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DNA Amplification - Science method

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How realistic would it be to amplify a 15Kb plasmid with no errors?
We are currently trying to amplify a plasmid needed for transfection. The problem is that the yield from ecoli amplification is poor since the plasmid is low copy. We have tried chloramphenicol in our culture and it helped somewhat but still not close to what we need. We want to try the PCR route, but I have read that it can be quite tricky. Any suggestions on how to go about this are appreciated.
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To amplified your plasmid : make the culture to 32°c with 12,5 ug/ml chloramphenicol. Strech your bacteria and pick one colonie to start your 400 ml culture.
For the extraction resuspend in 30 ml of 50mMTris/HCL ph8 10mM EDTA 100ug/ml RnaseA, then pour 30 ml of cold 200mM NaoH (fresh) 1% SDS. Mix gently until you obtain flakes. Leave on ice 2mn. Spin 30mn 12000g. 4°c with angle fix rotor. With the supernatant fill 4tubes (50ml) with 20ml/tube
Add 14ml propanol-2 (0,7vol). Mix several time, leave on 10mn. Spin again
30mn 12000g. 4°c. Wash the pellets with 5ml EtoH70%, spin again. After remove alcool and dry at RT. Dissolve the DNA pellet in 500ul TE/RNASE 20ug/ml, wait 5mn at RT. Precipitation : add 50ul (1/10 Vt) of 3M NaAc pH5,2 and 390ul propanol-2, mix gently until you see a medusa . Pick up with a tip. Wash with ETOH 70%, dry and dissolve in TE. This 2° precipitation allow to separate RNA from plasmid
I use tRNA as a carrier in the first step. With a kit you loose a lot of plasmid that stick to the membrane
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The purpose of denaturation is the break down of dsDNA, so what is the need of intial denaturation while denaturation can do the same.
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Use of the initial denaturation conditions for all cycles can result in deactivation of the polymerase (depends on type) which is why there is a separate initial denaturation stage for standard PCR conditions. The initial denaturation is sometimes carried out at a higher temperature and for longer than the melting carried out during subsequent cycles. This is because some templates can have a high GC content or secondary structures (like hairpin loops) which inhibit the PCR reaction. By having a hotter/longer initial denaturation it is possible to separate the DNA strands enabling the first round of PCR to take place generating shorter/less complicated products that can then be melted under milder denaturation conditions.
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Hello everyone!
I am using AB Stepone system with Qiagen QuantiNova PCR kit. After doing my first trial, my qPCR results were very strange, the plot are "hook"-shaped (attachment). Anyone know what happen? The machine I use is not calibrated (the last calibration was around 2012~2013), will that cause any problem like this?
Our plate centrifuge are out of order and I cannot centrifuge properly, but no visible droplets remains on the wall. There are still some bubbles on the top liquid surface, is it a possible cause for my results (I thought the bubbles will be removed upon heating)?
I am doing a quantification experiment with 5 standards (300,000 copies, 1:10 diluted to 30 copies). I am new to qPCR and really confused by the result, please help :(
Thank you for your help!!
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Hi there! I don't know if you solved this already, but just in case I write. I had recently this problem and came to this post to try to find a solution, I found it elsewhere in the end, so I am writing for all the future newcomers as well. The problem with this is that the machine is reading the ROX value on your endogenous control. If you didn't add ROX to your samples (like me), this makes the machine subtract the values of your endogenous to all the samples, resulting in no amplification. Go to "set up" and, on the "assign targets to wells" tag, select the endogenous and select "none" in the reference. Then go to "analysis" and click on the "analyze" button. Magically, all your amplification curves will suddenly appear!
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Component: Pure water, Bsm buffer, Bsm polymerase, primer, MgCl2, dNTP, DNA Salmonella
Heat 60 Celcius 1h and 80 Celcius inactivation 10min
Sometimes the results from gel electrophoresis appeared DNA bands but sometimes less band intensity. What happen? Not suitable primer or expired chemical?
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@Chayanit Thairat, Thanks for the protocol. I wanted to know if your are following a published protocol, then you may be able to compare your results with the reported observations. If you are optimizing your protocol, some parameters may have to be adjusted to get the desired gel profile
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Analyzing qPCR (Sybr-green) results from one of my last experiments I found that for few samples there was no amplification curve, however specific melt peaks and sharp bands on the gel were observed. What could be the reason for a lack of amplification signal (curve)? Is it because of low concentration of DNA (I used lysed bacteria as a template, so the concentration of DNA was unknown ) or technical problem with the sensor of the machine (Bio Rad CFX96)?
I would be really grateful to have suggestions from you!
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Hi Lubomir,
Can you explain more?. I'm facing similar issue at the moment.
Thank you
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When run a LAMP reaction, sometimes false positive appeared in agarose gel electrophoresis. What are factors to create contaminations to false positives? What contaminations do in LAMP?
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Are you sure that you did not have any smpling problem such as insufficient amount of primer or different sources of genome! If not, you might optimiza the amount of enzyme, temperature and Incubation time. Also, i highly reccommend you in order to observe better amplification in LAMP assay do not forget using betain and MgCl2, I used it, it works😊
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I've been trying to amplify a plasmid that I'm running low on, but every time I complete the purification step and try to run the products on an agarose gel, the samples float out of the wells. I've tried making sure to remove any remaining wash solution (containing ethanol) but that did not help, and I've run the samples on a nanodrop to make sure they contain DNA.
I've heard that using a glycerol-based loading buffer can help weigh down the samples on the gel, but I was wondering if I could just add glycerol directly to a pre-existing loading buffer (this one here specifically: www.neb.uk.com/products/neb-catalogue/other-products-(cloning)/gel-loading-dye,-purple-(6x)).
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You should prepare the DNA loading dye in-house. The gel loading dye has to be mixed with the DNA samples before loading in the wells for use in agarose gel electrophoresis. Each component of the gel loading dye has a purpose.
Gel loading dye contains xylene cyanol, bromophenol blue and glycerol.
During agarose gel electrophoresis, xylene cyanol makes the lower dye front and is often used as a tracking dye during agarose gel electrophoresis. It has a slight negative charge and will migrate in the same direction as DNA, allowing the user to monitor the progress of molecules moving through the gel. The rate of migration varies with gel composition.
The bromophenol blue makes the upper dye front. These two-dye front help the user to monitor the rate of migration and to prevent the over-running of gel. In addition, sample loading to the gel is much easier with colored solutions.
Glycerol present in the loading buffer is used to increase the density of a sample so that it will layer at the bottom of a sample well.
So, you need to prepare the gel loading dye and mix it with the DNA samples properly before loading in the wells to get a good electrophoretic run. I have attached the recipe for preparing 6X DNA loading dye.
The pre-existing loading buffer which you have mentioned from NEB cat# B7024S already contains Ficoll which performs the same function as glycerol namely, providing density to the DNA sample. Hence, DNA samples will not come out and diffuse in the buffer. It makes DNA settle at the bottom of the well. There is no need to add glycerol to this loading buffer. You can use this product directly at a concentration of 1X by mixing with the DNA samples before loading on the gel.
