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We will collect mouse fecal samples and perform 16S RNA analysis to monitor germ-free status. Looking for any protocol or resource recommendations. Thanks in advance.
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Hi Rahman
I have performed V3/V4 microbiome sequencing in my study. I used to keep individual animal in a freshly cleaned and sanitized cage without the bedding material. The animals should defecate in 2-3 minutes, collect the fecal pellets (2-3) sample using a pick and store it in labelled containers. Keep the samples in dry ice or -80 till further analysis.
Hope it helps..!!
Thanks
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I'm in the initial stages of planning a miRNA seq experiment using human cultured cells and decided on TRIzol extraction, Truseq small RNA prep kit, using an illumina HiSeq2500. The illumina webinar suggests 10-20 Million reads for discovery, the QandA support page suggests 2-5M, and I wrote the tech support to ask, who suggested I do up to 100M reads for rare transcripts. Exiqon guide to miRNA discovery manual says there is not really any benefit on going over 5M reads. I was hoping to save money by pooling more samples in a lane, so I was hoping someone with experience might be able to suggest a suitable number of reads.
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i am working on cardiomyopathy patients Blood samples . and wanted to do miRNA sequencing can some one please suggest how many millions reads i need to sequence 20 millions or 30 millions and also please suggest the platform as well .
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I am trying to isolate neutrophils from mouse bone marrow with percoll (62%). The goal is to stimualte the isolated neutrophils with LPS and afterwards lyse the cells for RNA analysis. I wonder what is the best temperature to work with during isolation (on ice or room temperature) since I read that neutrophils are not easily responding when put on ice? However working on room temperature comes with a cost of activation during isolation?
I also wonder who is experienced in isolating RNA from these cells?
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Hi,
Isolation of neutrophils using percoll should be done at RT for sure. Once you isolate, put the cells in media with FBS if you want to stimulate them with LPS. Within 2-3 hrs, cells get adhered. After you are done with the stimulation, you can use this kit for isolation of RNA (considering these cells are sensitive and low life span):
PicoPure™ RNA Isolation Kit, Catalog number: KIT0204.
Good luck!
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I have seen many papers and discussions here about various techniques for isolation and culture of microglia and astrocytes from adult mouse brain. Isolation of these cells by Miltenyi magnetic beads is supposed to yield a great purity and have been used for short-term culture (6-24h?) and RNA analysis. Can anyone give me feedback from their experience about separation of cells using Miltenyi beads and culturing these cells for a week (or at least up to 72h) , as described in the Miltenyi protocol?
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I am trying to culture mixed glial from adult mouse 2-3 months old. I can only get microglia but not astrocytes. But when I culture from p0 mouse I can see astrocytes and microglia. I am using DMEM -f12 media with antibiotics. Using PDL coated dishes. Also I am using papain for digestion for the tissue. Does anybody have insight on this?
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which RNA should be studied to make a comparison between cells grown in 2D and 3D constructs?
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Either I'm not clear to you or you can provide more details abt the context of the question. What you asked abt which RNA, I answered MESSENGER RNA (mRNA) should be studied. Very sorry, couldn't able to what I have misunderstood.
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I am separating neuron populations from mouse brains using FACS. I am trying to find a fixation technique for intracellular antigen staining and FACS that will allow long term storage afterwards for RNAseq. I have looked at ZBF which allows long term storage of cells with little change in RNA integrity, but they have to be taken out of the fixation buffer for flow, so RNA integrity will likely be affected during flow and I am not sure if they can be 'refixed' after. For any other fixation methods I have read that mention the shelf life of cells, they refer to how long they can be stored BEFORE staining and FACS, and do not mention any storage AFTER facs.
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After the FACS, you can centrifuge the cells down to remove the solution from the FACS running, and then add the RNA later onto cell pellet and then store your cells at negative 80 for later RNA isolation. For RNA isolation, just do the same thing centrifuge the cells down after thawing, then remove the RNA later and go on with RNA isolation kit instruction on the cell pellet.
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Hello,
I am going to perform some saliva's RNA analysis. I found in the litterature several protocols to collect and prepare saliva for RNA/DNA analysis. For main of them we have to use kits and there are a bit long. I also saw one using an extraction buffer containingTris HCl EDTA SDS for a whole night at 56°C.
I would like to know if there is a quick and simple protocol to prepare saliva for RNA analysis.
Thanks for your help.
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Select a viral kit specific for the virus that you are using for your experiment.
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Recently our lab has started a project where we are trying to capture non-coding RNAs (<200nts) by 4SU labeling. This will be my first time working with RNA or doing any sort of labeling. I am trying to find out what step has gone wrong.
I am labeling adherent cells with 1mM 4SU for 5 minutes. I extract using Trizol and then purify either with ethanol precipitation or Qiagen columns. Then I later biotin-HPDP label and pull down with magnetic streptavidin beads. I QC after all my steps and I have RNA going into the beads, but nothing is eluting out with BME or DTT despite various lengths of time/concentration/temperatures. 
I then wondered if the biotin labeling was efficient so I did a dot blot with 4SU labeled and not labeled RNA that both have been exposed to biotin. I see faint dots for all samples at 1ug concentrations, but nothing lower than that so it might be residue biotin and not actually labeling. I then went to see if the 4SU is actually being incorporated by use of nanodrop, but I don't see a peak at 330nm. I'm not sure if there just isn't enough 4SU labeled RNA out of the total RNA (I have tons of RNA). 
Can anybody see where I might have gone wrong or what I can do to fix this? Any more QCs I can do to try to figure it out? 
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Hi Ivana,
yes we did, we used INSPEcT pipeline (De Pretis et al. 2015) on RNA Seq performed on total and newly transcribed (4sU label led) RNA to infer transcriptome wide data. If you would like to test by qPCR (quality control before RNA Seq) you can use the method presented in Doelken et al.2008, that is how we optimized the protocol and confirmed what we were enriching for in the pull down step really was newly transcribed RNA
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I am starting a new RNA-seq based project. I will be looking at both coding, and non coding RNAs. To enable me to look at both of these RNA species simultaneously, I will be ribo depleting the samples to remove rRNA. My question is will my RNA-seq data have enough coverage if I want to look at splicing changes. I realise when people usually do this sort of analysis they Poly A enrich samples for mRNAs. I could split the samples in two and do both ribo depletion and poly A enrichment independently but this would cost double for RNA seq. Any help or ideas much appreciated.
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follow
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I'm a molecular biologist, and i have a few projects coming up in transcriptomes and small RNA analysis. Can i get by without knowing any programming using user-friendly software such an Geneious Prime or another program you can suggest or is it absolutely a must?
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Hi,
To be an efficient bioinformatician, you need to learn at least any programming language. You need not[ be high-end developer, but at least know how to do your bits. Also, you can be comfortable and can use various user-friendly GUI, but it would need more time and space, whereas in coding you can customize, according to your needs.
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RNA Analysis
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Based on what I have read above, if we are collecting cotton swabs from patients, they can be place in 1XPBS with 1% TX100 for transport? And then the swab can be placed in the first solution of a RNA isolation kit? Would that work? Will dead viruses still be on the swab in the presence of 1% TX100?
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I performed agarose gel (1%) for PCR products of COX-2 from saliva samples of patients suffering from periodontitis. I could not see any bands and therefore in order to check if the RNA was isolated appropriately, I also performed agarose gel electrophoresis and again could not see even a band of RNA. Can anyone suggest where we are making mistake? 
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Revise the whole experiment from the first like sample collection and the final stage. Depend on other person is not good.
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I'm currently investigating a lncRNA and i'm very curious to see what the subcellular location is due to silencing strategy and prediction of lncRNA function. Could i somehow separate the nucleus from cytoplasm and isolate/qPCR to find out where expression is present?
I was looking at fractioning protocols, but these are mostly aimed at proteins, if at all possible, could you recommend me a protocol for RNA analysis?
Huge thanks in advance!
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Erik van Butselaar hi?did it work well for your cell?
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I am doing RNA immunoprecipitation, using the RNA TRAP technology, with which you can capture actively translating ribosomes with the L10 ribosomal subunit fused with GFP, using GFP antibody coated magnetic beads. Right now I'm testing it on HEK293T cells transfected with L10-GFP and I am having trouble finding good reference genes / controls for qPCR.
