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I use the MEGAscript T7 Transcription Kit (Cat. no.: AMB13345, ThermoFisher) for in vitro transcription of linearized plasmids. However, I have measured a high concentration of DNA in my mRNA after in vitro transcription. I use 15-20 ng of plasmid for the reaction, and I have already incorporated a second DNase treatment step in my protocol, but it doesn't seem to remove the DNA. I use DeNovix for measurement of concentrations. Anyone who has experience with this? If yes, is your mRNA still functional even though DNA is present?
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Hi Amy! Thank you for your answer :)
I use the RNA Clean & Concentratior^TM- kit (R1018, ZYMO RESEARCH) for RNA clean-up. Do you have experience with that? If yes, is that sufficient enough to remove DNA and DNases?
I will use the transcripts to develop an mRNA vaccine
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I would like to design a few couples of primers to amplify several define transcript region of IgG gene. With forward primers in the V or J region and reverse Primers in the constant region. However, I can not check the primer specificity using NCBI primer blast. Do you know which platform or sofware that I can do primer blast for IgG transcript? Thanks and regards.
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Hi Thuy
you can check on the UCSC database, there is a tool named "in-silico PCR", but if it's for human, the issue is well documented in papers since of huge variability in this region, degenerated primers are often used. on UCSC, in hg19, you can also check for qPCR primers designed in former studies (optional track in the expression toolbar).
all the best
fred
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  1. Observational Analysis: Researchers observe and record conversations to study how individuals from diverse linguistic backgrounds interact.
  2. Transcription: Spoken language is transcribed into written form for detailed analysis, including pronunciation, intonation, and pauses.
  3. Coding and Categorization: Linguistic patterns and sociolinguistic variables are identified in transcripts, such as code-switching and language choices.
  4. Quantitative Analysis: Statistical techniques may quantify sociolinguistic phenomena like code-switching frequency or linguistic feature distribution.
  5. Qualitative Analysis: Researchers explore the meaning and context behind linguistic behaviors and language choices.
  6. Questionnaires and Surveys: Self-reported data from participants, including language preferences and attitudes toward languages, can be collected.
  7. Corpus Linguistics: Large collections of texts or spoken data are analyzed to uncover linguistic patterns.
  8. Experimental Studies: Researchers design experiments to manipulate variables related to peer interactions and sociolinguistic competence.
  9. Interviews: Semi-structured interviews provide insights into participants' experiences and perceptions.
  10. Audio and Video Recordings: Recordings capture spoken and nonverbal aspects of communication, such as gestures and facial expressions.
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Dear Chisom Paula,
Thank you for the answer. It will definetely help me to choose for the right participants for my research.
Best,
Zafar
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Hi everyone,
I got an exon level counts matrix (obtained via the Bioconductor recount3 package) and I would like to transform the exon counts to an estimation of transcript abundance.
Does anyone knows a way of doing this?
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Dear Susanta,
as far as I know, htseq and featureCounts take only sam/bam files as inputs (i.e., alignment files).
What I got is a count matrix on exon level. What I need is a summarization/quantification on transcript level. Or do you know a way to make these counting tools taking count matrices as input?
Best regards
Markus
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Hello,
We have a gene that is modulated by the binding some transcription facors (TFs). We recently found that one of these TFs is mutated and we want to know if this TF can still bind to that gene. Is there any software or tool to figure this out?
Thank you in advance.
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Dear Andrew J Spiers and Susanta Roy,
Thank you very much for your answers.
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from where the transcription of the gene in the vector starts , is it always from 5 to 3 or it depends on the direction of the promoter .
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Ok, are you trying to clone a new gene into pET29a?
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I did a knockdown, after checking the level of protein expression with Western blot, I got some percentage let's say like 60 to 70 % of knockdown, but when i tried checking with qPCR to check the transcription level it was as if there was a complete knockout, what could have been the reasons, I am confused can some help me with an explanation? You help will be highly appreciated.
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Are you using siRNA? If yes, then you are most likely to face this problem.
You need to consider protein stability which is highly variable. It may so happen that your target protein may be highly stable with a longer half-life. In such a case, you may consider choosing shRNA-mediated gene silencing method over siRNA. While making a choice, you will have to consider the length of the assay as well as the half-life of the target protein.
siRNAs are transiently expressed in cells, while shRNA is stably integrated into the host cell genome. As cells divide, the shRNA is passed on to daughter cells. Using lentiviral vectors for expression of shRNA, provides permanent knockdown without needing to transfect the cells multiple times.
The protein levels subsequently go down over time because the shRNA constantly keep suppressing mRNA.
So, if you are trying to knockdown the expression of protein that has a long half-life, then stable expression of shRNA may be required.
Best.
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Analysis of qualitative data requires intensive reading of the transcripts, field reports, diaries, journals, and other documents. It is a continuous and to-and-fro process. What changes do you, as a qualitative researcher, face during data analysis and how do you overcome those challenges?
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I have recently been experimenting with artificial intelligence for the interview of qualitative data via ChatGPT, and I am very impressed with the results. In particular, I started by re-analyzing data from two of my previous studies, and I was surprised by how rapidly the program produced the main concepts from those studies.
Just asking a few general questions produced the important key dimensions, and asking follow-up questions gave more detailed information about each of those dimensions. Of course, the program cannot literally "interpret" the results for you, but it certainly could replace a laborious coding process as a tool for locating the core content that you need to interpret.
Like any other approach to qualitative analysis, it does require familiarity with your data (you can't just throw anything at it), but beyond that, the program has a strong potential for being an alternative to existing techniques for the initial stages of working with qualitative data.
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Dear colleagues,
I'm currently running some in-vitro transcriptions with the HiScribe T7 RNA Synthesis Kit by NEB. As Templates I'm using a purified PCR product as a run-off template. I properly thought about my Promoter sequence, bearing all the necessary bases not only for the binding but also for the initiation site. I'm expecting a ssRNA of roughly ~750-780 nt for my transcripts. However, my Urea-PAGE looks like displayed here. I can't really explain all the additional bands...
After the in-vitro transcription I'm using a RNA clean up kit and purify my transcript prior loading on the gel. I don't think it's unspecific degradation, the band look to sharp for that.
I'm looking forward to your suggestions and expertis
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quality control, just to be sure samples are not degraded too much that could disable the best amplification.
on the other hand, you can also use DMSO at 5% to enhance PCR
fred
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I know many websites have simple tools like transcription and translation available, but are there any analysis tools that researchers need that either do not exist or are not publicly available? It could be anything from algorithms to visuals. Thanks!
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Abhijeet Singh Thank you for your response and mentioning my earlier post! My belief is that researchers would know tools that are missing based on the fact that they would run into such problem often during their research. If there is some manual analysis task that researchers can automate, I believe that PeptiCloud can be the perfect platform to develop and make those tools publicly available. (For instance, PeptiCloud has a unique feature that allows users to further alter codon sequence of each amino acid after codon optimization with respect to a specific bacterial strain). With that being said, if you could check out PeptiCloud for yourself and see if anything could be added or improved, that would be greatly appreciated!
