Science method

PCR - Science method

PCR is an in vitro method for producing large amounts of specific DNA or RNA fragments of defined length and sequence from small amounts of short oligonucleotide flanking sequences (primers). The essential steps include thermal denaturation of the double-stranded target molecules, annealing of the primers to their complementary sequences, and extension of the annealed primers by enzymatic synthesis with DNA polymerase. The reaction is efficient, specific, and extremely sensitive. Uses for the reaction include disease diagnosis, detection of difficult-to-isolate pathogens, mutation analysis, genetic testing, DNA sequencing, and analyzing evolutionary relationships.
Questions related to PCR
  • asked a question related to PCR
Question
4 answers
Hi everyone,
I was running a PCR and, unfortunately, the thermocycler died around the 7th cycle. It felt like a waste to simply throw away the PCR, so I put it on another thermocycler and lowered the cycle number to 27 (it is usually 34). Has anyone had experience with changing thermocyclers during PCR and what were your results? Should I even use this PCR for downstream steps or should I just re-do it entirely?
The thermocycler settings are:
1) 94C, 3 min
2) 94C, 30 sec
3) 55C, 30 sec
4) 72C, 45 sec
5) Go back to Step 2, 34x (again, I lowered this to 27 when I changed thermocyclers)
6) 72C, 20 min
7) Hold on 4C
Thank you.
Relevant answer
Answer
If you finished the PCR you could let us know if it worked. In principle, there should be no problem to continue with the rest of the program on a second cycler. Remember, in the early days of PCR there were no thermostable polymerases, so they had to add the enzyme after every cycle of the reaction, so interrupting the program for a few minutes should still give you enough product.
  • asked a question related to PCR
Question
2 answers
Hello. I'm looking for suggestions on troubleshooting index (or barcode) PCR. Specifically, my problem is that some of my sample DNA concentrations dropped after adding the indices/barcodes. My ideal concentration is no lower than 9nM. One 96-well plate had 5 samples and another plate had 28 samples that were below 9nM.
For context, I'm adding Illumina indices to my mixed amplicon (combined ITS and 16S) sequences. After the index PCR, I cleaned up my samples and quantified them with Qubit. I had some samples that dropped significantly in their DNA concentration after this step (index PCR). For example, one sample had 133nM (measured after 16S amplicon PCR), but then dropped to 8nM (measured after index PCR).
Prior to indexing, I diluted the separate ITS and 16S plates to 20nm then combine 2uL of each of the 20nM diluted plates together. From the now mixed 4uL, I then use 1.5uL of each mixed amplicon sample for index PCR. For the index PCR, I typically use Kapa HotStart ReadyMix (7.5uL for 1 reaction), water (3uL for 1 reaction), and 1.5uL of each index (for each sample).
The thermocycler settings are:
  1. 95C, 3min
  2. 95C, 30sec
  3. 55C, 30 sec
  4. 72C, 30sec
  5. Go to Step 2, 7 times
  6. 72C, 5 min
  7. Hold on 4C
I've already increased the DNA concentration input for the index PCR. Originally, it was 10nM, but 20nM worked better. I'm unsure about increasing the DNA concentration again as many of the samples lowest concentrations (prior to indexing) were around 20nM.
Another suggestion was to toggle with the thermocycler settings, but I'm unsure how to optimize these as other samples worked fine with these exact settings.
Thanks for reading this very long post. I'm open to any suggestions on how people have troubleshooted indexing!
Relevant answer
Answer
@Raghad Mouhamad Thank you for the detailed answer. I plan on increasnig the cycle numbers.
  • asked a question related to PCR
Question
1 answer
Vectors which can accept PCR products up 700-800bp
Relevant answer
Answer
700-800 bp is not that big. So pretty much any vector you would want to use should be able to handle that size PCR product without any difficulty.
  • asked a question related to PCR
Question
6 answers
I ordered a primer when I received it where one nucleotide is different in the forward primer.
the original sequence is: CTCTTTGGGCTCAGAGTGAGTCTGG
the sequence which I get: CTCTTTGGGTTCAGTGTGAGTCTTG
Three nucleotides (Bold) are different
This primer will work?
while I have tried so many times but there is no result. If it is working, how can I fix my PCR protocol?
Relevant answer
Answer
Just buy a new primer! It's not worth your time or reagents to try and get a faulty primer to work. Primers are cheap.
  • asked a question related to PCR
Question
4 answers
Tetra ARMs PCR
Relevant answer
Answer
Hello, in your question, it was my first time encountering the word T ARMs PCR. From the article, I found that T-ARMS PCyR (Tetra-primer ARMS-PCR) is a single-step genotyping method for detecting single nucleotide polymorphisms (SNPs). It involves a single PCR reaction using four primers to amplify specific DNA fragments, which can reveal the presence of different alleles. The four primers include two outer primers (Outer Forward and Outer Reverse) that amplify a larger region around the SNP as a control and two inner primers (Inner Forward and Inner Reverse) that are allele-specific, amplifying smaller fragments to indicate the presence of either the wild-type or mutant allele. Depending on the genotype, the PCR will produce a distinct pattern of three possible products: a large fragment for the control and smaller allele-specific fragments that help identify whether the sample is homozygous wild-type, heterozygous, or homozygous mutant.
After PCR, the amplicons are separated by gel electrophoresis to observe the characteristic band patterns: the wild-type homozygous genotype will show bands for the control fragment and one allele-specific fragment, the heterozygous genotype will show bands for the control and both allele-specific fragments, and the mutant homozygous genotype will display the control fragment along with the other allele-specific fragment. The choice of polymerase affects the reaction's accuracy and efficiency. Traditional Taq polymerase, with 5′–3′ exonuclease activity, can cause non-specific bands and often requires the use of DMSO for stabilization, especially in GC-rich regions. In contrast, SD polymerase, which has strong strand displacement activity but lacks exonuclease activity, minimizes non-specific bands and performs well across a wider range of temperatures (50-60°C) without the need for PCR enhancers like DMSO. This flexibility makes T-ARMS PCR a cost-effective and straightforward genotyping method, although careful optimization is still required depending on the specific conditions and polymerase used.
Reference: Alyethodi, R. R., Singh, U., Kumar, S., Alex, R., Deb, R., Sengar, G. S., Raja, T. V., & Prakash, B. (2018, February 15). T-arms PCR genotyping of SNP RS445709131 using thermostable strand displacement polymerase - BMC research notes. BioMed Central. https://bmcresnotes.biomedcentral.com/articles/10.1186/s13104-018-3236-6
  • asked a question related to PCR
Question
1 answer
What do you do when you have tried everything to transform your yeast strain and you see a lot of colonies but most of them are false positives.
Regarding confirmation of transformation, I isolate the gDNA and then perform a PCR but for some reason I do not see any bands even with the positive control. My technique is fine and I tried every possible change with no results. HELP ME!
Relevant answer
Answer
If you aren't getting your positive control to work, then the issue is with the PCR. My first guess is that your primers have degraded.
Make a new dilution from the stock tube or order a new tube.
Make sure your controls are working before you try to do any PCR of your samples.
Good luck!
  • asked a question related to PCR
Question
3 answers
I have been doing PCR genotyping for mouse samples for 2 years. It worked well before, until I ordered new gotaq polymerase last month. I kept the polymerase at -20C. When I used the new polymerase, I did not see any band, but when I increased the amount of taq pol, I could see the band. I then used it again for current genotyping, but it did not showed any band, even when I increased the taq amount per reaction. I used positive and negative control that previously worked, but they also did not showed up. I tried changing the taq, PCR water, dNTPs, and diluted fresh primers from 100uM stock, also dilute DNA and increased PCR cycle. All cannot solve the problem. Then I bought new primers and collected new tail, and suddenly it worked and showing band nicely. However, when I repeated the experiment (only one day apart; using the exactly same reagents), I could not see any bands.
Can anyone help me with this issue?
i have been struggling with this for 3 weeks.
Thank you!
Relevant answer
Answer
I would start by diluting a new primer stock in TE (not water). Set up a pcr with positive control using normal amounts of Mg and primer and also tubes with 2x as much Mg and separately 2x as much primer. Before using all ragents thaw eack reagent and them mix by flicking the reagent tube. Sometimes when thawing frozen materials you get water layer on top and concentrated salt/oligo /protein layer at the bottom and you can pipette out water if you take up the upper layer of reagent. Be sure to dilute your primer in TE in case the water has nucleases in it which chews up the primer. run the samples with a dna ladder to check that this is not a gel problem...if you can see the ladder all is well with the gel
  • asked a question related to PCR
Question
10 answers
Hello
I am having a trouble with the result of agarose gel electrophoresis of the PCR product of Entamoeba gingivalis
DNA
Samples:
Paper point samples from the sulcus(between tooth & gingiva)
Stored in a PBS transition solution
DNA extraction:
1-samples heated to 37˚for 10 mins and mixed well on vortex.
