Science method
Polymerase Chain Reaction - Science method
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 Polymerase Chain Reaction
I am working on a paper for my bioinformatics class, where we are running an analysis of transcriptomics on A. thaliana col-0 about the difference in gene expression in the temperatures of 21 and 28 degrees celcius.
In order to draw conclusions from the analysis we are asked to design primers for the differently expressed genes and run a PCR.
I am attaching also the galaxy pipeline used for the purposes of the assignment.
These are the genes we are asked to compare on NCBI:
We prepared a PCR buffer and assessed it with approximately 12 primer pairs, all of which resulted in successful amplification. However, there was a specific gene region that failed to amplify. The primers work well and efficient with Ampliqon 2x red PCR buffer resulting in the aplification of our desired targets.
The picture shows the PCR products attained from both Ampliqon and our custom PCR buffer for two DNA samples (performed in duplicate).
Various additives including DMSO and PCR enhancers were used, yet none proved effective.
I welcome any suggestions that could assist in resolving this matter.
What component or agent can the TE buffer contribute to having an unsuccessful PCR result
Trouble 1: I ran a PCR reaction and found a low molecular weight band (100 bp) when doing electrophoresis. 1ng plasmid (OD260/280=1.84) was used as template. Final concentration of primers was 0.2uM. I thought that band might be primer dimer. So I ran primer solutions on a gel again without PCR, but i can still see the band. Everything was newly prepared. Length of each primer is 20bp.
Trouble 2: ddH2O was used as negative control but an obvious band (with my target size-1100bp) can be seen. So I ran water directly without PCR and there was nothing on the gel. I freshly prepared water and did PCR again and the 1100bp band appeared again.
How could the above happen? What should I do?
Trying to identify the isolate from the high saline area . morphologically isolate is pigmented. extracted the DNA and tried to amplify the 16S. but repeatedly unsuccessful.
1) DNA dilution is done
2) change of primer concentration is done.
3)different primer combination is done.
4)change of PCR cycle is done.
Looking forward to get an answer and successfully amplify the isolate microbe and identify it.
Thanks in Advance
Hi all,
Hoping for some troubleshooting/advice. Currently setting up a virus typing assay for routine use. I have the assay working with good sensitivity (regularly amplifies samples that are above 30 Ct in detection assays) using Superscript III/Platinum Taq One Step RT PCR (https://www.thermofisher.com/order/catalog/product/12574035) using the reaction in the in the photo and following conditions: RT 50 deg 30 min, 95 deg 15 min, 40 cycles of 95 deg 30 s, 50 deg 30 sec, 72 deg 30 sec, then 72 deg for 10 min. prior to starting T, i relax RNA by 65 deg for 5 min hen cool on ice (with reverse primer in mix), before adding enzyme. Primers are gene specific.
However, majority of our samples are converted to cDNA upon arrival to facility (using random primers), and so it would be easier to adapt this to a PCR assay. On a cDNA plate, I tried the reaction in picture 2 using the same cycling conditions as previously described but obliviously without the RNA relaxation or RT step (starting from denaturation). This was totally negative.
To troubleshoot, I ran a known positive using the OneStep assay as described above in duplicate, however, I stopped the reaction after RT and for the second replicate I took 5 ul and added to PCR as described in 2nd paragraph, whilst leaving the first replicate untouched, and continued the reaction to the PCR. In this case, the 1st replicate (One Step - untouched) worked perfectly whilst the sample split into a PCR didnt work at all (completely negative). This indicated an issue with the PCR itself since the RT must be working fine.
I tried to bring the reaction down to 25 ul with a 3 ul input to see if this would help however this was also negative.
Any ideas or help would be greatly appreciated.
Thanks!
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.
I need your help with optimazing my PCR reaction. My PCR product should be 850bp. Starters Tm is 65, they do not form dimers or secondary structures and they’re specific (Blast doesn’t show any unspecific product that may occur). I've tried gradient PCR with typical mix and with Hot start polymerase and every time I receive strong band that is 150bp. What can I do to get rid of this unspecific product and what can it be?
I have a plasmid. This plasmid has 2 promoters, the first one is T7P to express araC protein. And the second promoter is pBAD to express lacZ protein. Both promoters are on the same plasmid and in the same direction. What I want is to do a PCR reaction to make them express the 2 proteins in the opposite direction. How to do that?
Good day! I extracted bacterial DNA from (rotted) apple fruit samples and am planning to run a 2-step PCR. This result was from the 1st PCR. This was from non-diluted DNA samples of about (20 - 50 ng/ul), I used primers with overhang attached, and used DMSO for the reaction.
I have previously tried (not shown) to dilute these samples but the target DNA just keeps getting low and the 2nd band is not eliminated. I am at a lost now how to troubleshoot this problem. Will MgCl help? Any and all advise is highly appreciated. Thank you!
Is a 5` phosphate needed for homologous recombination?
I am a bit confused whether phosphorylated DNA is needed to achieve efficient homologous recombination in e coli. I believe there are methods that use ssDNA oligos or PCR products as inserts for recombineering so these would usually not have a 5` phosphate but there also seems to be some literature that makes it sound like it is beneficial.
Can anyone clarify this, please.
Thanks
Hi , I have run my PCR products on gel electrophoresis, but I have seen excessive smear , do you have any suggestions for that?
Recently, I have been attempting to use fluorescence PCR to detect UGT1A1*28. I utilized the primers and probe reported in the paper "Rapid Allelic Discrimination by TaqMan PCR for the Detection of the Gilbert's Syndrome Marker UGT1A1*28."
