Science method

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.
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Hi, 
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).  
Protocol: 
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.
Thanks!
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Hello
Emma Zwilling
, did you ever find some solution?
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Dear All,
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!
Best regards
Binbin
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This background noise seems to be an intrinsic part of smFISH. The probes seems to bind to the protein or lipid.
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Hi,
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.
Thanks!
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In our experience with the smFISH probe, the concentration is 10nM. since DNA hybridization also has an affinity like several nM, you can also have several-fold lower concentrations and it will still work.
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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. 
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Our experience with DsRed shows a significant signal loss after 4% PFA fixation
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Hi all,
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?
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It doesn't sound like a good idea to use a fluorophore-labeled mRNA for imaging purposes inside cells. Instead, you would try a hybridization probe against your mRNA with their related amplifiers to make it visible. Otherwise, labeled single mRNA molecules will likely be stay below the signal detection threshold under a microscope even with 63 or 100X objectives. I have been using the RNAscope system to visualize endogenous mRNAs at single-cell resolution, which I think what you need. In your case, you may request your vendor to use synonymous codons for certain amino acids if need in order to avoid cross-labeling endogenous mRNA. But sample processing is quite tedious, though could be very accurate.
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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
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Hi Matthew,
How is your experience with AUM BIoTech? Does customer need to provide probe sequences? Thanks
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See the FISH protocols
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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
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
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Good question
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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?
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Alguien ha probado el Xmatrx® NANO de Biogenex?
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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.
Thank you.
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Hi Sreemukta. Thank you for your interest in my question and for your reply.
I did try fluorophore attached as well as DIG/Biotin attached probes but the problem with DIG/Biotin probes is that it has lot of background signal and fluorophore attached probe does not have strong signal. So I was curious if something like I proposed in my question is available so I can use primary antibody to detect the protein attached to the probe and then do conventional assay to detect the signal.
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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!!!!
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Hi Chrisopher,
I had the same problem with a couple of probes that worked separately but not together. I solved the problem by increasing the dithiothreitol (DTT) concentration in the hybridization mix up to 1M. Maybe this can help...
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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.
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It looks more like a speck which has acquired the fluorescent. Did you confirmed whether its amplifying the right DNA segments, probably doing a sequencing or restriction digestion of the probe sample. Another thing you could be possibly tune the hybridization condition. If its GC rich region try doing till 80degree and 37 degree ON incubation. Further, did you tried adding any control probe, CEP are the best to go.
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Following are probes required for fish analysis
1. AMX368
2. EUB338, EUB338II, EUB338III
3. NSO190
4. NIT3
5. Nsv443
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I used in my research the 16 Rna technique. There are many papers that you can see it.
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Hi
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). 
Thanks.
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Maybe, you can try the longer and more specific RNA probes for your target mRNAs.
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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!
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The most likely issue is that your nucleic acid is crosslinked to much - this is normally due to excessive length in formalin. You may want a more aggressive antigen retrieval, you could heat for longer if needed - we heat for 10 mins @ 121C - as long as morphology doesn't suffer you could be okay. You could also try using a high pH buffer like Tris instead of sodium citrate. Another approach would be to use a microwave or formic acid to uncross link your samples.
Fixation time may vary for sample to sample, but your target probe will also vary by how much it is affected by fixation, its annoying but you need to try and find a good balance.
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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!
Thank you!
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Something that is not mentioned yet is that you actually can underfixed tissue as well. I have tested small biopsies that were fixed 16h or less, which did not stain optimal in ISH, even they worked fine in IHC. It gave higher variability and more unspecific staining. I discussed it years ago with the old Ventana technical staff, and they had similar experience. For ISH, 1-2 days is optimal, >2d leads to diminished signals, <1d leads to higher background and uneven staining.
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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?
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I think the below reference is helpful
Malanovic, N., & Lohner, K. (2016). Gram-positive bacterial cell envelopes: The impact on the activity of antimicrobial peptides. Biochimica et Biophysica Acta (BBA)-Biomembranes, 1858(5), 936-946.