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can a DMRT( Duncan's Multiple Range Test) result in 11 columns for which i get compact lettering till 'k' (starting from 'a')?
Software used : SPSS v25.0
After i did the post-Hoc with Duncan, i got an output for a specific dependent factor with 18 treatments (3 transcripts of each) as 11 columns.
To put this DMRT result as compact letter display in my table, i'm getting letters 'a' to 'k' for the respective columns.
Is this possible/correct?
significance values for few columns exceeds 0.05. and few are below 0.05.
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Hello Ushnish,
If your intention is to show a summary of all possible pairwise comparisons among 18 conditions (= 153 comparisons), that will be a very full tabular display no matter what you do!
If only a few of the comparisons meet your criterion for statistical significance, you might restrict your display and summary to just those comparisons, and note that all others were nonsignificant.
One relevant point is that the Duncan test is likely not the best choice for several technical reasons. Consider something along the lines of the Benjamini-Hochberg method or the Ryan-Einot-Gabriel-Welsh method.
Finally, to address your question about letters, if the Duncan output showed 11 "homogeneous subsets," then yes, you may attach letters a-k as appropriate to show instances of nonsignificant differences (subject to the implied concern above about density of your display).
Good luck with your work.
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I'm working on a qualitative study that will include 30 interviews conducted online in Arabic. Can you please recommend reliable transcription software that supports Arabic? It doesn't have to be free
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There are many but Sonix and Rev are recommended
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Get more than 80% - 90% of transcription efficiency
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1) Check mycoplasma contamination.
Only after number should you proceed to optimizing transfection efficiency because mycoplasma contamination inhibits transfection (having experienced it myself). It is more common than thought.
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Transana is a software for qualitative research that enables researchers to work on authentic data that is recorded or videotaped. If anyone has employed this tool, I would be interested in learning more about their experience with the tool and its effectiveness in managing and analyzing complex qualitative data in their studies. Furthermore, the software is with different versions, so I am confused about which one to use.
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Dear Mohammed Azam,
Have you employed tranzana in your research? If so, which version please. You mentioned a free version that I have bot yet found, do you have access to it? Many thanks dearest friend.
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I want to produce 29kb mRNA using the in-vitro transcription method. It will be used for protein production in mammalian systems. I found RiboMAX large kit that can make a 27kb transcript. However, I did get any information about its application on protein production in the mammalian system. Could anyone suggest the best method or kit to produce this long-size mRNA transcript?
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Can you provide a bit more detail? Because...I am really not getting what you're trying to do, or why.
You have a 29kb mRNA that you want to produce via in-vitro transcription, so is your gene currently...on a IVT-compatible plasmid?
Are you proposing to clean up your transcripts after IVT? If so, how?
And then are you proposing to use them in an in-vitro translation system, too?
And if so...well, why? Could you not just express it in mammalian cells and let them do all the work for you? Mammalian cells would have no issues transcribing a gene 29kb long (they can handle genes 2.3Mb long, after all).
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Hi all, I am trying to extract RNA from 12 well plate and so far the yield obtained from trizol/chloroform protocol is OK.
After that I tried to do DNAase step (using NEB DNAse I) but I found that it does not allow cDNA transcription.
I don't know if it was the inhibition step 70°C x 10min of DNAase activity or smth else.
Do you have any similar experience? if yes, any trick to share with me that can help to do the DNAase step without inhibiting the reaction downstream? Thanks a lot
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Have a look at the ingredients in the buffer supplied with the DNase and compare it to the Reverse Transcriptase buffer composition. Are they similar? Each enzyme will be optimised to work in specific buffer conditions and it's possible that the DNase buffer will be inhibiting the Reverse transcription reaction.
Similar to the poster above, I would recommend an additional cleanup, using a Zymo column to remove the DNase buffer, and any residual DNAse. The columns have a good RNA recovery rate, and I think you can ask for free samples ;-).
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Dear all,
Thanking you in advance for your attention, I would like to ask you two questions regarding an electrophoretic run on 2% agarose gel:
1) I did an electrophoretic run of the Real-time PCR products (with Sybr green). Although the band of my target product is evident, I notice some very slight sub-bands (indicated by the arrows) which, according to the molecular scale, could have a size of 500 bp. I designed the primers by myself, and making a blast they are specific for my gene of interest . What do those sub-bands represent? Can they affect the analysis of fluorescence?
2)The same primers gave me products of a different size (in the photo, the circled bands). The primer targets a gene that has 3 transcript variants. Each well has a different individual (animal). Could those two circled products be a transcript variant?
Thanks a lot for you precious help,
Best Regards
Matteo
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Thanks for the help and support! Yes I have done a blast of my primers and they match with my target gene.
Do you think I am "safe", or do I have to sequence the dna in the band to be 100% sure?
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In allotetraploid B.napus plant transcriptome, Based on pacBio 3rd generation sequencing, one gene has evidence of Alternative splicing (AS) in it. This gene has four exons and produces 2 distinct transcripts. The structural analysis of two transcripts shows that transcript 1 is produced by using only exon 1 and 2 of gene, whereas, the transcript 2 is produced from exon 3 and 4. Therefore, these two transcripts have no overlapping sequences between them. I have to verify the presence of these two transcripts. Is there any other other strategy  to amplify the these two transcripts at once and show the evidence that these two transcripts are definitely transcribed from the specific gene, except using the transcript specific primer for each transcript (using transcript specific primer will just help me detect that transcript but will not provide enough evidence that it is originating from this specific gene.)?
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I come a bit late but maybe this could be useful for others.
You totally can amplify both transcripts with the same set of primer.
For this, you need to design a forward primer targeting the 5'UTR of your pre-messenger RNA, and the reverse targeting the 3'UTR. If, as you said, both your transcripts are spliced from the same pre messenger RNA they have to share those features.
Run the PCR products on an agarose gel.
Assuming that both your transcripts isoforms don't have the same lenght, you can extract both band with a gel cleaning kit and then send them for SANGER sequencing as a verification.
Hope it helps,
Philippe.
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"during both the sample preparation and computational analysis phases, at which imperfections and biases may be introduced. These limitations may affect the ability of the experiment to address specific biological questions, such as correctly identifying and quantifying which of multiple isoforms are expressed from a gene8 . This example is particularly relevant to very long, or highly variable, transcript isoforms such as those found in the human transcriptome; 50% of transcripts are >2,500 bp long in humans26, with a range from 186 bp to 109 kb" the Author of the Artical
RNA sequencing: the teenage years - PubMed (nih.gov)
31341269
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One specific limitation mentioned is the difficulty in accurately identifying and quantifying isoforms expressed from a gene, especially for very long or highly variable transcript isoforms. This is particularly relevant in the human transcriptome, where transcripts can vary greatly in length, ranging from 186 base pairs to 109 kilobases. The presence of such long and variable isoforms poses challenges for proper annotation and quantification.