Primer forward
5′-AACATTAACTTGGTGTATCGATTGGT-3′
Primer reverse
5′-AGCAGGCCCAGGACAAGT-3′
Probe (TA)6
6-FAM-5′-TTGCCATATATATATATATAAGTAGGA-3′-MGB
Probe (TA)7
VIC-5′-TGCCATATATATATATATATAAGTAGGA-3′-MGB
Initially, capillary electrophoresis revealed that the amplification efficiency of the primers reported in the literature was low, so I redesigned a pair of amplification primers.
RT-UGT1A1-28-F2
AACTCCCTGCTACCTTTGTGG
RT-UGT1A1-28-R2
GCTCCTGCCAGAGGTTCG
When testing samples using the new amplification primers and the probe from the literature, I noticed an abnormal baseline phenomenon. Instead of the baseline being a straight line, it inclined upwards. I would like to ask for possible reasons for this phenomenon to optimize subsequent experiments.
I have been trying to get a good band for a while. I am using the Phire Tissue Direct PCR Master Mix (Thermo Fisher) and my DNA sample is from a mice tail.
I am using a TRPM8 Primer (This is the 3rd primer that I am trying):
- Forward 5' to 3': GGT CAT GTT CAC GGC TCT CA
- Reverse 5' to 3': TTA GAT GCC CCA GTC CAC AC
I did the gradient test for the primers and I chose 64.4C. After setting that temperature I ran a new PCR (Fig.1), before preparing the PCR samples I always checked the amount of DNA, and for 20ul of reaction I used 2.5ul of my DNA sample solution. I got a band but it does not look big enough. So, I tried again (Fig. 2) and this time I used a gel at 2% and increased the annealing cycles to 45 (usually I set up at 40 cycles), for this DNA sample solution I used 2ul for 20ul of reaction.
What can I do? I am planning to use my PCR product for Sanger sequencing.
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.
Kindly help me out in this
A friend did PCR and then loaded it on agarose gel. All samples were prepared in the same way; the PCR was identical, with the same template, only primers differed, the same amount loaded on the gel... but 5 of them did something weird. It reminds me visually of the front of an SDS-PAGE elfo. Any idea, what could be wrong with this?
I have been preparing NGS Library, where the samples input volumes, conditions followed and the PCR Cycles are same but still the concentration obtained was uneven and the fragments size where also differ from sample to sample. What could be the possible reason for this uneven results.
Hi everybody,
I have tried to make my home-made master mix for our laboratory. I have used two type of dyes , cresol red and Bromophenol Blue(BPB). I see when I use BPB my PCR is inhibited but no inhibition is observed for cresol red.
Have anyone had the same experience? Do you think the pH of BPB need to be adjusted before use? when I add BPB to my colourless master mix in the proper concentraion it return to blue so I think pH readjustment of master mix buffer is not needed. How do you think ?
primers are to be used for RTPCR
Hello :)
I am looking at the presence of toxin biosythensis genes in lake sediments. Nothing fancy, since I only want to verify the specificity of PCR results for primers related to marker genes for cyanotoxins biosynthesis (e.g. mcyE for microcystins); I am using the Sanger results to confirm which organism in a given sample is responsible by searching for the results in BLAST.
I have so far done both forward and reverse Sanger, created a consensus sequence after trimming and then run the search. I can see the benefits of having the Sanger done both ways for some samples because one or two stopped abruptly, so then I could rely on the opposite read and a few bases had low quality scores.
But beyond quality control and upping my chances of getting a longer read, is there any benefit for doing it twice for a fairly simple "whose genes are these?"/PCR product verification approach?
Thanks :)
I know that to many of you this might seem like a really stupid question, but I am a beginner with PCR. If I have a DNA sequence (for example, a plasmid) into which I want to insert a small DNA fragment (about 25 bp) using PCR, how can I design the primers? Is it enough to design 2 primers (one forward and one reverse) so that both contain the DNA fragment (that I want to insert) at their 5' ends (see the picture)?
If the sequence of the primers are given as:
Forward: ATATAGACCAATATTCCTGTTAGCA,
Reverse: AAAAATTAGCCGGGCGTAGTGGCG
and size of the PCR product is 2500bp, then what will be the PCR profile for the given reaction
while using Phusion Flash High-Fidelity PCR Master Mix with 0.5uM primer concentration?
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?
I have been trying to get my PCR to work and amplify the correct region but can not figure out what is going on with it. I seemed to have nailed it down to something with the primers. One of the primers is fine, but the one that I need to use seems to hinder amplification of the amplicon and create a brighter band below the amplicon and slightly above where I expect primer dimers. I have tried adjusting template and primer concentration but it has not seemed to help too much. The results are attached below. I am having trouble with the primers labelled V1-V3.
The best online resource for conducting in silico PCR simulations of bacterial DNA.
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?
I have 6 bacterial isolates, I have checked their band on agarose gel electrophoresis, I am getting a genomic band but when I am performing the PCR and after running the gel, I am not getting good results. I have tried several times still the same results... I have checked the purity of all 6 gDNA it is between 1.6 - 2, the primers I am using are IDT universal primers (27F and 1492R) with 1 microgram final concentration. Isolation I did use Promega extraction kit and PCR using Promega PCR mix I don't know where I am going wrong. Kindly help. I have attached gDNA and PCR images for reference
I am doing my undergrad thesis that requires me to do PCR-RFLP analysis on DNA that I extracted from some blood samples. Initially, I was using the Qiagen mastermix for the optimization of my entire process using just 6 samples out of 100 samples. I successfully completed my optimization step, and got my desired results for those 6 samples. Once I have the measurements of the entire process i.e the amount of mastermix, forward priner, reverse primer, DNA template, and nuclease-free water along with the optimum annealing temperature for my selected primer, I proceeded on to doing the exact same thing on the remaining 96 samples serially. But I started encountering an unexpected error. For some reason, I started getting a faint band in my negative control(which only has nuclease-free water and no DNA template). At first I thought any of the reagents were contaminated, so I conducted a series of PCR reactions where in one I would use a freshly new mastermix, another time a fresh made working solution of primers and also an intact nuclease free water. In every PCR, I am getting a band in my negative control. Today I also tried using a mastermix of an entirely different company i.e. Takara Bio, but got the same results. I am just frustrated at this point. Can anyone help me figure out what is happening? How can I troubleshoot this and complete my thesis!
