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Can you please advise?
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Do you want advice about the software or about qPCR primer design or both? I can help with qPCR design considerations, but not the software.
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Hi everyone, I am working on in-vitro transcription to generate mRNA for LNP fabrication. The kit I am using is the HiScribe® T7 mRNA Kit with CleanCap® Reagent AG from NEB. The first time I was doing it in October, the yield was very high. But the yield has been getting lower and lower since the second time.
Even with the control template the kit provided, the yield is always lower than 7 µg/20µL reaction.
Then, the technical support sent me a new kit, but the control template generated an even lower yield.
I've been talking to the technical support for a whole month. I am sure that I always put the T7 polymerase mix on ice, add reagents in the same order as the protocol, and set the reaction in room temperature. I am always aware of the RNAse. There was no RNA degeneration diffuse under my RNA band. The electrophoresis always shows very fainted bands even from the aliquot of unpurified 20µL IVT reaction.
Now, I suspect that the enzyme just started degenerating since I opened the T7 polymerase tube. Does anyone encountered this situation before?
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Thank you again for your help! I hope my experience can help. did it by improving the quality of the plasmids. I was using the phenol-chloroform method as described in the manual to purify it after linearization, which has a process of freezing the purified plasmid with NaAC and Ethanol for precipitation. It was time-consuming and harmful to my plasmid. Because the plasmid template could somehow inactivate during the freeze-thaw cycle. I finally used the purification kit, which is easier and allows me to use fresh plasmid every time. I am kind of sure thatan this freeze-thaw cycle damaged the plasmid (or maybe by other mechanisms) because once I freeze the plasmid, it won't work for IVT again.
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I have a list of genes and would like to find transcription factors that regulate them based on experimental evidence. I have used some online tools, but they provide different results. Can anyone recommend a reliable tool or database for this purpose?
Thank you!
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Hi Muhammad,
The relationship between a transcription factor and its target gene can also be determined by reviewing the literature. The Alphafoldserver website can be used to analyze the interaction between transcription factors and target genes.
First, access the Alphafoldserver website and select two modes:
  • Protein: Corresponding to the transcription factor sequence (can be retrieved from the UniProt database; species information should be noted).
  • DNA: Corresponding to the target gene sequence (can be found in the NCBI database), typically selecting the first 2000 bases of the gene for analysis.
After submission, the results will display the binding affinity between the transcription factor and the target gene.
Hope this helpful, Feel free to reach out for more details.
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I want to find out different isoforms of a gene. I know microarray or RNA seq can be used. But I need clarification or any other methods that can be used.
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According to me,
To find transcript isoforms of a gene, you can use:
  1. RNA-Seq (Best method) – Provides high-resolution transcript isoform identification and quantification.
  2. Microarray – Not ideal, as it mainly detects known transcripts and lacks isoform-level resolution.
  3. qRT-PCR – Can validate specific isoforms if primers are designed accordingly.
  4. Isoform-Specific Databases – Use resources like Ensembl, RefSeq, or GENCODE for known isoforms.
Microarrays are not suitable for discovering novel isoforms but can be used to detect known ones. RNA-Seq is the preferred method.
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I found the phosphorylation level of ERK1/2 in my cell sample increased when I treated the cell with a medicine, but the mRNA level of my target protein decreased obviously. Is that normal? Coz when ERK is phosphorylated, it usually bind to the transcriptional factor and promote transcription, right?
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Thank you very much! Kristina Sharlo
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I created a stem cell cell line with BCR-ABL1 fusion transcript. The Sanger sequencing at DNA level confirms fusion but could not be detected by RNA seq. The fusion in DNA level is located at the intronic region for both BCR and ABL1. Cell growth is different (higher) in the mutated cell ine with BCR-ABL1 fusion compared to The wild type counterpart. This is very likely because of the presence of such a cancer fusion gene. Is it possible that the fusion transcript is not possible to be detected in this case?
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In case the reference genome used for reads alignment does not contain the fused gene sequence,You may generate the reference genome using the expected fusion product and try aligning the reads to them.
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Generally, CRISPR-Cas9 editing leads to the formation of indels at the target site in the coding sequence of a gene. This often results in an altered reading frame (frameshift mutation) which will then lead to the formation of truncated/ non-functional protein. So, in this scenario, transcription should continue as usual and we should see a similar gene expression level as WT in the qRT-PCR. There is no doubt that transcripts will produce the truncated/ non-functional protein. Am I correct?
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It depends. There are "non-sense mediated decay" pathways that can remove mRNAs in the cell. But sometimes the mRNA expression levels are not changed. You'll have to figure it out empirically for your gene of interest.
