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Hi, Has anyone done dissection (e.g.,brain dissection) using the laser capture microdissection, and done the qPCR analysis on the samples? How was the RNA quality and which kit did you use?
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I do LCM of mammary epithelial cells. Total RNA of harvested cells was isolated using Qigen's RNA easy Microkit and quality and quantity was evaluated using Bioanalyzer 2100. Use RNase inhibitor in solutions while staining to protect RNA degradation. RIN of RNA was suitable for qPCR and RNAseq.
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the recent researches have shown that blind people react(smile) to the smile of a person in front of them. how this connection occures?
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Professor Zeashan Khan,
Thank you very much.
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I would like to take rat kidney tissue and isolate from that the proximal tubule cells for immediate use in a Clark-type oxygen electrode to study mitochondrial respiration. I do not want to study isolated mitochondria (for reasons I will not go into here), and using tissue sections (e.g. 200 micron thin slices) does not allow full respirometry studies to be performed, only crude O2 consumption rates (even with permeabilisation).
I have seen lots of protocols for isolating PTCs for use then as primary cell lines however I will not be using the cells for this purpose. In my experiments the cells will be discarded after the short respirometry studies have been completed. I'm unsure of how much of these protocols I should follow if I just want to obtain a cell suspension for immediate use and not prolonged culture. Ideally I would like something quicker and simpler than most of the protocols I have found so far seem to be.
Prior to respirometry studies the naive kidney tissue will have been sliced to 200 um thick and exposed to certain insults (e.g. LPS, septic serum, etc.) for around 90 min. I would like a method that is as quick as possible to get the PTCs from the tissue in a state in which I can add them to the Clark electrode. This is in order to observe any acute changes that may have occurred to the mitochondrial function during incubation which may also revert relatively quickly once removed from the insulting environment. Once I have a cell suspension I intend to permeabilise to allow addition of the respiratory substrates, ADP, uncouplers, etc. required for respirometry studies.
If it's likely that this potential method will encounter too many problems I may first isolate the PTCs and then expose them to my various insults immediately before washing and beginning the respirometry studies. In that case would I need to culture the cells, or could I just expose them in suspension? Ideally I would however like to be able to expose slices to the insult (for reasons of consistency with other methodologies I am currently employing) as initially suggested.
Many thanks.
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I want to do rat brain immunohistochemistry, and I find that rats were perfused transcardially with PFA under deep anesthesia in all published papers. But now some people in pathology told me that rats can be decapitated and immediaterly to get brain, after saline washing, the brain fixed to 4% PFA is OK. Should I to do perfusion at first?
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Some antibodies won't work after several days in PFA 4%. You should wash and put PBS after 48 hrs and cut as soon as possible.
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Hi, we got 15mm x 15 mm x 0.15 mm Mica substrates. But after cutting by scissors, we found many cracks, as shown in the image. Does anyone know how to cut it gently without introducing those cracks? Thank you very much.
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No, really no, but I have the intution that the cutting with the cutter asisted by a rule may be good. It wil be important to make pressure with the ruler, previous to make the cut.
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I have to isolate microglia from brain tissue for my research. Almost every protocol about microglia isolation suggest a perfusion with ice-cold PBS/HBSS .
I want to know why perfusion is necessary before brain dissection.
Thank you.
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One justification for doing so, is to remove as much of the blood that's in the vasculature which will have leukocytes and other cell types that may express biological markers similar to brain-resident microglia.
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I am worried of its DNA, that it may be damaged....
