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.
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I am using nested PCR to determine the alleles present in my parasite population for a specific gene. This has gone well, but one gene has given me an issue.
The initial PCR for gene MSP1 resulted in a product of 477bp which was expected. The following reaction should produce products between 100-300bp, however, I am seeing those amplicons in some samples as expected but there some samples have a band around 500bp.
The primers are from literature and I have not designed them.
Is it likely the primers are non-specific? In this case where the expected amplicon size for the allele is known do I then ignore these larger amplicons?
Any help would be appreciated
Edited to add: I'm using the PCR conditions specified for the mastermix with the annealing temperature as determined by the supplier's annealing temperature calculator available online.
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did you gel purify the first PCR product? If you are just diluting the reaction you could have WAY too much of the primary primers in the reaction and they are continuing to amplify in the nested reaction.
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the primers are going to be used to detect Klebsiella pneumoniae specifically, so I made it from specific sequence after looking on NCBI. The sequence was from Korea, while I tested the primers using samples from Indonesia. The PCR settings I used was pre-denaturation at 95 degrees Celcius for 5 minutes, and then 30 cycles of denaturation at 95 for 1 minute, annealing for 30 seconds, elongation at 72 for 30 seconds, then polymeration at 72 for 10 minutes.
once a friend suggested to try using the same settings, but instead of 30 cycles, we used 40 cycles, took half of the sample's volume for electrophoresis, and I got single band on the right spot but too faint. So we used the remaining volume that is still inside the PCR tube, for another 40 cycles, and I got a single band and very visible.
I ran 3 samples on 3 different degrees (56, 57, and 58 degrees Celcius), and the result at 56 was very visible and on the right spot, at 57 was very visible but below 100bp, at 58 was very visible but a little bit higher than the 56 one.
What does it mean ? Does my target sequence has a very low copy number that it requires a very long PCR time for it to be amplified enough to be detected ?
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1. first check you primer specificity on https://www.ncbi.nlm.nih.gov/tools/primer-blast/ if you are setting amplification using cDNA use RNA reference or if you are using genomic DNA use genome as database blast with your organism. analyze the results, does this results have your fragment of interest?
2. make sure your primers have G or C at end.
3. check the species and strain of your samples.
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hii,
I have one group in six different ZTs --> ZT0, ZT4, ZT8, ZT12, ZT16 e ZT20.
I performed gene expression by real time PCR using the 2- deltaCt method
I have used graphpad to represent the curve but I don't know what statistical method to apply to obtain the p value.
Thanks in advance :)
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Hello.
I tried to get bands from a few samples using the nested-PCR method, and I tested the 2nd round PCR with 0, 10, 100 and 1000 dilutions of the first product. Only in 0 dilution, smear is seen. Where is the problem with this test and how can I optimize this test?
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Hi! I hope you are well.
Did you manage to get an answer to this? As I have been having a similar issue.
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Hi all, I am struggling to transform my BY4742/1 cells using the Gietz protocol along with a linear PCR fragment. I understand that using a PCR gene disruption cassette is not as efficient as using a plasmid for homologous recombination however this is the situation I find myself in. My fragment is ~ 2kb and I have used a high fidelity pol for amplification. I'm wondering if anyone has any experience in doing a gene deletion this way and how much DNA & ssDNA carrier you used for a successful transformation. Would it possibly be better trying electroporation and if so how much DNA would be suitable in that situation?
Thanks!!
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Hello Jessica, when I use PCR cassettes, then after PCR I check the accurate size of the amplified products on gel, later I purify and concentrate the PCR products (not from the gel). There is no homologous sequence between the cassette plasmid and yeast genomic DNA, so no worry to get an inaccurate transformation. When I use integrative plasmids, I have to linearize before adding to the transformation. If the restriction enzyme is able to linearize the total amount of integrative plasmid, you don't need to purify from the gel, but you must run a gel to check the linearization status before purification of the linearized plasmid. I would recommend to purify from the gel if the restriction enzyme doesn't work efficiently, but consider that you may loose some yield of DNA.
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Hi everyone,
I am trying to knock out a non-coding region downstream of my gene of interest. I looked at a publication which did exactly this but in a different species.
They seemed to use two sgRNAs flanking the region of interest and didn't mention any additional components to the knockout in the method section. They show almost complete knockout in their paper (using PCR, the bad after KO is 500bp shorter than in the mock)
My question is; is this a common approach? Just using two sgRNAs a few hundred bps apart without any HDR template and hope that the NHEJ leads to a loss of the region between the two sgRNAs?
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Yes, this is very common method for making deletion/KO using a pair of sgRNAs. The efficiency is length dependent and also requires each sgRNA to be efficient. Check out the publication bellow https://www.jbc.org/article/S0021-9258(20)47516-7/fulltext The repair by NHEJ will not be perfect with some random indels, and occasionally the deletion may spread further away from the predicted cut site. You could add an HDR template if you need a very precise deletion with specific length.
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Does the DNA remain stable or degrade at this temperature? Would there be any difference in thermal stability between supercoiled and linear forms of say, 3 kb plasmid.
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The question of the structure of DNA is not a question of the origin of the Universe. It must and can be solved experimentally. But for this you need to have a desire to know the truth. The problem is lack of desire.
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Dear Researchers,
I did stable transformation in passion fruit and now doing detection of mutants, I run the PCRs using the mutants DNA (Target), control plants without target gene DNA ( CK), Agrobacterium/ plasmid (positive), H20 ( negative control) with the target gene primers (around 1700 bo product length), later run the 1.5-2 % Gel, but the bands of mutant samples were above the positive band, mean above than the target product length which is around 1700 bp, as the positive (Agrobacterium showing exact bands) but the samples showing around 2000 bp or above, marker 2KB, I have attached the picture, underlined are all mutant samples, both side 2kb marker, and +ve mean the known and confirmed Agrobacterium with around 1700 bp of the target gene.
so what possible problem and solution about these having bands above the target?
I shall be thankful
Regard
Rizwan
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Could it be that you have homozygotes at the right size but in the last round of pcr in heterozygous samples the dna strands re anneal with a bubble at the mismatch in the dna and this causes the shape of the dna to change and the dna runs slowly so just looks like it is larger.
If this is a random size effect then possibly some of the amplimer has some polymerase attached so is running slowly as suggested above in which case adding 0.1%sds to a slow runnung sample may cahnge its size to smaller dna only
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I have been dealing with degenerate primers for paramyxovirus virus detection. The detection assay calls for a heminested PCR with the same reverse primer used for round 1 and round 2 of the PCR reaction. The issue arises when trying to perform round 2 using the round 1 PCR template.
I wanted to ask if there is some specific way to dilute/ purify the round 1 PCR products to get proper bands in round 2.