GADPH & b-actin seem to be differentially expressed between the IP & the unbound (the supernatant of the IP) fractions, which result in very different & unreliable results when I look at enrichment of GFP between IP & unbound. For example, b-actin sometimes shows Ct value difference of 5 between IP & unbound fraction, of the sample.. Actually b-actin seems to be 'enriched' in the IP sample (so I suspect it binds to the beads..)
What do you suggest I do? Should I compare with the input sample? n
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Hi, I know this topic may be a while ago. But I recently gained some knowledge from somewhere else about what should be used as RIP-qPCR controls.
Theoretically, there is "no" proper controls when assessing RIP-qPCR, since regular internal controls such as gapdh, actin, tubulin, they should normally not be bound by the RBP of interest, which in theory, should not be in your IP-ed RNAs. I got a calculation chart downloaded from Sigma to show how to calculate ChIP and RIP-qPCR fold change, which is, you have your input, anti-RBP, and anti-IgG RNAs, input here just serves as one kind of control that amplicons will be amplified using IP-ed RNAs can be amplified in your input. Then, the comparison should only be done between anti-RBP RNA and anti-IgG RNA (none of them should contain gapdh, actin or tubulin), but you know they came from the same amount of cells (you control this), then the only thing you should do is to directly test your amplicons of interest, and calculate the (fold compared to IgG) of your anti-RBP IP-ed RNAs. Since anyway in a regular qPCR, internal control genes are only used to normalize RNA amount due to different cell numbers to start with.
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We are doing a pilot study and are stuck on the workflow needed. The starting material is plant and fungi material in RNAlater and end goal is making libraries with the Illumina Nextera Flex.
We are doing the RNA extraction with the Qiagen RNeasy Plant mini kit. We have a couple of options in mind for the first strand cDNA synthesis but which one is best is probably dependent on how we'd get second strand cDNA.
So in short how do we go from RNA to second strand cDNA that we can use for library prep?
Also at what point do you do quality control and how? We have most machines available to us (Bioanalyser, Qubit, Experion) but no kits for RNA analyses. Ideally we'd want to keep costs as low as possible.
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The kits and method/protocol for the double strand from mRNA is easily available and commonly used, you can find it easily.
I would recommend to use Bioanalyzer but if you are making cDNA, quality control is not that critical as if you were doing RNAseq. You can simply check the quality on agarose gel (MOPS). I guess, for the cDNA double strand library, just qubit (DNA) quantification would be sufficient, and generally there is not much quality issue in DNA libraries, provided standard protocols have been followed.
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Hi,
I am trying to purify synaptoneurosomes for RNA analysis from 4-week mouse brains. I am using a Percoll density gradient followed by washing 2X in buffer to remove residual Percoll (Dunkley et al., 2008). From the pellet, I have tried Trizol extraction as well as extraction using RNeasy Mini kit, and neither have been working. Does anyone have any tips or tricks for RNA extraction from synaptoneurosomes?
Thanks!
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hope these articles might be a help. Ill have to agree with Dr. Mickey Trizol method works perfectly, the overnight trick would work indeed.
Best of Luck
Kat.
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So I've been doing research on PBMCs isolated from buffy coat, infecting them and then extracting RNA. I've been getting very low amounts of RNA (maximim I've had is about 16ng/ul, but mainly it is below 5) despite using up to 10 million cells per well and then checking the number of cells on the plate by the microscope to check they've been removed. Does anyone have any ideas on how to increase the RNA yield? Myself and 2 previous phd students have found the same. Thank you!
I've used both the Qiagen RNA extraction kit and the Nucleospin RNA extraction kit
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Could you please too send me a trizol method protocol you use? Thank you! Valeria De Arcangelis
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Dear colleages
We have carried out a species identification by 16S RNA analysis, and the result is indicating a coccus bacterium, yet our microscopic observation shows that the strain is a bacillus (we have repeated the sequencing 2 times).. We are wondering whether there are reported examples of such discrepancy? Many thank in advance.
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Hi Alexis,
There are three things I can think of:
1. Bacteria was misidentified when sequence was entered into the database. You can check in using NCBI blast and asking getting the taxonomy tree output. If even if its misidentified, those branches closest too it should be correct.
2. Bacteria can have morphological plasticity, especially when grown in stressful environments (high salinity?).
3. Mutations can give rise to cell well structural changes that result in changes in morphology.
Regards,
Frank
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Since MOPS is one type of Good buffers, can it be substituted with any other? Can I be using TAE/TBE for RNA gels?
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Don't use TAE/TBEin case RNA as these buffers loose buffering capacity below pH 8.
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My study is on using microbial DNA present on fingerprints as identity markers of humans. In this case, how high is the accuracy of identifying the phylogenetic DNA of bacteria present on a fingerprint with the use of crude DNA extraction? I am only familiar with the protocol based on other papers - how do I acquire enough DNA for PCR
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I was referring to just the basic swabbing technique.
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I would like to do neurite outgrowth analysis and live-dead viability assays with Calcein/Ethidium Homodimer-1 on PC-12 cells. I would like to know if anybody has tried these stainings prior to RNA isolation with TRIzol reagent. I seed my cells on a dark-colored scaffold, therefore I'm planning to use Calcein-AM to make visible the neurite structures while neurite length quantifications.
 I am wondering if it is possible to isolate RNA after calcein staining and if the dyes cause decrease in the RNA quality/stability or puts them under stress (so that the RNA profiles of the target gene would be disturbed). I'd be glad to get any comments. Thanks.
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Dear Sarah and Jasmin,
Eventually I did try this but the scaffold I seeded my cells on was quenching fluorescence, so it was of no use for me. Now I don;t work there anymore, but would be interested to know the answer, if someone else tried.
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Hello,
I am just learning about PCR-DGGE to identify microbes in food samples. I am not from microbiology background, so I have difficulty in understanding some points.
I have some doubts when it comes to interpreting the results.
For example, in ice wine sample
KT323272 Penicillium sp. is identified - in RNA and + in DNA
and
JX681079 Coniothyrium cereale is identified + in RNA and - in DNA
The paper I am reading now is Novel insights into microbial community dynamics during the fermentation of Central European ice wine. The pdf is attached here.
I hope some one can help in understanding this concept regarding DGGE.
Thank you in advance
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Welcome....
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Hi,
We are looking for some advice on improving our method of detection of Mycobacterium species from fish tissues. We have fish tissues with granulomas and can see presence of acid-fast bacteria in the granulomas. We can extra total DNA and but when we run a Eubacterial PCR we get either very faint bands or nothing. I wondered if the problem was inhibitors at some stage since we were working with formalism and alcohol fixed tissue but when I add in Mycobacterial DNA to the recovered total DNA (ie essentially spike the sample with a known concentration of high quality bacterial DNA) we get a nice positive band. This makes me think that the assay itself is working but we are unable to detect any Mycobacteria in our sample either because its not mycobacteria (possible but unlikely given the pathology) or its at such low levels we are unable to get a quality result. I was hoping to send the PCR product for 16S r RNA analysis to confirm if we have Mycobacterium but am struggling. I have done this successful with Gram negative bacteria including from formalin fixed wax embedded tissue sections but really struggling with the Myco - any suggestions gratefully received!
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you could try a second round pcr with a nested primer or primers with the 3' end of the primer 10 bases inside the existing primer sequence. dilute the first product 1in 100 and do not run too many 15-20 cycles of second round pcr
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I have a RNA sample with 300 ng/uL of concentration and 260/280 as well as 260/230 ratios higher than 1.9. I ran an agarose gel to test the quality of my RNA sample and my loading buffer had a denaturing agent formamide. I followed the protocol mentioned in one of the articles about RNA electrophoresis. It says that I can run my RNA on agarose in normal TAE provided I have formamide in my loading buffer. However, I do not see any sort of band which might correspond to my RNA nor anything in the wells. What problem could this be that I am not being able to understand?
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Hi Anand,
I am facing the same problem now. I get good RNA concentration about 300ng/uL with 260/280 ratio from 1.8-2. However, when I ran it on 1.5% agarose gel in 1X TBE with a positive control, the control can be observed but none of my samples are visible. No smear or any bands. Very clean lane. New gel and buffer was used.