I would recommend you prepare the 6X DNA loading dye yourself if you have all the necessary chemicals namely, xylene cyanol, bromophenol blue and glycerol in the laboratory.
Good Luck.
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I'm trying to prepare the LAMP reaction without the Loopamp kit, with the components separately. I am having many problems because sometimes I get positive results and sometimes, in the same conditions, I don't. I changed polymerase (large fragment, 2.0 and warmstart), water, primers, Buffery and temperature. Does anyone know what can be my problem? Is the order of addition of reagents important?
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There is no ladder like pattern but two or three bands were obtained near the wells along with primer dimer at the base in the positive control. Earlier, the results were clear cut. I have changed all the reagent, amplification conditions, volume of template DNA, etc. though nothing helped me out. Pl suggest what should i need to change
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I have a DNA sample which in previous step induced Brij 35. can I directly input this sample to PCR reaction? Will existence of Brij 35 interrupt the taq enzyme and the PCR reaction?
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I am usiing LAMP kit, specifically designed for liriomyza huidobrensis.
I have a random samples of differet insects and LAMP seems to amplify insects from other Genus as well. Although, other species from liriomyza are not being amplified which is what I want but why other genus get amplified and annealed.
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Syed Rizvi I recommend with their answers.
1. I would suggest that you firstly align all your sequences (target seq, same genus and other genus) through a multiple alignment tool (I use MEGA X). This will help you to see which area that your primer is targeting.Perhaps your primer is targeting the conserved regions of the gene which maybe similar in all these samples.
2. You can also redesign your primer (I used PrimerExplorer) to target another sequence.Do make sure to always cross check your primers through BLAST to ensure that only your target samples are amplified. I would suggest to re-design your primers if a majority of your primers amplify other non-target species.
3. It is also possible that during the experimentation, cross contamination occurred causing false positives. LAMP is highly sensitive to cross contamination.You might have carried over DNA from your previous assay onto your non target species.Ensure your workspace is always sterilized, and you should make sure to open the reaction tubes at a different place compared to the incubation area to prevent aerosol contamination.
4. Lastly, the false positives can be contributed to high reaction temperature or high primer concentration (as mentioned above) . Do try to adjust these parameters and see if it helps.
Goodluck!
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Hi everyone,
I need to clone in pGEM-T vector the result of amplifying a metagenomic library by PCR with a high fidelity polymerase but I only want to clone the fragments of an specific size (1-4kb). The rest is not interesting. I was thinking about:
-Purifiying this PCR product
-Do a A-tailing
-Run an agarose gel to cut the band of the desired size
-Perform pGEM-T clonning
I am not sure if A-tailing will survive the whole gel process. Any idea or help with this?
Thank you so much beforehand.
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Yes, A-tailing will survive after electrophoresis. Purify the PCR product and perform the A-tailing reaction at 72 degree for 10 mins. Perform ligation with pGEMT vectore only after purification.
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Dear researcher,
I am extracting DNA from IV plants which contain a lot of phenolic compounds and secondary metabolites, due to which the DNA extracted always contains impurities (I tried several ways to remove them).
Please tell me the ways to either purify the DNA or such PCR conditions/programs which could amplify the DNA.
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I'm not sure the methods you have used for DNA purification, elaborating a bit would help us in finding a solution. I assume you're using phenol chloroform extractions? I can advise you to be very careful in not disturbing the aqueous-phenol interface when removing the aqueous phase. Don't be greedy, it's okay to leave some of the aqueous phase behind. If that doesn't work you can try commercial DNA purification columns, but I would advise trying the phenol chloroform extraction again as the DNA purification kits can be expensive. Even if the plant has lots of phenols they would be trapped in the organic phase and with proper centrifugation should not massively contaminate the aqueous phase. The problem with PCR is that your phenol contaminants can inhibit the reaction, so having pure DNA is essential for PCR.
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When I use RPA (fpg kit), I see a lot of primer noise on a gel. I need a clear band for a melting curve analysis. I tested different combinations of primers with litte result. If you have any suggestions on how to avoid this, please let me know.
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Yeah, SAMRS primers do cost a lot. The primers I used in my current assay (TwistAmp Basic Liquid) was designed by Primer-Blast with some parameter modification, however, primer dimers still appeared.
Recently tech support of Twist Co. provided a primer design software for RPA primer design, named "PrimedRPA". I'm thinking if this could solve the dimer problem. While this need a python backgroud......we do not have the technician. If you do, maybe you could try to design a few primers to run a test.
The primer dimer is also a big problem for our team, if you have the method to improve this situation, would you like to share it?
Thank you!
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My lab is working on a simple library prep and are running into an issue with our amplification PCR, our samples do not successfully amplify. After preforming the reaction (using 2X Phusion HF Polymerase, water, and our primers) for 12 cycles, the samples very low concentrations according to our qubit. In addition, when run on a gel we see nothing (including no primer dimers).
However when we use this product and run a standard QC PCR, the DNA amplifies successfully and can be seen on a gel.
For trouble shooting we have tried running the reaction with KAPA HiFi HotStrt ReadyMix and running the reaction for both fewer cycles (in case it was being overloaded) and for a greater number of cycles.
Has anyone run into this problem before or have other suggestions for trouble shooting?
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The primers that we use in the pre-hybridization PCR are the same ones that we use in the QC PCR. While we do not get amplification in the pre-hyb PCR we do in the QC PCR indicating that they are working.
The pre-hybridization PCR is run with water, 2X Phusion HF Polymerase, our primers and our template DNA. The QC PCR uses product from the pre-hybridization PCR (the same DNA) and is run with the same primers but using taq, PCR buffer and dNTPs.
It seems that the Phusion may be the problem however I ran the same reaction with KAPA HF HS in place of Phusion and saw the same results.
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The size of mature miRNAs range between 22 to 26 bp. So, to design primers for the amplification of these sequences by using traditional (general) primer designing protocol is not applicable because PCR methods require a template that is at least twice the length of either of the specific forward or reverse primers, each typically ∼ 20 nt in length. Thus, the target minimum length is ≥ 40 nt, making miRNAs too short for standard RT-qPCR methods.
So stem loop primer is the way to achieve the task and I used several tools for the same including miRNA Primer Design Tool.
In this case forward primer may include mature miRNA itself and may vary, but in most of the cases reverse primer is always found as universal primer, can anyone explain this to me?
I would also like to know if, to design primer for pre-miRNA is not beneficial over mature miRNA sequence? 
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Reverse primer remains same ,it is complementary to a sequence within the RT stem loop Primer.
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I am working with fungal DNA amplification from plate culture at ITS and 28s rRNA regions by using ITS1/4 and 5.8SR/LR7 primers. from previously, I have done to amplified the fungal DNA from both regions before and now I got the problem with my pcr amplicon from ITS region. Because at the same sample and same times, I got smeary bands from ITS unlike the results from 28s. I attach a photo of an agarose gel in order you could visualize those weird smeary bands! Thank you in advance.
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is one of these samples a no template control sample?
These smears may be contamination of the reaction mix with previously amplified material. This will then amplify very well and become more concentrated than the primers and will then join with each other to form randomly long concatomers of amplified product. Can we see apicture of the amplimer with at least one sample being water not dna in the reaction mix please?