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transcription and translation of eukaryotic and prokaryotic cells
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This applies to not only this question but for all the question you asked...
Please focus on your class/notes/lectures. Get the basic and fundamental knowledge. Asking random questions here would not help.
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Hello,
I am designing a plasmid with an SV40 promoter-driven antibiotic resistance. Does expression from an SV40 promoter require a TATA box upstream of the transcription start site? The original vector had a TATA box at -30, however this is lost in my cloning strategy. With my current plan, the transcription start site is just 8bp from the end of the SV40 promoter. Will this allow for expression, or is a TATA box needed?
Thanks!
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The SV40 (Simian virus 40) promoter is a strong viral promoter commonly used for driving gene expression in various experimental systems. While the presence of a TATA box upstream of the transcription start site is a common feature in many promoters, the SV40 promoter is unique in that it lacks a canonical TATA box.
The SV40 promoter utilizes an alternative mechanism for transcription initiation called the "TATA-less" promoter. Instead of relying on a TATA box, it utilizes other elements and transcription factors to initiate transcription. The absence of a TATA box in the SV40 promoter does not necessarily impair its ability to drive gene expression.
Therefore, in your current cloning strategy where the transcription start site is located just 8bp from the end of the SV40 promoter, it is likely that the expression can still occur without the presence of a TATA box. The SV40 promoter contains other regulatory elements and transcription factor binding sites that can facilitate transcription initiation.
However, it's worth noting that the exact transcriptional activity may depend on the specific context and the downstream sequence elements present in your plasmid. Experimental verification, such as measuring the expression levels of your gene of interest, can help confirm the functionality of the modified SV40 promoter in your specific system.
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I am working on the transcription of an RNA strand. More than knowing all the characteristics of my RNA, I would like to know if the transcription was successful or not. What methods can be used for this? My RNA strand should be 21nt long. Is it possible to use gel electrophoresis or spectroscopy?
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There are several characterization techniques for RNA that can be used to assess the success of transcription:
  1. Gel electrophoresis: This method can be used to visualize RNA size and purity. RNA is separated based on its size using an agarose gel, and the bands can be visualized using staining agents such as ethidium bromide or SYBR Green.
  2. Spectroscopy: This method can be used to measure the absorbance of RNA at specific wavelengths to determine RNA concentration and purity. The absorbance ratio of A260/A280 can be used to assess RNA purity.
  3. Northern blotting: This technique involves the transfer of RNA from a gel to a membrane, followed by hybridization with a labeled RNA probe specific for the target RNA. This can be used to assess RNA size, expression level, and specificity.
  4. RT-PCR: This technique can be used to amplify and detect RNA using reverse transcription followed by PCR amplification of the cDNA. The products can be visualized on a gel, and the RNA expression level can be quantified using real-time PCR.
  5. RNA sequencing: This method can be used to sequence RNA molecules and provide information on their sequence, expression level, and splicing patterns.
In your case, gel electrophoresis and spectroscopy can be used to assess the success of your RNA transcription. Gel electrophoresis can help confirm the size of the RNA strand, while spectroscopy can be used to assess RNA purity.
These video playlists might be helpful to you:
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Hi! I'm planning on using methyl-3-nitro-1-nitrosoguanidine (MMNG), for inducing transcriptional mutagenesis. From my understanding MNNG causes the formation of O6‐methyl guanine (O6MeG) inducing thymine mispairing during DNA replication. However, is it likely to for MMNG to to lead to other mispairing that may occur in low abundance? Or are there any disadvantages to be mindful of? I want to ensure that I can accurately quantify mutation patterns.
Thank you for your insight :)
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No alkylating agent (with the possible exception of those containing a targeting moiety) can be considered selective it is more of a general preference (see Coles, B. Effects of modifying structure on electrophilic reactions with nucleophiles. Drug Metab. Rev. 1985, 15, 1307−1334.) MNNG in addition to targeting the O-6 position of guanine, will cause depurination, strand breaks, alkylate phosphate etc. etc. For a more detailed discussion see
MNNG will do what you seek, but you must be aware of it's other targets, which are not restricted to the O-6 position of guanine in DNA, or even nucleic acids, or macro molecules.
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Can anyone recommend a software that could be used to help in Arabic interviews transcription/ translation? I am currently using Trint, but unfortunately it is not accurate.
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Bita Faraji thanks a lot for these recommendations! I will check them out.
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I'd like to know that what are the different ways to know/identify whether a particular Gene is expressed or not ?
Few points from my side are :
1) identifying it's corresponding m-RNA transcripts level.
2) identifying the protein that was produced by the expression of that particular Gene.
Any other points ?
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Hi,
You can do qPCR to check the expression of the target genes.
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I was reading a research article where I found this term but unable to understand. AK5 gene in 2009 was reported to have two transcript variants i.e. AK5p1 and AK5p2.
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Dear Mohd,
the term transcript variants refers to the different mature mRNA products from the transcription of the same gene. On the other hand the term isoform refers to highly similar proteins that are products of the same gene (usually).
During transcription there can be different mRNA products due to the utilization of different start codons within the sequence of the gene and also due to the utilization of different sets of promoters upstream the coding sequence. Additionally during pre-mRNA maturation (splicing) different mature mRNAs may occur due to alternative splicing sites. Furthermore, mutations can create new starting codons, and also influence the splicing and the maturation of pre-mRNAs. Altogether, these mechanisms (the major mechanisms) produce the different variants (transcript and splicing variants) that you see in the bibliography.
Different mRNA products (transcript and splicing variants) of the same gene can produce proteins with high similarity that together constitute the different isoforms.
I hope that was helpful
Kind regards
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in transcription process why doesnt need primer for bulid mRNA
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That is a complicated question that comes down to the mechanism used by RNA polymerase to initiate transcription. Fundamentally RNA polymerases do not need primers because they have evolved to be able to initiate transcription at specific sites (promoters) that are thermodynamically favorable for initiation.
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Hello everyone!
I have interesting question asked by my professor and I could not find relevant answer anywhere.
Why are we seeing up and down pattern on transcript abundance? Example RNA seq data for a gene from a rice transcriptome data base is attached. LOCUS ID is highlighted in yellow and transcript abundance is in below three samples after drought treatment.
The question is ,why the signal level is not uniform on Exons? is it low signal reads? Why there are gaps or sudden fall in signals? ( which are Marked in Red arrows) How to read and understand this? and I know this is the common pattern in RNA-seq data, but I don’t know why? It’s an interesting question asked by my professor! can any bioinformatician help me understand this? Thanks in advance.
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I am not an expert in this by any means, but I have read a lot and have seen this type of data and interpreted it before as well. I can give you what I know from my experience and others may chime in.
It is a read out of transcripts that correspond to that particular site. it might be referred to as base resolution expression of the particular sequence. Essentially the higher number of transcripts that coincide with that particular sequence the higher the score. It could be areas that were difficult to resolve due to all kinds of aspects. 1) Sequence has a lot of repeats if that was the case you would see the same resolution in the other two samples but that does not seem to be the case. These areas might be resolved better if you increase the read depth of the study.