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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.
Yiyu
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Hello! Were you able to find a solution? I'm experiencing similar problems.
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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!
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Hi Cheng,
As I recall BACPAC offers construct design.
Well, maybe PNA FISH would be the best option for you then? In general, peptide nucleic acid probes enhance the signal greatly. Check out this paper and this resource for custom PNA synthesis http://www.panagene.com/_ENG/html/dh_product/prod_list?cate_no=3
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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?
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We use a more specific kit called RNAscope. It gives a higher signal amplification. They promote teh use of secondary staining such as H&E and we have used Toludine Blue O. I have not combined immunofluorescence, but as long as there is not an ethanol step used on slides developed with a kilometric kit, you should be fine. The alcohol decolorizes the slide. They do make fluorescence RNAscope kits as well. It is worth a try but you will likely at least have great FISH results.
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Does it impact the hybridization on the DNA target ?
Does it impact its stability ?
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Dear Amraoui,
The fluorochrome Cy5, when positioned in 5'end, can slightly improve the stability of the duplex as compared to the 3' end. Here is a reference explaining this property:
Best regards.
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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.
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You also can try F-ara-EdU to avoid toxic effect. FISH definitely must be used before click-IT chemistry!
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Can I only label mltiple intern sites in the probe?
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Hello, normally FISH technology not require quenching but oligonucleotide probes labeled with fluorescent dyes are used in a variety of in situ applications to detect specific DNA or RNA molecules and in some cases probe fluorescence might be quenched upon hybridization in a sequence specific way.
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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.
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You can try to add some Triton X 100 before you do FISH,
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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).
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As a target is typically bound by several antibodies, it might help to enlarge your "detction sandwich". For example, if you have a human-anti-XYZ antibody as primary ab, use an unlabelled rabbit-anti-human in the second step and a labelled goat-anti-rabbit tertiary antibody for detection. It might help, but might also result in more background signal as all existing signals get multiplied.
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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.
Thanks
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Hi Sebastian,
I would have immediately suggested analysis of 16S rRNA from environmental DNA by employing NGS, e.g. Illumina MiSeq. However, this is a PCR based technique. Nevertheless, any sequencing laboratory will do downstream operation after DNA has been isolated from the sample. I think you could even send a raw sample to them and they would do DNA isolation and the rest. In the end you would get a qualitative and quantitative report on the microbiome of interest. Of course, this kind of complete workflow and analysis comes with a price, you should ask for a quote to calculate your expenses and to check whether is it in line with your budget. Alternatively, you can perform part of the workflow alone to lower the costs, e.g. isolate DNA, and analyse sequences coming from he sequencing laboratory.
Another method that you could use for a rapid quantitative and qualitative analysis of your microbiota are the microarray techniques, which don't require PCR at all. Does a Phylochip ring a bell? If not, check it out (https://ipo.lbl.gov/lbnl2229/)!
  • "I am not sure about FISH as the technique seems to be more about looking at the occurrence of specific strains that you already know before-hand (and thus know the complementary DNA)."
Exactly, if you want to assess total microbial community composition FISH technique is not the way to do it.
  • "The abundance itself could be achieved by DAPI/Hoechst staining and fluorescent microscopy."
Again correct, but you will lack compositional information.
To obtain both, qualitative and quantitative information of total microbial community in certain environment, in my opinion, your best bets are molecular techniques mentioned previously. I would refrain myself from staining approach in this instance.
Hope this helps.
Deni
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Hi everyone,
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!
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Dear Meng-Ju Wu, Are you dealing with human constitutional samples, is the information on origin of marker chromosome available? in that case you can select BACs corresponding to that region, make FISH probes, hybridize and observe, methods for this are on [https://www.e-c-a.eu/en/WORKING-GROUPS.html]. If you do not know the origin of the extra DNA material, and want to stain with FISH, see the link for marker chromosome on above link, and contact @Dr. T. Liehr for methodology details. Hope this is useful.