How can you regulate the number of random inserted mutations in error prone PCR?
Hello,
I am making some plasmid constructs via Inverse PCR and T4 enzyme ligation.
I started with a plasmid that housed nucleotide sequences for two fused proteins. My goal was to cut out the nucleotide sequence for just one of the proteins.
For my Inverse PCR primers, I designed the forward primer to be at the beginning of the desired protein. The primer was 35 nucleotide long
I designed the reverse primer to be 34 nucleotides long. It included the vector sequence just before the undesired protein.
After amplification with inverse PCR, I ran the sample on the gel and if the band was correct, I purified the DNA from the gel.
I then added I 2ul ligase buffer and 1ul T4 PNK and incubated it at 37C for 30 minutes. Then I let it sit at room temp for 5 minutes before I added 1 ul ligase.
Then I left it at room temperature for a few hours and before I stored it in 4C overnight.
I have done this process before and it has worked for me. However, I keep running into an issue where I continually have a one nucleotide deletion right at the site of ligation.
For example,
If I want this sequence: ATGCAC
I will get this sequence instead: ATCAC
That "G" is right at the junction. For some reason, it continually gets deleted during ligation.
I was wondering if anyone has any suggestions on how to prevent this from happening? Maybe there is something wrong with the procedure that I am using to ligate?
I have been trying to collect bacteria from human fecal sample to run on MT-PCR, however, for some reason, the concentration of human DNA is quite low and could not confirmed on PCR. My technique is making a solution of fecal sample with PBS in 1g per 10 ml, centrifuge at 500g for 3 minutes and collect supernatant, repeat three times and final centrifuge is 20000g at 4 Celsius degrees for 20 minutes. Beside it give very good concentration of bacteria, but I also need a good concentration of human DNA also. Should I lower centrifuge force or reduce time? Thank you for you help.
I am doing my undergrad thesis that requires me to do PCR-RFLP analysis on DNA that I extracted from some blood samples. Initially, I was using the Qiagen mastermix for the optimization of my entire process using just 6 samples out of 100 samples. I successfully completed my optimization step, and got my desired results for those 6 samples. Once I have the measurements of the entire process i.e the amount of mastermix, forward priner, reverse primer, DNA template, and nuclease-free water along with the optimum annealing temperature for my selected primer, I proceeded on to doing the exact same thing on the remaining 96 samples serially. But I started encountering an unexpected error. For some reason, I started getting a faint band in my negative control(which only has nuclease-free water and no DNA template). At first I thought any of the reagents were contaminated, so I conducted a series of PCR reactions where in one I would use a freshly new mastermix, another time a fresh made working solution of primers and also an intact nuclease free water. In every PCR, I am getting a band in my negative control. Today I also tried using a mastermix of an entirely different company i.e. Takara Bio, but got the same results. I am just frustrated at this point. Can anyone help me figure out what is happening? How can I troubleshoot this and complete my thesis!
The 260/280 ratio is ok between 1.8 and 2.0 however we were unable to advance due to the 260/230 ratio
The last couple of months I've been trying to replicate a small part of the DNA by using PCR. While everything was working great at first the PCR just stopped working one day. I haven't changed anything regarding the components nor the procedure and I really can't figure out what happened and why it's suddenly not working at all. Any help would be greatly appreciated.
Dear experts,
I am planning to generate a large numbers of Illumina libraries in an ongoing project. These will be amplified via a single PCR step, in which the indexed adapters are added to the PCR primers. Can I just order such primers as normal PCR primers including the indices or will their quality be insufficient for NGS application (regarding index hopping etc.)? The commercially available kits for indexing are very expensive.
I would be very grateful if anyone who did this already could share some experiences. Kind regards and thank you in advance, Matthias
PS:
In case more information is required, this is the PCR protocol which I am trying to run:
Hi!
I am to site-specifically label dsDNA parts using PCR with fluorescently labeled primers. For my objective it would be optimal if the primers could carry the fluorophore as close to their 3'-end as possible, preferably exactly at the 3'-end. I am worried however that perhaps this would prevent the polymerase's ability to elongate during PCR.
I have seen others label the primers as close as 1 position upstream of the 3'-end e.g. in:
Nazarenko I, Lowe B, Darfler M, Ikonomi P, Schuster D, Rashtchian A. Multiplex quantitative PCR using self-quenched primers labeled with a single fluorophore. Nucleic Acids Res. 2002 May 1;30(9):e37. doi: 10.1093/nar/30.9.e37. PMID: 11972352; PMCID: PMC113860.
Can DNA polymerase elongate if the 3'-end of the primer is modified with a fluorescent tag?
Thankful for any input!
All the best,
Niklas Eckert Elfving
Uppsala University
My fellow Academic colleagues!
I together with my lab mates have a PCR-related issues that we hope that some(one) of you might have encountered and hopefully solved.
In “short”, our initial PCR (MiniAmp Plus thermocycler) and electrophoresis protocol works like a charm – the latter somewhat modified. We obtain weak to strong band that yielding concentrations of 9 to 20 ng/µl following clean-up using the QIAGEN QIAquick PCR (& Gel) purification/Cleanup Kit (with an acceptable A260/A280 ratio). We obtain rarely, but from time to time, a positive electrophoresis confirmation. But as we are using the same protocol for the confirmation, as for our initial PCR, we should have no issue confirming our results (one band per week).