2- 0.3ml of the suspension was washed 3 times with distilled water and spin down between each wash at 14000 rpm for 3 minutes
3- the pellets were re-suspended in 100 microlitre nuclease free water.
Pcr procedure:
1- Primers forward and reverse specific for E.gingivalis were used
2- GoTaq green master mix was used
3- The master mix consisted of the following:
A- 150 µl master mix(GoTaq green)
B- 6 µl forward primer
C- 6 µl forward primer
D- 114 µl nuclease free water
Divided on 12 samples =23 µl
Add 2 µl of each specimen to be the total of the mix 25 µl
4- The reaction cycle was prepared as following:
A-Initial denaturation 95˚ 2minutes
Then 30 cycles of the following
B- 95˚ 30 second
C- 55˚ 30 second
E- 72˚ 35 seconds
Final extension 72˚ for 5 minutes
Question is the following image of the electrophoresis is typica
Thanks in advance
update:
the expected size of the PCR product is
135
in the previous image
today i changed the voltage to 80
and the running time to 40 minutes
and got the following result.
Named IMG_3106
the first 10 lanes are the pcr products
11th is the ladder
20th negative control
update:
20 oct 2024
thanks alot for all the significat paticipation
the agarose gel is prepared by adding 1.5 gram powder to 100ml of TBE (1X) and 5µl of gelred
attached a new picture named IMG_3249
is a photo of the electrophoresis
of a positive control by doing
pcr for RNA 16 s
thank you
the last row is the negative control
Relevant answer
Answer
The ladder and the samples are now running well in the agarose so well done . I would like to see a pcr run with 3 samples which have no added dna ( negative cntrol samples ) because the amplification in sample 20 worries me that you may be generating results which come mainly from only one sample and the end results may be impossible to interpret if you do have pcr contamination
Question
1 answer
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.
Relevant answer
Answer
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.
  • asked a question related to PCR
Question
1 answer
Hello,
I am using amaR one PCR mix. My anticipated amplicon size is about 500 bp. I need help understanding the darker band at the bottom of each well. Is it the primer dimer or the Amaranth stain that comes with the PCR master mix?
Please help.
Thank you
Relevant answer
Answer
what is your dna ladder and is well 2 a dna sample or your no template control please
  • asked a question related to PCR
Question
5 answers
Hello!
I am optimizing conditions of stem-loop RT-PCR to compare the expression of miRNAs in different cell lines.
Now I'm performing the reaction in one step: RT with stem-loop primer and subsequent real-time PCR with a pair of primers and SYBR Green - all in one reaction. So my reaction mix contains three types of primers, heat-inactivated reverse transcriptase, hot-start Taq polymerase, SYBR and all the nessesary components such as ddNTPs and Mg2+.
I use synthetic miRNA and total RNA from human cells as matrices.
When working with synthetic miRNA, everything goes well (Figures 1 and 2), amplification plot is ok and there is a single peak in the melting curve.
But with total RNA I observe a decline of fluorescence in the plateu phase of amplification (Figure 3) and 2 product peak in the melting curve (Figure 4).
Reference gene amplification plot doesn't have such issues.
What are the reasons of the decline of fluorescence after the 30th cycle?
What are the possible variants of the second product here? Primer-dimers observed in no-template control have Tm = 80 degrees C.
Relevant answer
Answer
I was a bit fast in my response... re-hybridization so to my knowledge the only known reason for hooking amplification curves - but this effect is understood only for hybridization probe formats. You seem to use SYBR Green I detection. Such effects have been observed for SYBR Green I detection, but the mechanisms here are not clear. It is hypothesized that photobleaching could play a role. However, the interesting finding in your case that the hooked curves occur together with a second amplicon, some triple-helix formation or the formation of other secondary structures during annealing may reduce the effective space for binding of SYBR Green molecules and the linear energy transfer trough the amplicon molecules, resulting in a lower net florescence.
Again, just a wild guess. It may help to have a look at a native and denaturing PAGE of the PCR products, or even at their sequences. If you are really interested in clarifying the mechanism here you might want to generate synthetic molecules of that sequence and analyze the net fluorescence and UV absorption for different mixtures.
For your qPCR of miRNAs, this effect is not relevant, as long as the wrong product is amplified late. You could stop the PCR a few cycles after the exponential phase and check a melt curve and gel there. If at this cycle there is only the correct product amplified, the Ct value is valid. Otherwise you will have to optimize the assay (vary MgCl2 and/or primer concentration; it sometimes helps just to use/test another brand [that often includes a different variant of the polymerase as well]).
  • asked a question related to PCR
Question
3 answers
Hi, I am doing qPCR experiments. When the reaction finished, I obtain the Ct value of my samples is zero, but they have a peak with my expected Tm. Furthermore, I run the gel electrophoresis, the result also shows that these samples appear a band at my expected target size. But why the Ct value is zero?
Relevant answer
Answer
Look at the file containig information about the increase in the fluorescent signal as a function of Ct. Check how the Ct threshold is defined - it might be incorrectly positioned. Set it within the exponential phase of PCR.
  • asked a question related to PCR
Question
2 answers
During cDNA synthesis, same amount (400 ng) of RNA was added for each sample. The volumes varied for each sample. With the synthesized cDNA we performed conventional PCR for beta- actin. When the gel was run we saw the sample in which the initial RNA volume (not conc.) was more than others had a strong band density as compared to the one in which less volume was used to synthesize cDNA. This has happened multiple times.
Can anybody tell what can be the reason?
The image is of PCR product (RNA>cDNA>PCR) plotted as:
Well 1: ladder
Well 2: PCR product with RNA volume 13.07ul
Well 3: PCR product with RNA volume 2.86ul
Well 4: PCR product with RNA volume 3.86ul
Relevant answer
Answer
How did you determine the concentration of RNA in your samples? If you used a spectrophotometer or nano drop, you are measuring all of the nucleotides (DNA & RNA). If there is a lot of DNA in your samples, then you are getting an artificially high concentration.
Did you run out a sample of the RNA on an agarose gel to check that it's intact? Degraded RNA & intact RNA will also give the same reading with a spec.
A Qbit can differentiate DNA & RNA in a sample. Reagents are a bit expensive, but it can be worth it.
  • asked a question related to PCR
Question
1 answer
Ask a question for the JAX Stat3/flow mouse and the primers (19436/19437) which was provided by JAX. The expected results of standard PCR showed that WT=146-bp and mutant=187-bp. From the gene blast, the biding sites in the 7 Intron region, how to explain that PCR can extended to 18 Extron to 20 Extron?
Relevant answer
Answer
I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?
The ability of primers to extend PCR amplification from the 7th intron to the 18th and 20th exons can be attributed to the placement of the primers flanking a larger genomic region. This allows for amplification across multiple exons and introns, capitalizing on the continuity of the genomic sequence from the primer binding sites. The primers are likely designed to cover a substantial genomic area, which facilitates the amplification of an extended sequence that includes the target exons, despite initial binding in the intronic region. This approach can be common in genomic studies where spanning multiple regions is necessary for comprehensive analysis[1][2].
Reference
[1]
Song, X., Liu, Z., & Yu, Z. (2020). EGFR Promotes the Development of Triple Negative Breast Cancer Through JAK/STAT3 Signaling. Cancer Management and Research, 12, 703-717.
[2]
Zou, L., Yu, L., Zhao, X., Liu, J., Lu, H., Liu, G., & Guo, W. (2020). MiR-375 Mediates Chondrocyte Metabolism and Oxidative Stress in Osteoarthritis Mouse Models through the JAK2/STAT3 Signaling Pathway. Cells Tissues Organs, 208, 13-24.
  • asked a question related to PCR
Question
2 answers
Could you kindly guide me through the steps to design primers for the "OneStep PLUS qMethyl™ PCR Kit" to study the methylation status of specific genes?
Relevant answer
Answer
*Primer3 software
  • asked a question related to PCR
Question
4 answers
I have run colony PCR. I was supposed to get amplicon of 300 bp size. However, after running the PCR, i am not getting 300 bp amplicon. The bands are just stuck in the well. What should I do to get exact sized amplicon after PCR?
Relevant answer
Answer
Are you sure, that the bands which just stuck in the well are not genomic DNA, only? -> No amplification.
  • asked a question related to PCR
Question
6 answers
I am conducting DNA extraction from the juvenile leaves of cherry tomato (F4 generation) for molecular studies. I would appreciate suggestions on the best protocols or methods that provide high-quality DNA suitable for downstream applications like PCR. I am particularly interested in methods that work well with plants that have high secondary metabolites or are prone to contamination. Any specific tips on handling these issues would be helpful!