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Since i already have factors that im studying. and im using Quanti quali as my approach. Can i precode factors in Atlas.ti, upload transcripts and look for supporting quotations to the results i obtain in quanti?
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Noor Fadhzana Mohd Noor Got it!!! thank you so much!!!
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Just sharing my experience of doing thematic analysis for my in-depth interviews conducted:
When conducting in-depth interviews for a qualitative study on school stakeholders, i tried to compose the responses in spoken language -verbatims and then to transcripts. Well ,with a little of my experience of qualitative research, can suggest a few approaches handle large datasets in qualitative studies, please note that it requires the following for effective management:
- Guys methodically classifying unprocessed data (e.g., coding transcripts in qualitative data analysis programs such as Atlas.ti or NVivo)- I have personally used Atlas ti and personally loved it. (becos of the web version)
- later considered dividing data into digestible categories using early coding frameworks.
The next challenging phase needs more focus- please note how it was handled..... The categories were iteratively refined to prevent redundancy and data saturation.
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Ok. But, how many focus groups? How many interviews? Length of each? Get a total--an exact number and length if possible How many researchers on the project?
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Dear all,
I am trying to label TPM3.1 in live cells. I was wondering if there is a Sir-Dye specific to this ? if not, is there an existing plasmid designed for IVT (mRNA) of TPM3.1-GFP or RFP or else tagged which I could inject to my cell?
Thank you in advance.
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Thank you so much for your suggestions ! Have a great day
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When I used 95bp PCR product (full-length tRNA with promoter), the T7 polymerase RNA synthesis kit (NEB) gave a lot of large bands (more than 200bp). Can you explain the reason and the way to remove all the bigger RNA bands.
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First, you need to confirm that apart from the target band, these is no any other bands observed in your PCR product which is larger than 95 bps. If the PCR product is pure, you may need to purify the tRNA transcripts using denaturing gel and verify its molecular weight by Mass spectrometry after purification.
Good luck.
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Hello, according to published papers: https://www.nature.com/articles/nprot.2013.143/figures/4
It is recommended to have an extra guanine before the 20 bp gRNA sequence, for my efficient transcription from the U6 promoter. If my gRNA already starts with a 'G', do I still need to add an additional guanine?
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Mouse U6 promoter transcription starts at the +1 position (23 nt after the TATA box), with G as the preferred initiation nucleotide. See this article:
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I was just wondering as we can (quite easily?) isolate both RNA and proteins from the same sample, why can't I find much info about sequencing transcriptome and proteome "at the same time"?
It seems that single-cell multiomics is in trend now, but looking at transcripts and proteins from the same samples looks like a simplified multi-omics from my perspective. What are the limits to that? Even companies don't seem to provide such services. Why is that?
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Simultaneous RNA-Seq and proteomics are uncommon due to several technical challenges. First, RNA and proteins require different extraction protocols, and attempting to isolate both from the same sample can compromise yield and quality. RNA and proteins also differ in stability, turnover rates, and abundance, complicating simultaneous extraction. Proteomics is more complex, as proteins undergo post-translational modifications (PTMs) that require specialized mass spectrometry techniques, unlike RNA-Seq. Additionally, quantifying transcripts and proteins involves different methods, making it difficult to correlate their levels. Limited sample availability can further reduce material for both analyses, and the lack of standardized protocols for integrating RNA-Seq and proteomics data adds to the difficulty of adopting this approach.
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Hi everyone,
For a qualitative research study, we are scoping for potential transcription software to transcribe our interviews. We have the following criteria in mind:
  • We are looking for affordable software (maximum cost of $200 for 16 hours/960 minutes of transcripts).
  • Software should be good at transcribing East-African accents in English (specifically, Ugandan accents).
  • Software should have high data protection mechanisms in place. At minimum, it should be compliant with GDPR legislation.
I already came across Otter.ai, Trint, Sonix.ai, and Rev.com. I am wondering if you have used any of this software before and can provide feedback? Other suggestions for software that meets the aforementioned criteria are also welcome.
Thank you in advance for your responses!
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For transcription software suitable for research in Uganda, consider the following recommendations:
  1. Otter.ai: Offers real-time transcription and is user-friendly, with a free version available.
  2. Rev: Provides accurate transcription services, though it requires payment for human transcriptions.
  3. Sonix: An automated transcription service that supports multiple languages and offers a trial period.
  4. Descript: Combines audio editing with transcription and allows easy editing of audio based on text.
  5. Trint: An AI-powered transcription tool that provides editing features and collaboration options.
These tools can enhance transcription efficiency for research projects.
