Fluorescence In Situ Hybridization - Science method
Fluorescence In Situ Hybridization is a cytogenetic technique that is used to detect and localize the presence or absence of specific DNA sequences on chromosomes. FISH uses fluorescent probes that bind to only those parts of the chromosome with which they show a high degree of sequence complementarity. Fluorescence microscopy can be used to find out where the fluorescent probe is bound to the chromosomes.
Questions related to Fluorescence In Situ Hybridization
I am interested in doing FISH on leukocytes in a single cell suspension so that they can be analyzed via FACS. If effective, this would allow for more efficient analysis and of many more cells than traditional FISH-fluorescent microscopy (a method I have previously used with success).
1) Cells fixed with Carnoy's in a drop-wise manner. Spin, decant supe.
2) Permeabilize cells with 0.1% Igepal in 2X SSC. Spin, decant supe.
3) Resuspend cells in hybridization buffer (70% formamide 2x SSC).
a) Co-denaturation: Expose cells and probe to 75C for 5' together. 37C o/n
b) Separate denaturation: Cells 72C for 5' and Probe 90C for 10'. 37C o/n
c) Separate denaturation: Cells 37C for 5' and Probe 90C for 10'. 37C o/n
4) Wash with 0.1% Igepal in 2X SSC. Spin, decant supe.
5) Preheat 0.3% Igepal in 0.4X SSC to 72C. Add buffer to cells expose to 72C for 2'. Add equal volume ice cold PBS. Spin, decant supe.
6) Resuspend in PBS.
At this point I'm not attempting FACS, I am cytospinning on to slides until I optimize the hybridization conditions. I have given it a couple passes and I am routinely running into the same issues:
1) Enlarged nuclei, (the nuclei appear to be large and have a more diffuse DAPI signal). The nuclei seem to denature/enlarge once exposed to 70% formamide 2x SSC independent of heat. Possible problems: inadequate fixation, absence of graded alcohol dehydration steps like in traditional fix.
2) Ineffective hybridization. Since the conditions used have successfully worked for traditional FISH it would seem they should work. Also probably a inadequate fixation or denaturation issue.
3) Cell clumping. Cellular denaturation is resulting in the 'netting' of cells by DNA strands after exposure to 70% formamide 2x SSC.
I have some ideas of what to try next, but if anyone has had previous experience with Flow-FISH and could provide insight that would be great.
I did a quite interesting experiment in smFISH.
I use a short probe (10 base long) to incubate with 4% PFA fixed (15min) Hela cells. I cleared all the RNAs with RNase If (NEB) for 30min at 37C. The hybridization is carried out in 2XSSC at 37C for 30min. Finally, I washed with 2XSSC at 37C for 3 times, each 5min. However, I got very high background noise from the experiment, not only in the nucleus ( This can be expected since my probe is short) but also in the cytocol.
This quite confused me because I have removed all the RNAs with RNase If, so theoretically, there should not be background noise from the cytosol.
Does anyone know how I can handle this type of background noise?
I will appreciate for any help!
I am looking to perform a FISH experiment using a DNA probe labelled with fluorescein-12-dUTP. The probe has been labelled using a nick translation kit. The input of the kit was 1ug/12ul, and hence the output should be 83.3ug/ml.
A paper suggested using a concentration of 25ug/ml in the hybridisation solution. Is this a good concentration? To make such a high concentration I would have to repeat the labelling process.
I would like to visualize the RFP signal after performing the FISH protocol, but I am thinking that the paraformaldehyde fixation would not work with RFP.
I would like to fluorescently label mRNA that we will be delivering to MoDCs. I'm going to perform microscopy (Axios Imager D2 if that's relevant) to see localization of the specific mRNA within the cells. This mRNA is from a vendor so I can't transcribe it myself. Any thoughts?
we are working on FISH fluorescence in situ hybridization ,so we want to purchase DNA FISH probes sequence give any suggestion where i can purchase DNA FISH PROBES OR give any international vendors email or other. thanks
I am using specific probes for detecting and quantify bacteria with FACS, however in spite, it is supposed to work (Previous studies and In silico analysis of the probes), I only get high fluorescence but unspecific hybridization.