Also, i did not experience this problem when dealing with Nested PCR primers that are designed differently for the forward and reverse reaction.
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You are probably getting a smear of many size dna because of over amplification. Transfer of the outer primers does not help so dilute 1ul of first round product in 100ul of water then re amplify 1ul of this diluted dna using a 24 cycle pcr but remove one pcr tube at 14, 16, 18......up to 24 cycles . Some of these tubes will have clean and strong amplification and will be a guide to second round amplification in the future/ Remember in pcr every 10 cycles is 1000 times more product so over amplification is very easy in nested pcr
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I was trying to use Gibson Assembly to fuse 2 PCR products (although it is not really used for this purpose). Upon Gibson reaction of the gel extracted PCR products, I tried to amplify them with Q5. On the test gel, there was only short bands slightly above primer dimers. I was curious if Q5 is faulty and I tried to amplify only one of the PCR products with Q5 (I know for sure that the primers work since PCR from cDNA works and forward primer was successfully used for Sanger sequencing), the result was negative: Q5 did not amplify anything from the gel extracted PCR product. Did anybody experience the same/similar thing?
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Leonard Koolman thanks a lot for the answer. I checked the purity on Nanodrop, A260/280 ratio was around 2.00. It should be pure enough (maybe 1.8 is better), but I am afraid the concentrations were too low, like around 8 or 10 ng/ul and there was no clear peak on Nanodrop around 260nm. I am suspicious at this stage if I had good enough DNA for PCR. I will try PCR column purification. If you have any experience on Gibson assembly, whether it works from unpurified or column purified PCR products instead of gel purified products, that would be great to know.
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Hi guys,
Attached herewith is a 1% agarose gel picture of my PCR reaction (triplicates). As my total reaction volume for the triplicates were each 50 ul, 25 ul of the each reaction was loaded into one well. The profile of the triplicates was different. Only one reaction was showing the band of interest (indicated with a red box), while the other two were smeary. This would mean that the PCR reaction is inconsistent. I used 10ng/2ul of cDNA for my reaction, 0.3 uM of the primer pair, 0.3mM of dNTP, 0.5 U of KAPA HIFI DNA polymerase.
My PCR cycling conditions are as follows:
Initial denaturation: 95 celsius for 3 min
Denaturation: 98 celsius for 20 sec
Annealing: I did a 2 step cycling conditions, 52 celsius for 15 sec for 5 cycles, and 62 celsius for 15 sec for 30 cycles.
Extension: As the fragment that I want to amplify is around 2 kb, I used 2:30 min at 72 celsius
Final extension: 72 celsius for 5 min.
I am planning to add in DMSO, but would like to hear from you guys first before I proceed.
Please advise.
Thank you.
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I would definitely want to run a temperature gradient to optimize the reaction as those are broad smears. If the replicates consistently provide different results, you may want to check your PCR machine as some of the wells may not heat/cool properly.
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As stated, I would like to know what is the difference between NCBI blast and EzBioCloud?
I have sequences of multiple unknown bacteria and am trying to do the first identification on them using only the 16s sequence by sanger sequencing. Once they have been screened, I will pick the one that looks interested to move forward with the characterization.
However, when I tried to use these two services, I have gotten a bit different results.
For example, in NCBI blast, the sample I have generated more than 10 hits that have < 99% identity. However, in EzBioCloud, I can only get one hit up to 98.99% of similarity and it doesn't have a strain name or taxon name. The highest one with strain name and taxon name only has 97.91% similarity. Therefore, I don't know if this one can be considered a potential novel strain or if someone might already have isolated and identity it.
I was wondering what's the difference? Thank you.
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1. May I ask When I should click "type materials only" ? or should I do this every time I want to identify the sample?
-> If you want to know exact species name, should click “type materials only” but want to know distribution of similar strains do not click.
2. I just changed it and it turned out the best hit was at 98.66% similarity!! -> It’s on the border, however, this means need to confirm by genome indices such as ANI or AAI. Usually that value resulted new species.
3. I am also wondering how do I know if these sequences are misidentified? In the original result (without clicking the type materials only) the top 2 hits were new isolates from one paper that were published just last year. so I'm not sure how do I determine if this is reliable or not even when this sequence result was from one of the published papers? -> If top 2 hits have new name, need to think more but have previous species name but newly isolated strain, don’t mind it. Species is determined by comparison with type species.
4. In terms of EzBioClouds, how do you know it is an uncultured type? is it because it doesn't have a taxon name?-> Only NCBI accession numbers, its uncultured type.
Also circling back to Blast, the hit strain name of the first hit on EzbioClouds happen to be the first hit on Blast when I clicked "type materials only". Does this mean this one has been cultured but EzbioClouds has not updated its database? I quickly searched for the name of this first hit and it was only recently been proposed to a new species. So I guess that is why. But please let me know if anything I say doesn't make sense. Thank you!
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Hello,
I am using the Qiagen PCR clean-up and purification kit. I am purifying a PCR product of 750 bp.
I have added the buffer PB as described in the manual. The color of the mixture was violet (too high pH) so I added 10 microliters of 3M Sodium acetate (pH 5.0) as advised by the manual. However, the mixture remains pinkish in color.
is there any other way to regulate the pH and lower it?
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Mu-Shen Chang I am using PCR product directly.
i kept added more acetate yet the color became a more diluted orange rather than yellow. (pH indicator gives yellow for pH <= 7.5 and orange/violet for >7.5)
Is there any other way to lower the pH?
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I have used 10 microlitre forward and 10 microlitre reverse sequence stock to make 100 microlitre working primer solution is that the reason for these kind of bands what is dimer ?
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You need to change the annealing temperature for the specific amplification.
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Dear All
We are trying to detect Wolbachia in Culex and Aedes spp (adult, larvae and pupae). Can anyone please tell what would be the preferred approach to start with considering adult moquito samples. For instance, some articles refer to use to whole body while others have isolated gDNA from organs such as ovaries. It is also mentioned that removal of head improves detection due to presence of inhibitors. It is also proposed somewhere that Wolbachia could exhibit tissue tropism. Kindly share your experience and expert advice on said query.
Looking forward
Thank you
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maybe you can start by MLST?
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I got a problem in my PCR product sequencing. The agarose gel electrophoresis showed that the band of PCR product is single and strong, though a smear which is smaller than the desired product can be seen. I cut gel and purified the PCR product and send it out for sequencing. However, I got bad sequencing results. Has anyone else run into this issue before? Could you give me some suggestions? Thank you.
Attached are some example file that were giving me trouble.