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What instruments might measure empathy of adult students in asynchronous online learning discussions? We would have access to the transcripted text of academic conversations between peers without having access to the students who engaged in those discussions. Thanks for your ideas!
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As an academic in social work, measuring empathy in asynchronous conversations requires both a qualitative and quantitative approach to analyzing written texts. Since asynchronous conversations are text-based, we can use a number of tools and measures that rely on content analysis.
Sentiment Analysis: Natural Language Processing (NLP) techniques can be used to analyze texts and infer the level of empathy based on the linguistic patterns used. The focus here is on words and phrases that demonstrate emotional understanding and recognition of others’ feelings.
Qualitative Frameworks: A qualitative framework for analyzing written texts can be developed based on empathy theories in social psychology and social work. This is done by identifying indicators of empathy such as expressing appreciation for others’ experiences, or offering emotional support.
Social Interaction Scale: This scale can assess the quality and extent of social interaction between participants, including how responsive they are to others’ feelings and opinions.
Frequency Analysis: Measuring the frequency of use of empathic phrases such as “I understand how you feel” or “I agree with your experience” can be an indicator of the level of empathy.
Additionally, we could design a custom tool to measure empathy in academic discussions based on existing models such as the Davis Interpersonal Reactivity Index and adapt it to fit the analysis of asynchronous texts.
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I would like to know if T7 in vitro transcription can be effectively terminated by pol II terminators (specifically bGH poly (A) signal)? And in reverse can a pol II promoter (eg CMV promoter) induced in vivo transcription be effectively terminated within eukaryotic cells by a class I terminator specicially T7 terminator (TΦ large terminator)?
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I was wondering if you found a good answer to this question?
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The target transcript was clearly detected in my Northern blot results. Howerer, two " white" bands which are the same band sizes and locations as 18s and 28S rRNA were shown, what's the reason for this? thanks in advance.
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don't worry, this happen when there is a lot of material in one position (here 18 and 28S): there is no room for the probe to hybridize so there is no background at this position ... hence the white bands. It happens also with proteins in western blot ....
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It is said that the transcription is started by using a start codon.. my question is that why all those are necessary?
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Dis you mean start nucleotid (any of A,T,C, G bases) and not codon? Start base in transcription can be well upstream of start codon.
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I was wondering if it is possible to form a permanent open "ssDNA bubble" similar to a transcription bubble (>13 nucleotides) within E. coli. These criteria are important:
1. Open ssDNA bubble within replicable (in E. coli) genetic element. So no C-Traps under force.
2. No proteins, nucleic acids, or other toxic chemicals supporting the bubble. Can help during nucleation, but bubble has to be accessible for protein interaction.
3. Stable in bioorthogonal conditions. Physiological pH, salt, 37 °C, etc.
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Well, creating a semi-permeable transcription bubble can be challenging in the context of the structural stability of DNA as it tends to reanneal to its double-stranded form. Next is the concern of replication, which involves the fact that semi-permanent unwinding can potentially hinder the replication machinery from proceeding with DNA replication. Lastly, to maintain the transcription bubble to its semi-permanent unwound state, RNA polymerase is required to be halted in its activity at the transcription site, which could, in turn, lead to instability and interference in the replication of the plasmid. Considering these aspects, I believe three probable yet theoretical strategies can be adopted in this regard. First is genetically engineering a modified RNA polymerase, which can maintain the plasmid DNA at its single-stranded state by getting associated at a precise plasmid location without hindering the transcription process. Second is implementing genetically engineered single-strand binding (SSB) proteins, which can keep the plasmid DNA at its unwound state without interfering with RNA synthesis. Lastly, chemical molecules such as intercalating agents are introduced, which can develop proximal unwinding by being inserted at the nitrogenous base pairs of plasmid DNA; Molecules that are enhancers or activators of helicases; Hydrogen bond destabilizers like Di-Methyl Sulfoxide (DMSO), Urea or Formamide which can perform denaturation of double-stranded DNA; Cross-linking agents like Psoralens which forms covalent cross-linkages between single-stranded DNA molecules and DNA or RNA polymerases; Ligands which associate with single-stranded DNA such as Peptide Nucleic Acids (PNAs) and nucleic analogs which stabilizes the single-stranded structures of DNA; Alkylating agents such as Nitrogen and Sulfur derivatives of Mustard gas, Ethyl Methanesulfonate (EMS), Methyl Methanesulfonate (MMS), N-Nitrosoureas and Temozolomide. Nevertheless, besides being hypothetical, all these strategies have cons of their own.