You had mentioned the same problem a year back. How did you solve the issue? Any help would be appreciated.
P.s. Previously I have extracted RNA from cell cultures and ran it on the same gel composition. I never faced this issue.
Thanks,
Varsha
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Hi!
I have mRNA after in vitro transcription. Nanodrop shows conc of 4150 ng/ul, (measured at 10x dilution, 415ng/ul), but Qubit and Tapestation show 10 times lower concentrations. The gel from Tapestation shows very weak band. I do phenol chloroform purification after IVT which is suppossed to remove most of the free nucleotides. The A260/280 ratio is 2,05 and A260/230 ratio is 2,34. I have no idea where does this difference come from. I tried different Nanodrops and different Qubits...
Maybe you can help?
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This is an old question but for the sake of people searching for this answer, here is the reason:
Nanodrop is just a UV spec reading. It is only looking at absorbance at 260 nm. Plenty of things absorb at 260 nm other than the RNA you make in your IVT reaction. This includes the T7 protein, pyrophosphatase (if you use it), your DNA template, the NTP monomers, the abortives, truncates, and even salt. Your nandrop is giving a signal for ALL of these things.
Qubit is better. It gives a reading based upon an intercalator that is specific for RNA. This means that it will (likely) only give signal for your full length RNA, abortives (above a certain length), and truncates.
This explains the disparity between your nanodrop and Qubit readings. And btw, a reading of 4000 ng/uL on the nanodrop is too high to trust. 10 fold dilute that and try again.
Better than Qubit would be to actually purify your RNA (DNase followed by EtOH precip/dT/etc., ultrafilter, recover) and then run the final material on a gel or capillary electrophoresis or HPLC and then compare against a standard curve.
Pain in the ass, but it's the best way.
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Hello everyone.
Right now i am working on differential expression of miRNA using next generation sequencing. analysis post sequencing was done using "Small rna analysis" pipeline in a integrated analysis software. I have disagreement with my supervisor regarding the results of dysregulated miRNA in terms of normalization method used. I use small rna analysis pipeline, because it was explained in the software about this pipeline if i want to analyse miRNA differential expression. Many journals also use this pipeline to find differential expression of miRNA. Regarding normalization method, i also read that method of normalization for RNA seq can be used for small RNA seq, for example DESeq/edgeR, TMM etc. but my supervisor need more assurance because he wanted normalization method specifically for miRNA sequencing. Since i am new in this field, i need some confirmation from experienced people in this forum.
1. Is it right for me to think that miRNA sequencing is the same as small RNA sequencing, and almost the same with RNA sequencing (just different on enriched small RNA annotation)? if
2. if previous question's answer is yes, so is it okay to use normalization method usually done for RNA seq,for example DESeq etc, and what is the reason for this?
if the answer is no, so what is actually the difference between those sequencing, and what kind of normalization can be used specifically for mirNA sequencing?
Looking forward for any link or elaboration from experienced people here in this forum.
Regards,
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Hi Dr. Ar,
To briefly answer your question, yes you can think of miRNA differential expression analysis as the same as RNA-sequencing, but only as long as you verify the distribution (i.e. if using a parametric fit-type, the data must fit a negative binomial distribution).
I have used DESeq2 in R to analyze miRNA-seq data, to both normalize the miRNA levels within each sample and compute differential miRNA expression. Because of the small sample size within each group, I calculated dispersion estimates via maximum likelihood, assuming that genes of similar average expression strength possess similar dispersion. These gene-wise dispersion estimates were then shrunk according to the empirical Bayes approach, providing normalized count data for genes proportional to both the dispersion and sample size. Once the aligned count matrix was normalized, I could then compute differential gene expression with these normalized read counts (as described above), the log2(fold-change) according to the Likelihood Ratio Test (LRT), and a Bonferroni-adjusted p value (i.e. q value) were generated for each aligned and annotated gene.
I have attached the R script for you to reference if interested, which outlines part of my miRNA-Seq analysis workflow.
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I want to make some single-cell RT-qPCR or scRNA-Seq experiments. But, I'm affraid, that during tissue dissociatation and FACS sorting are cells stressed, and it's activate expression of some genes, which we want to study.
So, in my idea, I want to first make tissue fixation, after that dissociate cells and continue with sorting/analysis. Have anybody experience with protocol like this?
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I agree with your second issue. For droplet based methods you require cells in suspension.
I also have to admit that I heard rumors that RNA quality is low for microdissected samples. However, MMI is using a low-power laser to avoid any DNA/RNA damages. Importantly, there are plenty of publications using laser microdissected samples for RNA analysis (spatial transcriptomics, RNA seq, ...)
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Hi,
I want to isolate exosomes for RNA analysis from blood serum. Blood samples take about 5 hours to reach the laboratory. İn which conditions can I transport blood samples properly????
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Do you want to get RNA from whole blood? Then it should not be a problem. But consider that you will most likely have emolisis and that many circulating RNAs (not associated to your vesicles) will be found free in that suspension. In order to separate the vesicles from contaminating RNAs the best technique that you can use is chromatography and then use some hemolisis controls (microRNas). Finally, at least once, you should check that in your exosome pellet you don't have Ago2, since it is a protein often found circulating and associated to microRNAs.
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Hello All,
This is the image of Total RNA non- denaturing gel electrophoresis. The 2 rRNA bands (28s and 18s) are prominent with 2:1 intensity but there is no smearing in between them that indicates presence of mRNA, so was the mRNA lost during extraction or what could be the possible reason for the disappearance of mRNA? Can I proceed the remaining Total RNA extract with In-vitro Transcription?
Thankyou.
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Elar: I would caution against replying to 3 year old posts, but since you asked: RNA can be run successfully under normal (non-denaturing) conditions provided you denature it first.
Prepare a 1% agarose gel as normal (for DNA, etc).
Prepare a loading mix with 1-2ul of your RNA, 2ul of 5x DNA loading buffer, and 6-7ul of formamide.
(formamide is a denaturant, so you want to aim for 60-70% final)
Heat this to 65 degrees for 5 mins (this denatures secondary structure: 65 degrees is the hottest you can really get RNA before it begins to degrade).
Remove from heat block, place IMMEDIATELY on ice. This prevents secondary structures reforming.
Load these samples on your gel, run it.
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Hi everyone,
We are planning a RNA analysis of expression of mRNA of an interest protein. The primers that appear to be the most reliable are Taqman primers, suggested to work better with Taqman Polymerase Mastermix.
However, we already have a master mix from Agilent (PfuUltra II HS DNA polymerase). Has anyone tried using the Taqman primers with other polymerase types with success? We are afraid to ask the manufacturers as, of course, they will prefer us to buy their own polymerase.
Thank you in advance.
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You can choose a wide range of compatible polymerases with hotstart feature in-built. However, the most important point to note is the content of the master mix especially Mg2+, Na+ and other molecules being used as PCR enhancers. Else, you can do a Mg2+ titration using the same master mix, your new polymerase, taqman primers and your positive template to achieve the desired RFU.
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I am having a bit of trouble trying to obtain good quality and decent amounts of mRNA from ~5,000-10,000 FACS sorted cells.
Tumours are being dissociated to a single cell suspension, stained for the markers of choice, and sorted by FACS in to media containing 20% FCS and spun down. I've also tried sorting in to lysis buffer directly. Using the QIAGEN RNeasy micro plus kit I get low yields and fairly poor quality RNA. Same goes for TriZOL LS.
I seem to be fine with cell lines which have not been sorted. But I fail to see why FACS sorting would change things so dramatically.
Any suggestions very welcome! Thanks.
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A. don't bother trying to see RNA from that amount of cells. with the exception of certain cell types that are very sturdy and large like TAMs - even the pico chip won't help.
I validate that I have workable RNA with logical results by RT-qPCR.
B. Try the RNAqueous-micro kit from intvitrogen. 
C. the choice of library preparation kit is a significant one. Look into what Takara are offering. 
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hello i am trying to do  qPCR for a specific microrna from a virus and i want to confirm the mature microrna and also the pre-microrna. so i want to know how to designer primer to amplify the pre-microrna?