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Your sequencing is good Bin Wen . Strong signal,no big unincorporated dye peak and all the run parameters are typical for capillary sequencing. You report a single strong band on agarose gel so the problem here is your sequence not your sequencing. The interesting parts of your sequences are the start and all of the parts of the sequences that have a run (4 or more) of the same base. In all of the monomeric base sequences the middle bases are clean and strong but at the ends of the motifs there are 2 0r more bases overlying each other. You are sequencing a pcr product which has insertions and deletions ( probably single base changes) so the sequencing starts well but at the first indel you now have 2 basecalls at each position because you have 2 bases at each position. In sequence 1 there are multiple overlays so I think that this amplimer has more than one indel and you are reading 3 or more similar sequences. Cloning the pcr product and sequencing clones will confirm this. If you have a sequence of this product which has sequenced cleanly it should be possible to see what the nature of the indel is by subtracting the known sequence from the double sequence to see what ,and where, the indel base is
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Hi,
I wonder if anyone knows how to use sample release reagent from Sansure Biotech ?
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Hi,
The supplier claims that their "sample release reagent" (nucleic acid release technology), can lyse pathogens at room temperature very fast, with no need to heating, centrifuging or replacing tubes, the sample DNA/RNA can be extracted quickly through simple operations. The reagent is applied for the pretreatment of nucleic acid molecules, to release them from specimens, then the released nucleic acids can be used for clinical in vitro diagnosis or for the detection through appropriate apparatus.
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I electrophoresed 6 ul of PCR product for 90 min at 85 volt in 2.5% gel and TBE buffer. my PCR product length is 490 bp.
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It seems that too much PCR product has been uploaded. Have you run the controls along? Please check with the PCR product conc. and load only 1ul if you dont want to dilute. You can lower the cycle number in PCR reaction in future as well. Good luck
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I want to do MSP for InsR and slc2a4 gene. My template DNA comes from skeletal muscle and blood of diabetic Mus musculus. After DNA extraction I treat the DNA with bisulfite treatment. I'm still in optimization stage. Before MSP I do nested PCR to my bisulfite treated DNA. I tried to change primers concentration (10mM, 5mM, 2mM, 1mM, 0.5 mM 0.25 mM, 0.1 mM), used temperature gradient for annealing temperature (for both nested and MSP), and adding more template.
I uses PowerPol 2x PCR mix by abclonal as my mastermix. My friends tried to use the same DNA and they got their target band (Their target genes are FTO, KCNJ, PDK4, and PPARG). I set the PCR program exactly the same as the mastermix user manual suggestion. My mix contains 25 microliter PowerPol 2x PCR mix, 1 microliter primers (I have tried 10mM-0.1mM), 1-5 microliter of template(bisulfite treated DNA for nested PCR, Nested PCR product for MSP), and adding Nuclease Free Water until the final volume 50 microliter. What should I do to make my target band appear? The only things that appear in the gel are non-specific bands.
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my feeling is that it is unfortunate to have a long (600bp) amplimer using low temperature annealing primers because the template dna at low temperatures will
re anneal quicker than the primers can anneal so you have a low efficiency pcr. Ideally longer primers or a shorter outer sequence at first amplification could work better . Still these are the primers that you have so you could try running the pcr in presence of 1M betaine final concentration to try to stop the template reannealing for as long as possible or try adding 6%dmso to the pcr mix to stop secondary structure formation again to hold the strands open for as long as possible . For slc i would run a gradient from 45 to 55 annealing for the first amplification even if this only provides a dirty amplification because it is hard to troubleshoot a complete failure but relatively easy to clean up a dirty amplification
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Hello everyone.
I am working on Aspergillus sp., so I ordered primer specific to the Aspergillus genus targeting ITS1 and ITS2 markers. I tried the primer on a positive control and one of the isolates that confirmed by morphology that they are one of the Aspergillus species.
Unfortunately, the band (595-600bp) was seen in the positive control but not in the chosen isolation.
PCR conditions :
94oC -5min
94oC-1min
58oC-1min
72oC-1min
72oC-10min
35 cycles
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180 ng in a pcr is rather high for template dna and you run the risk of having pcr inhibitors present in the dna Manal Abdullah Al-musa but more worrying is the high OD ratio. 1,8 is the maximum for dna but a value of 2 suggests that most of your dna is actually RNA
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I have done electrophoresis of my PCR product 70 V for 40 min and the bands were moving away from the ladder range what is the correct way to do electrophoresis?
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Can we see a picture of the gel. A good pcr band should work at many times and voltages and I wonder if you just have primer dimer and no amplification
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I am trying to use PCR to amplify my samples with barcoded primers for 16s sequencing. I used Phusion Hot start II DNA polymerases kit. My primers are supposed to target the V4 region and amplify the products around 330bp. However, besides the target products, I found some bigger products (about 550 bp) appeared. I haven't came across the issue before and the negative control was fine (which exclude the contamination from water or materials). The PCR protocol I followed is : The 10-20 ng of DNA was used as template in the PCR reaction (50µL), which contained 10µL HF buffer (Thermo Fisher Scientific), 1µL dNTP Mix (10mM; Bioline, London, UK), 1U of Phusion Hot Start II DNA Polymerase (Thermo Fisher Scientific), 500 nM of each barcoded primer. PCRs were performed with an Alpha cycler 1 (PCRmax, Staffordshire, UK) using an adaptation of the cycling conditions of Caporaso et al. (2012). The cycling conditions consisted of an initial denaturation at 98◦C for 3min, 25 cycles of: 98◦C for 10 s, 50◦C for 20 s, and 72◦C for 20 s, and a final extension at 72 ◦C for 10min. The size of the PCR products (∼330 bp) was confirmed by agarose gel electrophoresis using 5 µL of the amplification-reaction mixture on a 1% (w/v) agarose gel.
Few thoughts from myself:
1. the DNA template I add was too much, I add about 100 ng DNA per reactions, which might increase the unspecific products. However, my cDNA samples also had the bad as well.
2. the initial denaturation is too short
3. the annealing temperature was too low. the lowest Tm for my primers is 59 degrees.
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If the above good advice does not work then try using half the amount od primer and run a dmso gradient. prepare tubes with exactly the same reagents but use 0.1%, 2%......8% final concentration of dmso and hopefully at some concentrations the non specific band will disappear and you will get a clean amplification. Run 2 more cycles of pcr to get a strong amplification
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I got primer dimers for my PCR. Then I tried to use hot start master mix for PCR and ran the gel. I got multiple bands for hot start PCR product. What is the reason for this.
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There are several reasons you can end up with multiple bands on PCR. Common causes are too many cycles, annealing temperature too low, annealing or extension time too long, too much primer in the reaction, incorrect ramping speed, or too much MgCl2. I would double check the calculation on the concentration of primers and MgCl2. Then I would try to increase my annealing temp. Not sure how many cycles you went but if you go much above 35 you increase the chances for multiple bands.