2) It may be more suggestive of a difficulty to read them. These results may be affected by post modification of the RNA as well.
This paper describes this is clinical samples but that does not restrict the affect only to humans post-modification
Sci Adv. 2021 Aug; 7(32): eabd2605.
Published online 2021 Aug 4. doi: 10.1126/sciadv.abd2605
PMCID: PMC8336963
PMID: 34348892
Judging by the fact the title on the samples say drought I might think a more epigenetic effect (Post-modification of the RNA maybe due to stress)
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Hello scientists, I was hesitant to ask such a question because it is somewhat simple, but in the end there is no shame in the learning process, so, What consumes more energy, the polymerization of coding region or its translation?
and how to calculate the energy required for transcription?
Thanks in advance.
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If by "polymerisation of coding region" you mean "mRNA synthesis", then honestly, there is no fixed answer to this question (for eukaryotes, certainly).
Many genes have enormous lengths of intronic sequence, all of which must be transcribed, only to then be spliced down to a comparatively small mRNA for translation. Dystrophin, for example, is 2.3 million bases long, but is spliced down to ~14,000 (i.e. 99.5% of the transcribed nucleotides are spliced out and recycled).
Here transcription clearly requires a greater energy input on a 1:1 mRNA:protein basis, whereas other shorter (or intronless) genes might be closer to parity.
However you also need to factor in energy requirements for recruitment of initiation factors, unwinding of the transcriptional start site, scanning, abortive initiation, etc, which will also vary from gene to gene. Some genes are permissive, other are less so.
You also need to factor similar variables to translation: targeting of mRNA, recruitment of translation initiation factors, unwinding and scanning of 5' UTR, etc.
And finally, this all assumes you are comparing mRNA and protein on a 1:1 basis, which is entirely inaccurate. A single mRNA can be translated many, many times, so a large energy investment in one transcript might ultimately pale in comparison to the vast energy investment in making 10,000 proteins from that one transcript. Alternatively, a single mRNA might be degraded without ever being translated even once (some 'immediate early' genes are transcribed continuously, 'just in case', but then degraded if not needed).
If you want to know (more generally) 'what uses more energy in a cell, transcription or translation?', then the answer is almost always translation: translation (i.e. protein synthesis) consumes about 50% of the energy budget of a proliferating cell. Ribosomes are not very efficient, and cells contain LOTS of them (~80-85% of cellular RNA is just ribosomes). They are always in use.
I wouldn't worry about asking questions like this, by the way: it's an excellent question and exposes quite how many complexities there are to even such an ostensibly simple consideration.
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I performed an RT-PCR on my gene of interest hoping to see which isoforms are present on the mRNA level. Literature suggested there should be 4. To my surprise, i see lots (too many to count) of transcripts of various sizes. I isolated about 50, sequenced them, and aligned them to the original gene cDNA. Majority of the transcripts I isolated have chunks of sequence missing at seemingly random places, with random chunks of exons would be missing here and there. some has majority of exons missing.
I am wondering if transcription often produce these aberrant transcripts or is something unique going on here?
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Hi
the main point is to know wether you used total RNA or polyA+ RNA. Since in total RNA you'll get mature and non-mature templates, it's not surprising. on the other hand, in regards to a previous work where I sequenced a targeted gene after 3'RACE-PCR (full length sequencing), it's not surprising to find new exons and transcripts. you just need to count each molecules and see if it has sense in your study to go further.
best new year
fred
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I will be interviewing clergy members.
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ELAN, software allows you to store video/ audio file, annotate and transcribe files. at the end it can do language analysis as well. parts of speech etc...
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Why are mouse chromosome Y transcripts (avg) significantly shorter than its other chromosomes' transcripts? The calculation & comparison of the average lengths were done with t test according to the entire UCSC mouse genome. Any ideas?
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The average length of transcripts on the mouse chromosome Y is generally shorter than those on other chromosomes due to the unique characteristics of the chromosome Y.
Chromosome Y is the smallest and least gene-dense chromosome in the mouse genome, with only a few hundred protein-coding genes. Most of these genes are involved in male fertility and the development of male characteristics. In addition, chromosome Y lacks most of the repetitive sequences and transposable elements that are present on other chromosomes, which can lead to the production of longer transcripts.
It is also worth noting that the gene expression patterns on chromosome Y are different from those on other chromosomes. Chromosome Y is generally expressed at lower levels than other chromosomes, and the genes on chromosome Y are often expressed in a tissue-specific manner. This may contribute to the shorter average length of transcripts on chromosome Y.
Overall, the shorter average length of transcripts on chromosome Y can be attributed to the unique characteristics of the chromosome and the patterns of gene expression on the chromosome.
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Hi to all.
My question is how can I optimize my RTqPCR if the cDNA dilutions ended up in similar Cq?
I synthesized my cDNA from 350 ng total RNA, assuming 1:1 production I should have 350 ng cDNA in 20 ul right? Then I did a dilution of 1/2, 1/5, 1/10 and 1/20 (I know the first three are consider quite a lot to be used in the run) and used them in a 20 ul run. The gene is a ref. gene: GAPDH. Interestingly the Cq values aren't that different between the dilutions (~29, ~30, ~31 and ~30). Obviously these aren't good values but I don't know what can I do to optimize the run.
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Alright, I will try to share what tips/tricks I can.
Honestly, while RNA is vastly more labile than DNA, it isn't really some sort of mystic-grade vulnerability, and you don't need utterly RNAse free environments to isolate perfectly viable RNA. They will help, obviously, but just starting with RNAse-free stuff, using careful pipetting and not making obvious mistakes will usually be sufficient.
So: use filter tips. Here the filter is primarily protecting your sample from whatever gunk might be hiding up in your pipette barrel. Use filter tips for everything (1000ul, 200ul, 10ul).
Use RNAse-free microcentrifuge tubes (most prepacked tubes should be certified RNAse free): keep a dedicate bag for RNA work, keep the top sealed/folded over when not in use, and only fish out tubes with gloved hands. If you put an ungloved hand into the bag, then assume the bag is now no longer good for RNA work (or use at your own risk).
Use RNAse free water for everything: either buy it, or make your own using DEPC or DMPC: add DEPC to 0.1%, shake vigorously and leave at 37degrees overnight with the lid of the bottle slightly loose. Autoclave, then close the lid tight.
Take small aliquots for working (I tip out 50ml at a time into a falcon tube) so you're not constantly dipping in and out of your stock. If an aliquot gets contaminated, or you suspect it's contaminated, throw it away, make another.
Use a bench area you trust: this doesn't mean you need a dedicated area, but use common sense (if a genomic DNA extraction protocol involves 'add 100ul of RNAse H', for example, go do that protocol somewhere else).
Use common sense in general: just be aware that the primary source of RNAse activity is the investigator: we are covered in bacteria all the time, and all of those are robust RNAse sources.
Wear gloves. Wear them basically all the time. If you think the gloves are dirty, change the gloves.