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exiqon LNAs might help.
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Hi, 
I extracted RNA from Penicillium infected plum and nectarine tissue using the Qiagen RNeasy Plant Mini Kit. Concentrations from nectarine samples were good (67.6 - >200ng/ul using Qubit Fluorometer) but low for sample from plum (4.4 - 34.6ng/ul). I presume phenolics were the problem since the darker (red) the tissue samples the lower the concentrations.
Would it be worth sending the low concentrations for analysis on the Bioanalyzer or Experion or will it just be a waste of money? The budget is tight and another kit to counter phenolics/sugar/salts will add to costs. I can load maximum 66ng RNA from the lowest concentration for cDNA synthesis. I will probably end up loading 50ng for all the plum samples. Would that be okay considering we need to publish? 
Thanks for your input,
Pieter
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Hi,
So I submitted 24 samples to the Bioanalyzer. The first 12 samples had low concentrations whereas the second set of 12 had moderately low concentrations based on the Qubit. 19/24 of the samples had acceptable RIN values for qPCR (RIN>5). Looking at the "Electrophoresis File Run Summary" the samples seemed fine. I was glad to see that I don't need another kit. The 5 samples with low RIN values will be reisolated and resubmitted. 
I know calculating concentrations using different methods yield different results but some of the samples had >3 times higher concentrations on the Bioanalyzer. Maybe I did something wrong? I assume the Bioanalyzer is more accurate so will use the new consecrations for cDNA synthesis. 
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I grew up bacterial isolates, then did DNA extraction using spin kit for soil, which I have used many times.
This time I appear to have got more bands than just the genomic DNA, why would I have got this and what can I do?
I think there was a lot of starting bacteria scraped off the plates, the nano drop showed an error message so I couldn't get readings but I suspect they are high.
Thanks
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I agree with Eric Marques, you should contact the kit manufacturer. Further before contacting, kindly check expiry date of RNase. I too used many kits such as Qiagen, Banglore  Genei and Himedia etc and did not got promising results. So, based on experience in bacterial DNA isolation, I recommend basic CTAB method for the best quantity and quality.
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My problem is that i'm trying to precipitate RNA with a Ethanol RNA Precipitation protocol and the yield is bad (almost half of initial concentration and half 260/280 purity). My initial volume is 30 uL and 70 ng/ul of concentration approximately and the precipitation gives me worst concentration and purity than the initial one. Is there a minimum volume for RNA precipitation ? because i'm using the entire 30 uL for the precipitation protocol.
Thanks
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Hi,
Your elution volume is probably too high. If your column is not so broad like in the case of Zymo kits, you can try a lesser volume of RNAse free water for a better yield. Also make sure you get rid of ethanol completely before eluting. However, you can also try magnetic bead methods.
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Any advice on sonicators v homegenisers and brands/types that will get the highest and purest RNA concentrations? My old lab used the diagenode biorupter (sonicator) which worked really well, but it is a bit out of our price range. We have a hand held homegeniser we use for protein extraction/isolation but I'm worried that too much of the tissue is lost in this process for reliable RNA analysis, and I'm wondering what other groups might use? 
We will be primarily working on rat spinal cord/brain tissue.
Any wisdom would be appreciated!
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Using the RNeasy mini kit and the QIAshredder from QIAGEN have always given me sufficient RNA for downstream analysis. 
I would use RLT buffer to lyse the tissue then put it through the QIAshredder to homogenise it then use that flowthrough on a spin column with the RNeasy kit. You can Qubit/nanodrop/gel electrophoresis to determine purity and concentration. 
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Hello, 
I am sorting monocytes from spleen for RNAseq. I pre-purify my cells using Miltenyi isolation kits and then I FACS sort the cells to obtain a high purity. Since I had problems with the RIN values (below 8) I added RNAse inhibitors to the sorting buffer and the recollection medium since the spleen contains a lot of RNAses. After this I increased the integrity of my RNA to reach values over 8 but not in all samples. Also I directly isolate the RNA after the sorting.
Does anybody knows how to further improve the quality of my samples? is the perfusion of  the mouse with PBS with RNAse inhibitor a good option? it would be toxic for the cells?
Thank you in advance for your help.
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Thank you for your answer. 
The viability of my cells is really high (>98%) since I add a viability dye to sort only those cells that are alive. However may some still die during the steps after since when I added RNAse inhibitors to the recollecting buffer I much improved the integrity of my RNA. My main issue is that still it is not a reproducible method since some samples have a RIN of 6-7 which is not suitable for RNAseq while others are above 8.
You are right, Adding RNAse inhibitor to the perfusion buffer may affect the expression of my surface markers. Do you have any experience with that?
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Hi all,
Looking for some advice.  I have a small number of iPSC-derived oligodendrocytes (approx 10,000 cells/sample) that I'd like to do RNAseq and whole genome wide methylation analysis (e.g. Illumina 850k).  Can anyone give me any insight as to whether this is feasible?
Does anyone have any experience of doing such work on small number of cells (and perhaps can recommend kits for extraction) and/or amplification steps if required?
Many thanks for any advice,
Vicky
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Hi,
      I have some experience of EWAS and RNA-Seq before but I seldom do expriment. I would do both RNA-Seq & EWAS on the same sample if I wanted to explain the difference of RNA level by DNA methylation. Make sure you have enough sample size to do EWAS first(34 paired in my case but less is ok). However, that's gonna cost too much since sometime only a small number of genes overlap in both EWAS and RNA-Seq. My point is, if you want to focus on RNA level, you can select the top interested gene by RNA-Seq on small sample size and the just use bisulfate sequencing to detect the DNA methylation of those gene; If you are interested in epigenetic you might do EWAS first and select the top CpG site to detect the mRNA level by PCR. 
     BTW, if you want to use 850k, make sure your DNA concentration > 3ug (in my case, 60-1000 ng/μL,   OD260/280 1.7~2.1) and minimize the batch effect.
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Recently, I am working with a newly isolated yeast RNA for RNA sequencing. I have sent my RNA samples for RIN analysis using Agilent Bioanalyzer. However, I have received error results for all 12 samples. Based on the results attached, the 18s/28s ratio and RIN cannot be calculated. The retention time for 18s/28s for all samples were not within the start time/end time mentioned in the report. Is that why the RNA ratio and RIN cannot be calculated? What are your suggestion/troubleshooting for these results. Is the manually adapted results accurate? Can I use this results as reference for RNA sequencing
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Looks like the samples themselves are OK, but retention times across wells are inconsistent. The most common reasons for such an issue are the gel (freshly prepared? correctly centrifuged?), priming (setting of the priming station? duration of priming?) and sample concentration (very high in your case; try diluting next time). Have you checked the lower marker peaks for each lane? If these don't appear at the correct time/position, sample peaks will also be off. Before proceeding to RNA-Seq, I'd definitely dilute these samples and run another chip using fresh reagents and paying extra attention preparing the chip - good luck!
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I have RNA extraction from mouse kidneys. I had very good RIN values for that RNA extractions (around 9.0). Since I had bad experience with bacterial RNA contaminations I decided to measure the RNA conc. (and RIN) on the same Agilent Tape Station, but chose the "prokaryotic programm" instead the usual eukaryotic detection.
Now my question is if anybody ever tried that and if so would be open share his/her conclusion from that? Is that a possibility to check contaminations at all? Otherwise I don't know how else I could test my samples for bacterial contaminations.
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Not sure why it would be necessary since you have a very pure RNA sample and, I'm assuming of course, you're probably going to do some sort of transcriptomics (Reverse Transcriptase PCR, RNA seq, etc.) on that sample.  If you are going to do RNAseq on total RNA then bacterial RNA will present a problem, however if you're doing RNA capture, you'll be hybridizing with specific eukaryotic sequences, same with RTPCR in a sense.  I think the main question would be what is you downstream application?  As for determining if you have bacterial contamination in the sample you could simply do RTPCR using a bacterial primer.  The best bet would be to use a primer for the 16S ribosomal subunit, it will be present at a much higher concentration than other bacterial RNA's, therefore if that's not there than you have no need to worry about bacterial contamination.  Not familiar with the Agilent Tape Station, but I've used the Agilent Bioanalyzer 2100, and your RIN number is great, what were your concentrations like?  Anyway, hope this helps, and best of luck.