Thank you in advance for your insights!
Relevant answer
Answer
I would like to extend my gratitude to all the contributors Uditha Maduwantha Dissanayaka Mohini Kajla Sakshi Balyan Park Sowon Harish Chandra Singh who provided valuable recommendations regarding DNA extraction protocols. After reviewing the various approaches, I conducted the CTAB method for DNA extraction at our molecular lab. However, based on my observations, I noticed that prolonged storage of the leaf samples negatively impacted the DNA concentration. Specifically, when leaf samples were stored for extended periods before extraction, the DNA concentration was significantly reduced, likely due to degradation over time. Therefore, I recommend immediate or timely processing of leaf samples to ensure optimal DNA quality and yield.
Once again, I sincerely thank the community for your insights.
  • asked a question related to PCR
Question
11 answers
I'm trying to clone some large pieces of cyanobactrial gDNA (sometimes even more of them) into a plasmid via fusion PCR. Then, I transform it into E. coli TOP10, as my PI says it works better (for HiFi DNA assembly).
However, I quite high percentage of my positive clones had some problems in the plasmid elsewhere (like missing I think 2.3 kbp of the plasmid; or some mess in another case etc.).
It is true, that when I compared TOP10 and DH5alpha, TOP10 had many more colonies (more than 10-times). However, I think that DH5alpha should be less prone to DNA alterations, right? So should I use rather DH5alpha and hope that I will get at least few colonies, but they will be positive?
Relevant answer
Answer
I have tried both methods. Sometimes PCR cleanup is enough and gives good results but if the fusion is problematic, gel purification is much better.
  • asked a question related to PCR
Question
6 answers
I am using PCR product as my template for in vitro transcription and then loading the Rna samples on 8M urea gel. I am observing multiple bands of both low and high molecular weight(my intended product is of 550bp). What could be the reason. Any suggestions?
Relevant answer
Answer
Didier Poncet I tried lowering the temperature to 4°C, but the yield was very poor, and it wasn't sufficient to load onto the denaturing acrylamide gel.
  • asked a question related to PCR
Question
4 answers
Hello ResearchGate group,
As M.D.-Ph.D. scientist working on molecular biology, I am wondering if anyone is interested in joining efforts for creating PCR standardization statistical tools.
Best regards,
Alexios
Relevant answer
Answer
Alexios-Fotios A. Mentis I am in no position to join your project at this time. I would love to see the results you and your team come up with! This is an area that needs some standardization. Best Wishes to you and your team.
  • asked a question related to PCR
Question
3 answers
I have synthesized a gene specific primer for amplification of my target gene. However, I am getting amplicon of only 100 bp rather than 210 bp (target amplicon size). So far I have tested so many optimization conditions for it but still getting non-specific band. I have cross checked the primer pair for non specific binding in my extracted DNA, it shows no complementary with any other region except for my target region.
Relevant answer
Answer
Hi Moein,
which is the melting temperature (Tm) of your primers? Which are you using to amplify your target gene?
Tm is key for primer specificity. You should adjust the Tm so it is very close to the lowest primer Tm. I usually try to use 62ºC, although up to 55ºC should work.
Another chance is the appearance of primers dimers. An amplicon of 100bp could be the two primers dimerizing. If they are long enough, they could appear as primer dimers. You can use any on line tool to ceck for primer dimers, you can check OligoCalc, for instance.
Good luck
  • asked a question related to PCR
Question
4 answers
Could DNA from relatives allow polymerase chain reaction (PCR)s to replicate offspring for the deceased? How exactly?
Research Proposal Honor Kirk Aanes
Relevant answer
Answer
The genome is huge and recombination takes place almost randomly and also pcr can only amplify tiny lengths of dna. Also there are vast numbers of viral sequences scattered around the genome so as only half of the parental dna goes to the child and a quarter to the next generation and one eigth to the next generation etc I am sure that getting an exact unknown sequence by inference will be impossible
  • asked a question related to PCR
Question
1 answer
,?
Relevant answer
Answer
rRNA-based studies, to assess microbial communities, rely on the accurate amplification of the corresponding genes from the original DNA sample. Here we present an analysis and re-evaluation of commonly used primers for amplifying DNA between positions 27 and 1492 of bacterial 16S rRNA genes (numbered according to the rRNA of Escherichia coli). We propose a forward primer formula (27f) that includes three sequences that are not commonly found. We compare our proposed formula with two common alternatives using linear amplification—providing a reverse primer-independent assessment—and in combination with the reverse primer 1492 (1492r) under appropriate PCR conditions for generating community rRNA gene clone libraries. For analyses of DNA from human vaginal samples,
  • asked a question related to PCR
Question
1 answer
Since we work mostly with genetic models, we usually genotype our mice after ear punching them for further breeding purposes.
Recently our breeding has taken up qutie a bit resulting in a high number of samples and quite a bit of time "wasted" on something that doesn´t lead to "valuable" results but is, nonetheless, essential for following experiments and breeding.
Does any of you know of a way to facilitate and speed-up genotyping? Someting like a machine that performs the entire workflow?
Our current lysis-PCR for 4 different genes-gel loading-electrophoresis and imaging as well as annotation takes well over half a day to a day for 100 samples and is simply no longer efficient to perform.
Thank you in advance!
Relevant answer
Answer
To enhance the efficiency of your genotyping workflow, consider adopting automated technologies that streamline various stages of the process. Automated DNA extractors, such as the Qiagen BioSprint and KingFisher Flex, can significantly reduce manual handling by automating DNA extraction and can be seamlessly integrated with automated PCR setup systems. For a more comprehensive solution, the Fluidigm Biomark HD System offers an integrated approach for high-throughput PCR amplification and analysis, minimizing the need for multiple instruments. Additionally, capillary electrophoresis systems like the Applied Biosystems SeqStudio and the QIAxcel Advanced System automate the electrophoresis and imaging steps, drastically cutting down processing time. For even greater efficiency, consider utilizing Digital Droplet PCR technology, which provides precise, absolute quantification and rare allele detection, thereby eliminating the need for post-PCR electrophoresis. Adopting these advanced technologies could significantly expedite your genotyping operations, allowing for quicker and more reliable results.
  • asked a question related to PCR
Question
3 answers
Details of the experiment:
We aim to clone 3 gene segments into a vector.
Details of the gene segments:
  • The first segment has a concentration of 34 ng/µL and is 1445 base pairs long.
  • The second segment has a concentration of 30 ng/µL and is 125 base pairs long.
  • The third segment, is 4758 base pairs long, with a concentration of 150 ng/µL, and is a PCR product.
These segments were designed to have an overlap of approximately 30 nucleotides with each other.
The vector has a concentration of 29 ng/µL and is 1850 base pairs long.
I have repeated this reaction using various volumes, but sometimes I did not observe any colonies, and other times, when I sent the colonies for Sanger sequencing, I found that one fragment had been inserted while the other two were not, resulting in different combinations.
In the last reaction, I used the volumes recommended by the NEBuilder site:
  • Vector: 4.5 µL
  • First fragment: 3 µL
  • Second fragment: 0.3 µL
  • Third fragment: 2.2 µL
  • NEBuilder HiFi DNA Assembly Master Mix: 10 µL
  • Each are = 0.114 pmoles
I incubated the reaction at 50°C for 5 hours, then performed a chemical transformation into E. coli Stbl3 strain.
I only observed 2 colonies, but after performing enzyme treatment and sending sanger sequencing, I saw that the desired segments were not inserted.
Could you help me understand what the problem might be?
Relevant answer
Answer
You will get much lower cloning efficiency if the overlaps are 30 nucleotides. You should aim for an overlap of 15-20 nucleotides; the shorter the overlap the better, as long as it has a melting temperature >48°C.
I've never done Gibson assembly where one of the inserts is ~3 times bigger than the vector, but it should still work. I use 50 ng of vector, and then use 3:1 insert:vector molar ratio. You could try these amounts instead. For your small insert, you might want to use 5:1; this is recommended by NEB.
Make sure you have checked all of your fragments and your digested vector on a gel. Make sure all your DNA is good quality. You can check the fidelity of your Gibson assembly mix using the positive control that comes with the kit. If all of these checks look good, then the problem is the design of your overlaps (in my experience, overlap design is the cause of poor Gibson assemblies >80% of the time).
  • asked a question related to PCR
Question
4 answers
I am a new person, want to understand the process of learning PCR. It is best to be simple and easy to understand with pictures. Among them, I want to first understand the process of standard PCR.
Relevant answer
Answer
Go to YouTube and search for videos introducing the polymerase chain reaction and/or search the internet for a A beginner’s guide to PCR.