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Does anybody know a lab in Australia or New Zealand who carries out in situ hybridization, FISH, GISH with polyploid plants?
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I think in Wellington Regional Genetics Laboratory, CCDHB New Zealand
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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?
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Thanks all of you to yours contribution. I would try to put in application all these suggestions.
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Hello all,
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. 
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Hello Cody Sharp,
I'm working with RNA probes for Mycobacterium since last year.I use clinical samples of swine lymph nodes, we use tissue paraffin-embedded, that are PCR positive for Mycobacterium avium, or M. bovis, or M. tuberculosis. I already used Lyzozyme, Achromopeptidase and Tripsina 0,1%. Nothing worked.
The Miguel's protocol worked?
I'm sending to you a picture of our FISH, and this is the only fluorescence that we got. We think this are the macrophage granules. not Mycobacterium. The fluorescen is in the health tissue, not in the necrosis.
Do you have some news, and better results?
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Hello,
I'm going to start my journey with Fluorescent in situ hybridization. Do you have some good protocol to FISH for miRNA on frozen human liver section?
Thanks for all!
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Dear Małgorzata Blatkiewicz
From the ResearchGate archives/Publications:
[just always search full text via "SEARCH" Upper menue line in ResearchGste (insert keyword, search phrase ore sentence(s) - click the loupe-icon and - after some second of waiting - choose from the sections: "Authors" or "Projects" or "Publications" or "Questions" ]. Naturally you can search also in Scholar Google (publications as well as recipes) or GOOGLE and any other data base (like PMC/NCBI/PubMed)
Here you go - to start with:
Standard Protocol
  • Cut 5um sections on silanized or positively charged slides
  • Dry at room temperature for 15-30 min
  • Fix in 4% formaldehyde in PBS at 0 ºC for 15 min
  • Wash with PBS and start the Paraffin Pretreatment kit protocol at the .2N HCl step and follow through to the end
Alternate Protocol
  • Cut sections as before and dry
  • Fix in Carnoy's fixative (3:1 methanol: acetic acid) for 5 min.
  • Begin the Paraffin Pretreatment kit protocol just prior to the Protease step (Wash Buffer for 5 min.)
and last but not least:
Preparation of Paraffin Sections and Frozen Tissue for FISH 2006
(no warranty for simplicity, easiness or guaranteed success, but - as always - confident in orthers' former tremendous work and scientific spirit).
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Dear Researchers,
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.
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Hi
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?
Thanks.
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Hi
Below is the entire protocolmy colleagues and I use now for X-gal and that seems to work. As far as I can tell, it was the xylene steps during the embedding. I have now shortened those to 5 minutes each, as opposed to the traditional 15-60 minutes (depending on tissue size, I mostly do mouse embryos). Still seems to work, albeit that you will have to check for possible artefacts, because if the tissue is not incubated long enough in xylene, they may not take up the paraffin as well and then they won't section as well.
For the few in situs I tried after my initial posts it seemed to have worked too, but I cannot be 100% certain since I only did a few of them. But since both procedures produce crystals, I suspect that the xylene affects them more or less similarly.
I have attached a link to a 'lacZ bible' I found online. Could not trace back the original author, but I did find that Andy Groves at Baylor College of Medicine had used the same protocol (https://www.bcm.edu/research/labs/andrew-groves/protocols). He was very helpful when I contacted him. Other than that, please see http://wmc.rodentia.com/docs/lacZ_bible.html, PMID: 28669819, and PMID: 1742021.
Hope this helps.
Best,
Piet
- fix embryos in 2% pfa, 0.125% glutaraldehyde in PBS on ice (5' for E9-10, 10' for E11-12) - wash cold PBS - I then keep them in PBS with 0.025% sodium azide at 4C if stored for more than a couple of days
- prepare and stain the whole mounts as described in the 'Lac Z bible' link: http://wmc.rodentia.com/docs/lacZ_bible.html  I haven't been able to track down the original author, but did contact Andrew Kelton Groves, who mentions it in one of his publications. As an alternative to the subsequent paraffin embedding, they do gelatin/sucrose embedding, which avoids the xylene requirement (protocol and additional explanations attached below). Unfortunately, our cryostat does not go below -22 C, and you need -30 to keep the blocks frozen, otherwise they become more like jelly and are impossible to section).