Usually, when we try to confirm our cut-out electrophoresis bands, running a PCR on our cDNA, something fails. We utilize the same primers and protocol, as for the initial PCR, but nothing shows up in our gel, our at best a streak. We’ve tried renewing our primer mix(s), new isopropanol, new buffers, using both RNAse-free water and the included buffer, modifying temperatures (thermocycler), number of cycles, and using the original non-modified protocol. But nothing results in an electrophoresis band when we try to confirm our initial band.
Thank you for your insights and help!
// Eriksson et al.
I want to do PCR amplification with my full-length gene and the addition of P2A fragment at the end of my gene. However after doing PCR, I run gel electrophoresis for analysis. But it doesn't include the band for my gene. I run the same template DNA with other primers for smaller fragment, and it has the band. I tried to redesign my primers for full-length fragment, but it still don't have the band for my gene. Can anyone help me? Thank you
Hello.
I have a problem with our in-house designed rt PCR for Cutibacterium acnes. Primers and probes were designed as part of mPCR. While testing each set of primers (monoplex) for LOD we are constantly getting false positive negative control. We repeated reaction many times. We change all reagents for new (to exclude contaminated reagents) and still the negative control in late positive in around 38 Ct. We tested negative control with 16S PCR and was negative. I was thinking to set a Cutoff/threshold? Does anybody has experiance with setting it? Thank you.
Anja
I use a Q5 polymerase to amplify a 7 kb fragment from a genomic DNA but get no results.
I use the stated protocol in NEB website. Any suggestions to modify the PCR protocol so I can get the amplification?
I have the PCR products of two bacterial genes (gyrA=1024 bp and rpoB=808 bp) and I want to know how many microliters I should add in the agarose gel?.
Which of this techniques can give us the best result in detection of chromosomal abnormalities
I am running a DNA PAGE after PCR (samples 6-15 are run in duplicate with the second sample digested) to determine serotonin genotypes. The ladder (well 1) is on the far right of the attached image). I would greatly appreciate any advice on how to enhance band brightness and definition, thanks.
Additional information: 5 uL ladder added, 10 uL PCR product per well, PH of the buffer is correct. Temperature of the room ~75F with Gel container NOT on ice.
Hello,
Please is goat genome not listed on UCSC In-Silico PCR website?
Please could someone assist on what to do if working with goat
Thank you
when I insert my primers in primer blast site and push the button "get primers" I just reach different errors and I don't know what the problem is!
Hi all
earlier I have seen in some papers people go for DNA extraction and normal PCR using 16S rRNA primers for the identification of bacteria. However recently I have seen few papers particularly dealing with Uncultured “Candidatus” bacteria, researchers go for RNA extraction, reverse transcription RT-PCR and real-time RT-PCR ? Molecular biology experts can you please tell me …..
1. what’s the key advantage between the two ? is there any particular advantage of RT PCR for the identification of Uncultured “Candidatus” bacteria ?
2. Is it because of the possibility of “relative quantification” of the bacterium by real-time RT-PCR by targeting the 16S rRNA gene of the bacterium?
3. Is there any advantage when (RT PCR) used for uncultivable bacteria?
4. what is this Cycle threshold ? what is the significance of this in the above reaction ?
5. Also “The eukaryotic elongation factor 1 alpha from the host was used as a control of the RNA amount, and a good extraction was expected to give a Ct-value around 15 (the cycle threshold was set to 0.1). ? all results with Ct-values above 45 were considered negative !, what does it all mean?
My aim is just to identify the unculturable bacteria from tissues! Can I go for just normal PCR (16s rDNA) and sequencing the PCR products? Please
thank you
regards,
I'm having trouble obtaining clear PCR bands for DNA fragments ranging from 907 bp to 655 bp. I've tried various methods, including:
- Testing different brands of Taq DNA polymerase (Takara, NEB, Vivantis, HF Pfu DNA pol).
- Using the appropriate buffer each time.
- Isolating fresh plant DNA using the CTAB method, followed by RNase treatment, ensures the template DNA concentration is not less than 100 bp (800ng/ul - 1500 ng/ul).
- Initially, conducting gradient PCR to determine the optimal annealing temperature in the range of 51 to 60 degrees Celsius.
- Using a new vial of primers (taken from stock primers).
- Running a positive control (Actin gene, 250 bp) alongside the PCR reactions. However, no amplification was observed in the positive control, with only smear and faint bands detected in some plant samples.
- Conducting in silico testing of the primers, which indicated they should work correctly.
Please provide suggestions on how I can obtain clear PCR bands for my products.
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?
I ran PCR using COI universal primers on DNA extracted from lice.
I added 25ul of 2x master mix, 5ul template, and 1ul each of 20uM F and R primers, with the remaining volume made up with DW to a total volume of 48ul, and ran PCR including a control group. However, no bands appeared on the gel after electrophoresis.
I then checked with a nanodrop, and all 5 PCR samples (including the control group) showed concentrations around 20000ng/ul, with A260 readings around 400, 260/230 ratios around 10-11, and 260/280 ratios around 37-47.
Where could I have gone wrong?
I would appreciate input from experienced individuals.
Hello,
Since a large part of my budget will go on NGS and I don't have the money to repeat it, I want to make sure that it will work.
The PCR products look good on the gel (nice bands of the expected length), but I was also wondering whether it's feasible to send a few samples for Sanger Sequencing to verify the product, before I spend all my money of an Illumina run.
Thanks :)
Dear Team Fungi,
We would like to identify root fungi from Vitis vinefera via metabarcoding. The methodology is established, we have had good results with other root samples using the isolation kit 'innuPREP DNA/RNA MiniKit' and the standard primers gITS7ngs/ITS4ngs with 49 °C annealing temperature. Unfortunately, we do not get any bands from Vitis roots, no matter what we try (e.g, adding BSA or a temperature gradient). Does anyone have any ideas on how to modify the DNA isolation or PCR to eliminate the potential interfering substance in Vitis roots? There must be something in there that interferes with the PCR...