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What recording and transcription apps/tools are best for historical research? I will be conducting interviews of about 60 - 90 minutes.
TIA
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I'd recommend Riverside.fm for transcription. Its quite more accurate than other tools I've used.
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Hi everyone,
I have DNA templates for T7 IVT mRNA transcription that are between 250 and 1050 bp long. I want to run T7 IVT mRNA synthesis using NEB2080 kit as recommended, but since tempates are so small, would you recommend to run IVT incubation overnight? For how many hours?
Any other recommendations would be greatly appreciated.
Thanks!
Kind regards,
Maria
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Hi Maria
We have been using templates of size 500-600 bp and we keep incubation time for 4 hrs. If you keep for longer time you can see degradation of finally synthesized mRNA
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As the title described, it is common that a target gene sequence contains motif that acts as the binding site of transcription factor rather then the upstream regulatory region. Does this affect gene expression? or transcription factor just release followed by subsequent upstream transcription? please share your thoughts.
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are you performing a gene reporter assay?
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Hi Everyone, I'm conducting research on culture shock using a mixed method design (Questionnaire + Interview). I have 60 Participants in Questionnaire and 10 participants (out of 60) in the interview. I have conducted my research regarding different factors causes culture shock (total 7 factors ). My question is that when writing the analysis section of my PhD Dissertation, how many interview participants should be quoted/mentioned in each factor analysis ? Is it appropriate to mention/quote the interview transcript of all 10 participants in each section or should I just mention/quote 3-5 participants? In most PhD Thesis that I have seen, the author mentioned/quoted a maximum of half of the participants.
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uhm maybe this paper could help:
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I was reading the T7 RNA polymerase protocol and noticed the following sentence regarding incubation time: "Incubate at 37°C for 1 hour. For shorter (< 300 nt) transcripts incubate at 37°C for 2–16 hours."
Can't think of a reason why short transcripts would take longer than long ones. Any ideas?
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"In general, transcripts that are shorter require a longer reaction incubation to allow for the highest yield (weight of RNA), as more RNA polymerase initiation events need to occur to reach the same weight of RNA compared to a longer RNA." - O’Donnell et al., Technical note "Scaling of High-Yield In vitro Transcription Reactions for Linear Increase of RNA Production", New England Biolabs.
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I will perform a apoptosis evaluation in cell lines using flow cytometry after siRNA transfection, and I want to evaluate the levels of BAX and BCL-2 to complement the experiment. However, I can only do a qPCR of both transcript, but not a western blotting. Can qPCR be used in this case, or it would be only informative if I measure the protein levels?
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Yes, qPCR can be used. It would be the most direct detection method to measure siRNA-mediated target knockdown. However, while using qPCR, the design of primers is an important consideration since using a primer set targeted to the wrong region may result in false negative results.
I would recommend that you should also perform Western Blot if you have the facility since it will show whether it is having the desired effect on functional protein as well. The qPCR results will show that the mRNA level of each gene is significantly lowered in the transfected cells than control cells. The western blot results can be used to correlate with the qPCR results where the protein band intensity reduction may be seen in siRNA-treated cells.
In this way, you could have valuable insights from protein as well that you will miss if you decide to just look at the transcripts alone.
Best.
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The desired gene region was amplified by PCR and the resulting PCR products were cleaned. Bands were observed in an agarose gel and measured in a spectrophotometer. Then, transcription was performed with the ABm Onescribe T7 transcription kit (E081). Of the PCR products, 196 ng for gene 1 and 133 ng for gene 2 were included in the transcription reaction. Despite the assurance that the kit was functioning correctly, the transcription products did not yield bands in agarose gel electrophoresis. No bands were observed with or without the optional DNase treatment following transcription in the protocol.
1. What could be the cause of the transcription problem?
2. What are the alternative methods to prevent DNA loss in the clearance of PCR products despite increasing the initial concentration?
3: How can the formation of dsRNA after transcription be determined? Can it be visualised with agarose gel electrophoresis? does the PCR product and the transcription product give the same gel image? Can it be measured in a spectrophotometer and what is used as a blank?
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There is one advice by @Victor G Stepanov that I would recommend too: "Actually, it might be useful to set up the T7 transcription reaction in small scale (~ 20-30 mcl), and then resolve the reaction mix on a gel to see if you had the mRNA synthesized in the first place."