I have tried different variables to increase the specificity:
-Formamide concentration in the Hyb buffer (20mM Tris 7.0, 0.1% SDS, 0.9M NaCl)
-Increase wash steps and change pH (SSC20x)
-Change hybridization temperatures (From 48° to 70°) 4 hrs with agitation
-Reduce the concentration of the probes
Universal GCT GCC TCC CGT AGG AG- 6FAM
Bifidobacteria GAT AGG ACG CGA CCC CAT-CY5
Lactobacillus ACA TGG AGT TCC ACT-CY3
In silico analysis: Mathfish
All the bacteria are pure cultures washed in PBS, fixed in Alcohol 50% or formaldehyde 3.7% and the permeability of the bacteria have been tested with PI
However, none of these changes had a significant change. Is there anything else that I could try?
Thank you in advance
The ThermoBrite Elite system automates the pre- and post-hybridization steps in FISH testing.
How efficient is it? Is it a closed system? Is it FDA approved?
Hi. I am looking for a FISH probe that does not have any fluorophore or DIG/Biotin molecule attached to it. Instead I need some inactive protein like Cas-9 (Or any other available protein) attached to it. If that's possible to do then Is it available anywhere? If do then please suggest.
Has anyone done FISH on cre-recombinased mice? I want to know if anyone has successfully been able to quench the autofluorescence from the those mice to successfully visualize the targeted genes? I have cre-mice crossed with ROSA26 tomato mice that I would like to do FISH on.
Any help or guidance would be greatly appreciated. Thank you!!!!
Hello. We are trying to make a DNA probe for FISH. But fluorescent signals are observed at the periphery of the nuclei (see figure). What could be the problem? The probe was made using DOP-PCR from a BAC clone. I give an example of the obtained product on agarose gel electrophoresis of 1.5%. After DOP-PCR (with Cy3-dUTP), the product was purified and a hybridization buffer (70% formamide, 10% dextran sulfate, 2X SSC, water) was prepared in which 5 μl of purified amplicons were dissolved. The resulting probe was applied to a glass slide with the nuclei of human leukocytes. Denatured at 75°C for 5 minutes, then hybridization overnight at 37°C. The next day, they were washed first in a 0.4x SSC in a water bath at 73°C for 3 minutes, then in 2X SSC with Tween 20 for 30 seconds, dried and examined under microscope with a DAPI /Antifade.
Following are probes required for fish analysis
2. EUB338, EUB338II, EUB338III
I've been doing whole mount ISH on E9.5-10.5 mouse embryos (with DIG-labeled RNA probes). For a long time, I've been struggling with getting a decent signal before the background starts to become too strong (NBT/BCIP). On gel electrophoresis, the RNA probe seems ok, so I'm assuming the probe quality is ok and I've included several steps to reduce background (levamisole, acetic anhydride in TEA, RNAse).
After sectioning some embryos that were hybridized for a cardiac specific probe and showed a signal in the heart as well as some overall background, I found some weak expression in the heart, but also a very strong signal all over the embryo surface.
Has anyone seen this before? I've googled google extensively, but I have not been able to find a convincing answer. I suspect it may be residual background signal, but am also considering overfixation/suboptimal protK treatment causing suboptimal penetration of the probe (even though I do a 9-10 minute incubation with 10 ug/ml prot K at room temperature).
I am currently trying to do FISH on human skin biopsies that have been routinely fixed in formalin and embedded in paraffin. I am using the CEN X/Yq12 probes from ZytoVision and their ZytoLight Tissue Implementation Kit. De-paraffinization was in xylol, isopropanol, 96% and 70% ethanol. The manufacturer's protocol performs a heat pretreatment for 15 min at 98°C in a citrate buffer and continues with a pepsin treatment at 37°C which should be optimized for the tissue and probes. Denaturation was at 75°C in the dark. Hybridization was at 37°C overnight. All (wash) buffers were used from the kit at correct indicated temperatures.
I saw a positive signal for the X probe with 6 minutes of pepsin in a female tonsil test sample. 6 minutes seemed to have also worked for the Y probe in one male skin sample, but no signal for either the X probe or a second skin sample from the same run. Since then, I have tried 2, 4, 6 and 8 minutes of pepsin in more skin samples but haven't been able to get any signal. A second tonsil sample (male) also failed to give a signal with 6 min.