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So, I've been doing DNA extractions and PCR of mushroom collections and cultures, and I'm having mixed results. Routinely, we've been diluting all our DNA extractions if it's above 50 ng/ul, but sometimes I'm not having good bands on gel after PCR, and I'm worried that my DNA concentration is not optimal, is there a certified optimal DNA concentration for PCR?
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It varies with the kits/polymerases you use. You can usually find this answer in the user manual of the polymerase you are using. For example, Thermo Scientific Phusion polymerase states using 100-250ng of high complexity DNA in a 50ul reaction (https://www.thermofisher.com/document-connect/document-connect.html?url=https://assets.thermofisher.com/TFS-Assets%2FLSG%2Fmanuals%2FMAN0012393_Phusion_HighFidelity_DNAPolymerase_UG.pdf).
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Doing SDM, getting PCR but not getting transformation colonies. What could be the issue? Comp cells are fine when used control.
-PCR: 10 cycles (40ng plasmid template), Can See right size PCR band on the gel.
-PCR digestion with DPN1 (1ul 2hr)
-Heat Inactivation 72℃ for 20 min.
-For transformation 0.5, 1, 5, 10 ul of above mixture in to 30ul of JM109 competent cells Kan+plates. (Control plasmid total 10ng in 1ul volume for 30 ul cells(same) gives colonies)
So no colonies on SDM mixture. What could be the issue?
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OK!! of course it is not possible to see the template plasmid. The PCR product is a circular DNA with two "nicks' at the 5' ends of the oligo; I don't know how it will run on a gel, not sure that it will run as a linear DNA (it got sticky ends the size of the oligos) . I think you have to look at the design of your oligos, maybe makes longer ones) do you have repeated sequence in your plasmid? ... good luck with your mutagenesis
regards
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Dear Community, I extracted DNA from yeasts and performed a PCR with ITS primers. The PCR bands are not intense and do not allow sequencing. I've tried to extract DNA with a lithium protocol, but the bands are still weak. Do you have any recommendations to have better yields of my PCR products?
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Mohd Imran I'll try to do it for the samples that didn't work at all thank you. I made PCR yesterday with dilution to the 1/2 and it worked better.
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I have done gel electrophoresis of my PCR product the results were shown like this there were no proper bands observed in the gel and some of the amplified DNA bands were stuck at the well region can anyone tell me what could be the reason for this?
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What is your DNA template? It is from genomic or plasmid? I am asking, because the top band, just below wells looks like accumulation of your template. And you have a lot of it, in my opinion too much. The same can be with primers, you can see them below the last band of the marker. It could be one of the reasons.
Usually I prefer to run PCR with as low amount of template as I can. So probably good point would be to lower the template concentration, I dont know, maybe 10 to 50 times?
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Hi, I'm Siska
I've do PCR product cloning for blunt end PCR Product. It amplified by using KOD FX Neo (Toyobo). But, the product PCR didn't ligate to pTA2 vector. I've try twice by using pTA2 cloning vector, but it's failure. What should I do with this problem?
Thank you
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No, I didn't. After A-attachment process,I did ligation by using T4 DNA ligase, incubation 30 min, and then did transformation
This is the protocol that I followed
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When I measure the concentration and quality of the purified PCR product, the value of 260/280 is within the range of 1.8 - 2.2, while the value of 260/230 is always close to 0. Why is this happening? Can it present any complication for subsequent procedures?, such as performing another PCR from that purified product. And finally, would it affect nucleotide sequencing?
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It depends on the absolute value of A260.If this is very low then you will need a large volume of dna to sequence and the guanidine salts may be a problem but if A250 is high there is a lot of dna so you will sequence a small volume so the salts will be diluted and the sequencing should work well
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I am using Step One real-time instrument and the Green master mix with ROX.
Thank you all
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Vipul Batra thank you, I will check it.
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Hi,
I want to PCR a 200nts sequence from DNA samples that are extracted from fixed cells. For DNA extraction, first cells were lysed in lysis buffer overnight along with proteinase K treatment followed by DNA separation using phenol:chloroform:isoamyl alcohol (25:24:1) and ethanol precipitation. DNA yield and quality were similar to unfixed samples, which was satisfying, but there are no PCR products using DNA from fixed cells. Do you have any suggestions on how to prepare a fixed DNA sample for PCR? I will test whether fixing with 1.5% PFA resolves the PCR problem. Do you think using FFPE DNA extraction kits could help?
Thanks
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yes, use FFPE DNA extraction kits. If it does not work design two shorter amplicons.
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I have recently obtained a mouse model B6;129-OoepGt(139A2-3)Cmhd/Mmucd from MMRRC
I am having trouble getting our genotyping protocol to work and would like to find the sequence of the gene trap insertion in context of the mouse genome. In other words I want the sequence of the gene including the insertion.
There are elements which could be targeted for PCR such as the LacZ or Neomycin cassettes however from an academic viewpoint this sequence should be available no?
I have found the sequence of the inserted plasmid Gep-SD5 but I would just like to explain differences in PCR amplicons from what we expect. There are a bunch of links on the MMRRC page but I find myself lost.
Any help or pointers would be greatly appreciated!
-Nick
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Look like this (one of the links in the page you linked) has a description of the insertion site you can use to design your genotyping primers
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I induced two cultures at 0.5 OD600: 1 with just the vector, and another with my ORF in the vector. However, following induction with 1mM IPTG with shaking at 140 rpm at 37 degree C for even 6 hours did not induce protein overexpression, even in the culture harboring just the vector. The vector and the clone are both proper (as confirmed by sequencing, digestion profile, PCR screening).
Help and suggestions would be appreciated.
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Dewald suggestion likely works because there is residual lactose in tryptone and yeast extract. A couple of points on that. Different batches have different levels. If you add glucose you will suppress this induction since glucose prevents lactose uptake by the cells. This is the basis of the Studier autoinduction media. We had noticed that ClpP, under the control of the pET T7 promoter induced to extremely high levels in LB (no glucose but lactose) but there was no leaky production of the protein in fully defined or minimal media such as M9 (glucose and no lactose). Bill Studier correctly used this information and his long history with protein expression to realize that LB has lactose and glucose suppresses uptake and then exploited this to develop the autoinduction media.
On the BL21(DE3), for pGEX it is probably better to use the original strain BL21 (with or without pRIL) for expression. DE3 introduces the T7-RNAP onto the E. coli chromosome under the control of the lac promoter. Thus the DE3 will add an extra cost, producing a significant amounts of T7-RNAP, which is actually translated very well. This protein is not necessary for protein production from pGEX but is essential to producing proteins from the T7-phage promoter on the pET and related plasmids.