Next up: practical tips/tricks and when to be most careful.
If you can, freeze tissue. Freeze everything until you need it not to be frozen. RNA inside a sample frozen at -80 will endure far better than RNA inside fresh tissue, and while its frozen, it cannot be broken down by RNAses (they're frozen too).
Try to keep tissue frozen RIGHT up until you lyse/denature everything.
Frozen tissue is safe.
Lysis: I use trizol (or trizol equivalent) methods for almost everything. Almost nothing survives the addition of large amounts of chaotropic salts dissolved in phenol: a frozen sample covered in RNAses can still be used for RNA extraction if you dump it straight into trizol, because the RNAses will unfold and denature right along with everything else.
I typically freeze tissue in liquid nitrogen, store it at -80, crush to to powder under liquid nitrogen (i.e. never let it defrost) and then add trizol directly to the frozen powder. The first time the tissue melts, it's melting in phenol.
RNA inside trizol suspension will endure, and can indeed be frozen at -80 for longer-term storage.
RNA in trizol is safe.
Once you add chloroform to initiate phase separation: THAT'S when you need to start being extra careful. The aqueous phase is RNA in solution, and it's essentially unprotected. Collect aqueous phases one at a time, tilting the tube to minimise stuff falling into it. Cap tubes as soon as you're done transferring.
I typically use isopropanol precipitation rather than columns, because I like to see the size of my pellets, but all downstream stuff from phase separation is extra-careful-time. Precipitated RNA itself is actually fairly safe, since RNAses can't really degrade a solid chunk of dry RNA (accordingly, you can also freeze pelleted RNA at -80 for some weeks).
If you're going down column-based preps, then all the on-column stuff is largely out of your hands. Keep the columns wrapped up and clean (most come individually wrapped, but if they're in a bag, treat that bag as for tubes, above: gloves for all the things, seal up when not in use).
Isolated RNA should be either frozen immediately, or kept on ice for spectrophotometry/bioanalyser, and THEN frozen.
Try to make it into cDNA as soon as possible, and try to minimise freeze thaw: better to make a lot of cDNA in one batch than to keep dipping into it for multiple one-step reactions.
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I’m going around in circles trying to find anything on APA (7th) recommendations for formatting supplementary materials such as an interview transcript. I am a student so one of the general recommendations for a student paper is double spacing for instance, but in an interview transcript, which is over 10 pages long with single spacing, I’m afraid the document will be unnecessarily long and hard to read. Are there specific rules regarding formatting an interview transcript?
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I would contact your chairperson or IRB representative and ask to be sure. They are the ones who decide what the requirements are, not APA. My requirement was 10 font, Times New Romans, single spaced. It does not matter how many pages it is, but double space would make your transcripts look like and encyclopedia.
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While trimming the adaptor and low quality RNA-Seq illumina paired end reads in Trimmomatic, I have got more Forward only survive of about 40 to 50%. This study is for estimate the transcript abundance (DEG) at various condition. How is the possibility to continue further...
1. USE singleton reads (R1-For only)
or
2. Only use both paired (survive) high quality reads (50% of the reads)
Any suggestion, Thanks in Advance
by, Ellango R.
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Thanks Abhijeet & Sonja.
After I deeply check the data. Forward has more over represented sequence of Illumina nextra adaptor/Index and Reverse has highly over represented by Poly (G) sequence. Finally we dropped this data and asked the vendor to redo the sequence.
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I am working with a non-model organism and we generated a complete genome that is annotated. How can you determine the transcriptions start sites in this genome? Is there any bioinformatic way to do it (software)? or Do I need to do an experiment to identify these sites in the genome? I know that CAGE seq and CHIP-seq are good techniques to do that. But I am not sure if there is a computational tool to identify these sites.
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If you have the genome sequence then I would do an RNA-Seq from your organism, trying to get as representative and as much RNA as possible. Map all the reads to your genome and the start sites should become clear as the most 5' end of a transcript.
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Hi there,
I'm trying to design primers to detect CRISPR-mediated KO using qPCR (as described in this paper: ).
I designed a few pairs of primers and neither of them show any signs of amplification.
As an example, this is a pair targeting the ITGA2 gene (exon 2):
Primer_FWD ("watching"): ATTGTTGTTTGGCCTACAATGTTG (target +/ 53026780)
Primer_REV: CTGCATAGCCAAACTGTTCACT (target -/ 53026816)
I used the following transcript for the design:
Imported from genome: GRCh38 (hg38, Homo sapiens)
Gene: ITGA2 (ENSG00000164171)
Location: chr5 52989326-53094779
Transcript: ITGA2-001 (ENST00000296585, CCDS3957)
This specific pair is a little low on GC%, but others have GC% within the recommended 40-60% range. However, neither primers seem to be working.
Apart from GC%, these primers comply with the recommendations for PCR primer design published on various websites. Yet, something isn't right - on qPCR amplification curves are dead flat after 40 cycles (I've tried a few template concentrations and annealing temperatures with no success).
Would anyone have some suggestions on what I do wrong? And what can I try?
Thank you in advance.
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Hi Oleg
performing in silico PCR on UCSC, your primers did well perform amplification at the good position. issues are therefore not coming for the primers (see attachment). maybe you could try a touch-down PCR in a range of 10°c and add some DMSO to get more accessible target.
all the best
fred
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What are the best assistive software that you're using to facilitate the transcription of qualitative data? I use soundscriber (a free tool). I am looking for better options.
Thanks.
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there is a new software on the radar
DOTE is a new type of transcription software developed by the BigSoftVideo team. It is tailored for transcribing social conduct, conversation and multimodal interaction for research purposes. DOTE has been designed to support two specific standards of transcription for qualitative research, which are commonly used in conversation analysis, for example. DOTE has some of the features commonly found in other software -- such as video playback, a timeline and a visual waveform, synced playback -- but these features are streamlined and easier to use in DOTE. Moreover, it has enhanced features that do not exist in any legacy software so far, including transcript parsing, smart auto-completion, transcript heuristics, 360 video support, video-cues, export to publishable document (and subtitles) and version control. There are many more features and enhancements planned for the future. Our motto is make transcription fun again!😜
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I conducted data collection (20 interviews in Sepedi Language) as part of my Ph.D. studies. All interviews were transcribed then translated to English.
So i would like to know that out of 20 transcripts how many should i back-translated to Sepedi to ensure/check accuracy?
Any literature recommendations will be appreciated
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I would recommend ten percent of all your interview transcripts. For example, 10% of 20 transcripts would be 2 transcripts, which to me is enough for the credibility and confirmability of your findings.
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Can someone recommend a plasmid that carries two transcriptional terminators ? We'd like to insert two adjacent terminators into a construct we have made to prevent any transcriptional readthrough. We are currently using one lambda oop terminator, but we'd like to improve upon this, if possible.
Unfortunately, we are unable to synthesize tandem terminators by gblock due to their structural complexity (IDT won't make them) and we have had problems amplifying terminators by PCR likely for the same reasons.