Nuclear counterstain was with ProLong Diamond or VectaShield. Additional CD3 staining worked fine in all cases. X and Y probes are both in the same tube (dual color probe), so it's not possible that I forgot to pipet one. Errors in microscope/laser/filter settings are rather unlikely. Thickness is 2 µm as recommended.
I am desperate for any advice on what could be issue here. I can still try to reduce the heat pretreatment or play around with the pepsin incubation, but I'm running out of time. Any theory or idea for troubleshooting will be highly appreciated!
Hello fellow scientists
I am planning to do fluorescent in situ hybridization work on fish intestinal tissue and skin samples. For which, I will be fixing my samples in 4% PFA for 12 to 24 hours.
The problem is; someone is doing the sampling on behalf of me and therefore I won't be able to process my samples right after fixing. They will be shipped to me which will take 2-3 days.
My question is, is it okay to keep the samples in formalin for 2- 3 days. Will this effect my hybridization work?
Any tips will be appreciated!
In the protocol of the book, FISH Handbook for Biological Wastewater Treatment, writed by Jeppe Lund Nielsen, the sample is usually fixed independently for both Gram negative and Gram positive cells. However, Eubacteria includes Gram negative and Gram positive bacteria, and I want to identify both of them in one microscopy observation. Now, should I fixed ram negative and Gram positive bacteria separately?
I am currently optimising a flow FISH protocol for an archaea species. The cells are fixed with and final concentration of 50% (v/v) ethanol before FISH hybridisation. The FISH protocol is carried out in cell suspension/pellet, and the resulted pellet is suspended using 0.5 mm needle in buffer solution prior to microscope check. There are several problems:
1) the cells tends to form aggregate;
2) the majority of archaeal cells are not labelled with the probes;
3) there is a very strong fluorescence background when checked under the microscope;
My assumption is that, the extracellular material of the archaeal cells is very stick, and this makes the cells get burst when physical force is applied. There might be other explanations. Can anyone offer some advice? Thanks in advice.
Hi! I'm new in the field of Fluorescent in Situ Hybridization (FISH).
I want to use DNA FISH to visualize a small region in the human genome (around 1kb). I'm not sure if this is too short to use probes generated by nick translation.
I guess I probably need to order a set of short probes that all anneal to this region to enhance my signal. Does the Stellaris® RNA FISH system from Biosearch Tech (https://www.biosearchtech.com/products/rna-fish) apply to my case?
Or do you think this experiment is doable? What is the best way to do it?
Thanks a lot!
I am trying to combine the two methods of RNA FISH and protein immunofluorescence to one protocol and apply it on E. coli cells.
Does anyone have experience in such procedure in bacteria?
Anyone has a protocol or recommendation?
I want to combine FISH protocol with EdU Click-it chemistry as well as immunofluorescence. Does anybody have a protocol or idea in which order to combine these different staining methods? Are there any specific treatments necessary? At least for immunofluorescence + FISH I know how to handle it, so the question mainly concerns the combination with Click-it reactions.
Hello, so I am doing an experiment where I am trying to locate specific transcription factors in the mouse brain. I have decided to use a dual probe fluorescent in situ hybridization kit that I ordered from a company. However, the protocol that the company provided starts at the hybridization step, but I need to know how to permeabilize the tissue. I am doing the procedure as free floating tissue at 30um thickness and was wondering if anyone had any ideas on how to permeabilize the tissue? So far I have not received any specific fluorescence and I believe it is because I am not permeabilizing the tissue enough.
I'm looking for solutions to maximise the signal-to-noise ratio in immunofluorescence and/or FISH. We are routinely using Alexa Fluor labelled secondary antibodies but now we need something more powerful. We primarily work with formalin fixed tissues, so the color should preferably NOT be green (due to autofluorescence).
Hi fellow researchers,
I have recently begun a PhD in an interdisciplinary team that involves some microbial ecology in field experiments on my part.
Correspondingly I am looking for a way to determine especially prokaryote diversity from soil and water samples (types of species and relative abundance basically) in a mobile setup.
It should be relatively quick and especially not too expensive to buy up-front.