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I am working on isolating Naive CD4+ T cells from human PBMCs using the Miltenyi AutoMacs. However, I am obtaining a higher percentage of Naive CD4+ T cells from the PBMCs than expected. I would like to confirm the identity of these isolated cells using transcriptional primers for mRNA gene expression analysis. Are there any specific primers that I can use for this confirmation?
Thanks!
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Naïve T cells typically express CD62L.
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I am planning to do invitro transcription and capping for an mRNA vaccine generation. Can anyone suggest whether it should be done on a bench top or inside a laminar hood?
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It’s prefer to do your experiment in laminar hood after sterilizing it, because its cleaner than bench
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I am performing PCR as a QC test to look for a transcription gene that should be negative after a CAR T therapy process. As we are comparing against a CAR transduced patient's cell, we require a used transduced ATCC cells. However, the ATCC cells have a low transfection titer, which makes the PCR band faint and when kept for long, it becomes fainter and fainter.
I was thinking of using another different grade of cells such as transduced research grade cells as it was observed that the bands tend to be much brighter than the ATCC grade cells.
Is it possible to use transduced research grade cells instead of ATCC grade?
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Using a different grade of cells for your positive control could potentially introduce variability or inconsistency into your experiment. It's generally recommended to use the same grade or quality of cells for both experimental and control groups to ensure reliable and accurate results. However, if you have valid reasons for using a different grade of cells for your positive control, it's essential to thoroughly validate and justify this decision to ensure the integrity of your experimental findings.
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Hi Guys
any Repository of Qualitative interview transcription??
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Are you looking for repositories that make qualitative data available for secondary analysis? If so, you might consider the Qualitative Data Repository at Syracuse University.
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Hello everyone,
I am working on the production of IVT mRNA. After transcription, the remaining DNA templates are digested by DNase I. I have tested transcription with uncut DNA (mostly supercoiled) and with linearised DNA as it is poorly described in literature, how the efficiency of transcription is reduced by using supercoiled DNA. What I ended up finding upon viewing my transcription products on agarose gels, was incomplete digestion of the DNA when it was in its supercoiled form. So I am now taking a closer look at that. Can anyone advise how the form of DNA impacts the efficiency of DNase I digestions?
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Agreed. After first cut in any accessible place on scDNA this DNA will not be anymore supercoiled and be digested as usual.
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Greetings, dear colleagues!
Our team conducts research on newly discovered SIRC elements in plant genomes ( , which are thought to be MITE transposons losing inverted repeats products, which could influence genome regulation) using bioinformatics, and we plan to conduct experimental molecular biology studies to elucidate the functions of SIRC. The problem is - our team is specialized in molecular bology experiments aiming to reveal the functions of genes, not non-coding DNA elements. That's why I want to ask your expert opinion - what experimental techniques would help to reveal the functions of abundant DNA elements of repetitive nature?
What comes to mind is the creation of mutant lines without several of these elements, but such experiments are too large-scale and can last for years, which is too complicated at the moment.
Another technique that comes to mind is the amplification of certain sequences and examination using circular dichroism spectroscopy to reveal whether given elements have unusual secondary structure like G-quadruplex of triplex DNA etc that could influence processes of genome transcription or replication.
And one more - we thought it could be possible to capture and identify plant proteins that specifically recognize SIRC via some modification of EMSA (electroforetic mobility shift assay) method. Unfortunatelly, up to date we didn't find any mentions of EMSA variant that uses not single purified protein, but whole DNA-free nuclear lysate, with subsequent identification of binding proteins via MALDI-TOF.
What other in vitro experiments could be useful?
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@Robert Adolf Brinzer The question is, what is the possible structural and functional role of newly discovered SIRC (Short Interrupted Repeat Cassettes) elements in plant genome. The point is that in bacterial genomes there are CRISPR cassettes which are looking similar to SIRC but have nothing common.
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What are the key differences and benefits over using CRISPR dCas9 system /CRISPR interference (CRISPRi) for transcriptional regulation over more traditional methods such as Structure-based combinatorial protein engineering (SCOPE) or others?
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Dear Esteemed Colleague,
Greetings. I hope this message finds you well and immersed in the dynamic field of genetic engineering and regulation. Your inquiry regarding the differences between the CRISPR-dCas9 system and traditional transcriptional regulatory methods highlights a pivotal area of interest in contemporary research. Below, I provide a detailed comparison that elucidates the distinct features, applications, and implications of these two approaches to gene regulation.