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I have GeNei PCR Master Mix of 2x concentration shall I dilute it to 1x before using for PCR or use it directly without any dilution
Can someone tell me what is the best way to use Master Mix
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(2X) x 10μl (2× master mix)/20ul (total volume) = (1X).
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Hi, i'm trying to cut the SARS-COV2 S protein from the plasmid pGBW-m4046887 to get een empty vector with small 6-His tag protein which I want to use for negative selection for SELEX.
I did PCR and now i want to make PCR clean up, then phosphorylate the ends and then ligate.
Could anyone correct me if my plan is wrong:
1. After PCR clean up meten de concetration of dna
2. Phosphorylate with T4 PNK in T4 ligase buffer:
100 ng dna
1 ul T4 PNK
2 ul T4 ligase buffer (10X)
milli Q water until 20 ul
30 min incubation , 37 gr. Celsius
20 min 65 gr. Celsius
+ add 1 ul ligase and incubate 2 hours at room temperature.
I'm not sure if could just add 100 ng dna, because i'm not quite sure how to calcultate the right amount.
The plan is to transform this vector to Neb-10Beta celles, isolate the vector and sequencing.
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Awesome!! I'm happy to hear that your experiments are working!!
Regards
Noe
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Hello!
I have black C57 DsRed & EGFP mice (from JAX) and need to quantify the expression of EGFP or DsRed at mRNA levels with qPCR. I am wondering if anyone knows about the primers?
Thanks !
P.S
I have the primers for the genotyping of these mice but I think those primers are too big for this purpouse.
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Hi Sina
designing primers for qPCR follow the same rules as for classic PCR, at the exception that amplicon must have 100-150bp length and one of the primer need to be across splice, at most 3' end of the gene..
take a look at the UCSC database, there is a genome browser and typically you can choose to make available the track presenting primers that have been designed and published on the gene (do not forget the rules for qPCR primers). after designing or choosing, you can also test them by in silico PCR on the same site.
all the best
fred
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Dears researchers,
Has anyone had the error below when calibrating the Step One Plus equipment (Thermo Fisher Scientific)?
"Spatial Calibration failed: Well locations are not evenly spaced.
System will revert to previous calibration.
Exit the calibration wizard and refer to the Hrlp to troubleshoot the calibration failure.
Error Code 1302"
I can't find the error code in the troubleshoot. Company support has not found a solution yet.
I already did the decontamination and the Backgroud calibration worked.
Regards,
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We also had a problem and after consulting with the experts of Kiagene Fanavar Company, who are based in our university, we finally decided to buy a MIC machine. I will send you the company link and email, maybe they can help you
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I'm doing 16s rRNA amplification using DNA extracted via the boiling method for several bacterial samples.
The quantities of each PCR reaction are;
DNA template - 1 ul
Forward primer - 1 ul
Reverse primer - 1 ul
Nuclease free water - 9.5 ul
Go-Taq Green master mix - 12.5 ul
5 ul of each PCR reaction is loaded into the 1.5% agarose gel.
Clear bands can be observed for 7 of the 9 samples, as well as the positive control. However no band is present for one, while another one is present but fainter than the rest.
What could be causing this?
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Some possible reasons for no band visible relate to there being no pcr product to see. This can happen with
1 too little dna in your sample.
2 poor quality dna with pcr inhibitors present in the dna that failed to amplify.
3 The failed dna is good quality but there is a polymorphism at the 3'end of one primer so one primer does not anneal so no amplification.
Similarly the faint sample can also be that there is too little dna or too much dna with pcr inhibitors present but in smaller amounts so there is some inhibition of the pcr reaction.
You can test for inhibitors by mixing a sample that works with one that does not work and if the pcr fails then the failing sample probably has pcr inhibitors. Alternatively try diluting your failed dna in water 1:2 ,1:4 and 1:8 and if the pcr works at higher dilution then the dna is probably dirty and contains inhibitors. Another possibility for dealing with dirty dna samples is to add mor magnesium as many inhibitors remove Mg from solution or to use a polymerase that tolerates dirty dna better
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I am trying to amplify ssDNA product using PCR. I am just wondering whether following the normal PCR protocol will yield a product since the starting DNA is not dsDNA. Have anyone tried amplifying a ssDNA or know a protocol that I should follow?
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Yes, it will work fine. This is what happens during all reverse transcription PCRs as the initial template (first strand cDNA) is single stranded. I would not skip the denaturation step because depending on the template, the ssDNA may form a secondary structure, which the denaturation step would melt.
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I bought a synthetic gene from a vendor, the gene is ligated in the pUC57 plasmid (with ampicilin resistance). the pUC57 plasmid have been tested using electrophoresis and it's all good.
I'm trying to transform it to the DH5 cells to store and propagate the plasmid, but it's failed (it's been 6 months trying). i have been using some methods such as CaCl2 and SMOBIO CK1000 transformation kit, heatshock. and using ampicillin LB plate (100 ug/ml and recently i use 200 ug/ml)
i have specific primer to detect the gene inside pUC57 plasmid, the colony i pick is shows positive result (PCR) but when i isolate the plasmid (from liquid culture using GeneJet Plasmid MiniPrep kit) and run it in agarose gel electrophoresis it shows smear (really thin) i also run it with positive control (also DH5 that have a known plasmid in it, so i think this could ommit the possibility of dnase contamination in the plasmid isolation kit).
does anyone have ever encountered something like this? or anyone have any advice? thankyou in advance:)
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Dear All,
The problem is fixed; it's caused by plasmid aggregation I guess
because when I try to linearize, it somehow shows a single band with the correct size in the gel.
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Hello,
I have a significant difference of the housekeeping genes between the control group and the rest of the groups in my PCR results. The reason for this is that I had to transcribe the control group individually (RNA -> DNA). Can I still evaluate the control group after normalisation or do I have to restart the experiments?
Many thanks!
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Dear Soner
That is the most obvious reason. I transcribed the samples in 2 rounds. The transcriptase kit had to be used in different concentrations than those from the original supplier, because the activity of the enzyme was too low. I therefore strongly assume that in the two rounds, which took place on two different days, the activity of the enzyme was different and therefore too much DNA was transcribed in the first round.
Best
Yasmin
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Hi,
I did this PCR using Platinum™ SuperFi II DNA Polymerase following the instructions in the protocol.
I tried multiple conditions (gradient PCR, longer extension time, less amount of DNA etc)
My expected fragment was around 250bp, and I did get it, however there is a double band that I have not seen before.
The band can be observed in all samples and I'm not sure if these are primer dimers or something else.
I've used the KOD polymerase before with the same primers and I haven't had these issues.
Any ideas?
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Thank you for your suggestions, however I don't see how there could be a problem in gel loading.