  • asked a question related to PCR
Question
9 answers
I use gradient PCR for single target with two different Primers set. both the set failed to get the band. 1st Set Tm is 62.9 and 2nd Set Tm is 63.2. anneling Temp. is as follows 55,57,59,62C 1st set give 76bp amplicon and 2nd set will give 147 bp amplicon
Relevant answer
Answer
I would suggest using this tool to determine the Tm of your primers
And try to use less template 5µg RNA in 20µl RT is at the upper limit; and using 2µl of this cDNA in a 10µl RT-PCR reaction is a lot. Dilute an aliquot of your cDNA 1:5 with water and then use 2-5µl diluted cDNA in your RT-PCR.
If the smear in your reactions still persists and the sequence you are trying to amplify is GC-rich, you may add 2-5% DMSO.
Good luck.
  • asked a question related to PCR
Question
7 answers
Kindly answer this
Relevant answer
Answer
The starting dna is very long and has a very high annealing and denaturation temperature so in the first 2 cycles it is essential to get complete denaturation . After thie first 2 cycles the amplimer is very short compared to genomic dna so melts much easier and reanneals at a much lower temperature so a shorter denaturation is ok. If you are wondering why the amplimer does not reanneal and stop the reaction the higher primer concentration and the law of mass action ensures that primer anneals to most of the amplimer before it has time to reanneal with its opposite strand
  • asked a question related to PCR
Question
7 answers
I have treated the cells with ligands with different doses and performed PCR to determine cell expression. Now I need to decide which dose I should include for my studies using EC50. Can anyone guide me, on how this should be done?
Relevant answer
Answer
To determine the optimal dose for your studies using the EC50 value, follow these steps: 1. **Data Collection**: Ensure you have collected data on cell expression levels for each dose of the ligand you tested. This typically involves measuring the expression of a target gene or protein after treatment with various concentrations of the ligand. 2. **Data Normalization**: Normalize your data to a control sample (e.g., untreated cells) to account for any variability in PCR efficiency or cell number. This will allow you to express your results as a percentage of the control. 3. **Dose-Response Curve**: Plot a dose-response curve with the ligand concentration on the x-axis (log scale) and the normalized expression levels on the y-axis. This curve will typically have a sigmoidal shape. 4. **Curve Fitting**: Fit your data to a sigmoidal dose-response (variable slope) curve using a statistical software package or graphing tool like GraphPad Prism, Excel, or R. This will generate an equation that describes the relationship between ligand concentration and response. 5. **Calculate EC50**: The EC50 is the concentration of the ligand that induces a response halfway between the baseline and maximum response. Your curve-fitting tool should provide this value directly. It represents the potency of the ligand. 6. **Determine Optimal Dose**: Based on the EC50 value, you can decide on the optimal dose for your studies. For most applications, a dose that is close to the EC50 is often chosen because it represents a balance between efficacy and potential toxicity or side effects. However, the specific dose may depend on the goals of your study. For example, if you are studying the maximum effect of the ligand, you might choose a dose higher than the EC50. Conversely, if you are interested in a more subtle effect or want to minimize off-target effects, you might choose a dose lower than the EC50. 7. **Confirm with Additional Experiments**: Once you have identified a potential optimal dose, it is advisable to confirm this dose in additional experiments to ensure reproducibility and robustness of the effect. Remember that the EC50 is a guideline, and the choice of dose may also be influenced by other factors such as the specific biological question you are addressing, the availability of the ligand, and practical considerations such as cost and ease of handling.
  • asked a question related to PCR
Question
5 answers
Where I work, there are three separate rooms for setting up PCR reactions. In room number 2, whener we use UV light for decontamination, there is a particular odor that remains for minutes after opening the door. I have been reading about this, and, apparently, it is ozone. Has anyone here experienced the same? Do you have any thoughts on why does this happen only in one of the PCR rooms? Is it harmful to be in the room smelling this? How can I overcome this issue? Thank you very much!!
Relevant answer
Answer
UV light @ bactericidal wavelength - 254 nm - does not generate ozone - rather it breaks down ozone. If your "UV" lamp is the source, assume UV-C, it's emitting at lesser wavelengths and may not be that effective vs. microbial contamination.
I
  • asked a question related to PCR
Question
5 answers
I am currently optimizing a RPA-based protocol. As everyone knows, this nucleic acid amplification technique is based on isothermal amplification (around 37-42˚C) in combination with recombinases and single-stranded DNA binding (SSB) proteins.
I have designed several candidate primers to optimize my RPA. All of them were searched in literature and designed considering the criteria to be used in an RPA reaction.
These same primers were verified by conventional PCR (At different Tm: 55-65ºC), obtaining a successful result.
The problem started when I used the RPA master mix (lyo version from Twistdx) and performed the RPA reaction with the same primers and samples used in the conventional PCR. ALL THE RESULTS BECAME NEGATIVE?!
Primer concentrations used in RPA reaction was 400nm (recommended by twistdx) and the reaction time was 30 minutes at 39 °C (conditions recommended for the set of primers tested). Visualization of the amplification was done on agarose gel (1.5%) and doing a posterior melting curve assay. In all cases, no amplification was detected.
Does any one have a clue on what is happening???
I don´t know which other variables I can change to obtain good results
Suggestions?
Thank you
Relevant answer
Answer
We manage to make it work with another kit! The problem emerges because we bought the exo kit by mistake which amplifies and degrades at the same time to produce signal with the prove. Thank you all for the replies. I am no longer working on that projeto neither on the company developing the method
  • asked a question related to PCR
Question
1 answer
I am trying to run a PCR to verify insertion of my construct into the AAVS1 locus in iPSC using CRISPR.
I designed three primer pairs to amplify the left and right insertion regions (one binding inside the insert, one binding outside; see image), and one primer pair to amplify a region inside the insert. The insert contains a fluorescent protein, which I can see expressed in the cells under the microscope, so I am pretty sure that the insertion has worked correctly; however, I cannot get any specific PCR product for sequencing. (Even if it has been inserted unspecifically somewhere in the genome, since I am seeing the fluorescent reporter in the cells, I would expect at least to get a positive result for the "internal" region.)
I used DNAzol to purifiy gDNA from the cells, and when I checked it on a gel I noticed two additional bands, which I thought might be rRNA (see image), and when I treated the samples with RNAse the bands disappeared, so I continued happily with the PCR.
For the "internal" region, I am able to use the plasmid as a control, and here I can see a specific PCR product with the expected size, however, for all other plasmid / gDNA template combinations, I get a huge amount of large-sized unspecific PCR products (see second image). Which is why I am currently suspecting that something is not right with the gDNA? But it looks pretty good on a gel.
I am using KOD1 polymerase (KOD1 master mix) according to manufacturer instructions when it comes to amplification from gDNA:
PCR: Total 25 µl per reaction
1.25 µl DMSO
1 µl primer fwd [10 µM]
1 µl primer rev [10 µM]
8.25 µl H2O
12.5 µl KOD master mix
0.5 µl DNA (= 25ng)
Init. Denat. 94°C 1.5 min
Denat. 94°C 5 sec
Anneal 58°C 5 sec
Extension 68°C 1 sec
and also tried
Init. Denat. 94°C 3 min
Denat. 94°C 45 sec
Anneal 58°C 45 sec
Extension 68°C 1 min
I have triple-checked the specificity of the primers, and compared with other primers used in literature for the same purpose (AAVS1 locus). I have re-designed new primers that bind in slighly different places. I have tried different elongation / annealing times and temperatures... It always looks the same (large-size unspecific products).
I a last-ditch effort, I cut out pieces of gel from the "unspecific" results around the size where I would expect the PCR product, and repeated the PCR with those as a template - I got some promising looking results on a gel, but when I sent them for sequencing, it was all unspecific.
I am currently at my wit's end and hope someone else has seen something similar and was able to solve it in the end!!
(Plan B will be to re-do the DNA extraction and try again from the beginning I guess...)
Relevant answer
Answer
Dear Anja,
If I understand your protocol in Image 2 (PCRproduct.png) correct. You are using 25 ng gDNA in your PCR. That is way to low. You should think about copy numbers. Of cause 25 ng plasmid has millions of of copys in there, while in one genome your AAVS1 knock in might exists only once. Please try to use 300-500 ng for your gDNA along with your low plasmid DNA concentration.
And than you should optimize your PCR condition subsequently. You should not really see any products in 1 and 2 both lane 1-3.
And after doing comparable stuff with TALEN in Hek293 cells. If your are planing to use single cell clones. You can use both outside primers to detect if your cell is homo or heterozygous.
Best wishes
Soenke
  • asked a question related to PCR
Question
2 answers
PCR is a technique used to amplify specific DNA sequences, and the quality of the DNA template plays a significant role in the efficiency and accuracy of this process.
How DNA purity impacts PCR and what can be done to improve it?