Our ideal strategy would be to cut an clone a DNA fragment bearing two tandem terminators. Any recommendations?
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Here is a plasmid source:
It carries 5 tandem copies of rrnB1 T1 terminator next to a MCS and might be useful for what you want.
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Hi all,
I'm currently in the process of working on DIG-labeled mRNA probe transcription. I've done this successfully plenty of times, but the process never fails to create new ways to stump me, and I'm having trouble solving this one. I ligated my fragment into a PGEM T Easy vector and digested with SpeI for T7 and SphI for Sp6. I checked to make sure neither recognition sequence is in my fragment. I sequenced my plasmid after miniprep, and the sequences were perfect. The digest looked fine (photo attached) with the SpeI and SphI fragments both at the same, correct size. Only weird part is that the uncut plasmid ran slower that the cut plasmids, which has never happened. Issue persisted with replication. But that's aside the point. My PI told me to transcribe anyway since the cut plasmids were all the correct size.
I first transcribed last week, and the T7 looked perfect. Sp6 did not. I know that RNA can take multiple conformations, but I've never seen the bands look like this when that happens. Our T7 polymerase is pretty new, but Sp6 is a bit older, so my PI had me order a new tube. I tried again yesterday, re-transcribing both T7 and Sp6. T7 still looked perfect, but Sp6 did the same weird thing, just more intensely. It's hard to see on the gel because the second Sp6 transcription is so bright, but the bands are the same sizes for the Sp6 probe on both runs.
If it's a case of multiple conformations of RNA, I don't really understand why only Sp6 would be displaying it in both rounds of transcription. I have a feeling that my PI will suspect issues with the restriction enzyme and have me order more, but I want to explore any other avenues.
Any thoughts about what's going on here will be greatly appreciated.
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Didier Poncet this would make a lot of sense. We usually use NcoI for Sp6, which leaves a 5’ overhang, but the recognition sequence is in my fragment. I have some plasmids that have my fragment backwards. For my sense probe, I’m just going to cut those with SpeI and transcribe with T7, but this is extremely helpful information moving forward. I don’t know how long it would have taken me to figure that out on my own. Neither my PI nor the lab tech in the adjacent lab knew that the overhang made a difference! Thank you so much!!
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We have a gene with two isoforms, one with a longer 5' UTR and one with a shorter 5' UTR. It's been demonstrated that which isoform is predominant changes over development.
We want to study how the UTR might influence the translational efficiency of the transcripts. In the past, we tried to create a luciferase fused construct that had our UTR, our gene, and luciferase. However, the results were somewhat messy and hard to interpret and we questioned whether the luciferase construct itself was affecting translation.
Does anyone have any experience with alternative methods of studying 5' UTR's effect on translation, or have better experience with luciferase constructs?
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Don't use luciferase because too many variables affect the assay. Transfection efficiency, reaction times, substrate availability, etc. Just do western blots to examine protein levels directly. Control for transfection efficiency by using a second control plasmid, such as GFP.
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I recently did a transcriptome assembly using Trinity and I got one Fasta file. I want to do further analyses on unigenes only. My question is how do I identify the unigenes from the transcripts and have a Fasta file of unigenes only?
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Pleasure to you Lungelo Khanyile enjoy the downstream analysis of transcriptome data.
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I have been trying to get RNA via in-vitro transcription of a G-rich DNA sequence after PCR amplification since 5-6 months. But i fail to observe any RNA bands under UV light after running the in-vitro transcription product in Urea PAGE. Sometimes I observe a faint or a smeared band after EtBr staining. Can I get any suggestions about the same?
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Are you using SYBR green the stain your PAGE gels? It is hard to help beyond the gel technicality because I do not have a sequence to look at. If you share your promoter sequence and maybe a small portion of the initial transcribed region, it will be easier to help
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Dear community,
does anybody have experience with cloning two genes in opposing reading direction so that the genes theoretically could share just one bGH polyA signal.
I think the inverted polyA signal for one of them should not make a problem, since the polyA has no "direction". I am just concerned that the polymerases might sterically "crash" when both genes are transcribed at the same time but wouldn't that also terminate the transcription? :D
prom->tDTomato->bGHpoly(A)<-PuroR<-prom
Best!
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PolyA signal is not directional. There are consensus AATAAA sequence, cleavage site (CA), followed by GT rich region.
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I am not able to visualize anything on agarose gel except primers and gene ladder.
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i have included a control as well and it is amplifying with a good intensity
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Hi everyone,
Please correct me if the information mentioned here is incorrect. In the animal mitochondrial genome, there are no introns coding sequences.
I was wondering whether the primers designed for a mitochondrial gene (DNA sequence) work for the cDNA for its mitochondrial transcript?
Looking forward to a discussion!
Thanks in advance.
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That's great Saurabh Tiwari
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I have a big problem in my transcription. After taking my cDNA and PCR of my gene of interest, I did purification and miniprep. My results shown everything are great (nanodrop and agarose gel). But I can't have my RNA. In my agarose gel, I see a trail.
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Well... This is impossible to answer because it lacks very critical details. I have a few questions.
1. which rna polymerase are you using?
2. did you check that the promoter sequence is correct?
3. what are your transcription reaction conditions (does it match the literature)?
4. Are you using a common, well studied promoter or a novel one? Are you sure it is a promoter? Did you clone enough DNA upstream region for RNAP to bind? Does the promoter need an activator?
5. If you are using a native promoter, sometimes they require specific reaction conditions to get them to work.
You should consider these questions when you are troubleshooting this.
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Why is there a bright smear below my RNA band from in vitro transcription?
I have only observed it for my longer transcript.
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Your reaction looks pretty good for 5 kb. I would not worry about that smear, it is less than 5%. Load less sample and you would not even see it.
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...
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Given the differences in your questions and codebooks, I agree that you should concentrate your comparisons in the Discussion section.
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I want to publish the (qualitative) data that is associated with my research paper and have explored journals like Data in Brief and Scientific Data published by Elsevier and Springer respectively but they could not consider the data (descriptor) manuscript I submitted to them due to technical and situational reasons. Other journals I have seen are not suitable as many of them either consider other types of data or are based subjects different from mine. The data based on social research and consists of the interview transcript on pandemic policing and public (compliant) behaviour.
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Extending the answer fron Mahdi Movahed-Abtahi , your work might be adapted according to various other publication types to increase chances of publication. A letter
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Are there any bioinformatics tools or software available that allow verifying if the processed transcript acts as a lncRNA?
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Hi! I think the LatchBio console tools should work for this. I've used the platform before, and it is really easy to use.
I hope that helps!
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It is about Molecular Biology. Please answer up. Thanks
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There are two mechanisms responsible for proper transcript termination: in E.coli intrinsic termination and Rho-dependent termination. The Intrinsic termination is mediated by signals directly encoded within the DNA template and nascent RNA, whereas Rho-dependent termination relies upon the adenosine triphosphate-dependent RNA translocase Rho, which binds nascent RNA transcripts and dissociates the elongation complex. Although significant progress has been made in understanding these pathways, fundamental details remain undetermined. Among those that remain unresolved are the existence of an inactivated intermediate in the intrinsic termination pathway, the role of Rho–RNAP interactions in Rho-dependent termination, and the mechanisms by which accessory factors and nucleoid-associated proteins affect termination as reported on Annual Review of Biochemistry about six years ago
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The primers designed should be allele specific so as to amplify either my WT or Mutant transcript and not both.