I always make the staining solution fresh and use X-gal (dissolved in DMF) that has not turned too yellow yet.
- I combine several embryos in PBS in one 5 ml glass vial (Fisher) on a rotator (individual embryos marked by removing e.g a limb or tail) - for E11 and older, make sure to puncture the thorax and abdomen to ensure proper incubation - rinse 2x 30'-1h and then overnight at 4C on a rotator - stain overnight at 37 C with pre-warmed staining solution (make sure the crystals are dissolved well) on a rotator inside an incubator - post-rinse for 10-15' at 37 C - the staining solution can be used multiple times when kept at 4C and filtered, but it does tend to stain weaker each time. - wash 3x PBS at RT - fix in 4% pfa/PBS overnight at 4C on a rotator (or longer, since overfixing does not negatively affect the paraffin sectioning). - wash PBS and dehydrate to 70% ethanol for long-term storage or further embedding. I also take pictures when they've been in 70% ethanol since it improves the contrast between embryo and staining.
For embedding: - wash PBS, dehydrate 30-50-70 on day 1 (if not already done so above) and then on day 2 80-90-95-100-100% ethanol/water (I normally do 20-30 minutes depending on stage) - max 5-10 minutes each (for up to E12, I haven't tried older stages): incubate 50/50 ethanol/xylene, 2x 100% xylene at RT and then 50/50 xylene/paraffin and 3x paraffin at 60-62C.
For sectioning: - I do 10 um sections, stretch well in waterbath, place on slides, dry well (overnight or longer).
For staining: - rehydrate: 2x xylene (up to 5 minutes seems to work very well) then 1 minute each ethanol and other step. I've found that most, if not all, of the remaining staining will keep during these steps. Gently dipping every 10-15 seconds helps too, but be careful because sections may partially detach. - counterstain with nuclear fast red (my preference) or eosin, and dehydrate again. Mount with Permount. Dry for several days.
If you just want to look at whole mounts, they do not need to be overstained, but just up to the point where it's strong enough and then fixed. I can get you a clearing protocol from a colleague who is doing X-gal followed by clearing to look 'inside' the whole mount. You will lose some staining however, so it's a bit of trial and error ( PMID: 27348591).
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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?
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Hi,
Possible reasons for these
1. Your ssRNA does not match with your probe
2. It may contain low GC and melt at relatively low temperature
3. No. of nucleotide that match with probe are not enough for fluorescence
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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?
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Have you looked at the ACD Company solutions. They do have quite "sick" in a good way things in their portfolio.
Best,
Ivan
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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.
Best wishes,
Astrid
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The slides were stored at RT in the lab. It is generally not dry in the lab- I would say it varies. If it would be too humid storing them, would that not also have an influence on the rest of the slide outside the hybridisation zone? The hybridisation takes place in a humidity chamber and there was no sign of the slide drying out.
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Hi,
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.
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same cy3 probe? which filter you are using for visualization?
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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?
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Dear Dr. Poczai
Please read the following article and I hope you find the answer in it.
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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?
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Hello Elisa,
Thank you for your comments. We are considering to test that suggestions.
Best regards
Marcos
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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!
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Dear Carlos, I have no experience in tge kind of samples you are assaying. There are some points however that you may find useful. Provided you are using water samples, it is possible you are handling a lot of photosynthetic bacteria in them. Photosynthetic pigments will of course be absorbing light in several portions of the spectrum and emmiting at some others. Getting rid of them through a selective culture step could help you. In the old times, people would get rid of some autofluorescence issues by incubating 5min with 1% evans blue. This would switch much of the greenish autofluo into the red channel, thus allowing them using Fitc-conjugated stuff.
There is still another option which is performing a lambda scanning of your unstained preparation and finding out which portions of the spectra do not autofluoresce. Once identified, you can choose some quantum dots as fluorophores with very narrow absorption and emmission spectra. Hope these comments would help you.