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suppose, say that purity of the control sample ( mRNA) is 2.10/1.06, how do you determine its purity? or, How do you know its impure?  
My mRNA concentration falls around
control- 3809.2ul
Injury - 8424.6 ul
 I have been told that differences are high, and  I know it's a  simple math, but what is the optimum difference, or what should be the optimum concentration? 
thank you for helping. 
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Pure (total) RNA has usually a 260/280 ratio of 1.9-2.0 and A260/230 of 2.0-2.2. You can measure these with the NanoDrop.Absorbance based methods however are not giving you reliable concentration values as they measure everything absorbing at 260 (including DNA). This is especially true at low concentration. For more reliable concentration values you can use the qubit which uses RNA specific dyes. The optimal concentration depends on what you need to do. As mentioned by others, you can determine the integrity of your RNA samples with the BioAnalyzer. Bear in mind however that all this is intended for total rna. After the extraction, highest fraction of RNA is ribosomal RNA. And the BioAnalyzer actually uses rRNA (and not mRNA) to calculate integrity. For mRNA concentration you first need to purify your samples (e.g.polyA selection)
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Hi all,
I'm working on T7 in vitro transcription. I've tried both Neb and Megascript kit, incubated at 37 degree for 2 hours, denatured RNA at 70 degree 5min, then run simply using agarose gel (I'm trying for denaturing gel now).
You could see the attached picture, lane 1 is positive ctrl (linearized plasmid); and lane 2 is my sequence of interest (pcr template).
You could see there are both strong lower bands around ~200bp; while expected full-length bands (1-2kb) are quite faint. I basically could rule out RNA degradation because RNase inhibitor is included in IVT reaction and RNase-away used in gel running and i'm very careful for any possible contamination.
I think it's more likely INT is not complete with T7 polymerase stop around 200bp. So what should I do? I heard people would first unwind/anneal double-strand DNA at 70 degree prior to add polymerase, but is it really helpful and necessary given the protocol of neither kit mentions this step?
Thanks
=================================
Problem solved.
Simply run denaturing gels, either agarose or polyacrylamide.
This is a lesson that RNA will form structures and hugely distort the size on native gel.
thx
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You should be able to get cleaner bands than you are seeing here after heating to 70˚ (especially on denaturing gels). You may want to try extending your IVT reaction to ensure that the 200bp band you're seeing isn't just incomplete transcripts. 
You can also try adding a little DMSO (5-7%) to the reaction to reduce any secondary structures in your DNA that may be causing a early termination of transcription. 
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I am looking for institution where miR-16, miR-132, miR-146a, miR-155, and miR-223 can be analysed. Some kind of colaboration or cooperation is possible. Please, answer who is working in this field.
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Hi Luca,
Sorry for late answer, I 'we just seen your message.
I am looking for genetic predictors of poor and good outcome in adult patients with juvenile arthritis. It os nown that some children who suffered from JIA had remission in adulthood but 40% had sever joint damages and ankilosing and disease activity in adulthood.
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I have some DNA samples for genotyping-by-sequence (GBS) extracted with the Qiagen blood and tissue kit without treated by RNase A. I've quantified DNA concentration with florescent dye (Promega Quantiflour). This dye is dsDNA specific with minimal binding to ssDNA, RNA and protein. However, the gel image showed that there may be RNA contamination in the DNA samples. Would the presence of RNA affect GBS in this case? If so, how?
Most institutions require that DNA samples submitted for GBS to be pre-treated for RNA, which suggests to me that the presence of RNA in the DNA sample is bad for GBS, but I do not understand why is it bad (given that DNA concentration can be accurately quantified even with RNA contamination).
Many thanks ahead for any insight!
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Hi Grace,
After consulting with our sequencing center, my understanding now is that the presence of RNA in the DNA sample can inhibit enzyme activity in library prep, so they essentially lowers your enzyme digestion efficiency a little bit. Very dirty samples will get cut less, and after PCR, you get less fragments from these samples to go into the sequencing lane, so it potentially affect both your sequencing coverage and depth. If your samples contain different amount of RNA, you may get different sequencing depth for different samples. Since the enzyme used in GBS should not cut RNA or RNA-DNA hybrids, I'm not too worried about bias in the allele frequency in the sequencing results. 
I now treat all my samples with RNase A during DNA extraction. It only adds 2 more minutes to the protocol. If your samples are already in TE or AE buffer and you don;t want to go through the whole clean-up process (which always leads to the loss of some DNA), you can simply add RNase A to your samples, let them sit in RT for 20 min. Your DNA can be used directly for GBS library prep without cleaning up for RNase A. This is suggested by our sequencing center but I've never done it. Alternatively, since RNA lowers the enzyme activity, you can also load a little more enzyme (but not too much) in your library prep.
Now with results from my GBS back, I can tell you that even with RNA in the DNA sample, it still worked just fine, but cleaner samples resulted in slightly higher read count number and much lower read number variance. Basically your get more homogenous QC results from cleaner samples. I hope these will help.
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Actually I'm trying to detect the new subtypes of Influenza A in bats ( H17N10 and H18N11) of the floodable savannas of Colombia.
I would like to know a recommended RNA extraction protocol for samples taking in RNAlater, we have had used RNeasy mini kit by with low yields in the measuring of RNA quantity and quality in NanoDrop.
Someone has experience in the subject or knows the sequences of primers and probes for real-time PCR detection?
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Thanks, those primers was design at the post-pandemia by de CDC in 2009, but the lack of information about Influenza A-like subtypes detected in bats don´t report the successful use of them.
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I work with RNA on the Bio-Rad Experion chip gel electrophoresis. Is it possible to determine strand length from the concentration results? I have noticed that other papers using this or Agilent don't usually report bp. Is this over reach when reporting total RNA quantity and RIN?
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@Sandeep
The reference markers come with the kit, and the manual gives marker concentration.  I will have to check again, but I don't believe the marker's fragment lengths are mentioned.  Here is an older gel I ran.  One is the approximate gel rendition, and the other is the absorbance of the marker lane. I also added a sample well electrophoretogram to my original question. 
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 These samples are showing very good quality on gel exhibiting two intact RNA bands, without any degradation; but RIN value is not good.
We are using tape-station for RIN value determination. Is this platform suitable for plant tissue, or bio analyzer is better for plant RNA?
Kindly help me to know why this is happening?
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Hi shweta, if the sample concentration is  high dont load directly normalise all the samples in same ratio basing on qubit reading or nanodrop.
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Upon using the TruSeq Stranded mRNA LT kit for RNA-Seq sample preparation I obtain the following Bioanalyzer traces using a Agilent DNA 100 Series II kit.
What is the additional trailing peak near the upper marker? Will it hamper sequencing?
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Hi everybody, sorry for my delay, and when asking for the Bioanalyzer image I actually meant the trace – I have seen only now that you have already posted the PDF in the first question.
I agree with Matthew that the library should be of sufficient quality for sequencing, and that 4µg is quite a lot. We usually use 1 µg of high-quality RNA (i.e. directly extracted from cells/tissue). Furthermore, we have recently begun to use only 60% of a reaction to save money, and can still amplify with only 8 PCR cycles.
Tushar, how many PCR cycles did you apply?
To end with the SPRI – Ampure XP are actually SPRI beads, so there are lots of size selections within the Truseq protocols, which is based on the ratio of beads to reaction volume. You might have noticed, that this ratio is different after the ligation step (in order to remove adapter dimers). See also http://core-genomics.blogspot.de/2012/04/how-do-spri-beads-work.html
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How many hearts from newborn mouse are needed for RNA-sequencing?
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In terms of amount of RNA, you only need 500ng RNA for deep sequencing. The amount of RNA/heart will run into the 100s of micrograms. Therefore 1 heart will be enough. However, for the deep sequencing, you should take 3 hearts from 3 mice of the same genotype/treatment as this will be required for your statistical analysis etc.