I am not sure about FISH as the technique seems to be more about looking at the occurence of specific strains that you already know before-hand (and thus know the complementary DNA).
PCR techniques use quite a lot of rather costly equipment and is pretty lab-bound to my experience.
The abundance itself could be achieved by DAPI/Hoechst staining and fluorescent microscopy.
Any way to get a sense of types of prokaryotes involved / general diversity in the field though?
May alternatively look at fungi in the field aswell so if that would work in a similar fashion that would also be great.
Any ideas would be much appreciated.
Is there anyone who has experience on staining extrachromosomal DNA by fluorescence in situ hybridization (FISH) ? I would like to stain extra oncogenes on extrachromosomal DNA and chromosomal DNA. Could you share your protocol? I really appreciate your help!
Does anybody know a lab in Australia or New Zealand who carries out in situ hybridization, FISH, GISH with polyploid plants?
Hello everyone, I isolated PBMCs for Fluorescent In Situ Hybridization but I also have more Background when performing FISH. can some one tell me what currently contamination sources can occured when doing Ficoll Extraction form PBMCs?. In my case I suspect a contamination with thrombocytes or Erythrocytes it that true? when true how can I purified my sample of PBMCs?
I am having problems with FISH using DNA probes for staining mycobacteria (specifically Mycobacterium avium subspecies paratuberculosis). It appears that about 25% or less of the time the probes are actually going into the bacteria and don't look that bad (but I have to put the smart gain on max just to see color). The other half of the time the probes seem like they are not penetrating through the cell wall. I have tried many times changing our protocol, but still only getting that percentage of actual success. Even when using the same protocol over and over, the results still come out different almost each time. Other bacteria species work pretty well, it's just Mycobacteria that are not working well. I know Mycobacteria have the thickest cell wall of almost all bacterial species, but the protocol that I use still should work.
Has anyone worked with bacteria (specifically mycobacteria) using FISH using DNA probes (alexa probes are added to the 5' end)? Please let me know your protocol and I will show you ours as well. Thanks.
I need a comprehensive guideline to follow to determine analysis cut-off points (e.g.: number of nuclei per sample, percentages to consider a deletion present, false +/- testing for each probe since the % of hybridisation is different for each probe).
I am using Leica: TP53, LEU1, +12 and ATM mutations for my FISH experiments. So far the only one I have found that is of any use is this:
Does anyone have any other papers that may be useful/helpful for my purpose? Would be better if it was specific to CLL.
Thanks in advance.
I've been doing WISH and X-gal stainings on E9.5-12.5 mouse embryos.
After obtaining a decent (ISH) or even strong (X-gal) signal in the whole mounts, I fix the embryos o/n (or up to several days) in 4% pfa, dehydrate them to 70% ethanol in PBS for a couple of days to improve signal/background ratio, take pictures, and embed them (ethanol dehydration, xylene, paraffin).
During sectioning, the signal still looks ok. Unexpectedly, the signal nearly dissolves in the water bath when I stretch the sections. By the time I place the sections on the slides, the signal has become very weak and sometimes nearly undetectable.
I've read some suggestions that incubation in ethanol or xylene may dissolve the color crystals, causing them to leak from the sections, but I have been unable to find exact times. Does anyone know how long is too long? Would any other dehydration method work better with regards to preserving signal?
I have been working with fluorescent in situ hybridization on paraffin-embedded tissue of papaya leaves in order to localize two RNA viruses.
After several adjustments in the protocol, the hybridization for dsRNA virus works well, but not for ssRNA virus.
My probes have 400bp and was synthesized by random primer labeling with fluorescent dUTP. The template sequence for the reaction was obtained by cloning PCR fragments from the viruses in pGEM-T easy vector.
Hybridization temperature, proteinase K concentration, and the denaturation process were adjusted.
Does anyone have a possible explanation?
We are setting up 16S FISH for animal tissues with low bacterial abundances. What is the most effective & specific system for probe detection & signal amplification?
The metaphase spreads look absolutely normal outside of the area where the denaturation and hybridisation took place. Within the area of the denaturation and hybridisation, however the metaphases look weirdly shaped and clumping but also the nuclei look somehow digested/degraded.