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Hello I am trying to amplify a stretch of Double stranded DNA (of 1.4 Kb) using A forward primer of 112 base pairs and reverse of 21 base pairs. The Tm value of the 112 basepairs long Forward is 91 deg. and that of 21 base pairs in 63 deg.
My question is which strategy would be better so that the 112 basepair long forward will get annealed properly. along with the 21 basepair long reverse.
and secondly i would like to know if I initially run the PCR at a temperature where only reverse will anneal, and later add the forward and increase the temperature nearly to 80 deg so that i get a full stretch.
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The difference between forward and reveres primer should be within 3 degree to done PCR, but if exceed you go to touch down PCR, But I think the best choice involve redesign primer and if you obligate to this product size the better way using nested PCR
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I have amplified a circular DNA viral genome using rolling-circle amplification (RCA) mechanism. I want to do complete genome sequencing of the virus directly without cloning. Can some body assist me? I don't know whether :– I need to purify the amplified products, which method/system of sequencing is appropriate and the primers to use in sequencing since RCA does not use primers.
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by using restriction enzyme you can do sequencing according to yr desire size of virus DNA.
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The pet28 is a very low copy plasmid. I cant get a pellet in isopropanol step. I have to use a mini spin kit Qiagen a to do fraction of 10 ml.
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I raccomend to use the omega biotek mini kit2 starting from 10ml of e.coli culture grown o/n or their midi kit from 100ml of lb culture. In my experience omega biotek kit ( supplied by vwr) supply from 2 to 4 higher dna amount respect the same kit of qiagen, invitrogen, promega or mchanagel.
Manuele
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I'm currently trying to run MNase digestion assays using zebrafish embryos. I have all my protocols down and working well, only problem is I don't seem to be able to quantify the nuclei/DNA so that I can normalise across tubes. Most papers/protocols seem to state that they quantified using A260. I've tried taking sub-samples of my resuspended nuclei and quantifying at A260 on a nanodrop, but don't get any consistency. There is also a lot of interference from other components. I also tried lysing nuclei in the sub-sample to make the DNA more accessible, but still problems. Any help or advice would be much appreciated.
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i just have the same problems.
Did you manage ti resolve it
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I am amplifying a fragment from COI gene of tiny fig wasps. When I used a new BSA solution (50 micrograms/ microliter) that I prepared, I noticed a white jelly-like precipitate in the tubes in which I used it. Amplification was fine, the band is ok and bright. But what should I do with the precipitate? I need to sequence those PCR products ! 
Thank you very much for your suggestions, 
Kindly 
Paulina
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Thameen's link suggesting that the residue is pyrophosphate is good and the idea is testable. Just run 2 pcrs with and without enzyme and if the enzyme is producing pyrophosphate only one tube will precipitate. See if it centrifuges down to get rid of it. Another possibility is too much BSA congealing like cooked egg white. Chech your dilutions of the new BSA since it did not happen with the old BSA there may be a dilution error here.
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Hello, I am using chickpea DNA to amplify a gene of size 1041 bp and I have a problem in the amplification. There are many repetitive bases in the starting and end sequence of my gene. Like ATGAAGAACAAAATATTATCATCAT is the starting sequence where you can find AAAAT in that. The end sequence is TTCTCCTCTTCCACCCTGCAAAACT and when you reverse compliment it for reverse primer you get TTTTG, which is 5bp compliment between both. so i tried with KOD High Fidelity , with DMSO and I am not unable to amplify it. But when I delete the last 6bp AAAACT and make a reverse primer without it, I am getting amplification very easily. 
My question is.
1. I need full gene for cloning and sequencing
2. Is there any method to amplify this kind of gene without deleting sequences.
3. I have to clone and sequence it, what is your suggestions. 
Thank you . Please suggest. 
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you could try amplifying first with the short primer that works then a reamplification with the longer primer when the increased amount of template means you can get product with a less efficient pcr. 10-15 cycles of a 1in 1000 dilution of first round pcr should be wprth a try
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We have been cleaning genomic DNA after a WGA step with the Zymo kit and the samples showed nice clean bands of high molecular weight on an agarose gel immediately after the DNA cleaning. We then kept the samples 2 days in the fridge before storing them at -20C. One week later, we rechecked the samples on agarose gel for selection of candidates for NGS and all samples were degraded - showing a smear instead of the earlier nice clean bands of high molecular weight. The only explanation we can see is that the Elution buffer was contaminated with a DNase (the eppendorf we are using are RNAse and DNase free).
Any recommendations for alternative kits that will not destroy my extremely valuable DNA?
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Actually, our lab is struggling with DNase contamination too & we also use the zymo spin columns. Is the degradation consistent with each sample? or is it more sporadic where one sample will degrade and the other will not? 
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I'm testing Hidroxynaftol blue like dye for DNA amplification type LAMP. However, I didn't get a significant difference between positives and negatives results using the concentration more recomended by papers (3 mM). If anyone is using this dye, please, I need advise about how it use.
Thank you!
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Thank you for the answers. Now, I gonna try to test it. 
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Hey guys, I'm looking into getting a clear and unique band of DNA >40kb or higher for Pacbio genome sequencing extracted from fresh blood. I know Quiagen and some other companies have kits you can use, but in my experience the columns break the DNA so that you get a smear of smaller sizes too.
Does anyone has experience and can recommend an ideal extraction method which yields mainly larger DNA fragments (>40Kb) ?
Thank you 
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Thanks a lot guys! It's been really helpful!
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I did PCR to amplify BCR-ABL genes and used three gBlocks as my positive controls. I used MyTaq HS Red Mix from Bioline. My sample generated a band representing internal control but all the gBlocks showed no band. All of the gBlocks contained 10^3 copies DNA. I wonder if anyone have experienced this kind of case because it's rare to happen that gBlocks are not amplified while the samples are just fine and amplified since the DNA amount in gBlocks are high (10^3 copies).
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I've experienced problems with gBlocks before. Apparently, IDT only guarantees that a certain percentage of the DNA is your actual sequence. In other words, the sequence you designed and ordered can be as little as 40% of the actual DNA. On the other hand, IDT has a very good policy or reorders/re-synthesis. So what I did was call them, explain that the gBlock was not working as it should (and I had many previous orders where it worked fine) and that I though that maybe something was wrong with that particular gBlock. They sent me another tube free of charge.
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Hi,
I'm trying to amplified and get the sequence of one gene (4kb) in 10 sorghum accessions and I divided in 3 fragments the gene (I fragment: 1.3kb, II fragment: 2.7kb and III fragment: 0.8 kb). I could amplified and get the sequence in the las two fragments but the first fragment I couldn't (it was possible just in one accession). I try difference set of primers, change the PCR conditions, cloning ratio. However, I can get a specific band with the correct size but at the end when I sequence the clone is not the fragment that I expected. I only can get the sequence of the first fragment in one accession (single sequence), but in the rest of the accession that it have 2 locus is not possible. 
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Hello Fiorella,
Have you included controls in your PCR? If not, I suggest you include:
- positive control: DNA that you have amplified before, so you know that it works, with primers that you have used before (ideally for a long amplicon, close to the 4Kb), so you know that they work.