Relevant answer
Answer
hunins and rna can look like dna if poorly purified away and lead to lower pcr yield due to low amount of statring material. Protein can interfere with magnesium concentartion as well as enzyme binding leading to failed or low pcr yield.
Many recombinant modern polymerases can amplify in the presence of pcr inhibitors or the purity of the dna can be improved using proteinase K digestion and phenol chloroform purification. Column purification of the dna will help dna purity and in exceptional cases PhiX enzymes and whole genome amplification folloed by pcr can work as phi is much less fussy about amplification than normal polymerases
  • asked a question related to PCR
Question
1 answer
Dinoflagellates often incorporate 5-hmU into their genome; does this modification cause issues with PCR, such that we'd need to use uracil-incorporating polymerases, or is a normal high fidelity polymerase sufficient?
Relevant answer
Answer
I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?
5-Hydroxymethyl uracil (5-hmU) can indeed cause problems with PCR because it can interfere with DNA polymerase binding efficiency and fidelity. Dinoflagellates, which often incorporate 5-hmU into their genome, may face amplification challenges, leading to potential misincorporations or incomplete extensions [1][2]. While normal high fidelity polymerases may struggle with this modification, uracil-incorporating polymerases are specifically designed to handle uracil and its derivatives. These specialized polymerases would likely improve the accuracy and efficiency of PCR reactions involving templates with 5-hmU, ensuring more reliable amplification [3][4].
Reference
[1] Rae, P. (1976). Hydroxymethyluracil in eukaryote DNA: a natural feature of the pyrrophyta (dinoflagellates).. Science, 194 4269, 1062-4 .
[2] Verma, A., Barua, A., Ruvindy, R., Savela, H., Ajani, P., & Murray, S. (2019). The Genetic Basis of Toxin Biosynthesis in Dinoflagellates. Microorganisms, 7.
[3] Orr, R. J. S., Stüken, A., Murray, S., & Jakobsen, K. (2013). Evolution and Distribution of Saxitoxin Biosynthesis in Dinoflagellates. Marine Drugs, 11, 2814 - 2828.
[4] Hudson, D. A., & Adlard, R. (1995). PCR techniques applied to Hematodinium spp. and Hematodinium-like dinoflagellates in decapod crustaceans. Diseases of Aquatic Organisms, 20, 203-206.
  • asked a question related to PCR
Question
3 answers
The bands of my PCR amplicons are mostly wonky, and I can't figure out why. When I did the non-touchdown version, the bands were straight, but results were stochastic (i.e. some samples amplified, some didn't). The chromatogram of the sequences also showed a lot of multiple peaks per position. I shifted to doing touchdown to lessen the chances of non-specific amplification. Touchdown seems to work well in terms of amplification success, but the amplicons are wonky on the gel.
I would appreciate any and all insights and advice on how this happens and how to correct it.
Below are my PCR parameters.
Master mix components (for 20ul reaction volume):
2.0ul 10x PCRx enhancer
2.0ul 10x PCRx buffer
0.6ul 50mM MgSO4
0.4ul 10uM dNTPs
1.0ul 10uM H3F/R primer
0.5ul 5U/ul Taq polymerase
1.0ul template (1:1000)
12.5ul ddH2O
PCR cycling parameters (touchdown): 95C/3min; 11 cycles of: 95C/30s, 65C/30s (-1C/cycle), 72C/1min; 25 cycles of: 95C/30s, 55C/30s, 72C/1min, 72C/15min
Gel electrophoresis: 1.0% agarose in 1x TBE, 1ul amplicon, 1ul 6x GLB + SYBR gold, 2ul Hyperladder 1kb, 120V for 20mins
Relevant answer
Answer
I would clean the comb, allow the gel to cool for much longer before removing the comb from the gel and be particularly careful that removing the comb does not break the bottom of the gel as some of the samples look like there is sample leakage
  • asked a question related to PCR
Question
1 answer
Hello Dear,
Im PhD student
I was using Gibsson assembly, and I cloned my gene onto the Psl18 plasmid . The plasmid was right when I sent it to Sequencing after geting a positives colonies.
However, when attempting to perform PCR and amplify my gene using my glyceral bacterial stock, I obtain a bande that is appropriately sized, but another bande that is nearly the same size as my gene (see gel picture below).
I don't understand why, as I should only have one bande! Can my plasmid exist in two different forms in the Glycrio Stokc 9 (one that is correct and the other that is not) or is there another explanation?
Or should I do the cloning all over again ?
It`s shloud be a primers issues ! (Althought I checked it in snapgene but it`s bande just in my gene of interest)
Also, I test 2 taq (polymerase and Q5 HF) but I got the same results.
I appreciate your help in advance.
Thanks
Ayoub
Relevant answer
Answer
Looks like the issue is the annealing temperature for your primers, based on the non-specific bands lower down. Try setting up a gradient of annealing temperatures.
Good luck!
  • asked a question related to PCR
Question
8 answers
Some context first:
on all my experiment I used a control sample labeled with PET. I have very little amount of this control sample so i used both primers for PCR PET-labeled. I thought that with both primers being labeled  i could use less PCR product and make it last more.
In every FLA i mixed together this control sample with other samples labeled with VIC, FAM, NED so i could compare these three samples against my control. Then i noticed that my control sample has 2 populations. Like if my primers were amplifying two different targets. But we are very sure that this is not the case.
In previous experiments I have used the same primer sequences. but this is the first time I use it with both primers (forward and reverse) labeled. So this issue only appeared when using both primers labeled.
In the last validation experiment i did the following test:
  • Amplify using forward pet-labeled primer
  • Amplify using reverse pet- labeled primer
  • Amplify using forward and reverse pet-labeled primers
  • Amplify using forward vic-labeled primer
  • Amplify using forward vic-labeled primer and reverse pet-labeled primer
so you can observe that when ever i used a reverse labeled primer the two populations appears.
technical details :
the target region contains tandem repeats, therefore the template has strong secondary structure. GC% is over 65%, PCR reaction was done with taq polymerase with 5% DMSO.
Relevant answer
Answer
Raul this is such an excellent rationale. Well done for finding the right people to help with this problem. The result shows that the product is running entirely single stranded....I should have had more faith that denaturation was complete and would not have thought of this solution at all since I was assuming that the dna was double stranded. Well done
paul
  • asked a question related to PCR
Question
3 answers
I have a PCR product that I've been using as a component that I'm ligating into a vector. Did sequencing it was confirmed to be correct and did a ligation it worked perfect and was sequence verified. I did two more ligations using the same PCR product into different vectors and now there is a mutation in the PCR product? How does this happen? I've essentially sequenced it twice(one time just the PCR product and then again to make sure the it was ligated correctly) and it was fine and now the sequencing is showing a deletion. I send to two different companies for sequencing and they have both come back with the same results each time. The first 2 times both showed it to be correct the last 2 times there is a mutation that both companies are picking up on.
Relevant answer
Answer
Sounds like you maybe had a mixed PCR product pool, some were fine and others had the mutation. You could try sub-cloning your PCR product (check the sequence in the sub-cloning vector) then using cut and paste to move it from there to your final vector. It does take time but can reduce the odds of a mutation in your final clone.
  • asked a question related to PCR
Question
10 answers
When we perform statistical analysis on qPCR data, do we use fold change or ΔCt?
Relevant answer
Answer
you mean -ΔΔCt?you could use either -ΔΔCt or the fold change ( 2 to the power of -ΔΔCt).
  • asked a question related to PCR
Question
3 answers
These two words are similar in the meaning of tech.or there is some different between them
Relevant answer
Answer
I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?
In the context of PCR technology, "elongation" and "extension" are indeed often used interchangeably to describe the same process, where the DNA polymerase synthesizes a new strand complementary to the template strand. During this phase, the polymerase adds nucleotides to the 3' end of the primer, thus extending the DNA strand. However, the term "elongation" is sometimes more specifically associated with the kinetics of the process, such as in measuring the elongation efficiency in various conditions [1][5][6]. On the other hand, "extension" is more commonly used in standard PCR protocols and guidelines [2][3][4]. Thus, while both terms refer to the same phase in PCR, "elongation" might be used in a more technical context to discuss the efficiency and dynamics of the polymerase action.
Reference
[1] Horton, R., Cai, Z., Ho, S., & Pease, L. (2013). Gene splicing by overlap extension: tailor-made genes using the polymerase chain reaction.. BioTechniques, 8 5, 528-35 .
[2] Hussain, H., & Chong, N. F. M. (2016). Combined Overlap Extension PCR Method for Improved Site Directed Mutagenesis. BioMed Research International, 2016.
[3] Montgomery, J., & Wittwer, C. (2014). Influence of PCR reagents on DNA polymerase extension rates measured on real-time PCR instruments.. Clinical chemistry, 60 2, 334-40 .
