 24 - 48 hours. After fixation, the tissue is cryoprotected in 30% sucrose solution for 48 hours. Once cryoprotected, I then section the tissue at various thicknesses depending on the assay I'm running. Previously I sectioned some heart and liver samples (sectioned both at 20um and at 8um) and ran a few H&Es. Unfortunately, the tissue was so damaged and under-fixed that we scrapped the blocks. This time around, the mouse liver tissue was carefully dissected prior to post-fixation into four quadrants to allow for better PFA permealization. Additionally, we post fixed in 4% PFA for 3 days instead of 2, and the tissue was cryoprotected for 2 days. To my surprise, when I sectioned this tissue on the cryostat, I still noticed severe artifacts. It is very difficult to see nice morphology, and there appears that there was an issue with fixation (in the liver the nucleus is flattened and the sinusoids are not clear, etc. the nuclei in the heart are also more flat and the muscle fibers have separated from one another). The staining was not as bad as the first tissue I had sectioned, but I was still unable to get a nice H&E stain depicting clear nuclear/cytoplasm. These artifacts appeared at both 8um and 20um. Does anyone happen to have nice fixation/H&E staining protocols for both liver and heart? I'd be happy to give an in-depth description of the protocol(s) I'm currently using. Additionally, is there anything that appears I'm doing wrong in terms of perfusion/fixation? Thanks so much! Allyse ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] H&E Protocol for Fresh-Frozen Brain and Spinal Cord
Hi Histonet! I was wondering if anyone could provide me with a reliable H&E protocol for fresh frozen tissue. Currently, we cut our slides on the cryostat and store them at either -20 or -80 (depending on the study director) until they require us to stain for morphology. Typically the tissue isn't always perfectly snap frozen, and often times there are freezing artifacts. However, I would like to preserve as much as the morphology as possible without further damaging the tissue. I have really only had experience staining perfusion-fixed tissue using the following staining procedure: Water rinse Hematoxylin (7211 Richard Allen Scientific) for 30 seconds Water rinse under running tap Clarifier 1 (Richard Allen Scientific) with three quick dips Water rinse under running tap Bluing reagent (Richard Allen Scientfic) for 1 minute Water rinse under running tap 95% EtOH for 30 seconds Eosin-Y with Phyloxine (Richard Allen Scientific) for two quick dips 95% EtOH for 30 seconds 100% EtOH for 1 minute 100% EtOH for 1 minute 3, one-minute exchanges in Clear Rite 3 (a xylene substitute, Richard Allen Scientific) However, today I was asked to run an H&E on a fresh frozen slide. I post fixed in 4% PFA for 10 minutes immediately (not allowing the slide to thaw), and then ran the above procedure as normal. When I examined the slides under the scope, there appeared to be terrible freezing artifacts. The study director happened to think it was both a post-fix and staining issue, so I re-ran the procedure without post-fixing, (per his recommendation) as follows: Immediately into hematoxylin for 30 seconds Water rinse under running tap Eosin for 2 quick dips 95% EtOH for 1 minute 100% EtOH for 1 minute 100% EtOH for 1 minute 3, one-minute exchanges in Clear Rite 3 I saw slightly better morphology, but the director still was not happy with the outcome. I have spent the remainder of my afternoon researching different H&E protocols for fresh frozen tissue, and all seem to be relatively similar to the two mentioned prior. Does anyone out there in Histo-land consistently run H&E on fresh frozen tissue, specifically on slides that have been stored for a while? I work particularly with brain and spinal cord, but we occasionally will cut other tissue and organs. I would like to optimize this procedure as soon as possible. Thank you in advance! Regards, Allyse Mazzarelli ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Autofluorescence on Retina Tissue
Hi all! Question for all you that may be more immuno-experienced than I: I've consistently run immunofluorescence on pig retina and I seem to have a severe case of autofluorescence/background. I've played around with primary and secondary antibody ratios, but that doesn't seem to help my case. The primary antibodies I use are goat-anti-FLT.1, and mouse-anti-rhodopsin. The secondary antibodies I use are AlexaFluor donkey-anti-goat488 & rabbit-anti-mouse555 (For my experiments, each slide contained only one primary antibody, and it's corresponding secondary. For imaging purposes, I did not double-label on these slides. E.g. FLT.1 was labeled with the 488 secondary, and rhodopsin was labeled with the 555 secondary).I re-hydrate, conduct antigen retrieval, and block as per normal IHC protocol. However, when imaging, I noticed that both slides, labeled with either rhodopsin or FLT.1 