Overview of CRISPR-dCas9 System
The CRISPR-dCas9 (dead Cas9) system is an innovative adaptation of the CRISPR-Cas9 genome editing technology. In this variant, the Cas9 nuclease is catalytically inactivated (dCas9), rendering it incapable of inducing double-strand breaks. Instead, dCas9 is harnessed as a precise targeting mechanism to modulate gene expression when fused to transcriptional activators or repressors, or when employed to physically block transcription machinery access.
Traditional Transcriptional Regulatory Methods
Traditional methods of transcriptional regulation encompass a broad spectrum of techniques, including the use of small molecules, RNA interference (RNAi), and overexpression or knockdown of transcription factors or co-regulators. These methods typically act by modulating the cellular concentration of regulatory molecules or by directly interfering with the transcriptional machinery.
Key Differences
  1. Specificity and Precision:CRISPR-dCas9: Offers unparalleled specificity in targeting gene promoters or regulatory elements due to the programmable nature of the guide RNA (gRNA). This allows for precise modulation of gene expression without altering the genomic sequence. Traditional Methods: While effective, they generally lack the same level of target specificity. For example, small molecules and RNAi can have off-target effects, influencing genes beyond the intended target.
  2. Flexibility and Scalability:CRISPR-dCas9: The system's versatility is evident in its ability to be repurposed for either activation or repression of gene expression, multiplexing to target multiple genes simultaneously, and facile adaptation to diverse organisms. Traditional Methods: Typically more limited in scope for simultaneous targeting of multiple genes. Each target often requires a unique set of molecules or vectors, complicating multiplex applications.
  3. Mechanism of Action:CRISPR-dCas9: Acts by physically targeting the DNA sequence, either blocking transcriptional machinery access or recruiting transcriptional modulators to the site of interest. Traditional Methods: Act through various mechanisms, such as degrading mRNA (RNAi), altering transcription factor activity, or modulating chromatin structure indirectly.
  4. Temporal Control:CRISPR-dCas9: Recent advancements have enabled inducible CRISPR-dCas9 systems, providing temporal control over gene regulation. Traditional Methods: While inducible systems exist (e.g., inducible promoters), achieving precise temporal control can be more challenging and less efficient.
  5. Potential for Off-target Effects:CRISPR-dCas9: Despite its specificity, there remains a potential for off-target binding and regulation, necessitating thorough validation. Traditional Methods: Off-target effects are a well-documented concern, particularly with techniques like RNAi, which can lead to the downregulation of unintended transcripts.
Conclusion
The CRISPR-dCas9 system represents a significant advancement in the field of genetic engineering, offering specificity, flexibility, and precise control over gene expression that surpasses many traditional transcriptional regulatory methods. However, the choice between CRISPR-dCas9 and traditional methods should be informed by the specific requirements of the research project, including the desired level of control, target specificity, and potential for off-target effects.
Should you have further inquiries or wish to delve deeper into the applications of CRISPR-dCas9 in transcriptional regulation, please do not hesitate to reach out. I am here to support your scientific exploration and contribute to the advancement of genetic research.
Warm regards.
Check out this protocol list; it might provide additional insights for resolving the issue.
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During my analysis, I observed that genomic position 77 is annotated with gene symbols F, M, NP, and P across various transcripts. Please find the example below.
I used the gff3 file from NCBI https://www.ncbi.nlm.nih.gov/nuccore/AF077761 for variant annotation that includes details about gene symbols and transcript types. But I'm not sure what it means biologically to have different gene types at the same position. Does anyone have an idea on this? Please shower some ideas.
For example
NODE_30_length_153_cov_24.473684_77_-/C NODE_30_length_153_cov_24.473684:76-77 C gene-F rna-F Transcript upstream_gene_variant - NODE_30_length_153_cov_24.473684_77_-/C NODE_30_length_153_cov_24.473684:76-77 C gene-M rna-M Transcript upstream_gene_variant - NODE_30_length_153_cov_24.473684_77_-/C NODE_30_length_153_cov_24.473684:76-77 C gene-NP rna-NP Transcript 5_prime_UTR_variant 21-22 NODE_30_length_153_cov_24.473684_77_-/C NODE_30_length_153_cov_24.473684:76-77 C gene-P rna-P Transcript upstream_gene_variant -
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I don't work in virology, so there may be some conventions for virus genome annotation of which I'm ignorant. However, in the eukaryotic setting where there are often broad and complex regulatory interactions at play in the transcription of a gene, a variant being within 5kb upstream of the TSS would be sufficiently close to implicate it as potentially relevant to the expression of the gene. With this in mind, I suspect the variant at position 77 is associated with gene-NP because it lies within its 5' UTR, while it is associated with genes-P, -F, and -M because their TSSs are all less than 5kb away from the variant. These annotations alone do not mean the variant at position 77 is at all functionally relevant, let alone relevant to the genes to which it is assigned as a "5' UTR variant" or an "upstream variant".