- negative control: the same as the positive, but without DNA to detect contaminations.
- sample negative: primers for the fragment you want to amplify, but without DNA (so you should not get a band here).
- DNA control: DNA from your accessions, but with control primers, so you ascertain that the DNA is amplifiable.
With these controls you will be able to tell if there is a problem with your accessions DNA or your cloning primers. If the controls give the right results, that would mean at least either of four things:
1. Your cloning primers are not good and you should redesign them, aiming at a Tm of 60-65 degrees. It's unexpected that in a nested PCR you don't get results, because they increase specificity and yield. Also, as Fred suggests, design them with primerBLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/). With this you will make sure they are specific, because it could happen that your are annealing elsewhere, reducing amplification efficiency and eventually giving you a faint band that is not the expected product.
2. Your PCR mastermix is not ideal. I suggest you use a robust enzyme like Phusion hot start II or Q5 Hot Start, for instance. For this kind of experiment a hot start will be really helpful, and both enzymes are very processive and would not have problems amplifying 4Kb.
3. The gene you want to amplify is tricky itself. The enzymes above will help, but if the GC content is high or secondary structures are troublesome, you can try designing primers with higher Tm (64-68 degrees) or adding DMSO. You could also try betaine.
4. If you did not have control primers for a long amplicon, bear in mind your DNA could be degraded. Run it on a gel to see integrity (expect a clear sharp band, no smears or ladders). If it is degraded it will be much harder to obtain long amplicons. Also, make sure the DNA is very pure (OD 260/280 ratio close to 1.8, OD260/230 ratio >1.7. This will make a difference when you try to amplify long sequences.
That's what I can think of, perhaps someone will provide more insight!
Best,
Ruben
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I understand that PFU taq have proofreading activity, therefore can reduce error during amplification, therefore it is recommended to use polymerase with pfu ability for fragments that will be used for cloning. 
I need to amplify inserts of 1kb and 2kb for cloning purposes, either using Gibson, Slice or Aqua method. But our lab do not have pfu polymerase but we have long range taq polymerase which contain a mixture of normal taq and also pfu. I used it to amplify my fragments but unfortunately, the pcr products obtained are low in concentration no matter how i optimize it. Is the long range taq polymerase not suitable to amplify small fragments? 
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Since Pfu acts at the any nonextantion moment, lower the Pfu concetration, rise the primers and dNTP conc at the same time
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I am comparing different extraction methods with LAMP and I want to maximize the sensitivity. At present only 2ul of extract (10-100ul total extract) goes into the LAMP assay. Is there a way to concentrate this DNA (without centrifugation) so that 2ul can still be added to reaction, but that 2ul is more concentrated?
Thanks in advance. 
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If you want to concentrate your DNA sample, go for vacuum concentrator to decrease the volume and increase the quantity.
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During thermal cycling, primers having self complementarity often tend to form primer dimers which usually renders the PCR reaction inefficient owing to the preferential amplification of shorter amplicons. However, from a thermodynamic, molecular topology and molecular crowding point of, the self complementarity in the oligo could also mean hairpin formation at appropriate temperature and could thus counter self dimer and the Polymerase occupancy of the self dimer. In case a hairpin structure is intentionally desired in the oligo, which of these two scenarios would be more prevalent over the other?
Thanks
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Hi Satpal,
You can use the following online software to assess the probability of hairpin formation of your primer sequences and their stability at the annealing temperature of the PCR. You can also check the stability of the possible primer dimers and compare the relative stability of the haripins and the dimers to assess which structure will predominate:
My hypothesis is that if the primer hairpin structure is strong enough, they will indeed prevent formation of dimers. I would recommend you to consult a 1997 publication on HANDS PCR (Brownie J et al) where 5' tails were added to the end of the amplimers to generate hairpin shaped structures for the primer dimers themselves, which prevented any further amplification of the dimers.
In your particular case, I think the key point here is the stability of your primer hairpins during the annealing temperature. If your primers do form a hairpin, their stability will depend on the Tm of the stem portion of the hairpin in case it's a perfect pan handle structure. If this structure is indeed stable at your annealing temp (stem Tm > or = annealing temp) then two things can happen depending on the primer-target Tm:
1. The hairpin stem Tm < the primer-target hybrid Tm: Then these primers can actually act as very good SNP discriminatory primers and lead to very specific amplification because the structural constraints of the hairpin will prevent the primer to bind to non-specific sequences (they will act more like molecular beacons). Very specific amplicons will be obtained and very less primer dimers.
2. The hairpin stem Tm > the primer-target hybrid Tm: This implies a very strong hairpin and the primer will prefer to remain in its hairpin conformation at the annealing temp. So no amplification will occur even in presence of a specific template sequence.
If the haripin stem Tm is lower than the annealing temp, then your PCR should work as desired and the hairpins might prevent formation of primer dimers which I think you already pointed out, but the PCR might be less specific if you are dealing with highly GC rich sequences. For further information on how hairpin primers can be efficiently used as SNP discriminatory primers, please consult the following publications
PMID: 15004082 and PMID: 15242954
Best regards and good luck with your research
Soumitesh
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I am trying to perform the control PCR protocol with the plasmids provided in the Surveyor kit, but have not been able to get successful amplification. I tried using a PCR buffer-MgCL2 in addition to a 50mM MgCl2 solution. The polymerase I am specifically using is Platinum taq polymerase. Any suggestions would be greatly appreciated. 
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Hi
The Surveyor Enzyme used for mismatch cleavage assay. I agree with Kaveh on the point to use good quality DNA. However, if you are only interested in surveyor endonuclease activity, you can try any other known variant sequence amplification as a control test cleavage activity like i used in "COTIP: Cotton TILLING Platform". I used indigenously isolated CEL1 instead of Surveyor But if you only want results from Kit's control PCR, check primer annealing temp. and concentration of control plasmids.
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I use vent exo- polymerase for amplification the template below but it seem not get high yield. Could you give me some advice?
5’-A (CTG)10 A AAG TTA GAA CCT ATG-3’ 
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if your primers exactly flank that sequence then your product is very small so will not trap much ethidium bromide. It will also diffuse into the gel by thermal diffusion so may look weak even if there is a lot of it but can you give details of primers and amplimer size please. How are you measuring the yield of product. If you are using UV/nanodrop are you purifying the product before measuring the amount amplified?
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We are using Lucigen Phi29 for specifically enriching circular ssDNA viral genomes in our samples, primed by Exo resistant random hexamers. The reaction is working fine. However, I wish to know if the same protocol can also cpy the linear dsDNA of the host? Or is there any other protocol for specific enrichment of viral linear dsDNA (5-10 Kb)? Any help with protocol or literature will be highly appreciated.
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Thank you Dr Mahmoudpour for your help. We will try with our samples.
SD
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I want to use one step RT-PCR to amplify human RB1 gene (mRNA-->cDNA). When I looked at NCBI nucleotide database (NM_000321.2), the gene has CDS on it. Which one should I amplify, the complete mRNA (4.7 kb) or CDS only (2.7 kb) ?