[4] Huang, M., Arnheim, N., & Goodman, M. F. (1992). Extension of base mispairs by Taq DNA polymerase: implications for single nucleotide discrimination in PCR.. Nucleic acids research, 20 17, 4567-73 .
[5] Yamagami, T., Ishino, S., Kawarabayasi, Y., & Ishino, Y. (2014). Mutant Taq DNA polymerases with improved elongation ability as a useful reagent for genetic engineering. Frontiers in Microbiology, 5.
[6] Cheng, B., & Price, D. (2007). Properties of RNA Polymerase II Elongation Complexes Before and After the P-TEFb-mediated Transition into Productive Elongation*. Journal of Biological Chemistry, 282, 21901 - 21912.
  • asked a question related to PCR
Question
3 answers
I have been working on a knockout in P. aeruginosa, however, after the final PCR check, I see both wild-type and mutant bands. Any suggestions?
Relevant answer
Answer
What do you see in your controls? Assuming those worked as expected, it sounds like you have a mixed population of cells.
Streak out to single colonies (again) and check a few of those.
Good luck!
  • asked a question related to PCR
Question
2 answers
Hello, I am currently learning to generate Agrobacterium deletional mutants by homologous recombination. I already prepared the construct containing the up-and-down- fragments of the gene from Agrobacterium in a suicide plasmid (SacB, GmR) in E. coli S17. To obtain the deletional mutant I did bacterial conjugation between Agrobacterium and S17. The result of the first crossover seems fine since the Agrobacterium colonies I obtained contain the up and down fragments after I did the PCR check. However, in the second crossover, I thought at least I would see some colonies going back to wild type, but no bands are shown in all colonies I picked following the PCR. This experiment was actually already done by someone else in my lab, and she obtained the mutant. I used her mutant and the wild type of Agrobacterium as control. The PCR result of the control seems fine. So what is the possible reason for this failure, and is there any suggestion?
Relevant answer
Answer
Hello Dr. Benedik, first of all thank you for answering my question. I actually stopped working on this bacterial strain for quite sometime, and just got back again by trying knocking out different gene with the same method. I already tried your suggestion to use few independent "first recombinant", and I still have the same problem (no PCR band). So, the gene I tried to delete is located in the tumor inducing plasmid (pTi), and apparently the colonies seem to throw away the entire pTi (I check another gene on the pTi and the PCR cannot detect it).
  • asked a question related to PCR
Question
3 answers
I'm now operating an experiment of MT genes amplification and sequencing. When amplifying the mt large subunit ribosonal RNA gene (16S), the agarose gel electrophoresis result showed me that there existed a smear bigger than the aimed band. Everytime when I was sequencing this gene I may encounter the same problem. So who can tell me what's the smear's component, and why it was mistakenly amplified?
(I use 16SAR/16SBR as the PCR Primers)
Relevant answer
Answer
I think that the weaker, upper band is not a mistake but indicates that your amplified sequence has at least 2 sequences (snps or small indels).
Your amplification is very strong and if we have 2 very similar sequences then at the last cycle of pcr a lot of amplified product will anneal to other product molecules. If we call these amplimers N and M where sequence M differs from sequence N by one base then when the 2 sequences combine we get NN and MM and some MN. In an agarose gel NN and MM are the same size so run as one band but NM has a mismatched base forming a bubble in the dna and this changes the shape of the dna so it runs as larger than N N and MM but the heteroduplex band just runs at a slower rate through the gel so is interpreted as a larger size band. To test this idea you can cut out the smaller stronger band and denature it,reload it and you should once again generate 2 bands just like the picture you posted. Sequencing either band should also show that you have 2 sequences correctly showing in both bands
  • asked a question related to PCR
Question
1 answer
My goal for this experiment is to knockout a gene of interest in budding yeast (Saccharomyces Cerevisiae) and replace the whole gene with a G418 resistance cassette.
As a broad overview, PCR was used to amplify the G418 resistance cassette from a strain from a deletion collection which was then used to transform into a WT strain. After transformation, the transformants were plated onto YPAD for recovery then selected for using a G418 media plate.
After going through the steps provided in the protocol (attached below), colonies were able to grow on the 2xG418 plates suggesting potential candidates that were transformed to have the G418 resistance cassette. However, after molecular genotyping using PCR, it was found that the cassette was not located relative to the desired gene loci.
We expect the issue is related to the transformation efficiency and was planning on increasing the heat shock time from 7 minutes to 15 minutes and also improve the homology of the PCR product by using a mixture of Taq/Pfu rather than Taq alone.
Any suggestions with troubleshooting would be greatly appreciated!
Relevant answer
Answer
Hello,
A few suggestions regarding the trouble with your transformations:
I use LiAc-ssDNA-PEG method for non-integrative plasmid, integrative plasmid and transformation with PCR cassette (deletion and integration both). All kinds of transformations work with this method. It is a fast and efficient method. It works with YPD media as well. I will attach the protocol with this answer.
  • LiAc based method is more efficient than CaCl2 in case of yeast transformation.
  • Heat shock incubation time at 42 degree Celsius can be extended till 1 hour. I use 20 degree Celsius which is optimum for most of the transformations.
  • It is recommended to use high-fidelity DNA polymerase (such as Q5 or Pfu) than the regular Taq polymerase.
  • Run a BLAST to check the homology of the gene sequence used to prepare the PCR cassette. Partial homology can cause nonhomologous recombination and it is an usual case in the deletion of a gene.
Best regards.
  • asked a question related to PCR
Question
7 answers
Hello
I am doing multiplex PCR for SCCmec typing of MRSA, but I am not able to get a band which is 1791bp and it is the largest band while all other five bands are present as intended. I used DNA extracted through boiling for PCR and the primer and PCR conditions are well optimized used by many researchers. Any suggestion in this scenario please.
Relevant answer
Answer
Dear Paul Rutland once again thank you for your valuable comments, I am going to try with these tips and will see the results.
  • asked a question related to PCR
Question
1 answer
Hi everyone,
I'm trying to perform ATAC-seq without using any commercial kit, but it's been challenging to find a protocol that aligns with this approach. I’d like to ask if anyone has experience with this.
Here’s the workflow I followed:
1. Transposon Annealing:
ME A: 5'-(index sequence)AGATGTGTATAAGAGACAG-3' ME B: 5'-(other index sequence)AGATGTGTATAAGAGACAG-3' ME rev: 5'-pCTGTCTCTTATACACATCT-3' By following the standard primer annealing protocol, I obtained two transposon oligos (ME A-rev, ME B-rev).
2. Nuclei Extraction: I followed the Kaestner Lab Omni ATAC protocol (file attached) and confirmed the extraction by performing MNase digestion. This step seems to be working fine.
3. Transposition:
I've attached the protocol for the Tn5 transposase I used. Following this protocol and the Omni ATAC protocol, I set up the transposition reaction with the following conditions:1 µl 10X Tn5 Transposase Buffer (final conc. 1X) 1 µl Annealed Transposon A (final conc. 1 µM) 1 µl Annealed Transposon B (final conc. 1 µM) 0.1 µl 10% Tween-20 0.1 µl 1% Digitonin 1 µl Tn5 transposase Add D.W. to 10 µl I added this reaction mix to the extracted nuclei and incubated at 37°C for 2 hours.
4. DNA Purification: I purified the DNA using the QIAquick PCR purification kit.
After these steps, I performed PCR for amplification and confirmed the product with gel electrophoresis, looking for bands at 300, 450, and 600 bp (corresponding to mono-, di-, and trinucleosomes, with ~150 bp added for index and adapter sequences).
This method worked once, but I haven't been able to replicate the results since then. Instead of the expected bands, I could only observe smears or odd bands after PCR (figures attached). I would really appreciate any advice or insights you might have.
Sincerely, Hyelin
Relevant answer
Answer
Tm
  • asked a question related to PCR
Question
5 answers
Dear Researcher,
I have used nested PCR to increase the specificity of Tetra Arms. Previously, I obtained results for another SNP using the same reaction conditions, and separate reactions (monoplex) verified that all primers are working efficiently. However, when I applied nested PCR, I observed no amplification; instead, the well glows. I have tried using a commercially available good quality ready mix and different PCR components, but the issue persists. I also tried changing the amplification temperature. Can anyone tell me the reason for this problem?
picture of gel is attached, well 1-6 = products from nested PCR, 7 = PCR product that was used for nestedPCR, well 8= ladder
Note: I am working on deletion polymorphism
Relevant answer
Answer
using 0.1ng of a 306bp amplimer is to start the pcr with 3x10exp8 copies of your amplimer. 25 cycles could be an amplification of approx 3x10exp 15 molecules which is too much amplimer. Try starting with a 1/1000 dilution as
Steve Baeyen suggests and try running 15- 20 cycles of second round pcr
  • asked a question related to PCR
Question
4 answers
Dear all,
I'm learning and taking the same genotyping PCR experiments and want to get / share some advice or experience on obtaining the right bands for the data.