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The best way to address this question is with ddPCR:
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I've been trying to do invitro transcription of 70 bp long DNA oligo using T7 RNA polymerase to figure out formation of G-quadruplex structure. The following components are added to the master mix for 20uL reaction volume:
1. 200nM ds DNA (prepared in 50mM tris and 10mM MgCl2),
2. Transcription buffer {trisCl (pH 8), 2mM spermidine, 50mM KCl, 10mM MgCl2, 10mM DTT} followed by
3. 40% PEG 200 and at last 2mM NTP .
I have successfully carried out transcription using this method earlier but now gradually the transcription seems to be having an issue.
The problem is when NTP is added to mixture having peg 200 , it immediately forms white precipitate. I tried repeating this in absence of peg and seen that there is no ppt formation. Not able to understand why is it happening. Is there any report of interaction of peg 200 with NTP?
After EDTA addition, the precipitate disappears. To check if DNA is precipitating due to Mg2+ chelation, I have also made the reaction mixture in absence of DNA , still I can see precipitate formation.
Can someone please help.
Note: I have assembled the reaction in room temperature.
Thanks in advance.
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It is the magnesium precipitation with phosphate or pyrophosphate. When the transcription happens, pyrophosphate is generated, if you don't add pyrophosphatase, then Mg will form insoluble magnesium pyrophosphate with it. if you add PPase, Mg will form Magnesium sulphates with the monophosphate. And the precipitation should be a sign for successful transcription.
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Hello,
We are currently evaluating software for the analysis of transcribed video recordings of dialogues. Coding units are episodes, and we want to quantify codings as duration of time. In previous studies, we successfully used Transana for similar analysis.
We are now evaluating to use MAXQDA instead, which does not assist reports of time duration of codings on transcripts, but only number of characters. For the export of duration data, coding has to be on the video, which does not meet standards for linguistic discourse analysis.
Does anybody know studies and/or have experiences with number of characters instead of duration as value for the quantification of transcript-based coding of dialogues?
Thank you for sharing your knowledge and experience!
Kind regards, Annelies Kreis
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I wrote the CLASS program to accomplish your goal. See http://class.wceruw.org/index.html & http://class.wceruw.org/class.html.
Good luck.
Martin Nystrand
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Im using mMESSAGE mMACHINE T7 ULTRA Transcription Kit to transcribe my DNA. It produced 30 ng/ul which is 750 ng although the kit promises to yield atleast 15-20 ug of RNA. Even the control DNA provided in the kit produced 6.2 ug RNA. I followed the protocol as it was instructed. Kindly share tips to increase the yield upto 10 ug atleast. TIA!
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As the previous answer mentioned, you can increase the time of the reaction. For T7 RNAP, I typically use 6-8 hours at 37oC. Make sure you have sufficient amount of NTPs for long reaction times. Likewise, you can increase the volume of reaction and consolidate
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I am trying to express a short RNA sequences in the nucleus in Drosophila. Are there any UAS-based expression vectors in flies that do not have PAS in the 3'? Or on the other hand, is it possible to just express a transcription termination sequence at the 3' of my transcript so that my transcript terminates before reaching the PAS? I don't want to have to mess with the vector backbone itself (removing PAS...etc.). I am quite new to molecular work regarding vectors, so if I am missing something important, under a misunderstanding of sorts, or if you have good suggestions, please let me know. Thank you!
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Talk with folks in your lab. Just because you want to do something doesn't mean it's possible or practical. Do what you know will work.
As my Ph.D advisor said "you can't rush quality".
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Hi, does anyone know of any post-transcriptional mechanisms that could explain no change of the transcript (mRNA) but still impact the protein?
so modifications that may show the protein's expression low in mRNA but the protein is still expressed in high amounts after translation?
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Malcolm Nobre thank you this helps
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I have in my RNA-seq quantification data from Arabidopsis obtained by mRNA-seq polyA enrichment library transcripts encoded by chloroplast and mitochondrial genes in significant DEG. How is it possible, if chloroplast and mitochondrial transcripts do not have poly-A tail? Are these data reliable or contaminants which should be ignored?
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I checked it and I probably found the response: Moreover, polyadenylation is often
a degradation signal in organelles, meaning that researchers
using poly(A)-selected RNA-Seq for measuring differential
expression in organelle systems may, in some instances,
be measuring the opposite: differential degradation (Smith and Lima, 2016).
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Hi guys,
I'm currently doing a research in BCR-ABL. I'm trying construct the Fusion transcripts that causes philadelphia chromosome. For example, presence of fusion transcript e1a2 leads to ALL. In order to create the fusion transcript, I designed primers to amplify the exonic regions of BCR and ABL individually and once amplified, to fuse the amplicons using BCR/Forward and ABL/Reverse primer.
Unfortunately, the individual gene amplifications hasn't worked and it just gives smeared appearance for a specific temperature. The template for the PCR is cDNA synthesized using MMLV.
Appreciate if anyone could help me with troubleshooting this issue.
The PCR cycle is,
Initial denaturation 95c/5min/1cycle
Denaturation 95c/30secs
Annealing 60c/64c/67c/ 30secs
Extension 72c/1minute
Final extension 72c/5minutes/1cycle
Hold 4c/infinity
Denaturation, annealing and extention went for 35 cycles in veriflex.
The gel image is attached for reference.
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How big is your expected product? The general rule is 1 min extension for every 1000 bp of DNA. You also might want to try a gradient of annealing temperatures to see what works best.
Good luck!
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Hello! I have been trying to produce IVTs from gBlocks, starting amount is ca. 100/120 ng of DNA.
When the reaction is done, I do a DNAse step, purification on column and then go to Nanodrop. When I measured the samples, I had a concentration of 1000-2000 ng/ul and a A260/A280 of more than 2.20.
I found this very weird, so I repeated the DNAse step on the IVT sample and purification again, but the situation didn´t change. I then did a third DNAse step, with a different DNAse, and again purification but even so I always have similar results.
My question is, can it really be that I have this much RNA? How can I check whether in my sample there´s RNA, DNA or a mix of both?
Thank you!
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Depending on the volume in which you eluted your RNA the concentration is not out of the realm of possible.
However, you should remember that nanodrop will also detect small fragments and possibly even nucleotides carried over from the template. You can measure your RNA using Qubit which is specific and will only detect the RNA.
You should also run a little bit on a gel to verify you get a clean band at the right size.
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I use some applications, but I seek more useful options. I wonder you're the applications that you are used to and your experiences with them. Thanks for your replies in advance.