I attached two pictures of the dapi stain to visualise this difference.
The probes I am using still work well on the bad looking cells (not shown), but I need clear images with recognisable chromosomes. We already tried reducing the denaturation step and decreased the temperature, but it didn't help.
Did anyone come across this issue and can help?
Thanks for helping.
I am using EUB338 probe ( 5'- GCTGCCTCCCGTAGGAGT -3' ) at 35% formamide concentration and 46'C hybridization temperature as described by Silva protocol(attached) for bacterial cells on membrane filters. I am also fixing the cells with 4% Paraformaldehyde in PBS prior to hybridization.I am not getting any results while seeing under the microscope. Please suggest in which step I could go wrong or how to troubleshoot.
I would like to design FISH probes for cytogenetic experiments for Solanum species. There are a number of Solanum genomes available so probes could be ideally done for each chromosome with known locations. I haven't done such thing before, so are there any probes already available for tomato or potato? What should I keep in mind to design probes for in situ hybridization?
We are trying to break the mycobacterium spp's wall to do FISH in formalin fixed tissues. We already use Lysozime and/or Trypsin, but didn't work. Someone has some experience with that?
I have been trying to perform FISH/DAPI staining of bacteria in water samples for cell counts and identification of certain bacterial populations. The issue I am having is that several cells show high autofluorescence in all channels I am using for the probes so I cannot be confident of the signal I obtain with my probes.
I have tried using pre-treatments with UV exposure (I read this can potentially damage the ribosome and affect the hybridization) and using Sudan Black B (although this was used in tissues, not planktonic cells) with little success.
I was wondering if what protocol/method you know or have used to reduce or at least minimize this issue in water samples? Thanks!
The goal is to preserve the mucous layer. The slides have been IF stained for a week now.
if i designed a pair of primers and examined their specificity to the exact gene and by PCR i obtained the exact band. can i use this antisense primer as oligonucleotide probe directly for performing fluorescent in situ hybridization or i should make gene cloning and sequencing before using it.
I want to see localization of mRNA in cell using DNA probe there r two alternative LNA probe and DNA probe,I want to know how we can design a DNA based fluroscent probe for mRNA detection ,if any one has used it kindly share protocol
I am struggling with The XL DLEU/TP53 locus-specific probe (metasystem) detects deletions in the long arm of chromosome 13 and in the short arm of chromosome 17 in Chronic lymphocytic leukemia (CLL) by FISH
Im trying some new reagents for FISH. Fast red dsnt seem to work at all, ant this product from "Vector labs" claims to be much better- But it looks like it works even worst then the fast red. I cant even get background signal.
During in situ hybridization of DNA usually detergent are used to make pores in the phospolipids containing nuclear membrane. However, in case you perform ISH on FFPE tissue sections containing cells with a nucleus of at least 10 micron (e.g. cancer cells), which are cutted around 3-5 micron, I would suppose that most cells are already open? Why is then supposed to be so necessary to pretreat your sample very well?
At the moment I am working with Fluorescence in Situ Hybridization to study microbial communities in activated sludge
Samples which I've recently studied posses a strong tendence to autofluorescence. I've tried to deal with this problem by making some procedural modifications or reducing the amount of some extracellular substances.
But I am wondering if there are more ways (maybe more effective) to minimalize the autofluorenscence of activated sludge.
I want to mix digoxygenin and Biotin together in order to get yellow signals In FISH ( Flourscent in Situ Hybridization ） assay. Can anyone guide me either I have to mix both together in nick-translation reaction or do nick translation reaction separately and then mix nick translation products together?
I am struggling to attach FACS-sorted PDX cells onto the glass coverslips, for further downstream experiments like immuno-stainings and RNA FISH. Can someone help?
I need expert opinions on this matter as my plant samples are detached during washing step of FISH. I used agarose as well as transparent nail polish for this purpose and to me agarose didn't work at all but nail polish works slightly. It provides a base to sliced plant tissue, when I put my hybridization slide into the washing buffer (50 ml falcon) and then in water bath; after heating up to 48 degree Celsius, the nail polish is no more stick and my plant samples swim in freely in the media. Though I doubt that it may not affect the hybridization but still I am not sure. Can you please share your practical experiences?