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I have gel images of the plasmid that has been digested with various restriction enzymes and an image of the transcription of digested plasmid and a plasmid map with the cut sites located. I understand that the smallest transcription product on the gel is closest to the promoter. I get a band of  transcribed RNA approx 100bp with EcoR1 so I know that on the plasmid map the promoter is either 100bp upstream or downstream of the EcoR1 cut site but how do I then know the direction of transcription. 
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dentifying the position of a promoter on a plasmid and determining the direction of transcription involves several steps, including sequence analysis and understanding of promoter elements. Here's how you can approach it:
  1. Sequence Analysis:Obtain the DNA sequence of the plasmid of interest. This can be done using various methods, such as sequencing the plasmid directly or retrieving the sequence from a database or literature source. Analyze the DNA sequence using bioinformatics tools or software packages that allow you to visualize and annotate features such as promoters, coding sequences, and regulatory elements.
  2. Promoter Prediction:Look for sequences on the plasmid that resemble known promoter motifs or elements. Promoters typically contain consensus sequences recognized by RNA polymerase and transcription factors, such as the TATA box, CAAT box, and GC-rich regions. Use promoter prediction algorithms or databases to identify potential promoter regions based on sequence features and known promoter motifs.
  3. Orientation of Genetic Elements:Determine the orientation of genetic elements, such as coding sequences or selectable markers, relative to the identified promoter region. The directionality of these elements can provide clues about the direction of transcription from the promoter. In most cases, the promoter will be located upstream of the coding sequence it regulates. Therefore, the direction of transcription will typically be from the promoter towards the downstream genetic elements.
  4. Experimental Validation (optional):Conduct experimental assays to confirm the predicted promoter activity and direction of transcription. This may involve promoter reporter assays, such as measuring the expression of a reporter gene (e.g., GFP) driven by the putative promoter sequence. Use techniques such as RT-PCR or Northern blotting to analyze the direction of transcription and detect transcripts originating from the identified promoter region.
  5. Annotation and Documentation:Annotate the plasmid map or sequence file to indicate the position and direction of the identified promoter. This information will be useful for future reference and experimental design. Document your findings in a clear and organized manner, including the sequence coordinates of the promoter region, the predicted direction of transcription, and any experimental evidence supporting your conclusions.
By combining sequence analysis, promoter prediction, and experimental validation, you can identify the position of the promoter on a plasmid and determine the direction of transcription, providing valuable insights into the regulation of gene expression in the plasmid construct.
l This list of protocols might help us better address the issue.
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I recently encountered an intriguing situation while examining a plasmid constructed by someone else for a eukaryotic expression system. This plasmid contains a unique arrangement of open reading frames (ORFs) that has sparked several questions regarding the potential outcomes of their translation.
In this plasmid, there is an ORF near the 5' end, where the translation initiation site is quickly followed by a stop codon, potentially resulting in a very short peptide. More interestingly, nested within this first ORF is a second ORF that begins inside the first ORF and could potentially translate into a much longer protein, consisting of 500 amino acids.
Given the common understanding that eukaryotic transcripts typically feature a single ORF, the discovery of this arrangement has led me to ponder the following questions about the translational dynamics in this specific scenario:
  1. In the context of this plasmid, will the translation machinery be capable of bypassing the short ORF to translate the longer protein, or will it prioritize the translation of the short peptide due to its proximity to the 5' end?
  2. If both peptides are indeed translated, what might be the expected ratio between the production of the long and short peptides?
  3. Is there a possibility that only the short peptide will be translated, effectively ignoring the translation potential of the longer, nested ORF?
Furthermore, I'm curious about how this scenario might differ if the plasmid were used in a prokaryotic system, which is known for its ability to translate multiple ORFs within a single transcript.
I'm seeking insights, experiences, or any relevant literature that could help shed light on the translational strategies employed by cells when faced with plasmids containing nested ORFs, especially in the context of eukaryotic expression systems.
Thank you in advance for sharing your knowledge and experiences.