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Good day
Best regards
Ph.D. Ingrid del Valle García Carreno
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I have ready collected several transcription data of tissues from patient(almost have metastatic region) with typical cancer . Is it OK for me to only compare the original cancer site v.s. metastatic sites for genes promoting metastasis? For that I have seen many researchers for similar purpose choose to compare original sites' data of patient with or without metastatic region, which is quite different from mine.
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Metastatic sites may share many of the transcriptome of the original tumor. I do not think that your comparison between primary tumor and metastasis will lead you to identify metastogenes. Remember that metastasis can also originate further metastasis.
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Hi all,
I'm working with an RNA-seq data set consisting of a large number of samples, sequenced at around 50-80M reads. There's a bit of uncertainty as to what the precise experimental workflow was for generating these data, but my best understanding at the moment is that the TruSeq RNA sample preparation kit was used (https://www.illumina.com/documents/products/datasheets/datasheet_truseq_sample_prep_kits.pdf).
This kit starts with total RNA, uses oligo-dT beads to bind polyA+ mRNA, then fragments the mRNA and carries out cDNA synthesis with random hexamer primers.
The data I've seen thus far show a very strong bias towards the 3' end of transcripts, in some cases so extreme that only the exons at the very 3' end are covered, with the rest of the regions having close to no reads at all. This bias is particularly pronounced in genes with long transcripts.
I'm aware that using oligo-dT priming is known to introduce a 3' bias into RNA-seq data as the reverse transcriptase will not always be processive enough to reverse transcribe in one go, but I'm at a loss to explain why the approach above might generate 3' bias if random hexamers were used.
Could anyone suggest any ideas as to what the possible causes of 3' bias in RNA-seq data might be? Are there any causes other than oligo-dT priming?
Would also really appreciate a link to a paper if one exists. Thank you!
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This could be due to the mRNA being somewhat fragmented even before the polyA+ capture. RNA is fragile stuff.
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In Sanger sequence, after having sequencer output (ab1 file), I would like to apply a free software to check the mutation on level of transcript. I was already working with “mutation Surveyor” software, but it is not freeL. Is there any free software? Thanks a lot.
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Check DNA Baser tool
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I am doing an experiment where I have to treat the organoids with insulin, harvest RNA and then perform RNA-seq. But I am not sure how long should the insulin treatment be to see an effect on transcription? Any insights on this would be helpful. Thanks!
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This depends on what you are looking for. At best, you could do a time course to look for early and late responsive genes. Are you interested in direct targets of the insulin signaling pathway or genes responding to changes in glucose or metabolism? For direct transcriptional targets you should consider using cyclohexamide to inhibit translation and thus any RNA increases are direct.
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Can you please suggest what could be the possible reasons for the confirmation of only 4 out of 13 genes (by qPCR), not all 13? Have anyone observed the same ChIP-qPCR validation issues? Any PubMed suggestion would be great. Thank you for your help in advance.
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Hi Amit
getting results from one experiment and failing to reproduce in an other experiment can allow you the differences and biases from both techniques. maybe you could look at the specificities of all your primers and see what's different in the 9 remaining genes. for instance you can check for specificity by testing your primers in silico at the UCSC website (http://genome.ucsc.edu/cgi-bin/hgPcr).
all the best
fred
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Hello everyone,
I am creating transcriptional reporters by cloning a promotor to a modified GFP in a plasmid. The GFP is modified by the addition of 13 amino acids at the end of it to render it less stable (if interest, see paper DOI: 10.1128/AEM.64.6.2240-2246.1998 ). I get the plasmid with the insert (promotor + GFP), but, there are always nucleotides' substitution in the 13 amino-acids tail added to the GFP. The mutations vary within the clones sent for sequencing. I have send the PCR product of the modified GFP used before the cloning, it does not have any mutation.
Where do the nucleotides' substitutions come from ? Any idea ?
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I had a similar problem with a protein that was apparently lethal to E coli. No matter what I did there were always indels and my transformation efficiency was super low. Try using a new bacteria strain with pLysS to prevent accidental transcription. Or supplement with 1% glucose in the media. Or do what I did and just screen a looot of colonies.
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I performed a CTmax experiment on Broad Whitefish at two acclimation temperatures and harvested liver and muscle tissue to perform HSP70 quantification in addition to acquiring a full transcriptome dataset for each tissue sample. After RNA-seq analysis, I quantified the transcripts using Salmon with Rainbow Trout and Northern Pike cDNA as the target transcriptome (there is currently no transcriptome dataset for Broad Whitefish). My goal is to find HSP70 specific transcripts and determine if there is a significant difference in upregulation between the two acclimation temperatures. I'm currently on Ensmbl and I have the reference names for specific HSP70 transcripts from the northern pike and rainbow trout, and I'm filtering the Broad Whitefish transcripts to find these reference names in the files and what the respective TPM values are. I'm wondering if there is a better way to quantify the HSP70 transcripts? Thanks in advance for nay help!
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You can map your reads against HSP70 directly, instead of a genome. You will not have a count per million, since you will only quantify this gene. But for comparison between samples, I think that there is no need!
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Dear all, I have found white precipitate in my transcription buffer which is expired 6 months ago (Promega ribomax sp6 kit). I used the buffer to synthesis mRNA and run a luciferase assay. Unfortunately, I didn't get any luminescence signals.is it because of my transcription buffer or any other issues?
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The precipitate was DTT. Just be patient to dissolve it. If you don't trust the buffer, make your own batch (you can find the buffer composition in the kit's manual). Have you quantified your RNA or visualised it otherwise prior to translation reaction?
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There are so many DNA polymerase and they have many activities like polymerase activity, gap filling, proof reading activity, and DNA repair activities. Which one DNA polymerase does not have DNA repair activity?
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I agree with Malcolm Nobre
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In a recent study, RLB (RICE LATERAL BRANCH) gene were demonstrated to control the plant height of rice. A loss-function mutation of the gene led to reduced stature. However, when the authors overexpressed the gene, the transgenic lines also showed reduced plant height. The authors further determined the expression levels of endogenous and transformed RLB copies, and found that both of them were decreased to very low levels.
Question: RLB gene was proofed to positively regulate the plant height of rice, we expected that overexpression of RLB may cause overaccumulation of the RLB gene transcription and increase to some extent, at least not reduce, the plant height. Why overexpression of RLB resulted in reduced transcription for both endogenous and transformed RLB copies and caused the similar phenotype with the mutant? Based on the knowledge we have learned about small RNA biogenesis and functions, please:
1) give the possible explanation of the phenomena;
2) describe the mechanism may cause the above phenomena;
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Use your class notes to answer your homework. This is a site to ask questions about research projects, not a place for students to cheat on their assignments.
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I would like to study correlation between four transcripts (fold changes of mRNA expression) at different time intervals (5 time points). How can I perform this analysis?
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Try Correlation matrix on R Programming. Try corrplot( ) package in R
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I want to differentiate between mutant transcripts and WT that differ by one insertion.
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If you have access to a qPCR machine, you can also try "high resolution melting" to look for the presence of both transcripts.