I ran 3 samples of Stra8-iCre and 2 out 3 samples were not wild type, which was different genotype compared to the actual data from our laboratory and all of them were supposed to be wild type.
There was no contamination nor mistakes on performance of the protocol and I also regulated annealing temperature into 58 or 60 degrees Celcius for this whole times.
I wonder how can I get the right bands for Stra8-iCre and what was I missing except for the accuracy of the protocol and regulation of the temperature.
Relevant answer
Answer
Hi Dayun Lee
choose an annealing emperature that does amplify and run a pcr set with 0. 1%, 2% up to 8%dmso and at some dmso concentration the pcr will probably be a clean band. If your gene has pseudogenes or is one of a family of similar genes then touchdown pcr or new primers might be needed.. You can perform primer gradient PCR to optimize the concentration of primers. Also, You can use pico molar concentration of primers. or use different gradient pcr such as with annealing and denaturation etc.
  • asked a question related to PCR
Question
2 answers
I'm cloning a fragment of 3200 nts into plasmid. The cloning was successful, however, 02 amino acids were mutated. Now I want to fix these 02 aa by site-directed mutagenesis technique using original DNA plasmid as template for PCR. The PCR product contains 02 kinds of DNA which are in the same length (original template and newly synthesized product). PCR product will be treated by DpnI to digest all the original DNA. The remain PCR product (my target DNA, linearized structure) will be purified and performed in-fusion to circularized into plasmid, then transformed to E. coli for propagation. Plasmid will be extracted from the E. coli and confirmed by NGS. I repeat some experiments, unfortunately, it seems original DNA was still partly remain after DpnI or the site-directed mutagenesis reaction was not successful (I think) making new plasmid still identical to the original template.
Now I want to check whether my target amino acid is fixed or not before sending sample to NGS optimizing cost benefit.
Relevant answer
Answer
I concur that you only need to do regular sequencing not NGS and this should much much less expensive. However if your clone picked up two mutations during the cloning, why not just look for another clone? That might be easier than doing two rounds of Site directed mutagenesis.
  • asked a question related to PCR
Question
1 answer
I am having issues with the plasmid pDONR-P4P1R. I first conducted a BP reaction with this plasmid and saw a lot of background. To troubleshoot this, I performed a PCR reaction with M13F and M13R which revealed that some of the plasmids in my stock were missing the region between the att sites which contains the ccdB. I have tried replacing my lab's chloramphenicol, making new stocks, and making antibiotic plates at different concentrations of chloramphenicol but it is not resolving the issue. I have attached an image of my gel from the PCR for reference. Does anyone have any suggestions on how to resolve this? thank you!
Relevant answer
Answer
I am not sure exactly what the problem is, but if your vector has a mixture of plasmids some of which lack ccdB gene, then the recombination experiment is probably not going to work.
I would retransform your vector (at low DNA concentration) into fresh cells but be sure you are using a strain that is resistant to ccdB to propagate (such as an F' strain).
  • asked a question related to PCR
Question
1 answer
I run PCR to checked presence of specific gene in shigella but I received 2 bands, one band showed product size expected and another one showed higher product size. I would like to check the my primer pair may have multiple binding sites or not. I did BLAST primer and got the result with 1 primer pair. How do I know that my primer pair has multiple binding sites through this result? Any other websites can check whether the exact position that my primers bind to the genome.
Your answers are really appreciated.
Relevant answer
Answer
Can you share a picture of your gel? How bright is the unexpectedly large band? Does it show up in your negative control?
One easy first thing to try is use a gradient of annealing temperatures. That might be enough to get your PCR condition specific to your desired product.
Good luck!
  • asked a question related to PCR
Question
5 answers
I am doing RAPD analysis to test for clonality in wild populations of three different plant species (Trientalis europaea, Cornus suecica and Rubus saxatilis). While I get quite good results for one of the species (Trientalis europaea), amplification fails for the other two.
I extracted DNA with the Qiagen DNeasy Plant Mini Kit and I tested 60 decamer operon primers of three different series (A, N and AF). I already optimised my PCR protocol in many ways (annealing temperature/number of cycles/PCR program, type of polymerase, amount of Mg, amount of DNA etc.), but without any success. Besides, since I had successful amplification for at least one of my species, the problem seems to be species-specific rather than protocol related.
Attached is a gel photo of my latest run which shows failed amplification for one of my focal species while my positive controls show clear bands, which suggests the problem is not due to PCR conditions. What am I doing wrong or overlooking?
I would very much appreciate any help.
Relevant answer
Answer
Our CLIQS 1D Pro software is designed to perform this kind of analysis, you can get a free trial from here: https://totallab.com/products/cliqs-1d-pro/
Best wishes,
Steven
  • asked a question related to PCR
Question
1 answer
I have done an extraction for DNA from amniotic fluid sample using Thermo Kit but when i measured DNA on the Nano drop I realized that it is contaminated sample as the ratio 260/280 is 0.7..I tried to make PCR with concentration 100ng in total volume 15 microlitre but no bands appeared. could you please tell me how can I increase the purity of sample and how to overcome this problem to get a successful PCR?
Thanks
Relevant answer
Answer
Does the kit use phenol? If yes, any carry-over will decrease the 260/280 ratio.
A high amount of proteins in your sample can also cause a low ratio.
100ng of DNA in only 15 microliters is a LOT of DNA. I know it sounds counter-intuitive, but try diluting your DNA (1:10, 1:100, 1:1000). Try all 3 dilutions + controls.
Dilution can lower the amount of PCR-inhibiting proteins, but the polymerase is sensitive enough to still amplify the DNA.
Good luck!
  • asked a question related to PCR
Question
6 answers
Hello everyone,
I performed a PCR yesterday, and the results showed no bands on the gel. Of course, I probably missed some crucial steps, like adding my samples to the PCR strips themselves, for instance. However, I seem to remember not missing any steps in my protocol. One thing that I didn't do, however, was vortex my primer + master mix solution together after adding them to a 1.5 mL tube. I doubt this would cause me to have no bands, but would this really affect my results? Thank you!!
Relevant answer
Answer
Even in the absence of template, you should be able to see the primers. In the presence of amplification you see some amount of unused primers as the concentration of primers is high (1X of the total pcr mixture) compared to the template which is generally very low.
in the absence of amplification, primers are not used at all so in that can also you should be able to see the primers.
So, according to me try to dissolve fresh primers and try pcr again. if it doesn’t work maybe try increasing the concentration of primers and try pcr again and if this also fails than the primers are problematic.
see in the first well, no amplification occured but template was added. You can see quite alot of primers. In the second well amplification occurred, still alot of unused primers are there. So primers have some issue.
  • asked a question related to PCR
Question
2 answers
Hello all,
I have been trying to follow a 2-stage PCR protocol used to amplify barcodes of a large yeast library, as per Nyugen et al. (2022) - https://link.springer.com/protocol/10.1007/978-1-0716-2257-5_22#Sec7 (section 3.3)
I have been working on this for the past 1.5 months now and have tried exploring all the variables from bead ratios to buffers to primers to template concentration. Although the 2-stage PCR seems to work every time now, the resulting yield after 2 cleanup steps is terribly low and cannot be sent for bulk sequencing.
Me and my advisor - both suspect cleanup steps as there wasn't anything wrong with the PCR itself. There are 2 bead cleanup steps after every stage of PCR:
1st cleanup - So the primers don't carry over to the 2nd stage of PCR.
2nd cleanup - remove excess primers and other contaminants.
The image attached would give some context to what I mean by low yield.
Relevant answer
Answer
Firstly, you need to optimize bead-to-sample ratios and elution conditions. Basically, a bead-to-sample ratio of 0.8-1.0x is recommended to ensure efficient binding of the desired amplicon size range while minimizing the carryover of smaller fragments and primers. Amplicon yield concentration can be maximised by using a lower volume of a buffer or water and ensuring thorough mixing. furthermore, control of incubation time to ensure complete drying of the beads before elution can enhance recovery. Systematic and consistent pipetting is also recommended.
  • asked a question related to PCR
Question
1 answer
I am preparing to run a lot of samples on a 384 well plate and I wanted to test if any well in my PCR machine is dead so as not to lose any data. I would really appreciate help with how I can do that. Thank you.
Relevant answer
Answer
Is this a new machine (as in, you just bought it)? If yes, it should have an extensive user manual. And you get in-person training/tech support from the company.
The quick & cheap way to see if all wells can collect data is to create one big batch of "positive control" and distribute it to the entire plate.
I hope that helps!
  • asked a question related to PCR
Question
5 answers
What does ‘preventing carryover contamination’ mean?