Best,
Daniel
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The goal is to preserve the mucous layer. The slides have been IF stained for a week now.
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Thank you Elliot!
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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.
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thanks a lot
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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
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if using human cells try smart flare by millipore
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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
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Dear Wojciech Witarski
I done Fish according my protocol ( metasystem)
but at last when i look slide under the microscope have not get result, I think Dapi didn't paint cells
can you help me?
What can be reason?
What could be the reason that the dipi can not contain cells?
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I need to stain cytoplasm with a fluorescent dye in order to see it along with a FISH reaction.
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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.
Any advice?
Thanks!
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Hi, thanks for the comments.
--- Well, It turns out the substrate was bed.
We got a new batch wich worked wonderfully.
Actually, It works much better than the ROCHE fast red tablets we used to work with.
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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?
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Pretreatment of FFPE sections is necessary because of formaldehyde fixation and its effect on nucleoproteins (histones) and DNA.
Nuclei are cut and "open" but DNA is in close combination with nucleoproteins, that "hide" the bindingsites of the probe and they have to be digested by the proteinase. So the probe can bind better. Heat pretreatment reverses the effect of formaldehyde on proteins. Methyloladducts are loosened again and proteins gain almost their native structure. This helps also the proteinase to reach its binding-sites. Digestion after heat pretreatment works more effective.
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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. 
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That's a tough one. I'm not a "sludge" expert, but unlike with cells in culture or tissue samples, you can't easily do a sodium borohydride treatment. So I think your options would be: 1) choose dyes that are outside the autofluorescent spectrum of the sample, 2) image with one wavelength left open (specific label in green, and an "empty" channel in red) then subtract one image from the other using imaging software to artificially subtract out the autofluorescent signal, or 3) photobleach the sample with very strong UV light prior to labeling in the hope that the autofluorescent compounds will fade and not come back.
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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?
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Hi Muhammad, I mix the nick translations together in the hybridisation buffer as a cocktail and put this cocktail after denaturation on the tissue sections for hybridisation. The detection of the two probes by immunohistochmistry (Anti-Dig and Strptavidin) I do seperatly. Depending on your protocol take care about the different blocking steps you need.
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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?
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Thanks Sabine Strehl and Pieter Louwe. 
These things have not worked for me. I must say I still need to give it a go with gelatin and collagen which I will do soon.
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Hi all, 
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
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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. 
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In my opinion there is no way around testing your probes by conducting FISH experiments.
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Hello,
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 !
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Hello Lola,
You are right in everything you just said. The 16S rDNA gene encodes the small subunit RNA of the ribosome (ssu_rRNA) in prokaryotes, while the 18S do the same in eukaryotes. However, eukaryotes are composed by more complex cells that includes organelles. One of those is the mitochondria, which has its own genome and synthesize its own proteins, so it needs ribosomes, hence it needs ssu_rRNA. Now, according to the endosymbiotic theory (or symbiogenesis), mitochondria has bacterial origins (in fact the eukaryotic cell was the product of the endosymbiosis of bacteria within archaea -- see attached publications for more details), and that is the reason why mitochondria has a prokaryotic ssu_rRNA (i.e. 16S). This subunit is purely RNA -- it does not encode any protein, which is the reason it can has stop codons within because it is never interpreted in terms of codons.. it is just not translated, it is simply folded as RNA and used as such as part of the ribosome complex.
And yes, only the egg cell has mitchondria, not the sperm cell.
Relevant scientific literature:
Williams, T. A., Foster, P. G., Cox, C. J., & Embley, T. M. (2013). An archaeal origin of eukaryotes supports only two primary domains of life. Nature, 504, 231-236.
Martin, W., Roettger, M., Kloesges, T., Thiergart, T., Woehle, C., Gould, S., & Dagan, T. (2012). Modern endosymbiotic theory: Getting lateral gene transfer into the equation. Endocytobiosis & Cell Research, 23.