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I would not expect any problems...cells grow at 37c and the dna is fine so it depends on whether your saline is sterile but for reasonable length pcr product I think that you would be ok. Freezing and thawing too often might be more of a problem but so long as we are not talking tiny ( single cell) amounts of tissue you will be fine
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I am currently performing a craniotomy/durotomy procedure on rats (Long Evans – Hooded Blue Spruce 450-600g) to expose brain tissue for optical coherence tomography. To open the skull, I am using an engraving bit (Dremel size 105 & 106) to cut a 4mm x 4mm area of the skull. Once dura mater is exposed I am able to lift the skull off cleanly without damaging the brain tissue. The dura mater is then carefully lifted with a tweezers and cut with a micro scissors. During roughly half of the procedures the underlying meningeal layers are removed cleanly along with the dura. However, if the pia does not come off cleanly (typically when it contains large perforating cortical vessels), any attempts to remove it results in significant bleeding. This obviously inhibits the imaging process by blocking the light aimed to penetrate tissue. Occasionally the skull bleeds significantly as well. Thus, if I am unable to avoid excessive bleeding, does anyone know of any non-pharmacological methods for addressing this problem (other than Gelfoam Sponge or Actcel hemostatic gauze)? Or any pharmacological methods that would not compromise hemodynamics? In addition, the exposed brain tissue becomes inflamed, varying the focal length of the lens and further distorting the images. What is the best way to control for this inflammation? I have been applying aCSF (stored in an ice bath) to control for this. I have also tried opening another area of the skull (left lateral the sagittal suture, posterior to bregma and anterior to lamda) to allow for a release of some pressure. So far, this doesn’t appear to be beneficial. Lastly, what is the ideal dosage of Isoflurane to administer a rat under hypoxia and/or hypercapnia? After collecting baseline data (at normoxia), the rats are exposed to a hypoxic/hypercapnic gas mixture (10% O2, 5% CO2, 85% N) in order to examine capillary dilation and perfusion of the cerebrovascular reserve. Some of the rats remain stable under this condition, others expire within a few short minutes. Does anyone have an explanation for the variability? The rats are all the same breed, age, and are close to the same weight.
My goal is to examine hemodynamic activity in real-time. Therefore, any solutions to these issues should not further compromise normal blood flow, or vascular restriction and dilation.
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Hi Jacob,
1. This is a protocol nearly for the same goal but applied for mice. Maybe this can help you.
2. In this article, you can find anesthesia protocol and surgical procedure in the Surgical Procedure section.
3. On the other hand, how did you remove the pia mater? What kind of tweezer do you use?
Kind regards
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I am trying to hole punch for RNAseq in specific subregions in the rat brain. In order to be more accurate I want to do 300 micrometer. I am wondering if I should buy a tissue slicer.
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The important thing here is to make sure that whichever technique you choose, you can do it quickly with the minimum amount of freeze/thaws, since RNA degrades very fast.  If you can use the vibrotome immediately after sacrificing the animal to isolate the region of interest, and then either prepare RNA immediately or flash freeze at -80C, that would be ideal.  You could also use the cryostat after the original flash freeze, but you would be thawing the tissue back up to -20 to use the cryostat which can also degrade RNA. 
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Hello, 
I have a problem with the cryoprotection of mouse hippocampal slices. The samples are PFA-fixated and cryoprotected in 30 % sucrose in PBS solution; they would normally sink to the bottom of their wells in the 96-well plate after a couple of days in the cryoprotectant, but this time none of the samples (there are five of them all in all) would sink. I have already encountered such a problem a couple of times before, and according to my experience it is not a good idea to proceed with samples that are still floating, as that implies that they have not been cryoprotected well enough and therefore are very likely to get damaged during sectioning.
I placed them on a shaker in a cold room now, as I hope that that would help the cryoprotectant to enter the samples; however, I am not sure that that would work. Therefore, any advice would be very appreciated. 
Thanks a lot in advance! 
Max
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Hi Maksims, to run a sucrose gradient raw is a good idea. You can also start with 10%, 20% to 30% or smaller steps like 10%, 15%, 20%, 25%, 30% and stoer in 30% over night. Use bigger vials like 1,5 ml or 2ml Eppis. I am working with slice cultures as well as with 300-400µm thick brain slices after electrophysiiological investigations. We put the sections after fixation in this cups for a couple of days and get good results. To control the quality I recommand a NIssl-stain with cresylviolet. Good luck.