I've been preparing agarose gels for more than a year and I'm quite careful when I add the product into the wells. Can you please elaborate?
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I am looking for to buy a RT-PCR machine-96 wells. Please mention your experience with the machine in your lab. Any pros and cons. Thanks
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Lightcycler (Roche) outperformes CFX.
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Hello
I used 2% agarose gel 1X TBE buffer (pH ~9) for gel electrophoresis. For electricity supply I used 60V and 300A for 3m min. I used my known PCR product in this gel. After 10-12 min I found my PCR product in the gel. However after 30 min my product was vanished from the gel. I also attached my gel picture for your reference.
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Thank you so much @ Paul Rutland for your valuable feedback. I viewed the gel 2-3 minutes while taking the picture.
Your second suggestion was very helpful for my experiment and finally solved my problem. Yeah I sued 300mA current for gel electrophoresis and faced this problem. Thereafter I reduced current supply gradually and 50mA was giving the good result.
Thank you again.
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I have designed a primer set for LAMP and couldn't make it work. I am using NEB WarmStart® Colorimetric LAMP 2X Master Mix and tried different types of samples. Only thing I am worrying about is that my inner primers don't work. I know F3 and B3 works fine because I checked them in Real-Time PCR machine.
Also attached an image of gel electrophoresis of 3 positive and 3 negative control sample.
I would appreciate any help.
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Yan Liu I have the same problem. I am getting amplification in the Negative control. Is it also possible because of the cross-contamination?
If anyone is having any suggestions, I will be happy to follow.
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I am not getting any amplification for my target gene in real time PCR. The cDNA samples are fine since the amplification in positive control is fine. I have also obtained amplification for my target gene in semiquantitative PCR using annealing temperatures 60 and 62. Does any one has any idea where the problem is.
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If your controls are working well, it's possible that your gene of interest has a very low level of expression. What's the lower detection limit for your standard curve? You could try adding more cDNA, but it's not practical to do more than a 2-fold increase.
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I would like you to help me solve some doubts about the PCR conditions to obtain good quality products for subsequent sequencing. I am going to perform PCR targeting 16S, and I want to know if there is a range in the concentration of universal primers that works best (I usually use 0.4 uM of each primer) as well as a concentration of DNA to add in the reaction (I usually use 100 µM). ng) and finally if the appearance of the PCR product in the gel indicates that it is optimal for sequencing. So far I have obtained chromatogram results that are not very good, with overlapping peaks or longer sequence lengths than expected (extremes with low peaks). If anyone has suggestions I would appreciate them.
Sorry for my English
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The amount of primer used is not the most important factor in generating sequenceable template. The 2 most important things are that you have a single clean band to sequence and that all of the pcr primers have been removed before adding the single sequencing primer. I would sequence 6-8 without question and I think that the larger smear is so weak compared to the main band that it will still sequence well. Gel separation will get rid of the remaining primer...what method are you using to remove primer?. Can you attach an original .abi sequence file so that we can have some idea of the extent of the problem.
All sequences look longer than they really are in Sanger sequencing because small peaks of random electronic noise are interpreted as peaks after the end of the real sequence but they are low intensity so can be ignored.
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I want to add dATP on the ends of to my PCR product (Phusion, Thermo). My question is if it's possible with use of DreamTaq polymerase (Thermo)? I find informations about Taq polymerases but can not find out if i can use this polymerase to do that (a-tailing, not PCR itself, i got insert from PCR with Phusion polymerase).
Thx for help, cheers
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Yes the annealing temperature is often 4c below the melting temperature so you could try 40c and if this works then try 42c
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I have extracted DNA from fecal sample using salting out method.The concentration under nanodrop comes more than 100 and the ratio A260/A280 ranges between 1.5 to 1.9 whereas A260(10mm) is morethan 5. The gel image has been attached here also. Can somebody explain what could be the possible reason? (Note: Primers, PCR master mix, PCR conditions are fine using control samples)
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Hi.
The protocol that you are using...has it worked before?
You have to start by elimination and testing various variables such as working with different DNA concentrations....HOWEVER, the major issue in a PCR not working is usually the primers...Thats the first thing you need to address.
Good primers are key, and should allow the PCR to work over a variety of conditions..so start with that...and never give up. P C R- Please Continue you will get it Right.
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Instead I am getting a smear with a brighter illumination near the well.
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Does your dna amplify with other primers? You may be adding too much dna or poor quality impure dna. How much dna in nanogrammes are you adding and what is a typical OD269/280 ratio for your dna?
Your annealing temperature may be too high and only one primer is annealing.
You may be running the gel at too high a voltage and the gel is overheating and the dna is smearing. Run the gel slower. There are other possibilities but without more detail about reagent concentrations and cycling parameters troubleshooting is mainly guesswork.
Your gel does seem to haave run a very long way.What are the time,voltage and current of your gel running and does the gel or buffer feel warm after running it
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In my experiment, I use a plasmid (~16kb). What I want to do is to delete Luciferase gene from that plasmid by using a specific pair of primers with PCR technique. Last time, I tried and I got no band at all. So I wonder that if it is possible to extend the whole plasmid except Luciferase gene by PCR as the plasmid size is pretty big. Thank you.
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As previously stated, restriction digest would be the easiest way to do this. Otherwise a poymerase with proofreading activity should be used to avoid mutations in the amplified fragment.
I do not know what you intend to do with the resulting plasmid. However, if you do not have suitable restriction sites available to cut out the whole luciferase gene from the plasmid, maybe cutting out a part of the gene resulting in loss of function might be a way to go.
Otherwise you can optimize you PCR reaction by modulating the MgCl2 concentration and try a gradient to find the optimal conditions for your pimers or insert different amounts of template (dilutions).
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Dear all,
I am planning to do in vitro transcription from a PCR template. However, I need a poly(A) tailed RNA product. If I add 30 nucleotide poly(A) tail before the in vitro transcription, would it stop the T7 polymerase reaction earlier? Thanks
Cheers,
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No. Transcription doesn't use stop signals like translation.
Translation is simple: UGA, UAA, UAG = stop.
Transcriptional termination is a fairly unwieldy process with multiple different methods employed in both eukaryotes and prokaryotes, often involving stem loops and termination factors and a lot of mess, and of course in euks this is then followed by cleavage at the polyA site and addition of a polyA tail by a polyadenylase.
If you just...add the polyA yourself, you short-cut all of this, and your transcription reaction is terminated by the very straightforward mechanism of "T7 reaches the end, and then falls off".
So no: it'll be fine. Just make sure the polyA is on the coding strand.