NB: I want to clone the gene to plasmid vector (vector size: 4.7 kb) in E. coli, then express the gene into protein
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Hi Afdilah,
You cannot use the entire mRNA sequence to express in e. coli. You need to include a Shine-Dalgarno box (ribosome binding site) a few nucleotides before the initiating ATG codon, so the 5' UTR will block expression.
Plus, it is a big fragment to amplify (I highly recommend NEB Q5 polymerase if you must), so I would investigate whether places like Dana Farber has it already cloned for you (Gateway system, using pET62 DEST (discontinued, but ask me if you need it)). You probably need an affinity tag for purification, so a version without a termination codon for a C-term tag might be needed. 
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the amplication curve was so strange, they are two duplicates. x axis is the CT, y axis is the fluorescence intensity. This upper image was liner. The lower image was logarithm. Does it have something to do with the machine?
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There is simply no signal form any amplification.
Have you checked the products on a gel?
If there is no product, well, then there was nothing amplified.
If there is a product but of the wrong length and you are using sequence specific detection probes, then your primers amplify something else.
If there is a product that seems to be correct and you used  sequence-specific probes, then it might be that you used the wrong probes or they don't work (for whatever reason; a mutation in your amplicon sequence could be a possible reason).
If there is a product and you use dsDNA probes,then the instrument has a problem with detection.
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I am planning to amplify a large DNA sequence of 40kb with PCR. Can I use a normal Taq Polymerase? Is there any specific one for large sequences amplification? 
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I agree to Artu Burzynski and Ziguo Zhang, there is no chance to amplify 40 bp of DNA by PCR. The 2 best Polymerases for Long-Range-PCR are Q5 and Phusion. But they deal only with sequence up to 20 kp in best case.
Maybe you amplify the sequence in multiple steps in parts of 5 kp. Afterwards you can ligate the DNA-parts together with seamless Ligation methods like SLiCE or something else.
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Trouble with decolorization of whole blood
Hi,
We are testing DNA amplification and detection from crude extract of whole blood, but the red color of the blood interferes significantly with fluorescence detection (both FAM and SYBR green). We tried various methods to decolorize blood (hydrogen peroxide, sodium hypochlorite, proteases, etc.) but none worked.
Does anyone have any suggestions for decolorization of blood?
Thanks in advance!
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at risk of sounding facetious,,,use less blood and more cycles?
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I am having a huge problem with LAMP. My negative controls all amplify without DNA or RNA samples. I perform regular PCR with external and FIP and BIP and the negative controls are ok, in other words, no amplification is detected. I tried to change all the reagents and the contamination still remains. Is there anyone with experience on that?
Sincerely
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Hi, 
Not sure if this is a still problem with Alexandre and Yuliet anymore, but echoing what Cameron said, recently we tested a primer set for isothermal amplification and found spurious amplification. For the very first run using this primer set, a no-template control (NTC) was positive along with the two positive reactions. This was the case with a commercial master mix as well as reaction mixes we prepared in-house.
Since the NTC was positive the very first time we used this primer set, the contamination (carry-over contamination) can be ruled out.
We have since  found that certain additives can selectively suppress the NTC without reducing the amplification in the expected positive reactions (that is, without slowing the reactions of expected positives),  a concept similar to hot starting in PCR. Without risking sounding commercial, we do provide some solutions to minimizing the blank background amplification. Those interested, please connect/contact me. 
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Advantage in increasing the number of cycles in PCR?
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It can vary depending on when the program was designed.  Older protocols with less efficient polymerase or on older machines often say to use more cycles.  Newer equipment and enzymes are more efficient and newer protocols will recommend fewer cycles.  It's pretty much a "we did it this way and it works so why change it" mindset.
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If you have used, can you share your experience with me? Thanks a lot.
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The paper describing the TruPrime procedure, and how it compares to other methods in the market is appearing tomorrow in Nature Communications. 
Amazing performance!
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I amplified pUC-19 vector at minimum quantity of 25 Nano gram but failed to get amplification through crude DNA of Plant infected with Gemini virus. I verified the infection of virus in plant through PCR amplification.
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Hello Vikas,
The size of fragment affect the efficiency of RCA.The follow article introduces it.
Hope this halp for you 
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qPCR was done to a gene called 36B4, 20ng of DNA and forward primer with concentration 300 nM and reverse primer of 500nM.qPCR machine used is lightcycler 480. Program used 95 c for 10 minutes, followed by 35 cycles of 95 c (15 sec) , then 52 c for 20 sec and  72 c for 30sec.
Resuspending 36B4 forward primers
o   Concentration in (nmol) = 52.2 nmol
o   Stock concentration = 100 uM
o   Re-suspend oligos in = 522 ul
·       Resuspending reverse 36B4 reverse primer
o   Concentration in (nmol)=44.9 nmol
o   Stock concentration = 100 uM
o   Re-suspend oligos in = 449 ul
·       Forward Primer working concentration calculations
o   Initial concentration = 100 uM
o   Working concentration = 2.5 uM
o   Working solution volume = 80 ul
o   I added 2 ul of primers (100 uM) to 78 ul of H20
·       Forward primer Final concentration
o   Working concentration = 2.5 uM
o   Final concentration = 300 nM
o   Final volume =25 ul
o   Volume of the primer to be added to 25 ul tube is 3ul
·       Reverse Primer working concentration calculations
o   Initial concentration = 100 uM
o   Working concentration = 2.5 uM
o   Working solution volume = 80 ul
o   I added 2 ul of primers (100 uM) to 78 ul of H20
·       Reverse primer Final concentration
o   Working concentration = 2.5 uM
o   Final concentration = 500 nM
o   Final volume =25 ul
o   Volume of the primer to be added to 25 ul tube is 5ul
DNA stock solution calculation
o   DNA concentration = 213 ng/ul
o   Required working DNA concentration = 10 ng/ul
o   Working volume = 40 ul
o   I added 1.8 ul of DNA (213 ng/ul) to 38.2 H20
·       DNA final volume concentration
o   DNA concentration = 10 ng/ul
o   Required mass = 20 ng
o   I added 2 ul
Calculation for 1 reaction of 25 ul volume
 
DNA 2 ul (20 ng)
Forward primer 3 ul  (300 nM)
Reverse primer 5 ul (500 nM)
Sybergreen mastermix 12.5 ul
 H20 2.5 ul
 
 
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@all: and this is a good example of the value of a local supervisor - trouble shooting through internet should probably be the last step, when the local team runs out of ideas. So all you students: bug your supervisors, it´s their job, they are paid to train you, and of course, you yourself should critically go through your data, procedures etc :-)
@Jean Neil: thanks for sharing the solution.
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I want to amplify a 3.8kb GC rich DNA for sequencing and cloning. I need a polymerase enzyme that will do a good job. Please help.
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Q5 from NEB works well for high GC templates (https://www.neb.com/products/m0491-q5-high-fidelity-dna-polymerase)
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Hi there,
In my experiment I need to ligate 3 ssDNA, 80bp (S123) with help of 2 other ssDNA 80 bp as bridge (NS12).
this assembly uses single-stranded bridging oligos complementary to the ends of neighboring DNA parts, a ligase to join DNA backbones to assemble these 3 DNA fragments (S123).