seem to "bleed" through to the next filter. For example, mouse-anti-rhodopsin labeled with the 555 secondary works beautifully at a 1:600 ratio. However, when I switch to the FITC filter on my scope, all the retinal tissue appears green on the slides, even though it has ONLY the 555 secondary and NO 488. I've noticed this for the FLT.1 antibody as well (i.e. switch to red filter and tissue fluoresces even though no slide saw the 555 secondary antibody). As I mentioned, I decreased the ratios of all antibodies, but that still doesn't eliminate the problem. If anyone has any ideas as to how I go about fixing this, please let me know. I've only been in the field for a very short period of time, so if I missed something in my description, don't hesitate to ask! Thanks for whatever help you can direct my way! ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Hematoxylin & Eosin-Y Re-staining
Hi all you histonetters! I had stained a couple of slides yesterday using H&E, but the staining came out a little too light for my liking. Any set protocols for stripping the stain and redoing it? (I use 7100 plastic resin). I've put the slides in xylene to remove the coverslip/mounting medium, and if I cannot get a decent restain, I'll just cut more sections. I'm just curious if anyone has a protocol that works well. This would save me a lot of time! Thanks! Allyse Mazzarelli Histologist Neurotech USA Inc. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Immuno Bed Plastic Question
Hi histonetters! I have posted on this forum a while ago asking for opinions regardnig new types of plastics that work well with IHC. It was brought to my attention that "Immuno-Bed" (Polysciences) works rather well. I recently purchased the plastic, infiltrated/embedded well, and sectioned great, BUT, when I removed the excess GMA/rehydrated and labeled with my antibodies, NOTHING WORKED! Nothing showed up! Only DAPI!! The mechanism I used for removing GMA and rehydrating were as follows: 1.) 3 changes of 30 minutes each in 60 Celsius xylene. (Note: I recently let the slides sit in 60 Celsius xylene overnight too... but it did not work!!) 2.) 3 changes of 10 minutes each in 100% EtOH. 3.) 15 minutes in 95%, 75% & 50% EtOH 4.) 5 minutes in water 5.) 3 changes of 5 minutes each, PBS. I continued to block and apply my antibodies as normal. (Antibodies are all brand new, not duds!) Since this plastic was formulated specifically for IHC, I am at a loss. If anyone has used this plastic before, any insight directed my way would be extremely helpful!! Thanks, Allyse Mazzarelli Histologist NEUROTECH USA INC. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Immunohistochemistry
To everyone in histoland that is familiar with IHC, I am a new histologist, unfamiliar with many techniques in IHC. Here in my lab, we section devices that contain cells, membrane and scaffolding. Unfortunately, due to the scaffolding, the devices do not hold up well when embedded in paraffin. Likewise, the other plastics I am able to cut nice sections with do not take well to immuno staining. Does anyone have any suggestions for a specific type of plastic resin that works well with IHC? (I could try something other than plastic too). For the time being, I use Dorn & Hart acrylosin soft... but the results are medicore. We will be doing a lot more immunohistochemistry in the near future, and I'm looking to experiment with different resins to find one that works well. Any help is appreciated! Thanks for your time! Sincerely, Allyse Mazzarelli Histologist, Neurotech USA Inc. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Specimens for TEM
Hi all, I have a question about transmission electron microscopy. Currently, at the lab that I work in, we use a basic H&E stain and image with a light microscope. However, it was brought to my attention today that there has been talk of sending some specimens out for TEM imaging. I currently embed my specimens in 7100 plastic resin. I was wondering how others out there process/embed their specimens for TE microscopy. I've never participated in TEM before, so any help anyone can give me would be great. Thanks! Allyse Mazzarelli ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
[Histonet] Specimen Blocks too Soft to Cut
This question pertains to anyone that uses the Technovit 7100 kit. My lab has seen problems with humidity recently… (lab is too big to use a dehumidifier). I embed with enough hardener and mix my embedding solution well, but when I take my samples out of the mold they are still soft. I have tried to store them in an air tight container in dessicant beads over night, but that only partially works. Any other suggestions? Has anyone else that uses this kit ran into a similar problem? Thanks in advance to anyone that can send me some advice!! Sincerely, Allyse Mazzarelli NEUROTECH USA INC. ___ Histonet mailing list Histonet@lists.utsouthwestern.edu http://lists.utsouthwestern.edu/mailman/listinfo/histonet