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I'm trying to look for a specific field which people makes fungi molds to identify the species. I'm planning to make it as a machine learning project. Like in the field of agriculture, medicine. Thank you
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If the purpose of research is just to identify the fungus,e.g Aspergillus or Macrophomina, you use microscope to tell you, but when you have a new strain of same fungus, that case you need PCT to identify nucleotides added or deleated off this strain. Regards.
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Rt-pcr related question
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I think this is not essential, but you need to use an optically transparent film to seal the plates when working with real-time PCR. Good luck!
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I want to expose my lung epithelial cells to different treatments in vitro and then detect pro- and anti-inflammatory cytokines/chemokines in the conditioned media - can I then do PCRs on the conditioned media? Any guidance would be very much appreciated!
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PCR is used to detect nucleic acids, not proteins. Generally, you would use either a Western blot or an ELISA to measure the amount of a secreted protein in cell culture media.
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Real time PCR related question
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Yes. The dye is made for qrtPCR. According to Promega, the dye has very similar spectral properties to those of SYBR Green (https://www.promega.com/~/media/files/products%20and%20services/islides/is022-gotaq_real-time_pcr.ashx). Any instrument capable of extiting and measuring Fluorescein and/or SYBR Green should work with BRYT Green as well. To my knowledge, this applies to all available qrtPCR instruments on the market.
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Hello I need help putting together annealing buffer. I was given the following:
H2O supplemented with 10 mM Tris-hCl pH 8.5 and 50 mM NaCl
We currently have 10mM Tris-HCL, nothing higher. Therefore I am worried that when I put together the buffer, the diluted Tris-HCL will affect annealing.
Would my annealing reaction still be okay and should I adjust the concentration of NaCl?
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You may add solid NaCl.
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I designed the primers by myself (with a Tm of 57.3 and perfect binding to the template, checked for no secondary structure) and the expected amplicon size should be around 7.0kb, but I could not get a 7.0Kb product at the end of each PCR, instead it was a band that did not seem to look like a PCR amplification.
The ladder used was a 1KB ladder. The DNA in the sample was extracted and tested by Nanodrop and the A260/A280 was 1.92, giving a DNA concentration of 1.3ng/uL. I referred to the paltium superFi ii handbook and used a 20ul PCR protocol (10ul mastermix; 1ul of 0.2um forward primer and reverse primer; 4ul of DNA template; 4ul water).
PCR run times were also referenced from the handbook recommendations. But every time the amplification came out strange and I set Negative control (no sample added) and still got the same result.
The attached pictures are my electrophoresis results, well1 and well8 are 1KB ladder. well 2 is undiluted sample(4ul——1.3ng/ul DNA concentration), well3 - well6 are PCR for samples diluted 10, 100, 1000, 10000 times respectively. well7 is negative negative control (no DNA template added). Please give me some suggestions, I have tried many times with this result
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Hi Rita
The DNA concentration actually is low. I do not know what is your sample is. If the extraction is done properly and you have any positive data that your target is there like "Positive PCR from the same sample with another primer", My suggestion will be:
1- Increase the sample volume make it 8 ul instead of 4 and remove water.
2- Be sure that extension time is enough (at least 4 minutes).
3-Increase your volume to 50 ul instead of 20 ul.
4-Try to have a positive control sample
Add the used thermal profile also and the type of sample, to get better help.
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Hi there, thank you all in advance for your help!
I'm having an issue with my bands being ~500bp too short in the second round of nested PCR.
Here's my protocol
First round of PCR
-amplicon is ~2.2kb
-40cycles of PCR
-I get very clean bands with when there is sufficient template DNA. For samples without extremely low levels of template DNA, I move onto nested PCR
second round
-amplicon is ~1.8kb
-30 cycles of PCR
Explanation of gel below
Lane 2 of the gel below is the expected length of the amplicon for these primers. In this well, I use my positive control template DNA that did NOT undergo a first round of PCR. lanes 3-11 all are a 1/10 dilution from the previous PCR reaction. Lane 3 is the positive control from the first reaction.
My problem/question
For lanes 3-11, the band is about 500bp too short. I can't figure out why... Your help is greatly appreciated
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Hi,
you can see that there is a 2nd band for the first samples in the right size. I would envision, that the 1/10 dilution is not sufficient and this might impact the reaction kinetics here. I would go for a higher dilution and potential run a gradient. What are your PCr conditions? And which reagents are you using?
Sven
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Hi all,
1. How do you identify if you have bubble library or just insufficient size selection?
2. How to deal with bubble library? Would reconditioning PCR work? Is it recommended to use single primer or both forward and reverse primers?
3. How to quantify bubble library for library pooling? qPCR? How?
Thanks.
Lux
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Heat the libraries for 3-5 mins in 90'c and recheck the QC. Concentration will decrease if it is bubble libraries.
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I’ve been extracting very low copies of viral DNA and RNA and performing a PCR afterwards to detect what I extracted. When I run PCRs for viral DNA, the PCR always works and my negative control is always negative. But for viral RNA (converted to cDNA), the negative control shows a band when I run an agarose gel. I fixed this problem a few times in the past by changing primers but now if I change everything the negative control still has a band.
The negative control band is the same size I expect from my samples meaning that there’s template in my negative control. I also ran a qPCR and the melting peak in the negative control is also what I expect from my samples.
I’ve tried changing the polymerase, water, primers, lab coat and using 70% ethanol but I can’t seem to find the source of contamination. I’ve changed my pipettes, tubes and bench but that didn’t work. I ran another PCR without any positive controls (to avoid cross-contamination) but my negative controls still show a band.
Any suggestions on how to fix the problem would be appreciated. Thanks.
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This usually happen due to amplicon contamination. The huge amount of PCR product make it difficult to completely get rid from the contamination. A rapid solution is to change your primers.
Ethyl alcohol does not work with DNA. Bleach may help or DNAse.
You should keep the area where you prepare your mastermix and primers (Pre PCR area) away from PCR and Post PCR area.
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Dears, I am doing qpcr of genes with cdna made of RNA from adipose tissue. In the efficiency curve of the primers, the cdna serial dilution does not promote adequate amplification. The less diluted points dots are having less amplification. Example: in the photo, the last amplification is the highest dilution (1:2). I believe it is the presence of an inhibitor. Has anyone dealt with this? Know what to do? The 260/280 and 260/230 ratios are fine. The profile on the electrofluoresis gel is also good. The RNA was extracted with silica glass beads (sterilized and washed with acid), Trizol according to the manufacturer, and then the material was washed with precipitation with 3M sodium acetate and 100% ethanol. Resuspended in nuclease-free milliQ water.
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I find a lot of people don't realise quite how dilute cDNA can afford to be.
cDNA synthesis buffer itself is inhibitory to PCR (it contains DTT, for example), and you can also inhibit your reaction by saturating the system with too much target (or too much non-specific target).