Experiment process:
1)     Ligation
2)     PCR
3)     Gel Electrophoresis
My concern is that hybridization bond between S123 and NS12 affects on PCR even with using no ligase.(attached figure)
In the attached figure you may see almost the amplifications are same between the samples does have ligase and the sample which doesn't have ligase.
The only difference is that at high concentration of DNA with no ligase some minor bands (100bp~120bp) can be seen (sample #8), while the minor bands hardly can be seen for the sample #7 which does have ligase.
I wounder if there would be a way to confirm the ligation or if there would be a way to denaturate NS12 and separate S123 prior to PCR.
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To separate S123 from NS12, maybe you can try to work with biotinylated NS12 oligos and remove them after ligation through binding onto streptavidin beads and denaturation to release the S123 which can then be analyzed separately.
To confirm ligation within S123, one possibility could be to label S1 and follow its mobility shift by gel electrophoresis (*S1+S2+S3 vs *S1 or *S1+S2).
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Hi there,
In my experiment I need to do PCR with below DNA and Primers, I also use RT-PCR.
But I don't know why it cant be amplified. The other information is available in the attached picture.
dsDNA:240bp
TTCGCGAGATCTATGAATAAAGACACACTGATCCCTACAACTAAAGATTTAAAAGTAAAAACAAATGGTGAAAACATTAATTTAAAGAACTACAAAGATAATAGCAGTTGTTTCGGCGTATTCGAAAATGTTGAAAATGCTATCAGCAGCGCTGTACACGCACAAAAGATATTATCGCTGCATTATACAAAAGAGCAACGTGAAAAAATCATCACTGAGATACGTAAGGCCGCATTACAA
S1-Primer-F: 22bp
TTCGCGAGATCTATGAATAAAG
S3-Primer-R: 22bp
TTGTAATGCGGCCTTACGTATC
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Hi Karimi --
Whenever I design primers, I like to run them through a site like OligoCalc or Benchling for analysis.  
Looking at your primers through OligoCalc there seems to be two issues.  First, the melting temp of your Forward primer is about 56*C and your reverse is about 59*C.  Your PCR program is set to 59*C for annealing, and this is too high for the F primer.  Always set your annealing temp a few degrees below the lowest melting temp for a primer.  In your case, i would set it to 51*C instead of the 59*C you currently have.
Second, and probably less of an issue, is that your Reverse primer has a potential hairpin formation (check Oligocalc-Self Annealing) which could be inhibiting annealing.  
First try adjusting your PCR program with a lower annealing temperature. Let me know if you have success with that!  Good luck.
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Hello to all.
For my experiment I need to transfer a double-stranded DNA molecule (in fact, a PCR product) into the nucleus. The method I will use is nucleofection.
I presume, that success of my idea is dependend on the time, that molecule will stay untouched.
I can use any reasonable addition on 5`-end of my primers, I can use them with or without 5`-phosphate group, I can use polymerases with or without 3`-additional nucleotide. Also I can restrict division of my cells for as long as, I think, that experiment will need to become a success. Maybe I can use some other methods, I didn`t think of, but they are available in our lab.
Are there some features I may use to protect my PCR product from degradation? What would be the best strategy? Thanks!
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Hi Maksim,
While I can't speak to the efficacy of the methods you propose (I've not tried nucleofection of linear dsDNA), I seem to remember reading that treatment of the cells with caffeine - i.e., a broad-spectrum inhibitor of DNA damage-activated PI3K kinases such as ATM, ATR and DNA-PK - increases the lifespan of linear DNA.  Maybe this is a simpler method for your purposes?  Be aware that such an approach may have off-target effects on your cells, or may preclude certain experiments you may be interested in.
Hope that helps!
Dan
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I need to amplify a 2kb gene from the genome. The details of the primers are as follows:
FP - GC - 70%, Tm - 76o C
RP - GC - 73%, Tm - 77o C
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do the gradient again but add betaine at a final concentration of 1 Molar to all tubes. Betaine destroys the hydrogen bonding beteen the strands so the template melts esier. If your templte is GC rich then it will anneal very quickly and easily so the pcr will not work unless you use something like betaine to keep the strands melted while the primer sticks and extends. It is very cheap
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What I have heard is the normal Taq Polymerase which we commonly use can amplify a fragment upto 1.5 kb. But yesterday someone told me that the normal Taq Pol can amplify a fragment upto 3kb easily and we do not need to use high fidelity taq polymerase for it? Please suggest.
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It will work more or less equally reliably up to 1.5kb.  Above 1.5kb the efficiency is going to decline.  Some sequences will work, others won't.  The workable conditions become narrower, and the yield is going to be poor.  Higher than 3kb basically nothing works.  So if I had a 3 kb amplicon, I would definitely start out with hi fi.  It will make your life much easier. 
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We are using phi29 to amplify a circular DNA template. This is coupled with a restriction enzyme and we observe that without addition of primer, there is still significant amplification. We have cleaned our circular template twice on denaturing PAGE, after exonuclease treatment in order to remove the splint.  Any suggestion to minimize the background amplification?
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Hi Swapneel,
have you inactivated the phi29 polymerase after the amplification, before restriction enzyme analysis?
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I have a query regarding how to avoid smearing pattern of amplification product after performing reverse transcriptase PCR.
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Naoki, Thanks for your suggestion.
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I amplified some DNA in a PCR reaction and when I ran it on a gel I found I had an extra band which was twice the size of my band of interest. I had designed my primers to have restriction sites plus 6 nucleotides at the ends so I could cut my PCR product and put it into a cloning plasmid for sequencing. I did this for my band of interest (~350bp), as well as the bigger band (~700bp). The sequencing results showed both plasmids contained the 350bp sequence of my band of interest. So I am trying to figure out what happened to the bigger band. I had extracted the ~700 bp band out of a gel (run on the same gel as my 350bp band) but ended up with a ~350 bp sequence in my plasmid.
The only explanation I could think of is that my primers are somewhat complementary (only the 6 nucleotide overhang though), which could result in two 350bp sequences being joined together with the restriction sites in the middle, so when I then digested the 700bp fragment it would return to two 350bp sequences which would ligate into the plasmid and explain how I got a 350bp sequence from a 700bp band. However this would require one PCR product to act as a primer for another and, after much confusion over directionality, I think (?) this is impossible….
In terms of my experiment this is not a problem as I have already got my band of interest in the plasmid now, but it just got me thinking as to whether it is hypothetically possible for a PCR product to prime another PCR product if they used complementary primers. As far as I can work out this is impossible for all combinations of primer (ie. Fwd - cRev, cFwd - Rev etc.) even if they are complementary because the directionality would be wrong and you would be having to add to the 5’ end- is that correct?
Also if anyone has any ideas as to what happened to my 700bp band I would be interested to know your thoughts!
Apologies for the rambling question
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Totally possible!  This is essentially what is done with SOEing (splicing by overlap extension).  You put two PCR products, which have a region of complementarity, into one reaction and that allows for annealing and extension of a spliced product.  In a "normal" PCR where this occurs by accident, it should be much more rare than the amplicon you want, but it could conceivably be cloned.