Thanks in advance
I have some FISH probes which have been expired about 5 years ago. How can I find if they are still usable? I am looking for some way except performing a whole FISH study.
I am getting confused with the 16S marker and would need help.
I saw that the 16S rDNA is mostly used in bacteria research and is the component of the 30S small subunit of a prokaryotic ribosome, 18S for eucaryotes. So why is it also used for animal phylogenetics (e.g. fish) ?
Also just to be sure, from what I read, 16S rDNA is maternally inherited and non coding, right ? I have many stop codons in my sequences, whatever the reading frame used..
If someone can help me I would be very grateful !
Hello! I am going to do FISH and I have to make my experiment soon, but the probes are not here yet. How can I better store the cells and for how long? I saw in one protocol that you can leave in the permeabilizing step with 70% ethanol for 1 week. Is this the best and longest way to keep it?
Thank you for the help!
I need to perform fluorescent in situ hybridization (FISH) in hypothalamic tissue sections.
I already have the oligo probes, but they were going to be used in radioactive ISH.
Is there a way me to label these oligos for FISH? Are these kinds of oligos only useful for radioactive ISH?
Does anyone have some expertise in this subject?
I'm trying to detect the expression of several 250bp marker RNA sequences using FISH. For this I'm free to design the marker sequence but are limited by the size of the construct (200-300bp).
My current plan looks like this
1) generate random DNA sequences
2) Design multiple 20bp probes targeting this DNA using the Stellaris probe designer
3) combine those probes into a 250bp marker sequence
4) check the probes and sequences for stable hairpin and blast on NCBI balst
5) stably integrate my construct containing this marker sequence in a cell line
6) detect expression of marker sequence using FISH (several 20bp probes or longer probes)
As I don't have any experience in FISH I now have the following questions.
-Is it possible to reliably detect the expression of a 250bp sequence using FISH? (just the expression, I'm not interested in the location)
- I'm currently using the Stellaris probe designer and some Microarray probe designer to design my probes. Is there a better alternative or even a curated Database of validated FISH probes?
- NCBI blast shows that all the probes designed by Stellaris have atleast 65% homology to random genes. Is this a problem?
Many thanks in advance.
I am working on gene loci 3-D DNA FISH in suspension cells(primary lymphoid cells), but it's not working and I am not able to troubleshoot.I have tried to execute the experiment many times but I am unable to get the result. Followings are experimental details and queries:
Q1- How to isolate BAC DNA. I have isolated BAC DNA by Qiagen large construct kit but genomic DNA and RNA contamination are still coming.Is any special treatment required for isolating BAC DNA?
Q2- I am getting signal along with so much background so just help me out to reduce the background.
Q3-I am preparing probe by FISH Tag Kit(Invitrogen) by nick translation but I want to know how much concentration of probe DNA, cot1 DNA you used for precipitation of DNA and in how much volume of hybridization buffer and how many washes is required to get rid of the background? It would be nice if anybody can share the protocol with me.
I am FAC sorting testicular cells and I would like to do FISH to tell the spermatogenic stage. Some populations are very rare ~1,000 cells, so I wanted to sort them straight onto slides and do the fixation and staining there (sorting in tubes and cytospin would mean sample loss and the cells would be very dispersed on the slide after that). I have gone through so many protocols that don't really work for the above conditions.. Any suggestion?
I fixed the tissues in paraformaldehyde 4%. The hybridization was made using Alexa Fluor 594 in labeling reaction but I can't confirm if the fluorescence is due to hybridization signal or chlorophyll autofluorescence.
I am looking for a vacuum pump to infiltrate solutions into plant tissues in order to fixate them (for molecular biology experiments like ChIP, in situs, etc…).I need something powerful, because it is not just for Arabidipopsis seedlings, but not too strong and there are plenty of options in the market ¿How much vacuum power do I need?