In almost all cases, cDNA should be diluted prior to qPCR.
I typically dilute all my cDNA 1/20 (so for a 20ul prep I add 380ul water, giving me 400ul of final cDNA), and using 2ul of this per reaction is _plenty_: it allows me to reliably detect even low abundance genes.
For reference I use ~1600ng of RNA in a 20ul cDNA synthesis reaction, so 2ul of diluted stock is ~8ng of cDNA, assuming 1:1 conversion.
In your case it may be nothing to do with the lipids and everything to do with just...generally using too much cDNA.
When running dilution curves of cDNA historically, I expect the extremes at either end to be unreliable: with too much cDNA you have all the issues noted above, and with too little cDNA you have stochastic effects and thus highly variable data. This is FINE. The whole point of a dilution series of cDNA is to establish the range over which your reaction is linear and trustworthy.
Find that range, use it, don't worry about the extremes.
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I have one PCR workstation.
I performed my RNA extraction/cDNA synthesis on a separate bench (outside hood).
I now have very concentrated cDNA.
In previous labs I had 3 work stations.
1: Prepare primer/probe working stock
2: Prepare serial dilution of concentrated template
3: Preparing master mix and plating serial dilutions
I now have one workstation.
Would you perform you cDNA dilution outside of the workstation?
Or would you do it inside and just turn on UV light/spray DNAzap before plating?
Any advice on PCR workflow would be useful.
Thank you!
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Hi, I'm trying to make random mutagenesis on my interest gene. I want to use error-prone PCR. But I only know how to amplify and set condition for best mutagenesis rate. I wonder how can I ligate these products after amplification to my desired plasmid? Can I use infusion or just ligation by T4 ligase?
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Hi, you can check the method published in this paper.
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In the last few days it was not possible to access Miner website. Does anybody knows what is happen with the website?
Thank you!
Camila Carvalho
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Thank you very much Sheng Zhao
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Why hybridization based targeted exome sequencing library comes very high concentration that is 53ng/uL (expectation 1-5 ng/uL).
No change in the PCR cycles and master mix
No change in Beads concentration during clean up step.
Also no change in elution volume.
Your kind answers are highly appreciated.
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Did the hybridization conditions change?
I had an experiment where the hybridization temperature was too hot. The sequencing library had a lower concentration than expected.
I think the opposite could be true in your case. If the hybridization conditions are not stringent enough, there will be more off-target DNA. The starting DNA concentration in the PCR reaction will be higher. After PCR, the final library concentration will be high, even if the PCR cycles are the same. This is because of the extra off-target DNA.
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I have ran and reran this PCR so many times and am at a loss for what to do at this point. I started by cutting off a small piece of ear (about the size of rice) using a razor blade and tweezers. I used 70% ethanol spray to sterilize by tools between ears.
I then performed an extraction using chelex beads. 5g of beads in 50mL of nuclease free H2O. I used 100 uL of beads per sample. I used a shaking heater block set at 95C and heated the samples for 40 minutes vortexing every 10 minutes. I then centrifuged the samples for 2 min at 20G. I pipetted off the supernatant leaving the beads and the remaining ear behind.
I then ran a PCR. I used 1x PCR buffer, 1.5mM MgCl2, 0.3 mM DNTP, 0.25 primers, 1U taq, and 1 uL of DNA.
That extraction method didn't seem to be working so I used Platinum Direct PCR Master Mix to extract DNA from the ears. I cut a small 1cm piece of ear and added 20uL of lysis buffer and 0.6uL of Protinease K to each ear. As per the instructions I heated the samples at 98C for 1 min then centrifuged them at 20G for one minute. I pulled off the supernatant and used 2 uL to run a PCR. That seemed to work but I saw lots of high MW products so I centrifuged again at 20G for 20 minutes and ran the PCR again. My bands dissapeared.
So I decided maybe I needed to lyse them for longer. So I repeated the extraction but heated for 30 minutes and centrifuged for 10 minutes. This didn't work and it looked like the DNA was trapped in the wells of the gel. The guide for the kit suggested adding 1uL of Protinease K post PCR. So I used the same DNA and ran another PCR but added protinease K following PCR. This didn't work either.
I am not sure what to do at this point. I have attached the PPT that has all the gels I've run and the conditions.
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I don't have an answer to your question, but I am having a similar experience. I work with grass (Miscanthus) in a plant breeding lab. I also use the Direct PCR kit. I have been working with a new vector and when I run PCR on my samples (bacterial plasmids and grass lysates), the results are inconsistent. Bands will appear on the first run and then they disappear/show up randomly in subsequent reactions and gels. Maybe it is a problem with the kit..... I don't know.
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I am seeing multiple bands in my PCR reaction. Is it possible that my stock is contaminated or issue with a gel run?
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or simply the primers are not specific enough to get only one amplification target.
try an in silico PCR (at UCSC or NCBI) to get sure yours primers are optimized.
all the best
fred
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Direct PCR lysis buffer is one of the fastest methods to do genotyping. By taking a piece of tissue and placing it in a small volume of Direct PCR lysis buffer you can then take a few microliters and do a PCR directly. I would like to do a similar approach but performing a restriction enzyme digestion instead of a PCR. The issue is that the DNA is in a buffer of unknown composition and I am not sure whether this will affect the restriction enzyme reaction. Have anyone experience with that? Thanks! Miguel
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If the buffer composition is similar to the requirements of the restriction enzyme, it might work, this is rather easy to test.
As you are lysing tissue, I'd expect there are proteases like e.g. Proteinase K, which usually require a heating step for inactivation. Otherwise they might chew up your restriction enzyme.
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Hello everyone
Is a column extraction kit superior to nanomagnetic beads extraction kits???
What are the advantages and disadvantages of nanomagnetic beads extraction kits compared to column extraction kits????
Do magnetic beads extraction kits interfere with the PCR process???
Thank you in advance for your help
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The major issue is the surface composition of the resin or beads. non-specific binding affects all solid phase separations. This is why we block ELISA plates with protein BEFORE adding the sample to the well. The same issue plagues both column resins and magnetic beads. The more bead surface per unit of sample, the more NSB will affect your results.
The next issue to consider is diffusion to the capture surface. The diffusional path in columns tends to be shorter than the interbead distance in a batch of magnetic beads spinning in a beaker. Therefore, a column format tends to be faster than a batch capture.
So you have a trade-off: More NSB with the column and slower capture with the beads. Pick your poison.
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My Melting curves in qpcr look like this and have among other plates as well little peaks after the first one. I am not sure what could cause this as Ive ruled out most technical issues. Could it just be due to operator errors or what might cause this. I will drop some examples below so any help would be appreciated thanks.
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