With desperate regards
Kai
I want to copy a target gene from cDNA into a plasmid. The primers were designed according to the CDS sequence from NCBI. But when I performed PCR reactions I could not get any target bands. So the CDS sequence was synthesized into my plasmid vector. When used the plasmid as template and the above primers to run PCR, target bands were quite clear which means the primers can work. I know the gene copy number in plasmid must be much higher than in cDNA. So I increased the amount of cDNA template and cycle numbers (from 35 to 45 cycles), no target bands showed. Could anyone tell me what the problem might be. Is that possible that the CDS sequence in my cells has changed? If yes, is there any other ways to get the CDS sequence except artificial synthesis.
I am trying to amplify my gene for cloning. The desired PCR product is 2652 bp. The Tm for forward and reverse primers is 68.4 C and 61.3 C respectively. The annealing temp I set was 60 C. These are the results I got. How do I reduce nonspecific binding of primers? What can I do to increase primer specificity? Is the high difference in primer Tms affecting my PCR?
We run Fungal Panel using Molecular PCR Technique. We extract Nuclease Free water in the same way as we extract the sample. This eluted NFW is used as negative control on our PCR Plate for quality assurance. If we need to store this eluted sample for about a week, should we store it in the refrigerator (2-8 C) or Freezer (-20 C)?
How to identify sizes fragments of ZWF1 PCR product.
Maybe someone knows why this happens. The situation is that after PCR purification of gel products (cut one band), on the next electrophoresis, instead of one band, two bands appeared, how could this happen?
I used the Intact Genomics FastAmp® Plant Direct PCR kit and when seeding the gel with the PCR product there was a lot of DTT smell. After running the gel, partially degraded dna was observed, even in the molecular weight marker. Could it be possible that DTT diffuses into the gel and degrades the DNA?
I am trying to use the overlap extension PCR to combine two linear fragments of approximately 1200 base pairs in size. My first SOE-PCR was successful using Taq polymerase, with annealing and overlap temperatures set at 60 degrees Celsius. It had smear with my desired sharp bond. after that when I trying to repeat the process, I only obtained a smear with no specific bonds.
I amplified my fragments with taq and also pfu, but I don’t get my desired bond. I had just smear.
Does anyone have such experiment and help me, please?
Hi, I'm trying to develop a KO cell line from an established cancer cell line. My gene of interest is present in 3 copies in this cell line.
I'm using a multi-sgRNAs technique to increase my chances of a significant deletion. I isolated 4 clones of interest which share a similar trait: they all show 3 different bands after PCR amplification an electrophoresis on agarose gel. This is not so much a concern, since it was one of the expected outcome (the CRISPR/Cas9 system can create three different cutting pattern, resulting in 3 different bands). FYI, the 3 bands are all different in size from the WT band (with the top band being around 100 bp bigger than the bottom lane).
I ran again the sample on a more concentrated agarose gel (2%) with a lower voltage to get nice bands and being able to cut them. I extracted DNA from each band and re-run a PCR on each of them to increase my DNA material. For all my 4 clones, the bottom band amplify to a nice and single band corresponding in size. However, the middle and top lane display the 3 bands again, and it doesn't make sense to me. Indeed, I could understand finding the middle band in the top band sample, or the other way around. But I would have never expected finding the bottom band in the top band sample, because the top and bottom band are clearly separated and shouldn't contaminate each other.
I made a mistake by not using sterile instruments to excise my bands, which could explain in part some contamination. However, if it was this issue, I should have multiple bands in the bottoms sample, which I don't have, and I should have cross-contamination through all the sample, which is not the case. I'm pretty lost so if someone has any idea, I would take the advice with gratitude.
(I attached the gel picture from where I extracted the bands (small gel) and the re-run PCR gel whit the unexplained bands. On the gel, T= Top band; M= Middle band; B= Bottom band).
Thank you all!!
Is it possible to conduct PCR to check for resistant genes in bacteria and have no bands? All have bacteria have no bands of the resistant genes
Hi everyone,
I'm in the process of creating a zebrafish Knock-in line. In order to verifying that my integration has worked, I've created a positive control plasmid with the fragment that I would expect to have in my transgenic line.
Typically, using plasmids as a positive control for PCR reactions would yield single bands due to the purity of the plasmid. My concern is that, once I optimise my PCR using the plasmid, the PCR might not actually work when using extracted gDNA from zebrafish as the template. Hence, I was wondering if it is sensible to mix the plasmid with wild type gDNA to create an unpure template. I could then use it to optimise my PCR reaction. Does this sound feasible?
Thanks :)
I am trying to stitch in a 38 amino acid tag to the N-terminal end of my protein (3200bp) to be cloned into a lentiviral vector (~7000bp). The forward primer for the same, along with the overhang and the restriction site, comes about 150bp long. The first round of amplication gives me a band close to about 3000-3500bp along with a lot of other non specific bands at the higher molecular weight range. I then gel elute this specific band and reamplify using it as a template with the same primers but i end up getting a smear on the gel. I have also tried using this gel eluted sample to proceed with the digestion and ligation with my vector but in vain.
My PCR parameters are as follows:
1. 98 degC- 2min
2. 98 degC- 10s
3. 65 degC- 30s (2-4: x25 cycles)
4. 72 degC- 2min
5. 72 degC- 5min
6. 4 degC- hold
I use Q5 polymerase (strangely, I do not get any amplification with Phusion). I have tried a gradient PCR and it generally works in the range of (58-68 degC). I use about 50ng of the plasmid template for amplification. I understand that really long primers hamper the quality of amplification but unfortunately, this is a necessity right now.