If they are used, is pre-treatment with UDG necessary?
Thanks :)
Relevant answer
Answer
In the context of PCR and DNA amplification, 'carryover contamination' refers to the accidental transfer of previously amplified DNA (amplicons) into new PCR reactions. This contamination can lead to false positives, where the detected signal is due to these contaminant DNA molecules rather than the target DNA from the sample being tested.
  • asked a question related to PCR
Question
2 answers
Why does BLAST results give 100% similarity to two bacterial species and only identified to the genus level?
I did BLAST of the two cellulolytic species and one lignolytic species of bacteria after 16S amplification using PCR. However, my results indicate that the lignolytic organism is 100% similar to two different bacterial species. BLAST have only identified up to the genus level in my cellulolytic bacterial species. Can anyone assist to explain what has happened?
Relevant answer
Answer
Some things to keep in mind when interpreting BLAST results are listed in
The 16S annotations in INSDC are messy. You could try a BLAST against type strains only: https://www.ncbi.nlm.nih.gov/refseq/targetedloci/
But BLAST does local alignment and so may not always give you what you probably want: similarity statistics based on a full-length global alignment. Download the (type) sequences of interest and use MAFFT to align your sequence to them.
bacteria[ORGN] AND sequence from type[FILTER] AND 16S[TITL]
gives you only type-derived sequences. Replace "bacteria" with the species name you have in mind.
  • asked a question related to PCR
Question
5 answers
I am trying to use overlap extension PCR to combine 2 linear PCR fragments around 1kb each. I amplified both fragments with overhanging primers with a 20 bp overlap between the two fragments. When I do overlap extension PCR, I just get amplification of the individual PCR fragments. I am doing a PCR reaction for 15 cycles without the primers, and then adding the primers that flank either end of the combined product for another 15 cycles.
Does anyone have suggestions for troubleshooting? The overlap region between the two fragments has a TM of 54, and the primers have TMs of 74 and 78. For the overlap PCR reaction I tried an annealing temperature of 50 and 55, and for the extension reaction I have tried annealing temperatures from 55-70.
Relevant answer
Answer
I did it before. Add more templates to PCR reaction, two fragments will form full length as well. You can test by increasing concentration of both overlap fragments. Primer conc is similar to normal PCR condition.
  • asked a question related to PCR
Question
3 answers
Hello,
We found three packages of Illustra™ MicroSpin™ G-25 columns in the cabinet of an unused lab. They are very old but have never been opened. I have never used this kit before, and I couldn't fully understand what it is used for from my internet search. Is it just a simple DNA purification kit, or is it something more functional? I am interested in recombinant protein production. Can I purify my ligation product with this before cloning into bacteria, or can I purify my PCR product with this before ligation? In which scenarios is this kit indispensable?
Thank you.
Relevant answer
Answer
These are sephadex spin columns and work on the basis of size exclusion. So dna of more than 10 bases long will flow round the beads and elute early while small salts will have to flow through the pores of the beads and will elute much later. They can be used for dna purification from most smaller molecules and have been used to remove radioactive salts from end labelling of oligos with the labelled oligo eluting first off the column. The G25 is an indication of the size range that can be separated on these columns
  • asked a question related to PCR
Question
2 answers
We are in need of primers for a PCR protocol for chicken genome. We are looking at the COI barcode, 648-bp fragment of the 5' end of COI. Specifically Primers FalcFA, BirdR1, CO1_ExtF, CO1_ExtR.
Could anyone provide me with a source of where I can purchase these from?
Relevant answer
Answer
Hi Vanessa,
I think these are the primer sequences you need. Get them synthesized at a specialist supplier (see answer by Ilze Skujina).
FalcoFA TCAACAAACCACAAAGACATCGGCAC
BirdR1 ACGTGGGAGATAATTCCAAATCCTG
CO1-ExtF ACGCTTTAACACTCAGCCATCTTACC
CO1-ExtR AACCAGCATATGAGGGTTCGATTCCT
Regards,
Vince
  • asked a question related to PCR
Question
1 answer
I'm working on RT-qPCR optimization for gene expression. In melting curve analysis, the 5x primer negative control has a peak that resembles the samples peak but looks shorter. 10x and 2.5 have no curve, and when I applied the gel for verification, no product was seen. Why does 5x primer have this melting curve but no product in gel?
Relevant answer
Answer
Despite no visible gel product, the melting curve peak in the 5x primer negative control likely indicates non-specific amplification or primer-dimer formation, detectable by the sensitive melting curve analysis but not by the less sensitive gel electrophoresis. This could be due to incomplete PCR reactions, primer-dimer artifacts from high primer concentration, or fluorescent dye binding to small, non-specific products. To resolve this, re-evaluate primer design, optimize PCR conditions (including primer concentration and annealing temperature), use more sensitive detection methods, and include additional controls to distinguish specific amplification from artifacts.
Nucleic acid bands can fade or vanish due to diffusion or degradation when running an agarose gel for an extended time. To avoid this, it's crucial to monitor the progress of the gel electrophoresis and stop the run at the appropriate time.
  • asked a question related to PCR
Question
2 answers
hi every one
I am making vector construction (for fusion proteins) and in this moment I wanna to amplification of ADAM17 prodomain with PCR. to yet, I couldn't amplified the ADAM17 prodomain with RNA extraction of Human fibroblast and PBMC (from Blood).
can you please help me how i can chosen the best source of this sequence for extraction of RNA?
thanks with best regards.
Relevant answer
Answer
Check the Human Protein Atlas for expression in tissues and cell lines.
You will see that highest expression level is in placenta and lung. I have no idea if you have access to these tissues.
The Human protein atlas also gives information about expression level in cell lines. Maybe, you have access to leukemia cell lines such as K-562. Lower expresison level is in SH-SY5Y, but this is a relatively common neuroblastoma cell line.
If you don't have access to these tissues or cell lines, you can get the sequence by gene synthesis. The region you are interested in is only approx. 200 aa, a gene synthesis for 600 bp is relatively cheap (should be less than 100 $).
Good luck,
Sebastian
  • asked a question related to PCR
Question
2 answers
How to assign allele numbers and sequence type after amplifying housekeeping genes by PCR for genotyping of S. pneumoniae strains by MLST using bioedit ?
Relevant answer
Answer
after having the sequence for each, ......
Take this example:
let's say that you did mlst for your bacteria, on of genes you amplified is named: ace gene ... you sent for sequencing and and you received the raw forward and reverse sequences for this gene.  then you have to trim the sequence to obtain the latest edited sequence....thats means you need to align them with reference sequence .....
go to https://pubmlst.org/  ==> chose your microorganism, ===> download allele===>chose your gene====> copy your reference sequence  ==> use allele sequence as a leader to align with your forward and reverse sequences using any bioinformatics software (like bioedit). Then you will get your allele sequence. ....... to know what is the code, go to mlst.net===>locus query ==> single locus ==> past your sequence >>>>>submit ....then record the digit.
Do these steps for the rest six genes ..so that you'll have allelic profile of seven digits
for example: (1,4,3,3,1,7,21)  ...... in https://pubmlst.org/, go to  profile query ==> allelic profile ==> enter your digits ==> submit ==> then you will get your sequence type (ST).
then contact with https://pubmlst.org/ curator to assign the ST.
Note: sometimes the ST is not presented in the mlst database, then it might be novel ST which is common .......
enjoy
  • asked a question related to PCR
Question
11 answers
Dear Colleagues,
I have the following problem: I’m trying to amplify a cassette with a resistance gene (about 2 kb) from the yeast genome, so that I can then insert it into a plasmid. With Taq polymerase everything works. With proofread polymerase (Tersus) there is no PCR product. Everything works with the positive control (with ITS primers, the product is about 800 bp). I increased the elongation time to 4 minutes, increased the primary denaturation at 95 to 3 minutes. There is no the product. I would be very grateful for any advice.
Relevant answer
Answer
I was finally able to amplify the desired sequence using Tersus. Ibrahim was right, an increased annealing temperature is needed here. But not only that, I got a product using a combination of conditions (I tried high annealing temperatures and before): a lot of DNA template, few primers (<200 nM), 4.5 minutes of elongation (for 2 kb) and 65 degrees annealing. Moreover, if Taq from 53 to 65 degrees of annealing simply changes the product yield, but it is there, then Tersus has nothing at 60, and a clear band at 65 (under the above conditions). By the way, Pfu polymerase turned out to be much less capricious than Tersus - it gave the product both with a diluted DNA template and at 60 and 65 degrees of annealing.
Thank very much for the advices.
  • asked a question related to PCR
Question
1 answer
I get two different bands from BCMV after PCR
Relevant answer
Answer
The PCR reaction may need to be optimized. Additionally, contamination may be occurred. I suggest you have it (both two bands) sequenced and checked.