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I work with Rat & Mouse brain tissue. I have hippocampal slices in a 6-well dish that I have fixed. We keep stored at 4 degrees. We have changed the fix to PBS for long term storage and parafilmed the plate for spillage. However, I noticed that I had a slimy mold (look like a tadpole swimming) in one of my wells. Worried about contamination what's the best solution to preserve brain tissue without damaging the tissue or contaminating it?
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Hi Heater, 
I guess that you want to preserve IgG binding so your best bet is adding azide in your PBS if your safety regulation allows it. Alternatively, after PFA rinse with sterile PBS several times and bring to a culture hood for additional rinses. it should be sterile enough (both bacteria and fungi) for long-term storage at 4C. Additional precaution if you see diminished IF quality would be to avoid further oxidation by bubbling the PBS with sterile N2 (after sterilization). I hope this helps. Thomas   
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I am Working on Primary Neural Cell culture from SVZ of the Postnatal Mice Can anybody Help me to Provide Some guidance how to Precisely cut this part. and How to recognize SVZ in Brain of Postnatal mice. If you have some Videos Illustrative Diagram I could be better for me?
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Hi Masood,
I wonder if you have seen this JOVE article?
This is a protocol for whole-mount immunostaining but you can follow it's steps of dissection. It is mostly applicable to all postnatal brains. For new born (p0-5) the brain shapes slightly differently, but the general gist is there.
Let me know if you need more info!
Xiwei
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I use 50ug/ ml Gentamycin in my culture media already. I have to dissect the sub-ventricular zone under a microscope which takes a while to do and all the steps upto the collection of region of interest are performed outside the hood. Please suggest if I should add the antibiotic in my dissection medium as well, for extra protection from the contamination.
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I suggest you to keep the microscope in a hood and do the dissection inside the hood because adding antibiotic will lead to death of cells and will also affect the health of viable cells. Therefore, you will get very less number of cells for your cultures and also you cannot negate the chances of contamination even after adding antibiotic.
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How can i cut slices of cerebellum in a good way? I think someone who studies neuroscience. 
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If you are sectioning mouse samples, use paraffin. For larger animals, e.g. Rhesus macaques, use cryo: dry ice and obviously a larger microtome.
AFIP has a very good explanation of the methodology. ISBN 1-881041-00X
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I need to dissect these nuclei of macroscopic manner.
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Dear Angela
Please find the attached file, hope it will do for you..
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I dissect the animal, intracardially perfuse with 0.9% Saline, then with few ml of 4% PFA intracardially. Then i remove the brain and fix it in 10 % formalin for 2-3 days after which i embed the brain in paraffin and process it. During paraffin sectioning, i consistently observed there is vaccum in hippocampal area, which is like a shrunken hippocampus. I could literally see a cavity while sectioning in a microtome.  Although i can get hippocampal sections, little beneath the actual area.. Can anyone suggest any reason for this shrunken/Vaccum, which i encounter in hippocampal region? Does this cavity arise due to the Circle of Willus, which directly supply hippocampus, which is situated just above the basal midbrain????
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if you are fixing using PFA by perfusion, why dont you perform cyropreservation and cryosectioning
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I have flash frozen whole honey bees, and would like to dissect out antennal lobes and mushroom bodies for gene expression work. Is the ICE reliably good for preserving RNA in small bits of tissue? Also, does it mess with structural integrity of the tissue (ie turning it to un-dissectable mush)? 
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hi
put in RNA later for 24h in tempreture 4 centigrade, after than extract RNAlater and transfer to refuger -70
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In term of solutions to use during or after dissection, dissection conditions, drugs to use, etc.
Apparently, mossy cells are very fragile/sensitive. How can I improve their survival rate to be able to record them later?
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Check papers from Ben Strawbridge group - Best luck, SV
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Dear friends, I need to use tungsten needles to deliver dextran-conjugated fluorescent die to the spinal cord of Xenopus tadpoles. I plan to label some reticulospinal neurons retrogradely. I've never worked with these needles, and so cannot quiet imagine them, both in terms of how they look, and in terms of how they would feel during the dissection. FST have 3 types of needles in stock: 0.5, 0.25 and 0.125 mm in diameter (link below). Which one would you have used for this task?