I would really appreciate if anyone with experience can help me out here. My molecular biology is not THAT strong so please point out if I am committing any obvious mistakes.
Thanks in advance!
I ran the agarose gel and cut the right band, then put it to -20C, I performed PCR purification next day, but there were two bands. After two days, I ran the gel, the PCR products were almost degraded. Anyone could help me? Thank you so much.
Hi everyone,
I've been struggling doing PCR for fungi using ITS1F/ITS2R. My positive control (yeast DNA) worked well. My templates gave faint or no band. It sounds like my templates have inhibition but the 1 6S PCR worked very well for all of my templates. Also when I switched to fITS7bF/ITS4NGSR, PCR works for all of my templates as well.
I analyzed this pair of primers and found that they can formed self-dimer and primer dimer. So I've been trying many methods from increase annealing temperature, reduce primer concentration, touch down PCR, adding BSA, DMSO, increase denature time, even increase number of cycle to 40 cycles. But none of these really helps. I use Q5 hot start master mix.
Any suggestion please!
Thank you so much!
Hanh
I want to check my designed primers by in silico PCR in Genome Browser. but always face with this message [ No matches to cagatgagtcagtgccgttag agtaggtgctgactggttcc in Human Feb. 2009 (GRCh37/hg19)]. Is there any clue? Thanks.
I tested gene expression by RT PCR followed by Western blotting to test protein expression. I get an inverse correlation with up-regulation at mRNA level and down-regulation at the protein level. What could be the reason. Please suggest.
Hi All,
I am trying to amplify mitochondrial 16S gene for marine snails (Calliostomatidae) and other vetigastropods, but I only get primer dimers or nothing on the gel. These primers have worked previously in my lab and in numerous other publications. The DNA concentrations are low, but they have amplified for COX1 using the Folmer universal primers.
I am using the Palumbi 16S universal primers (Forward: CGCCTGTTTATCAAAAACAT and Reverse: CCGGTCTGAACTCAGATCACGT). We bought new primers in December 2023. I resuspended them and have tried multiple aliquots. I've tried gradients 45-55 and 55-65 and a touchdown PCR starting at 59 (-1 C/cycle for 10 cycles) and final annealing at 48 for 20 cycles. I've tried the standard, ammonium, and combination 10x buffers. All reagents are from Apex (not a hot start taq), except for the dNTPs.
Our usual protocol is: 2.5 uL of 10x standard buffer, 1.25 uL of MgCl2 (50mM), 0.5 uL of dNTPs (10mM), 1 uL of both primers (10uM), 0.2 uL of taq (5 units), and 2 uL of DNA. This does work for Folmer. Denaturation at 95 C for 4 min, 35 cycles of 95C for 30 seconds, 50C for 30 seconds, and 72C for 30 seconds, and final extension at 72 for 10 min.
I'm desperate and would love to hear any suggestions/tips on how to fix this!! I also unsuccessfully tried ethanol precipitation to increase the DNA concentrations, so tips on that would be appreciated too. Thank you
I am doing microvolume extraction which include physical( freeeze and thaw) and chemical lysis followed by a pcr base metagenomic library prep. My samples contain phytoplankton cultures with their microbiome. The mthod worked for most samples and timepoints but did not work for samples of one timepoint with high loads of phytoplankton and bacteria. I tried 3X dilution for direct PCR , bead-clean up and 3X dilution of sample and then bead-cleanup in case some inhibitors were hindering.
Looking forward for your scientific advice!
Thanks.
Greetings to all!!
DNA is isolated from infected cotton leaf.
The image is attached. It looks kind of shearing.
What are possibilities to use it for PCR?
I am looking for help to optimize a nested PCR from Long Range (LR) PCR (DNA as template). The LR PCR product looks spesific on agarose gel. We have tried various dilutions of this product as template, but keep getting a couple of unspesific bands on the gel in addition to the expected product (the unspesific bands are a little larger in size than the expected product). When we add genomic DNA instead of the diluted LR PCR product, using the same polymerase and conditions, we get clean product of the correct size. What could be the cause of the unspesific bands? Is it necessary to clean up the LR PCR product before using it as template for nested PCR when we dilute it (1:50, 1:100, 1:1000)?
Hello all, I have a query in resolving close products in agarose gel electrophoresis: I have three different expected products in my samples: 460bp, 480bp (only in mutation), 323bp, and I run them in a 3.2% TBE agarose gel, thickness 0.75cm, running conditions: 75V, at 4degreeC, for rough 8hrs with paused interruptions for detection every 2 hrs. What I see is that I have a definite signal at 323bp and 460 bp, then between 500-600 bps I have various other products (picture here after 8hr 10mins). I had cut the band, eluted the DNA and Sanger sequenced the products, turns out the band at 500bp is the same as 461bps and the band at 550bps is the same as the 323bps. The band at 460bps of the mutants, is a mixed signal in the middle of the Sanger sequence, where I could see that it has the sequence of the 323bps and the 480bps with the main 460bp sequence.
With the a second PCR of same settings and cdna, I run the sample with 3.2% agarose in TBE, reduced the thickness to 0.5cm this time, run it for 2 hrs in 150V, then further at 25V for 12hrs at room temperature. Here I do not see, multiple products between 500-600bps but one single product around 800bps. Can PCR products heteromerise when they run through a high % agarose gel?
I would like to resolve the products, but still avoid the poor resolution of some portion of the products, in the agarose gel. I am using the biozym LE agarose. any suggestions to improve for this experiment please?
Hello,
I am currently attempting to obtain amplifications of SoxC from a species of onychophoran belonging to the Peripatidae family using primers designed from a sequence of its sister family, Peripatopsidae. The sample I am using is cDNA, and I have tried different concentrations of reagents and cycling times in the thermocycler. However, I have not obtained any amplification yet.