I hope that helps. Best regards.
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HI.I mean can we use it like FISH that with a software it can calculate percent of aneuploidic cells in samples like sperm?
thanks for your comments.
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Thank you Houshang Nouri.
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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!
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I have do FISH in lymphocytes stored at -20 in fixing solution for a long time without problems
luis
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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?
Thank you.
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Yes, it helped very much.
Gracias...
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Hi,
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.
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by FISH, you cant be sure  that the signals you found are from your probe as its length is 250bp, the big problem of Fish is the diffuse, nonspecific signals found after hybridization. you can overcome it by trying techniques for FISH pretreatment that can decrease these problem, also effictive post hyperdization wash method.
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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.
 
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I am also standardizing Immuno-RNA-DNA FISH and could help you after two weeks I think. But as ann answer of your third question, I would suggest using 10ng/ul probe and 100ng/ul Cot-1 DNA and dissolve it in 20ul of Hybridization buffer (this is for one reaction of FISH). This is not mentioned in the paper but I have got this information straight from the Lee lab. I will upload the detailed protocol tomorrow.
Hope this would be helpful.
thanks
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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? 
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Hello Chrysanthi,
Maybe you have to try which approach might work better, a fixation with paraformaldehyde before sorting or once your cells are on the slides. You can also do the fixation on the slides, just ensure your cells adhere to it. In principle the protocol for tissues and for cells (on filter) should be the same. Maybe you can have a look here https://www.researchgate.net/publication/246206760_Fluorescence_in_situ_Hybridization_FISH_with_rRNA-targeted_Oligonucleotide_Probes. Still find this work by Pernthaler et al. very helpful.
Best regards!
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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.
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I'll not help you, excuse me. Theoretically, chlorophyll could be washed out in ethanol or methanol (that was used several years ago to wash excess of DAPI from membrane filters) but we did not reach perfect results. Moreover, we had even worse experience with picocyanobacteria pigments, which are water-soluble.
However, our experience indicates that the fluorescence of such pigments was not the same using different FISH probes. You should take it into account - the error is not the systematic one.
The alternative fixatives that could switch off the fluorescence: acid Lugol postfixed with thiosulphate & formalin (Jezbera et al. 2005). But not every time (my tool - ciliates- could have more unspecific red fluorescence)
Bouin fixative -picric acid in formalin + acetic acid (Fried et al. 2002). But it stains larger object too much to be observed (it is usefull for small ciliates and their food). Generally, the fluorescence is less bright but the background is perfectly black if the preparation is made on membrane filters.
Do not hesitate to contact me if you are interested in
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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?
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Yes, usually it is used a rotary vane pumps. But I learn that maybe a stroger vacuum is better (about 2 x 10-3 mbar / 1.5 x 10-3 Torr)
Thanks.
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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!
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ohhh. Thanks a lot. sorry for my delayed response. But these are really helpful. I will check them! Thanks again
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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?
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Which concentration of serum do you use ? Try to decrease the number of cell seeded in order to avoid contact inhibition (even if you  treat cells before they reach confluence)and to boost cell proliferation. I disagree with your protocol of serum starvation. Serum is undispensable for fibroblast multiplication. (See papers of HAM et al. on cell nutrition types). In my opinion, your problem is linked to the absence of serum.
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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?
thanks!
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I agree with Sonal opinion
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Before FISH analysis, cells are fixed in methanol:acetic acid
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technically yes as they wrote before, just take care, your DNA might be dirty or in insufficient amount, good luck 
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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? 
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I agree with Denise. In my case It has been about twenty years since I isolated satellite cells to grow but as I remember it involved digestion and d damage to the fiber (myocytes). Can you use fluorescent or immune fluorescence microscopic approach to both distinguishing the satellite cells from the myocytes and also for measuring drug uptake?
You may consider using myogenic stem cells and creating in culture myofibers (myotubes). I have been inducing myogenesis in C2C12 mouse myoblasts quite some time, it is easy.
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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?