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I'm trying to clone 700 bp gene in pet28a vector. After transforming I selected colonies for colony pcr. In colony pcr 3 of the agarose wells were glowing as if dna is trapped in the well. I have also run positive and negative controls which gave correct result. Please anyone help. I'm stuck.
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Hi Farah,
This has happened to me before and it has likely happened to everyone at first. There is most likely too much template DNA in your reactions from the colony preparation. Your controls were from purified DNA? I usually lightly touch with a small pipette tip the colony and then place the tip in 50 uL of pure water, mixing a few times. You should not see any bacteria on the tip (trust me they are there). Then heat in thermocycler at 95C for 10 minutes. Now use 2-5 uL of this mixture for PCR. If you do all of this in 96 well plates, you can multichannel from the first plate to the PCR and save time. The glow in the wells is a dead giveaway that there is too much template. 
Good luck and have fun.
Alex
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First, I optimized the primers which were designed for specific exons, with different Temp (annealing Temp. of PCR) in gradient PCR with standard human blood DNA. After getting annealing Temp. did the conventional PCR with specified Temp. with standard human blood DNA.  I got the band intensity good and I did not get the any non specified bands . After optimization of primers performing the PCR with subject DNA samples and sent to sequencing. They said that All samples showed smear indication of degradation. So what should I do for getting good PCR product for sequencing.
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try to use the minimum amount of genomic DNA sample. If you get a single amplification product,  clean up the PCR reaction using PCR and gel clean up kit to get purified  product suitable for sequencing.
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I've been trying to amplify the products for quite a while now and I have difficulty in getting good bands or any bands at all, I'm wondering if it's the storage condition of the components or there is contamination, any thoughts?
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For those cases its always helpful to run a positiv controll alongside (a template under conditions that always results in bands).
For the DNA extraktion you can have a quick look via Nanodrop. Collect different flowthroughs to figure out if on any step are a higher amount of DNA get washed of (if the kit uses a column).
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I amplifly ITS2 DNA from old mosquito samples preserved on naphthalene using antioxidant, but the target sequence does not amplify, only nonspecific fragments, smaller and more stable. This DNA would be fragmented? Fragments of mitochondrial DNA does not amplify as well.
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Use of pure ethanol (99%) is most recommended as it preserves the DNA in most suitable condition. Unless you  cannot get fresh material of similar species, the naphthalene preservative might have allowed degrading of DNA of the specimens.
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   I need to send Fosmid DNA for sequencing, but they told me the concentrations were too low & there are multiple bands. I tried mini-prep Qiagen kit many times & also midi-prep. How can I make sure the multiple bands are not degradation or genomic DNA?
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Hello Laila,
Sometimes, the multiple bands represent the supercoiled and non-supercoiled forms of plasmids/fosmids. So, I agree with Weber; you should digest your fosmid with a suitable restriction enzyme. If it is completely digested, you should see only one band of linearized DNA if your fosmid is pure.
Good luck Laila!
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I had amplified before cloned and sequenced using expired green  mastermix. Then I got a fresh supply and it has been difficult amplifying same genome. I need help please. What else can I do
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Thanks Ali. Thank you Khan.
@Prabhudas. I had use same template and used varying concentrations. Just think there may just be something slightly missing. Think I better be thinking of another enzyme here. 
Thanks all
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I am trying to amplify a large DNA fragment (about 5 Kb) by PCR. I used a FX-PCR master mix (for long and fidelity PCR). The PCR condition is as follow: 95C 5 min, 35 cycles of ; 95C 30 sec, 53C 30 sec, 72C 5min. But after electrophoresis of my PCR product I have smear on the gel. I have tried gradient PCR and also changed the primer but still I did not get any band.
Thank you
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the important thing in long pcr is that the template does not re anneal to itself before the extension happens so all long pcr should have primers that anneal at very high temperature to minimise the time that the template has to become double stranded again. I would design primers annealing close to 72c and do the pcr in the presence of 1Molar betaine which keeps the template melted for longer by destroying the triple hydrogen bonding of C-G bonds so makes the template melt easier. A profile of 95c,70c,72c would work much better . 53c is very low for long pcrs
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I use StepOnePlus Real-Time PCR System for relative gene expression studies. I follow SYBR Green chemistry. Is it advisable to use 0.5X  SYBR Green Master Mix instead of 1X?
Thanks in advance.
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We tried it (0.5x SYBR) successfully with ABI system. But for Agilent system 0.75x SYBR gives flawless result than 0.5x SYBR. 
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I want to perform LAMP analysis. According to the procedure betaine should be added. Anyone knows the best betaine for LAMP. I found three from Sigma aldrich.
Betaine (Can no.61962) costs less than 40 EUR/50 g
Betaine (Can no.30056) costs less almost 60 EUR/50 mg so it is alnmost 1000 x more expensive.
does anyone know it the cheaper one suits LAMP?
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I have also found that my assay doesn't need betaine, but it does depend on the GC content of your target. If your target is GC rich then you may need betaine to aid in the separation of the strands.
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Does some know the protocol for 2-step gateway pcr experiments?
It shows 2 step to get pcr product.
at first step, we need to prepare 50ul reaction mixture including:
5 µl 10X polymerase buffer
5 µl dNTP mix (2 mM each dATP, dCTP, dGTP, dTTP)
1 µl 12attB1 primer (10 pmol/µl)
1 µl 12attB2 primer (10 pmol/µl)
0.5 µl DNA template
1.0 µl DNA polymerase (2.5 U/ml)
36.5 µl sterile water
 The primers have to be designed as shown below.
12attB1: 5'-AA AAA GCA GGC TNN-(template specific sequence-3'
12attB2: 5'-A GAA AGC TGG GTN-(template specific sequence)-3'
PCR amplification
denaturation 2 min 95°C
denaturation 15 sec 94°C 
annealing 30 sec 50-60°C 10 cycles
extension 1 min per kb 68°C 
STEP2 : 
Amplification of the product produced in Step 1 using universal attB adapter primers. 
50ul reaction mixture
4 µl 10X polymerase buffer
4 µl dNTP mix (2 mM each dATP, dCTP, dGTP, dTTP)
4 µl universal attB1 adapter primer (10 pmol/µl)
4 µl universal attB2 adapter primer (10 pmol/µl)
10 µl PCR product Step 1
1.0 µl DNA polymerase (2.5 U/ml)
23 µl sterile water
 PCR amplification :
denaturation  2 min  95°C; denaturation 15 sec  94°C;
annealing     30 sec  45°C 5 cycles; extension 1 min per kb 68°C
denaturation  15 sec 94°C; annealing 30 sec  55°C  15-20 cycles
extension 1 min per kb 68°C; extension 10 min 68°C
QUESTION 1 :
Is same PCR amplification procedure with any DNA polymerase?
For me, I use Phusion Master mix or Phusion High-Fidelity 
Do I need to do PCR amplification as the protocol from Phusion kit? For example, change denaturation,extension temperature and extension time.
QUESTION 2:
Usually, in our lab, for PCR amplification, we use 10mM dNTP.
Can I use this dNTP?
QUESTION 3:
After step1, do I need to dilute the PCR product, which will be the template in the step2?