Educated guesses with any kind of rationalization are very welcome, as my best other option is to order the middle one just because it's in the middle.
The link to the FST site:
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I have no eperiance with that
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I would like to study neurosecretory cells of the brain of ceratitis. Can I use the carnoy as a fixative directly on head without dissection of the brain?
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As co-author of these papers I recommend to follow the steps reported in the papers by Conforti E. et al (1999), and Roda E. et al. (2004), according to the suggestions by Wolfgang H. Muss.
Graziella Bernocchi
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Can anyone definitively answer the question: do fixed brain samples in PBS with sodium azide need to be refrigerated for long-term storage?
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You did not say what you would use this tissue for. If you have any wish to use it for EM, better save some samples embedded-- as the membranes etc will be degraded. If you have -80 storage (see comments above)  cryoprotection storage is the best all round method.  For immunocytochemistry and many special stains you can store it in the refrigerator (I  had monkey brains in the refrigerator for 10 years and was still able to stain for enzymes and neurotransmitters.  But if you wish to quantify such things different storage methods must be used.  See above , snap freeze).  If during your storage the pH and osmolality is off,  things like mitochondria will not be optimal. So if you are not thinking EM but may be confocal,  you may have an  issue. Good luck
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I want to dissect visual cortex tissue for quantitative RT-PCR and I found that it is difficult for me to get visual cortex tissue evenly from diffrent mice. Does anyone have any suggestion? Any advice is welcome. Thanks a lot.
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Hello, you may use a tissue punch of appropriate size rather than microscissors or dumont forceps. This should lead to quite reproducible results.Usally these biopsy devices are used by dermatologists.They are inexpensive and ususally for single-use.
Good luck!
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Can anyone help me in dissection of rat hippocampus and amygdala ? if possible with dissecting manual of rat brain?
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I've dissected out fresh rat and mouse amygdala.  I place the whole brain in a cold matrix on ice, ventral side up, and use the circle of willis as a guide in making my cuts.  (A thin slice is needed.)  I then place the slice on a petri dish on ice.  To make it easier to find the amygdala when working with mouse slices, I have a light source shining up through the plexiglass, but with rats this isn't needed.  I then use the corpus callosum/external capsule as a marker to punch out a small area of the amygdala.  With fresh non-frozen tissue, I find it easier to pull away the surrounding tissue while leaving the punch in place, then remove the punch with the tissue to collect.  This punch technique only gives a small amount of tissue, but we've been able to culture mouse amygdala and run qPCR on the mouse amygdala, so it's not too small of an amount.  The link below gives an excellent visual description, although they do take out more tissue than what I've described with the punch technique.
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As of late, I am having trouble picking any cells from brain tissue slices using immuno-LCM techniques. We have successfully picked single and pools of single cells from brain tissue up until recently.  Now it seems that we are consistently unable to pick up any cells. Nothing significant has changed with respect to the equipment, protocol, or reagents (to my knowledge).  I have been troubleshooting for a while now and am nowhere closer to any sort of solution for this mystery. We use the following protocol for tissue sectioning and immuno-staining:
slice OCT embedded brain tissue using cryostat at -20C (10um thick slices)
store slices on charged slides (for better adhesion)
store sections at -80C until staining and picking
we fix slides using ice-cold acetone (1min)
air-dry slides post fixation (~ 2-3 min)
block with 3% BSA solution
rinse with 1X PBS
add primary Ab solution (incubate for 3 min at room T)
rinse with 1X PBS
add secondary Ab solution (incubate for 3 min at room T)
rinse with 1X PBS
dehydrate in a series of EtOH solutions of increasing conc. (70%, 75%, 95% 100%, 200%)
final dehydration in xylene (100% 1 min, 100% 4 min)
air dry and store in 50mL tube with dessicant for 10-20 min
I have tested a variety of factors targeting various aspects our protocol:
longer acetone fixation times (5-10min), room T acetone (2min), longer EtOH baths (from 30s to 2 min), longer xylene baths (5min-10min), used new reagents (unopened bottles), new caps, uncharged slides, longer dessicant drying times (as long as 45 min), and have adjusted environmental conditions as best as I can in the LCM room (relative humidity ~mid-30%, which we have been able to pick cells successfully in the past). The IR-laser is able to wet the caps fine as well.  We are able to pick up cells from older tissues that have been used in the past, so we do not suspect an issue with the LCM itself. 