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Hi All,
I have datasets derived from RNAseq experiments,I did differential expression for coding and lncoding RNA. I want to construct a co-expression network of these coding-noncoding genes,what should I do? Can you recommend friendly software? I found  WGCNA package of R, but please give me some information about Process .
I should construct a co-expression network for each experiment OR I can select special genes and lncRNA from all of experiments(I mean after doing meta analysis ,set filter and choose key genes) for constructing a co-expression network?
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Hi, you should start from the normalized values of your genes and lncRNAs.
If you already know which genes and lncRNA to test for correlation, then you can arrange a table with the samples as rows and selected molcules as columns. If you, instead, wish to identify correlated molecules from expression values, you may want to put samples as rows and ALL molecules as columns.
                       molecule 1 molecule 2 molecule M
sample 1
sample 2
sample n
What's do the trick is as easy as:
library(Hmisc)
rcorr(as.matrix(x))
where x is your dataframe above. Hence, you get a matrix of r pairwise and a matrix of p values for the r's. It automatically ignores missing data. These matrices are symmetrical.
Finally, you can select the pairs of molecules with significant p-values and a satisfying r values. Hopefully, among these pairs, you will find some gene-lncRNA pair.
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Dear Scientist,
I badly need someone's suggestion who are working with RNA from etiolated arabidopsis. In my case I am working with 4day old etiolated Arabidopsis seedling to isolate RNA by using TriZol reagent.(<100mg/1mL
Here is the condition I am using:(<100mg/1mL trizol, 15 min incubation>200ul chloroform/1ml trizol> 15 min
1. 100mg/1mL trizol, 15 min incubation
2. 200ul chloroform/1ml trizol
3. 15 min centrifuse@12000rpm @-4CAdd 500ul isopropanol> incubate 15 min @room temp. 
4. Add 500ul isopropanol incubate 15 min @room temp. 
5. incubate 15 min @room temp. 
6. 10 min centrifuse @10000rpm
7.Wash pellet twice with 70% ethanol.
But this case I cant see any pellet... :(
Please I need your valuable suggestion to trouble shoot it out.
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Dear Torikul Islam!
I recommend you method for isolation RNA using 2-methoxyethanol and cetavlon. This method is widely used to obtain nucleic acid (DNA or RNA) highly purified from polysaccharides that are components of animal, plant and bacterial cells.
I propose to your attention many works in which this method was used, for example:
Jones A.S. The isolation of bacterial nucleic acids using cetyltrimethlyammonium bromide (cetavlon). Biochim. Biophys. Acta. 1953.10: 607-612; 
JONES A.S., MARSHG E. & RIZVIS B.H. The Isolation and Composition of the Nucleic Acids of Aerobacter aerogenes J . gen. Microbiol. 1957. 17: 586-595;
Biggin W.P. An investigation of RNA induction in amphibian tissues. B.Sc. (Honours), The University of British Columbia, 1966; 
Naktinis V.I., Maleeva N.E., San'ko D.F., Mirzabekov A.D. Two simple methods for isolation of DNA from various sources using cetavlon. Biokhimiia. 1977. 42(10):1783-90;
as well as my published work:
Tsygankova V.A., Zayets N.N., Galkina L.A., Prikazchikova L.P., Blume Ya.B., An unusual minor protein appearing in embryonic axis cells of haricot bean seeds following germination process stimulated by 6-methylthiouracyl. Biopolymery and Cell (Ukr.), 1998, vol. 14(5), pp. 438-448:
 Sincerely yours, ScD Victoria Tsygankova
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I've extracted RNA from plant tissues, and want to store it at -20 degree. Is it possible?
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Of course you can but better if you add 70% alcohol, sample will be safe for long time.
Best
Devesh
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I am dealing with Candida RNA isolation from TRI reagent (used qiazen kit for extraction).I am really concerned about the quality of my RNA. According to Nanodrop reading my yields were ~100 to 150ng/ul. The A260/280s were ~2.1-2.2. Although I have used glass beads and vortexed vigorously, the quantity of RNA was not good. I am planning to use SAB (Sodium acetate buffer) for cell suspension and Phenol:Chloroform: isoamyl alcohol instead of TRI. Has anyone experienced on Candida RNA isolation without mechanical disruption? Please suggest me which one is better?
Thanks in advance 
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Many thanks for your suggestion. I will look into it. 
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Hi I am depleting the ribosomal RNA of my samples with ribominus kit for ion proton libraries. After running a pico chip in the bioanalyzer I see a clear depletion of the main 18s and 28s (just look at sample 3). However in the bioanalyzer "gel" image I have a very clear low weight black band. I dont know if it should look like that or just not bands. Also I dont know if the black band correspond to the 5s....
Thanks in advance
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In my opinion, what you see is the lower marker of the gel dye, which somehow has been misplaced by the software. If you go to your electrophoregrams, check if the lower marker is not set to a small "peak" (even the smallest visible bump in the FU curve) before your strong band and then reset it to the "normal" position. Also, I think your RNA ladder needs changing, it is partially degraded and should have a better "look".
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Dear all,
I am currently trying to find the concentration of RNA present in a solution in terms of copies of RNA/volume of solution. The RNA molecules are 508 bases in size. I will be using the formula involving Avogadro's constant and the concentration of RNA in terms of ng/ul to find the concentration in terms of copies of RNA/volume of solution. The RNA strands are all + sense strands. Although the RNA molecules are not complementary, I'm not sure if they could potentially form regions of dsRNA.  
How can I know if my RNA is single stranded or double stranded in solution? This is important since dsRNA would require me to multiply the molar mass of the ssRNA by 2, which can cause the concentration in copies of RNA/volume of solution to differ greatly. 
Thank you!
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Lots of online tools to help with your question.  If they are all + strands, then you don't need to worry about the single or double stranded part of your question.
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Is there a way to analyse the data from an environmental sample at once, I mean prokaryotic and eukaryotic RNA at the same time, or do I need to run the samples on two different chips to obtain the respective quality?
I am analysing an anaerobic reactor sample containing bacteria, archaea and anaerobic fungi.
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Hi! I'm using the RNA Nano kit and I need to have information on both eukaryotic and prokaryotic RNA too. Did you guys finally find a way to do so? I appreciate your help!
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Hello,
I will be using this kit provided by Qiagen for isolation of viral RNA from stool samples. I was wondering if anyone had any input on how to tweak the protocol to allow for efficient isolation of RNA as opposed to DNA.
Thank you for your contribution!
Regards,
Kyle
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Hi Kyle & Phillip, Did you ever manage to sort this out? We're thinking of doing something similar and would be curious to hear what you found. 
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Dear colleagues,
I am working on a laser capture microdissection (LCM) project to collect vascular cells from Arabidopsis cotyledons for RNA sequencing. RNA of vascular cells obtained from LCM has been isolated by PicoPure, and we are planning to linear amplify the RNA from nano grams to micrograms for sequecing by 2 rounds of RioAmp HS Plus. I have a couple of questions about the assessment of starting RNA quality and quantity.
We are currently not feasible to use a Bioanalyzer or fluorometry (RiboGreen) so we tried Nanodrop. The concentration was estimated from 2-16 ng/ul varied from different samples. I am aware that the readings below 10ng/ul is not very reliable so I am planning to perform a qRT-PCR to assess the quantity and quality together.
Publications (eg., Kube et al (2007). BMC Mol. Biol.) and kit manuals (eg., Arcturus Paradise Plus Reagent System for FFPE tissues) has introduced the procedures of qRT-PCR for this purposes. Basically the ratio of the RNA yield obtained from different regions is used as an indication of RNA quality, and the RNA quantity derived from the 3’ primer set is used as the quantity measurement of the RNA, by plotting standard curve of control RNA amount vs. Ct. This assessment is based on the assumption that the actin mRNA in the sample represents the average status of other RNA molecules in the same sample. The total estimated RNA amount in a given sample is expressed as an equivalent of universal RNA that contains the same amount of actin mRNA. Most of these protocols are based on using ß-actin in human, would you think something like Actin2 would fit this assumption in Arabidopsis (plant)?
Actually, I have done a semi qRT-PCR by using a series of dilution of a control RNA (isolated from fresh frozen samples) whose quantity and quality has been assessed. See the figure attached.