I would like to ask how you would start standardizing reagent concentrations and the thermocycler program for a set of primers generated from the sequence of another species for a conserved gene like SoxC (700 bp amplicon).
I have COI as positive, and before using any cDNA for SoxC, COI was amplified.
Due to resource optimization, the primers I use for SoxC had to be initially designed with the sequences of the promoters SP6 and T7.
So these are my primers:
SoxC-F5: ACGCCAAGCTATTTAGGTGACACTATAGGCGGCTACGGATCTTACACA
SoxC-R5: CAGTGAATTGTAATACGACTCACTATAGGGGGGAAATCAAAGTGCGAGCC
none exceed 50% CG.
Additionally, I am conducting tests with cDNA extracted from different tissues and stages to increase the likelihood of finding my sequence, but I still fail to obtain amplification.
Hi everyone, I'm doing PCR for mycoplasma detection I'm using the primers GPO-3 and MGSO; the denaturation, annealing, and elongation temperatures and times were 95oC for 2min, 95oC for 30s, 57oC for 45s, 72oC for 1min, 72oC for 7min, for 40 cycles. The results were analized by gel electrophoresis using 1.5% agarose.
A band in 270pb is a positive result, but in some samples a band is amplified in 200pb. I was suggested that this band could be interpreted as a positive result for a different mycoplasma genus.
Polymerase Chain Reaction (PCR) testing is a molecular biology technique used to amplify and analyze DNA or RNA samples. So, To perform PCR testing, which list of laboratory apparatus and equipment is required?
I am trying to check the homozygous mutant for Arabidopsis (seeds ordered from ABRC). I have done genomic DNA extraction, using Edward's method and checked the it in agarose, the genomic DNA is there. And also I got quite a good concentration of about 1ug/ul. I designed primers using SIGnal primers design against the salk ids, but my PCR is not working. I have tried different temperatures and polymerases.i have kept the 1st denaturation time about 10 min, and checked the primers, its interacting fine with the genomic DNA using clustal omega. where am I doing wrong?
I have a question for the professionals. The essence of the problem: I fix the oligos on a plastic substrate, they play role of primers in solid-phase PCR. I carry out a one-step PCR with simultaneous labeling of the product with biotin (I add 10% labeled uridine to unlabeled T to the DNTP mixture), then I wash it in PBS and incubate it with the streptavidin-peroxidase complex and then incubate it with the substrate for peroxidase. Everything would be fine, but in the control wells, where the PCR reaction mixturedoes not contain DNA, I have a staining of oligo spots, weaker than in the experimental wells, but it is there. Moreover, in the control wells, where only PBS was added, weak staining also appears at the localization spots of the oligos. If I simply add the complex to the wells (without any PCR treatmen), then only the positive control points, that is, the initially labeled oliagos, are stained in them. So here's the question. Can streptavidin (or peroxidase) bind to something other than biotin or DNA oligos? I’ve been fighting with the problem for a couple of months now. I 've changed blocking buffers, polymerases, washing modes, but the result is still the same. Help, good people!
The Quantstudio 3 qPCR machine works really well with low-ROX SYBR premix PCR, I just concern about whether it can work in high-ROX premix or no-ROX premix? In the software, the reference dye can be chose as ROX or none, etc, but not having options like low-ROX or high-ROX?
ChatGPT claims that the following general invertebrate primers are often used for nematode barcoding:
- Forward Primer: JB3: 5'-TTTTTTGGGCATCCTGAGGTTTAT-3'
- Reverse Primer: HCO2198: 5'-TAAACTTCAGGGTGACCAAAAAATCA-3'
Having problems with dimers and an answer from Paul Rutland for another question made me think that the fact that I store my PCR mix in the fridge might be the problem. I must add that in this mix I add both primers, so maybe dimers are being formed prior to the reaction during storage. Does this make sense?
Bands are appearing very, very faint! I presume low DNA in gel, but my DNA concentration is over 1ug/ul. This is after my overnight digestion. I used a MaxiPrep protocol to extract DNA from Bacteria culture. (I inoculated from my glycerol stock and extracted the DNA after a MaxiPrep).
My DNA (pUH-dnvamp2-iGluSnFR) seems to show clear band digestion on the gel and accurate band size. My other plasmid (pUH-iGluSnFR) seems to show very faintly on the gel after digesting the extracted DNA plasmid. I screened my glycerol stocks and with a MiniPrep and found one that showed my genes. I proceeded to do a Maxi Prep with that clone. My concentration for this DNA was 1349.3ng/ul, and the A260/A280 was 1.9. I have repeated this digestion multiple times. Yet, the gel run after the Maxi is very low.
I am about to run a PCR, assuming that any little DNA present will be amplified to confirm my genes of interest. My purpose, in the end, is to utilize the plasmid for virus production. Hence, a high DNA conc. is important for high virus yield.
Any help troubleshooting will be appreciated. (Pictures: first one is Gel after MiniPrep, second one is gel after MaxiPrep
I am working with an allergen and i am working using PCR, the result that offers the kit is copies DNA, although i need to give a result in mg/kg. Is any possible way?
Thank you in advance,
Kiriakos
I have ran a gel to determine DNA products with the following base pairs, 745
590, 317 and 825. However, I got bands just below the ladder, my negative control came out negative and I do not know what conditions to change to address this.
Hi all. I have some primer pairs which always produce those horrible primer dimer bright smear! Increasing annealing temperature does not solve the problem. Any suggestiopn for a PCR enhancer or another strategy? So far I have used DTT and DMSO and amplification quality still poor! Thanks
Hello,
I am interested in performing genomic DNA extractions and subsequent PCR analysis on some human cells (HEK293T). However, I am thinking of using a "colony PCR", i.e., by taking a number of cells and putting them into the PCR conditions and hoping that the initial denaturation temperature at 95℃ is enough to lyse the cells and release the genomic DNA.