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A few points:
  • For RNA-in situ you MUST fix the fresh tissue first in a cross-linking fixative such as formalin to assure 1) that the RNA is intact and 2) it does not migrate/mobilize away from its source cells.  RNA-Later or similar products do not do this; they only "stabilize" RNA chemically from degradation so it can be released in a homogenate.  For in situ techniques immerse in an abundance of fixative overnight.
  • 5-10 microns is thin for cryosections on OCT.  (I have never used M1.) Try 10-20 microns; you will get better quality sections and not waste your sample or custom probe.  Later as you improve the sections can get thinner, but the  thinner the section, the less intense the RNA signal (you reduce the amount of target to which your probe can bind).
  • Did you cryoprotect in 20-30% sucrose prior to embedding?  This removes water and helps combat ice crystal formation, that can cause poor section quality.  This step is often used with delicate neural tissue.
Good luck with your experiments.
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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?
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Hello,
What is the commercial source for these oligo probes and did they work. 
Thanks 
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Hi everybody,
I plan to perform FISH (telomeric and sentromeric probes at the same time) in human peripheral blood lymphocytes. Do you recommend using pepsin (is it compulsory??) or can I perform without using pepsin?
Thank you very much
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Dear Erman Salih Istifli,
Pepsin is the very important component because it's a protease and acts to digest proteins into peptides and it helps to digest away cytoplasm on the slide. Digestion with pepsin can significantly improve probe penetration. Pepsin treatment is often useful when specimen preparations are suboptimal,e.g., in many clinical samples.
Best regards,
Maxim
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Dear All
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
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Hi,
No it's not necessary. See below. Which PNA do you use and what is your application?
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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
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      I wouldn't necessarily expect to see big, distinct peaks for G2/M or S. Most cells don't like to sit in G2 or M for very long. These phases can be pretty short, so you might not have a lot of cells in those phases at any given time. Do you synchronize the cells in your cultures so that they are all progressing through the cell cycle more-or-less together? This might yield tighter peaks.
   If you need to visualize cells in different phases of the cell cycle in the microscope, the following might be useful.  We've done experiments on cultured smooth muscle and fibroblast cell lines using Ki67 immunolabeling as a marker for cells in the cell cycle in combination with other cell cycle markers (cyclins) and DAPI to visualize nuclei and chromosomes for visualization in the microscope. The Ki67 and cyclin staining worked beautifully using fixation with 4% paraformaldehyde in 0.1 M PBS. (PI labeling also works under these conditions, although that isn't what we used for these experiments). EtOH fixation typically doesn't produce as a good a result as PFA does and can extract a lot of stuff from your cells. I would recommend EtOH fixation only if PFA doesn't work.  We had very good luck with a couple of different Ki67 antibodies from Millipore and from Cell Signalling and a cyclin antibody kit from Cell Signalling using PFA fixed cells.
   Another approach that can be used in the microscope is EdU labeling (EdU is a BrDU analogue that gets incorporated into the DNA when it is replicated in S Phase). Molecular Probes has some very good fluorescent EdU kits.
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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?
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this eel observed in lagoon in western Sri Lanka. longer than 3.5 feet 
if you have any idea please help to identify 
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I agree with Philippe as well; the Giant estuarine moray, Strophidon sathete (Hamilton 1822) (family MURAENIDAE) matches with having a small head and a very long tail. The species lives in burrows in mud bottom, usually in estuaries and river mouths, in the Indo-West Pacific from the Red Sea and East Africa east to the Philippines and New Guinea.
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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.
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That is correct. I have been able to FISH on slides that were 2 years old- but they were stored at -20 with drierite chips to reduce humidity. If you have spare slides you may want to try the following that I have used on nuclei extracted from paraffin blocks that were quite old. Treat the slide with 50%glycerol in 2XSSC at 90C for 5-10 min. Dehydrate the slides in thorough ethanol series of washes- 70%, 80% and 100%. Look at the morphology of the cells - in a phase scope to see that they are still intact. I had to vary the treatment time depending on how old the blocks were. Then use your standard FISH protocol.
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