  • Left, RNA from fresh frozen samples, it has been diluted and and input RNA for RT-PCR is 4.96ng, 0.496ng and 0.0496ng (as indicated by numbers above).
  • Right, RNA from LCM samples. The numbers above indicated the ng of RNA input in RT-PCR, as concentration estimated by nanodrop.
  • All samples were performed in the same RT-PCR procedures and program. However, comparing to the bands on the left, even input as low as 0.046ng, is even a little bit brighter than the LCM RNA (right) which is estimated to has a input of 40.5ng.
Apparently, the nanodrop estimated RNA concentration (and therefore the input) is not consistent with the band intensity. Is this because:
  • RNA in LCM samples has been degraded and the A260 read the degraded RNA as well so caused a over estimation?
  • Nanodrop reading is not valid, because it's so low. However, our samples are above the Nanodrop limit (2ng/ul) and the diluted control RNA cannot have a reading on the nanodrop.
  • Due to the efficient of PCR or something else, the plot between control RNA's abundance and their band intensity is not accurate. Instead, something like Ct value in real time PCR should be used?
Any comments will be appreciated.
Cheers, Thomas
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Thank you very much Enos, and apologize for such a late reply. Yes I agree with your point on RNA degradation and the weakness of semi-quantitative PCR on gel (endpoint product in the plateau phase). I am planning to perform a qPCR to plot the Ct of my samples on standard curve to assess the input quantity, but I again a have a few more questions.
Well a quick question is that is this quantification approach independent of genes expression? ie, what transcript should I target and if this will work for different samples? For example, as I said, the samples I wanted to quantify are coming from different timepoint and tissue (vascular cells from 5 days cotyledons and 10 days leaves). The RNA sample that I currently have with known quantity and good quality is from 16 days whole leaf. Can I use this as control RNA and create standard curve of Ct against series dilutions of this control RNA, and then plot Ct of the samples from vascular cells of different timepoint and tissue, and expecting a quantity although not that accurate?
I have also noticed your comments on reference genes, then another question is that how to determine a reference gene from a novel transciptome? We have performed a RNA-Seq analysis to investigate the event in whole leaf level from 4 different timepoints. Finding a reference gene for qPCR validation of this RNA-Seq is easy - I have used a couple of genes based on the top stable genes from the RNA-Seq analysis itself. However for this LCM-RNA-Seq project with vascular RNA only, to perform such qPCR based RNA quantification, do I need to use a very stable reference gene? The problem is that I don't know which genes will be expressed constantly acorss the timepoints in vascular cell, which is kilely to be quite different from the ones in whole leaf level. My RNA-Seq data tells me Actin2 is a bad chice since there is a 1.5 fold difference between two of my timepoints. I used my own selected reference genes for qPCR to validate the RNA-Seq, but for the quantity assessment purpose, do I need such stable reference genes?
Actually this reference genes issue bothers me a lot. As I probably mentioned somewhere, I need to perform linear amplification (IVT based) from nanograms of vascular RNA to micrograms for another RNA-Seq. There is no guarantee that the amplification will be identical between each sample, so many people told me I need to normalize the results to reference genes. Again however, how could I determine a reference gene in such a novel transcriptome of vascuar cells, before the RNA-Seq could be done?
Many thanks again!
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When we run total RNA samples on a 10% TBE Urea gel followed by SYBR Gold stain, we can easily tell the 5.8S rRNA and the 5S rRNA bands. We normally call the multiple bands below the 5S rRNA band tRNAs bands. Normally there are two weak upper bands (Band 1 & Band 2) and one super strong lower band (Band 3). I assume the strong lower band (Band 3) is where most of the tRNAs locate, around 70-75 bp. Does anybody have any idea what are the two weak upper bands (Band 1 &2), especially the one right above the lower strong band (Band 2)? Longer tRNAs? tRNAs precursors? Or tRNAs with unspliced intron?
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David, thanks very much for your detailed reply. It answers lots of my questions. I was also suspecting Ser- and Leu- tRNAs though based on their lengths only. That's why I cannot be sure of it. I agree that any intron-containing tRNA precursors might be hard to be detected simply by SYBR Gold staining. Also the aminoacylation should not be able to change the mobility so much on a Urea denaturing gel. 
By the way, do you think the extra sequence of Ser- and Leu- tRNAs create extra secondary structures as compared to other tRNAs?
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I want to do research to compare mRNA in two groups and want to calculate the sample size for the study. Most of the articles has given expression of mRNA as fold change. Please help me how can i use these values, and what will be the formula.
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Do the other articles report the fold-change of the same mRNA you are going to study?
If not, then their values will not be very useful for you.
If yes, the these values could be used as "resonable effect sizes" you should aim to detect with sufficient precision in your study. What you will need additionally is an estimate for the precision, too (i.e. how variable will the results be from different samples/individuals). If this is not given in the articles, you might either need a pilot study or you need to make an educated guess.
This is the procedure:
1) express the effect d as the LOG fold-change
2) get the standard deviation s of such log fold-changes from different samples. An educated guess is that s is about log(2) to log(5).
3) for a reasonable precision your study will then need about n = 16*s²/d² individuals.
Example: consider the folg-change is 2. So d = log2(2) = 1, and let s = log2(4) = 2, hence n = 16*s²/d² = 16*2²/1² = 16*4 = 64. Note: the log2-values refer directly to Ct-alues from qrtPCR. This means: d is nothing but the ddCt, and s is the standard deviation of dCt values!
See: STRUTS:Statistical Rules of Thumb, Chapter 2, page 3:
Why use log fold-chage? - Because the distribution of fold-changes is roughly log-normal, so the distribution of log fold-changes is roughly normal, and the standard analyses (e.g. using the mean as central value, all the standard linear model stuff, including the rule of thumb cited above) are all based on the assumption that the distribution is approximately normal.
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I have been doing TriZol RNA extractions of zebrafish embryos with and without glycogen as a carrier. I've found that using glycogen during the isopropanol precipitation step increases the size and visibility of my RNA pellet, but I am also getting a large peak in the 220-230 nm range in my NanoDrop spectrograph. My samples without glycogen look great.
I know that peaks around 230 nm indicate contamination (phenol, chloroform, protein, etc.), but it doesn't make any sense to me that only adding glycogen would cause this contamination.
I've attached graphs from some of my NanoDrop readings. The green line is my glycogen-free sample, the rest have glycogen in them.
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Do you have similar data from a negative control reaction? if not, repeating the experiment without template with and without glycogen and comparing the results to your positives should  allow you to specifically identify if glycogen is the cause. That said, I am familiar with research identifying glycogen as a source of DNA contamination, and it seems reasonable that it may carry other contaminants as well, especially if the prep is poor. make sure you are using molecular (or RNA) grade glycogen and always, always, run control reactions.
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I need to store a single cell after isolation for future DNA/RNA analysis.
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For RNA you can store it in a storage solution called RNAlater, it protects the RNA and immediately inactivates RNAses.
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I did a RNA isolation with a Polysome Pulldown + TriZol, followed by a purification with a RNeasy Kit. With the isolated RNA a transcriptome should be generated.
My samples on the Nanodrop looked quite clean.
(Exemplary sample value for 260/280 is 2,19 and for 260/230 is 2,48. I know that the 260/230 is a bit high but I couldn't find an explanation why. Also it shouldn't be a problem for generating the transcriptome I was told.)
Now I got the Quality control data, which were measured with a Agilent 2100 Bioanalyzer and they look quite strange. 
18s/28s rRNA bands cannot be clearly descriminated, although the sample itself doesn't look degraded. You don't see a smear but clear distinct peaks. See attached an example.
Does anyone has an explanation for this?
Also an additional note: If I digest my sample with RNases, I cannot find any bands afterwards. So it seems those bands are RNA and not DNA contamination or whatsoever.
Don't know if it is important but those are c.elegans samples.
Also the all the sample look quite the same (n=6). You can't see a big difference between days.
Has someone experienced this before or does know what happened in my samples?
I am happy about every help I can get. :) 
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Thanks for your answers!
The problem is that I just do the isolation of the RNA and then send it to a company for producing the transcriptome. They gave me the Quality Control results of the Bioanalyzer. That's why I can't easily repeat the experiments to check the error.