Is this possible to be done? Has anyone attempted this, and if they have succeeded, how many cells are required and what are the parameters of the PCR conditions?
Thank you very much in advance!
I designed a primer for gene sequence(1480 bp) and when carried PCR, the product was 150 bp, what is the problem? and how to obtain the correct fragment?
After transformation in DH5 alpha, I got positive colonies. I have confirmed it by PCR (using an isolated plasmid of positive colonies as a template to run PCR by Takara Taq) and restriction digestion. In PCR, I got exactly the same size of band as my desired interest in the gene but did not get fall out of my gene in restriction digestion.
I have attached a gel pic of PCR and restriction digested . 20 ul of restriction digestion was put at different amount of plasmid ( 5 ul and 8 ul of 140 (C1 )and 305 ng/ul (C2 )
hey I really need an urgent help
I'm so confused with the primer sequence of 1492R.
some journals said that 1492R is GGTTACCTTGTTACGACTT (and I use this as my PCR)
but the research company that will help me sequence my bacteria said that 1492R is TACGGYTACCTTGTTACGACTT
I need this answer as soon as possible because I have to send my PCR product to sequence service, thank you
Hi ResearchGate community,
I have been trying to learn more about the optical differences between block-based real-time PCR machines like ABI StepOne versus rotor-based machines such as MIC or RotorGene systems.
I understand that some systems rely on ROX as a passive reference dye while others state that it is optional to incorporate it and others do not need such a factor at all.
My question is if you add this fluorescent dye to your master mix, would it interfere with the detection when it is being amplified using one of the systems that do not need such normalization?
Highly appreciate any insight in this regard.
Best,
Negar
I am amplifying target sequence 450 bp. I get single sharp band in the control and faint in sample with another sharp nonspecific product. why?
I need to get one single band from my sample to sequence the target . what should I change?
I changed annealing and DNA concentration, time of each cycle and used different PCR master mixes.
Hey everyone,
my question is maybe strange at first glance, but simple: is the rapid 16S kit's only real advantage the significantly larger 16S data amount generation? Shouldn't I be perfectly able to collect necessary strain-level diversity 16S data on the data analysis level from a total nanopore metagenome, without the PCR bias, given enough sample input? If the above thinking is correct, would you consider triple-digit ng input (below 1ug) sufficient, at least for key players of a mixed microbial community?
Just trying to understand if I really need the 16S barcoding kit since I have the native one (which I will use for total metagenome anyway)
Cheers
A
Hi everyone,
I need your help with optimazing my PCR reaction. My PCR product should be 850bp. Starters Tm is 65, they do not form dimers or secondary structures and they’re specific (Blast doesn’t show any unspecific product that may occur). I've tried gradient PCR with typical mix and with Hot start polymerase and every time I receive strong band that is 150bp. What can I do to get rid of this unspecific product and what can it be?
I successfully amplified fungal ITS from soil samples, however after running the purified PCR products in an agarose gel they are barely visible and don't look like defined bands but rather clouds. The purification was done with the Monarch PCR and DNA Cleanup Kit. Why could this be happening?
Hi, I am using VASA-seq for RNA sequencing. The last two steps of the protocol are cDNA synthesis (via reverse transcription) and PCR. I had been using Superscript III for cDNA synthesis and was getting a lower PCR yield. Then I switched to Maxima H Minus Reverse Transcriptase. The PCR yield increased dramatically, but I am getting this weird around 1200 bp long fragments (see the attached figure). My expected peak is around 300 bp. I have attached a figure of the fragment size distribution of the PCR DNA (analyzed on fragment analyzer).
#fragment_analyzer #PCR #VASA_seq #Maxima H Minus Reverse Transcriptase #SuperscriptIII
Hello. I am having troubles with serum samples. I know that they are positive for Leishmania but when i do the PCR, most of my samples are negative. So, any ideas to have a better outcome? Maybe some extra step.... I use a MagMax kit for extraction, but i can also use Promega and Nzytech. Thank you
Hello! I've encountered some challenges with Traditional PCR.
I've successfully conducted RNA extraction, quantification, and integrity checks, all yielding positive results. (first image its from the integrity of the RNA)
Moving forward, I proceeded with RT-PCR, followed by PCR endpoint analysis using Actin primers. My experimental design involves four treatments, including Ctrl, Resveratrol, LPS, Resveratrol+LPS, with two samples for each treatment.
However, I've encountered an issue where only the controls are being amplified during the PCR endpoint, despite using the same mix for all samples in both the RT-PCR and Traditional PCR for Actin. I'm puzzled and unable to pinpoint the source of this discrepancy. Any insights or suggestions would be greatly appreciated.
The second image its the results of the PCR.
I ran a PCR reaction and it gave good result during the trial run. However, once I ran the same PCR reaction on all of the other samples, there are smears and appearance of non-specific bands. I'm not sure on what went wrong. Hopefully, I could get some insights to fix this issue. Thank you in advance!
Dear virologists; What is the PCR technique used in virology to detect viral nucleic acids? The steps involved.
I'd also like to know, since some viruses have a single strand of DNA and RNA, how does amplification work in this case?
Kind regards
Does anyone have any tips for optimizing PCR reactions with low-quality DNA samples? My interest is in the identification of some species of bacteria using specific primers for each species.
I am working with a DNA sample extracted from different tissues (kidneys, liver, spleen, muscle, cartilage) from carcasses of mammals run over on highways. The tissues were stored in ethanol at room temperature (not my choice) during the collection period, after which they were frozen. DNA extractions were performed with an Invitrogen extraction kit and treated with RNase.
Any help would be appreciated, thank you very much :)