I am working with pathogenic oomycetes in grasses which are asymptomatic and rarely produce sporangia growing out from the host tissue. The grass samples were fixed in 5% Glutaraldehyde in PBS (pH 7.5) and thus difficult to proceed with oomycete-specific probes for FISH (GA creates a very strong fluorescent background that even my negative controls reacts with the different filters). Apparently, I tried using multiple fluorescent stains and combinations (Aniline Blue, Trypan Blue, and Uvitex2b) although these stains also binds to chitin. The problem is the host plant may have both fungal and oomycete pathogens together. Likewise, it is already sure that my target parasite is present since specific primers were designed (sequences obtained were 99-100% identical and forms a clade with the same species with >95% support by RaxML) and the samples for microscopy where derived from the same batch. I had also considered morphological aspects like the presence and absence of septation, but reports of septation in the hyphae of oomycetes had been widely observed in different genera (Plasmopara, Peronospora, etc). Trying chitin degradation had been an option too, but the procedure is too tedious, likewise the chances of degrading cellulose and glucans of oomycete cell walls are highly possible. Considering my little amount of samples, this risk is also quite the least option for me. Sorry for the long detail, I hope someone can offer some suggestions. Thanks!
I am trying to do fluorescence in situ hybridization(FISH) on primary culture of fibroblast. I was working with 25cm^2 flasks. Around 80% confluence, I treated cells with colcemid for 4H and followed regular protocol for chromosome harvesting of adherent cells. After I died cells with DAPI just to make sure that I did everything correct. However, I got high number of interphase cells and very few mitotic cells (1-2 per slide). Can anyone tell me what I did wrong and how to increase my mitotic index?
Hi, anybody knows how fast a bone marrow sample for Fish or conventional cytogenetics taken from bone marrow and placed on McCoy medium needs to be processed?
I work 5 hours away from the Research facility that does our cytogenetics and the admin personnel tell me it needs to be at the facility the morning after it was taken.
Is that so? even in a culture medium? can there it be an interval of 36 hours between taking the sample an it reaching the facility?
I need to quantify the uptake of our therapeutic oligo drugs in the nuclei of muscle fibres isolated from adult mice. However these fibres have several satellite cells attached to them (see photo of an FDB fibre isolated from a 15wk old BL10 mouse) and therefore I need to remove them first in order to be certain that the oligo uptake measured in the nuclear fraction is from the muscle fibre nuclei only. Is there a way to remove or encourage migration of satellite cells quickly from the muscle fibres i.e. agitation, centrifugation etc?
I am doing an RNA-FISH protocol on mouse dorsal root ganglion tissue that contains cells marked with neuronal tracers. Mouse DRGs are extremely small and hard to see in OCT or Sub-Xero cryo medium but M-1 medium is more transparent. They have been cryo-sectioned between 6-10um for custom-made RNA probe binding but the thinner the section the worse the tissue integrity; there are often gaps or holes in the tissue section. Im thinking maybe the tissue should be a bit more sturdy when cutting, therefore pre-fixation (or at least stabilize the RNA before post-cutting fixation). Has anyone had any luck on binding their RNA probe to tissue that was fixed before cryo-cutting?
We obtained a Fluorescent end labelled oligoprobe for in situ hybridisation from a commercial source to detect viral nucleic acid. To prepare a stock solution from this newly synthesized probe what diluent is required? Can we use nuclease free water or any buffer is required?
Is it necessary to use RNase A in fluorescent in situ hybridization experiments with PNA probes? Could FISH be performed without RNase A?
Thank you very much
I used company's protocol for PI staining of human proximal tubule cell line. I fixed the cells using 70% EtOH and let them sit overnight at -20C, the next day followed staining protocol but the cell cycle histogram does not show a peak at G2M phase or there is a small scattered peaks that look similar to the S phase peaks. I have enough cell around 10^6 and I used 10ul PI for 100ul cell suspension. Could anyone suggest a good protocol where I can see distinct peaks for cell cycle phases. Also could someone elaborate the right setting on the cytometer for the PI detection like selecting A, W , H etc for appropriate analysis. Thank you
I'm now trying hard to map the specific localization of a certain fusion gene by In situ hybridization (ISH), but I don't know how long the probe should I take in detection of pathological tissue and what is the proper method of setting corresponding control groups. If, on the other hand, the specific signals were to be detected, what should I do to exclude the possibility that signals are come from the separate genes?
I have archived slides which I want to detect cytogenetic abnormalities by FISH and would like to know the number of years of storage for which the slides will still be ideal for use.