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There are a myriad examples of this sort of ORF structure in eukaryotic viral systems (particularly RNA ones), where it has evolved to diversify/expand coding capacity and/or regulate viral gene expression. At first pass you can predict relative expression levels by examining the Kozak consensus around each AUG codon. We usually consider the -3 and +4 positions to categorise them as strong, middling and weak. If the first AUG is strong, you typically won't get much of the second product. If the first AUG is medium or weak and the second is strong, you'll get lots of the second product. Confounders to look out for; 5'-UTR length and protein stability. AUGs very close to the 5'-cap are often not translated well. Shorter proteins are harder to detect and if they're too short (or the wrong sequence) to fold up into a coherent structure, they'll be turned over rapidly and can be hard to detect unless (or even if) the proteasome is turned off.
After that, it can get properly complicated with all sorts of ribosomal gymnastics depending on the exact sequence of the transcript.
Bugs - need to consider Shine-Dalgarno (sp?) sequences or lack of, I think. But my undergrad lectures on this were many years ago :)
Cheers
Paul
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Hi ALL,
I am using a pair of primers to amplify a region in my gene of interest from cDNA samples. The cDNA samples are extracted from tissues of mouse of different ages. The gene is known to have decerased expression level when mouse ages. However, I did not see any change of the RT-PCR amplicon band intensities on agarose gel, indicating no change for the transcript level. I did not saturate the PCR products as I tried different cycle numbers (from 23 to 30 cycles). What could be the possible reasons? Should I design new primers targeting a different regions in my gene? Thank you for the help!
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Sequencing amplification product is always a good option to determine the specificity of the PCR. In addition, you should at least run an end-point PCR with a low number of cycles for an appropriate house-keeping gene as control for your input.
Could it be possible that your primers also amplify genomic DNA, which basically always contaminates your RNA unless you conduct an DNAse treatment step. If so, design exon-exon spanning primers.
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By using CRISPR-Cas9, we can insert any sequence into any locus we want, right?
Base on that,
What I'm actually curious about is that,
Is it possible to regulate the transcription of endogenous gene by inserting short sequence ( for example, binding site of specific TF ) at TSS-proximal region?
I'd be appreciated to hear examples of any studies that have a similar concept to "put a short sequence upstream of a gene".
Thank you for your interest in my question :)
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Unfortunately, this is not novel. Many have already done that. You're looking for is CRISPRi (silencing) and CRISPRa (promoting).
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What tools or methodologies are available to comprehensively analyze RNA expression profiles across various human tissues, specifically to discern and compare transcript variants of a gene
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Dear Prof. Mukaram Bhat
Genome browsers often provide links to external databases that contain additional information about transcripts and genes. You may find links to resources such as NCBI Gene, Ensembl, or RefSeq.
Regards
Jogeswar Tripathy
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We are looking to confirm transcript expression in cells that are positive for a reporter gene (tdTomato) in mice. We are able to locate the transcript in question via RNAscope. However, following the RNAscope protocol, we are unable to locate *ANY* tdTom neurons.
We know that tdTom is expressed in these mice, as we are able to visualize tdTom neurons in alternate series that were not exposed to the RNAscope protocol. Thankfully, the observed tdTom neurons overlap with transcript expression in alternate series. however, we need to be able to colocalize these signals with cellular resolution.
Any experience, suggestions, and/or references would be greatly appreciated.
C
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We leave out the ethanols then tomato ist still present
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I have cloned my genes in a CMV vector and transfected them in HEK293 cells. After harvesting the cells, the expression level at the transcriptional level is increased compared to the control well. However, the peptide sequence is not seen to be expressed as analyzed by mass spectrometry.
Can you please give an insight into the possible reason for the same?
Thank you.
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Hi,
some expression vectors also carry sequences for in vitro cell-free translations. If your vector does not have it, try different one with these additional pieces. It will allow you to have your peptide or, at least, will show that problem with expression (on any level) in original setting. Good luck.
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Hello!
This is my first time analysing qualitative data and so I have chosen to use QDA miner. I have six transcripts to analyse.
I have watched multiple tutorials on YouTube on setting up a project and followed them exactly. However, when I select the files first of all I noticed I am unable to click the remove text formatting option. The box is there but I cannot select it. Then when I import the documents they all have no text in them. The file names have imported but they are blank
I have tried converting my files into pdf and rtf versions in case this helped.
Any advice welcome!
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Rosie Hhh, Did you get a solution to this problem? I have a similar situation - be it PDF or Word or RTF, they all show as blank.
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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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It should be fine. Just know that if you didn't do a cleanup step to remove the DNAase etc. then the DNA is still present (just chopped into pieces). What are you planning to do with the transcripts?
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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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I intend to use open-ended questions and target linguistics students, although I haven't yet begun to formulate the questions.
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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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It is also possible that your gene is transcribed at a very low level but translated at a high level from those transcripts.
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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".