At this point, my colleagues and I believe it has something to do with the dehydration of the tissue (we note that in tissues that yield successful microdissection, they are typically opaque and white upon fixation and/or xylene dehydration).  However tissue that prove difficult to pick from are not nearly as opaque (more a translucent white - faded).  I would appreciate any insight from the community as to how we can improve consistency in tissue dehydration and/or techniques that will improve success of tissue microdissection. Any insights or thoughts would be helpful, thank you! 
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I had the similar problem and it was most probably related with ambient humidity. LCM has stopped working in summer when humidity in the lab raised up significantly. Desiccant did not help. I could not control humidity in the lab and had to wait till October when humidity dropped down and I could continue with LCM. You can control humidity in the LCM room. Is it the same room where tissues are sectioned and slides are prepared? However, you have mentioned that old slides are possible to process now, I am not sure if my case is relevant to your situation.
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I want to extract RNA from Fly larval brains. I had problems of degradation before. I am planning to use RNA later this time. Is it advisable to use RNA later while dissecting the brains or dissect the brains in water and transfer them to RNA later. Please provide me the details?
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Hi, RNA later is fine for brain tissues, not so much for other fatty tissues. I don' t know if you can manage to dissect in it, it is very very viscous. Try to dissect in 0.22um nitrocellulose filtered PBS (high protein binding capacity, to remove any contaminant RNase you might have on your solutions), keep the dissection plate (cleaned with RNAse OUT solution) on ice, and transfer immediately each dissected brain in TRIZOL or any other buffer of extraction that you use kept in ice. We are always doing like this.
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I want to micro dissect the brain of rats and mice to isolate hippocampus, prefrontal cortex and other brain parts. Can help me out to find the best video on micro dissection of brain either in youtube or any other website? Thank you.
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Hi Lakshmi Narendra,
You could refer to a book by Palkovits on brain maps. The title is : Maps and guide to micro-dissection of the rat brain.
If you would prefer videos as suggested by Ramirez-Franco,  Jove is a good place to look. Take a look at these as well:
Hope this helps!
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I am trying to collect the hippocampus, prefrontal cortex and amygdala from adolescent rat brains (unfixed).  Is there a way I could validate the accuracy of the dissection to rule out contamination from adjacent brain regions? For instance, I have been searching for molecular markers of the hippocampus that are absent in adjacent regions, and vice versa.
Thank you
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Take the slice from which you dissect your regions of interest, fix the remaining tissue and slice using a microtome/cryostat. Then stain the section using e.g. cresyl violet and validate based on anatomy
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I will be removing whole mouse brains to be later used. So, I need to know the best methods to store the brain.
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depending on what you need to do after....i.e.
-western blot: store the whole brain (or the different areas) @-80°C
-mRNA extraction: store whole brain (or different areas) in trizol (or RNALater) @+4 short term
-immunohistochemistry-fluorescence: fix it with PFA 4% then cryoprotect it with increasing sucrose,then store @-20°C
just an example..just specify what use you will do after...
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I always end up poking the brain to get rid of the vessels and butcher the surface.
This is proving to be too difficult. Any help would be appreciated.
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Hi,
I would suggest using the fine-tipped tweezers (see below for image) and removing this tissue starting at the olfactory bulb. After decapitating the animal and removing the skull, you will see like a black connective tissue around the olfactory bulb. Use the tweezers to break it apart and pull it off (carefully). You should then repeat this on the other side and the brain should be free.