(Agarose Gel and Nanodrop I did myself but they looked good, as you said.)
I thought someone here might know what went wrong just because by looking at those very defined pattern.
So if there is someone, I'm still open for suggestions.
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I'd like to know the features of single stranded RNA molecules such as diameter and charge, but I can't find any experiment on the subject. There are some works on dsRNA and hairpins, but nothing on ssRNA; I know it is a very difficult experimental matter, but I find strange that there is nothing at all about it.
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Thank you,
it is a very interesting article, but if the DNA fibre is 2 nm in diameter, how much would be that of a mRNA molecule? 
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I am planning to perform RNA seq using a MiSeq Reagent Kit v3 600 cycle, mean insert size of ~600bp, 2x 300bp reads, paired-end. I have had some divergent advice regarding the no. of reads required per sample to perform differential expression analysis on mammalian cells engineered to produce very large quantities of therapeutic product. 
How many reads will I need to allow me to perform differential expression? Does the length of my read or the fact that they are paired-end reads effect the no reads required per sample. I have been advised that 5 million reads per sample or 15 million reads per sample are suitable depending on the expert that I speak to. 
Any advice would be much appreciated.
Thanks
Clair
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5 Mill sounds few for me but I am not an expert and I am do not work with mmamalians genomes.
Any case, more than focus on the depth try to get as much  biological replicates as possible. To identify DEG, It is better 3 RNA seq of 5 Mill, than one experiment of 15 Mill.
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After a few practice runs on extra samples, I was achieving a 260/280 of about 1.9, a 260/230 of about 1.6, and a yield of about 80ug.  These were not perfect results, but I felt confident enough to begin isolating RNA from my real samples, and after the first run with the real samples, the 260/280 came out at a respectable 1.8 average, but the 260/230 hovered around 0.6 and the yield around 35ug. 
While I was performing the extraction, I felt that this was my best set until the nandrop results came back.  I am at a loss as to why the results were much lower than expected and would appreciate some feedback before I move on to isolate the rest of my samples.
I used a Qiagen kit to isolate RNA from human fat samples ranging anywhere from 5mg to 100mg in size.  I noticed no strong correlation between amount of sample and yield; however, I did notice that overall the samples were much 'uglier' than the extra samples I practiced on (meaning they were much more bloody and something about them was off).  I made sure to homogenize very effectively, I did not grab any interphase layer (and made sure to leave some supernatant in the tubes so as to not disturb the interphase layer, and I followed the protocol to the dot.  Oddly, one of the samples returned respectable results in the nandrop analysis (260/280 of 2.01, 260/230 of 1.64, but the lowest yield of about 27).  I doubt that I ran every sample incorrectly except this one, so that leads me to believe that my samples were not good in the first place?  
I am thinking about extracting the RNA from the column with a much larger quantity of H20 than the perscribed 40 uL, then speed-vac to concentrate, as this will yield more RNA and evaporate residual ethanol to possibly improve the 260/230, but I have to clear this with my PI first.
Any thoughts would be nice.  
Thanks!
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I prefer to isolate RNA using the Trizol protocol instead of a kit. I also clean the RNA on a bench without a kit or columns and usually get great results. I think for RNA, 260/280 is not very important, it's the 260/230 you want to see around 2.0. 
Remember too that those ratios don't indicate the quality of sample, only whether or not they are contaminated with proteins or reagents used in the extraction, like guanidine hydrochloride. You can run RNA samples on a gel (or a bioanalyzer) to get a look at quality.
As for 'cleaning' the sample, I hate those column kits. They are expensive and unreliable. I switched to this method (after Trizol extraction)
Adjust RNA volume to 500µl with (DEPC treated or nuclease free) H2O
Add 50µl of 3M NaOAc  (+ 10ug of glycogen if you want to maximize yield)
Add 500µl of RT isopropanol (Room Temp is important: cold isopropanol will just co-precipitate your guanidium again). Mix well (vortex is ok)
Incubate @ RT for 20m
Spin for 10m at 12500 rpm at 4ºC
Dump supernatant
Wash pellet by gently adding 400 µl ice cold 70% ethanol & spin at 10000g for 5m @4º
Repeat wash step once
Air dry 2-5m; resuspend in water, in a volume you think is appropriate (20-50µl based on pellet size).
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Hello everyone, I´m using the Agilent's Small RNA chip for detection of microRNA, but I´m not sure of the indicated concentration by the bioanalyzer (18% of miRNA), because I didn't observe the "hill like" pattern showed on the manuals (fig 1), instead I observe "peaks" (fig 2) on the miRNA region  (from 15 to 40 nt). Is that enough evidence of the presence of miRNA in the sample?
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Looks like a bad run to me.  The marker (at 4nt) is barely detected.  How did the ladder & other wells look?  If there's any surfactant in one of the samples, that can ruin runs for the entire chip.
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I have been doing RNA-EMSA with Biotinylated-RNA. The problem is that the complex band always show in the side of the lane, especially in the low concentration of protein. Does anyone know the ways to fix this problem?
Here is brief condition of my experiment: My protein is 55 kDa with histidine-tag. Predicted pI of the protein is 8.85. My RNA is 174 bases. Binding buffer contain 50 mM Tris pH 7.6, 10 mM Mg(OAc)2, 65 mM NH4OAc, 1 mM EDTA, 0.2 mM ADPNP, and 5% glycerol. The gel is 5% non-denaturing polyacrylamide gel (with 5% glycerol) in 1X TBE. Run at 4C at 120V for 2h (pre-run 30 min 80V). I also tried tris-glycine buffer which had similar result.
If you need more information, just ask me. I will answer as soon as I can.
Thanks.
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Hi Jacob and Jiulia,
Sorry for the delay. I have tried many things to improve my EMSA gel, but the complex bands still show in the side of the lane. Some of them look just a bit better. I tried adding 0.1, 0.5 mg/ml yeast tRNA, BSA, running the gel with less voltage and shorter time, reducing salt concentration in the buffer, changing the salt (to NaCl and MgCl2), adding NP-40, changing buffer pH, not putting glycerol in the reaction. The gels still do not look much better. By the way, adding 0.1 mg/ml heparin to the reaction kills the interaction (no shift band at all). If you have more suggestion, I'm totally open.
Attached is the example of adding 0.1 mg/ml yeast tRNA and 0.1 mg/ml BSA into the above mentioned buffer.
Thanks.
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Hello, I ‘am looking for information about bacterial DNA purity from Magna Pure 96. I will use the MP96 for purification of bacterial DNA for 16S RNA analysis on Illuminas MiSeq. The purity of DNA has a very big influence on the results of the NGS. Does anyone have experience with MP96 and Illuminas 16S NGS? Has anyone compared the purity of bacterial DNA from the MP96 with other instruments?
Thanks!
Olaf
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Hi.
MagNapure 96 will works quite good, but is not the best choice.
With best regards
Olaf
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Actually i have done RNA sequencing for my experimental purpose where they shows how much genes got up regulated by conversion the data through Log2 conversion by comparing with control and drug treatment sample but my question is that is possible to Calculate the P value and FDR value  from this data format here below i have attach a Picture.  if possible  how could i do this any instruction or procedure will be great helpful for my research 
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Only fold change values and not p-values are given in your table. If this is the only information you got, then you should import the counts into e.g. edgeR make a contrast matrix and from here you can get p- and FDR-values. 
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Has anyone seen Total RNA profiles like this? There is degradation but also some other contaminant. The lab told us that these are from mouse colon and bacterial RNA can be a contaminant. They repeated RNA extraction and got similar profiles. Tissue was frozen on dry ice and wasn't subjected to free-thaw. However, it's clear that these are partially degraded. Could the spikes be due to bacterial RNA contamination? Thank you for your input.
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you may find this publication useful for your sample prep. It appears to be a known issue.
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I am using the TriZol method for extracting total RNA from adult zebrafish tissue, whole Xenopus larvae and whole zebrafish larvae. Considering that my 260/280 and 260/230 values are near about the prescribed values, do I still need to do a DNAse treatment. Some have suggested that RNA extracted from tissue can never really be genomic DNA free. Any advice, please?
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