QUESTION 4:
If I prolong the cycles to 30 for  PCR amplification in the step 1 and I can get the bands, can I use that kind of PCR product as the template?
Thanks a lot!  
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I used 2-step PCR for and got the right products for BP reaction.
A1: I have used  Phusion 5X HF buffer and Phusion Taq,  I think you should adjust the PCR TM and cycle based on the enzyme and the extension time based on your pcr product size.
A2: I have used 10mM dNTP for both of first and second PCR, it works well. 
A3:  After electrophoresis of 1st pcr product, I used 3ul (50ul total volume)as input for the second PCR, it works well.
A4: Sure, you can use it.
GOOD LUCK!
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Guys,
I made a cDNA with High capacity kit and ran an agarose gel (1%, 1ul of cDNA) to check, BUT...there was nothing!!! I didn't had a single band, and not a smear either! Anyone have any idea what may be happening?? Because after that I ran a qPCR with housekeeping gene and another gene of interest and I had amplification! The results were ok, with good amplification and a single melting curve.
Any ideas?
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As Satti says: cDNA would appear as a smear, if it appears at all.
You've essentially copied all your RNA to single-stranded DNA molecules of wildly varying lengths. You wouldn't expect a band from this, unless you used total RNA and random priming, in which case you might (possibly) see two smeary ribosomal bands. 
Remember: DNA of sufficient concentration to be visible on a gel represents a truly ridiculous number of molecules, whereas PCR can amplify as little as a single transcript. You don't need to be able to visualise your cDNA to know it's there, nor would I recommend doing this anyway (it doesn't tell you anything useful).
If your qPCR is working fine, then your cDNA is probably fine.
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I want to amplify a cDNA (1400bp) using Pfu polymerase. I amplified it using Taq polymerase but I can’t amplifyit by Pfu. Can anyone help me to solve my problem?
My PCR program is:
95 5min
95 1min
58 1min
72 2min
35 cycles
Thanks
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thank you all
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I m trying to use my DNA extract from colonies on a culture plate as a standard for my qpcr so I want to determine the size of my DNA that is within my specified primers.
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Hi Marusa,
You could identify the position of your primers in a sequenced strain using BLASTn e.g. at the NCBI. However, the sequenced strain might not be exactly your strain, but a close relative. The result could be satisfying depending on your specific question.
Good luck!
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I have extracted soil DNA that are a lot of humic acid using FastDNA MP-Bio. at the first extraction i got problem that the DNA can not run in amplification. but after optimization using metal substance, the DNA available to amplify. and now i got new problem, PCR product appeared unspecific band otherwise the negative control (no DNA template) also show the desired band. I suppose that it caused the contamination, then i have make all a reagent in the new fresh reagent and also i have tried using gradient temperature about 50 C - 64 C, in that case i still get same result.  I need some help in this crucial problem, this is my first experience in methagenomics analysis.
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It is maybe also a silly addition to Annemaries good advice but: Don't overload your gel pockets! Not that the "contamination" is just sample that got carried over because the pockets were to full of sample.
If you prepare the PCR mastermix try it next time with stuffed filtertips and see if thats work.
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I am having trouble with my PCR amplification. I am using species specific primers for Mycosphaerella species from banana. I am getting amplification with the fungal pure isolate DNA but not with DNA extracted from infected banana leaves which symptomatically have the fungus. Any suggestions?
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Dear Janet, 
The problem might not be that you don't have enough targeted DNA, but instead that your CTAB extraction fail to remove some PCR inhibitors. Did you consider using diluted DNA at e.g. 1/10 as PCR template? This often solve such problems for DNA extracted from soil samples. One thing you might consider to confirm whether your DNA is ok for PCR amplification is to amplify the banana with some common plant primers located in the chloroplast. HTH
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I need to amplifying 3'UTR of a gene from cDNA for the reporter assay. In the first round of PCR, I have got the specific amplification at 584 bp. However when i have re-amplified the PCR product , I am not able to succeed with it. I have used 2ul of the amplified product as the template for 20ul PCR reaction. Kindly suggest me to rule out the error and the best way to amplify the 3'UTR successfully. Below is gel picture of second round PCR. Lanes followed by ladder are negative control and two different samples
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Hello Nalini,
I expect you have the answer by now but for others reading this thread:
your result is as expected. You must realise that pcr doubles the amount of product every cycle so every 10 cycles you have 1000 times more product than you started with. You  have used too much product for the second round of amplification and the product is soon at very high concentration and self annealing to other product molecules so the product is getting randomly longer and producing the smear of high molecular weight  material that you see on the gel.
You should dilute 1ul of product in 100ul of water and run about 6 tubes of pcr with 1ul of this dilution as template. remove 1 tube from the pcr machine every 5 cycles and probably one of the tubes at 10-15 cycles will have lots of quite clean amplimer and the 20-30 cycle tubes will look like your result with just a smear. The best way to get clean product in this example is to us e2 nested primers just inside of your first round primers.
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Hello! I have been trying to amplify the promoter region of the gonadic aromatase (800 bp) to analyze its methylation status. The genomc DNA used have been extracted from embryonic turtle gonads and treated with bisulfite using the Zymo reaserch methylation kit. I designed three pair of overlapping primers amplifying at about 300 bp fragment each using a 40 cycles Touch Down PCR protocol ann performng a second round. I came up with this protocol using embryonic brains of the same turtles and it worked perfectly, but when I switched to the gonads, the results are very inconsistent, sometimes I have amplicons but the majority of the times I dont, even if I do a secind and third round... and sometimes using the same samples... I would really appreciate any advise. Thanks!!
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 Hello guys! thank you so much for all your advises, I finally could amplify the promoter of my gene.. not complete but Im still working on the design of new primers and hopefully I can get it complete. But indeed repeat frozen/thawing cycles dont help. And the best way to amplify this Bisulfite treated DNA it was with nested primers, it work really well!! 
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I want to measure the production of mRNA of pro-inflammatory cytokines in an experimental colitis model, so I expect an upregulation with colitis while controls levels should be very low. 
Previously I had issues with amplification since I use dextran sulfate sodium as a colitis inducer and it has been reported that it interferes with RT-PCR.
After performing RNA purification with LiCl I was able to obtain amplification of beta-actin so I started with the cytokines. Unfortunately while I obtain a nice band for beta-actin in all my samples, is not the same when I want to measure the different cytokines. I attached the image of the gels I just obtained.
I would appreciate some help since I am lost on what to try next. 
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Hello Karla,
I am very familiar with this model - (DSS-induced colitis in mouse intestine).
Even with qPCR, the cDNA samples had to be diluted 1:350 prior to adding 6 uL of sample to the 25 uL qPCReactions to get rid of the inhibitory behavior of the cDNA samples on the qPCR.  But, since you are dealing with RNA (instead of cDNA) as template going into a 2-enzyme mix, your carryover DSS could indeed hurt even more. So, your conventional reverse transcription PCR followed by gel analysis in this model gives what I would expect - I am even surprised you got beta-actin signal - but your LiCl 'clean up' probably helped a little there.  What has eventually happened in this model is that the project ended up using RNAscope to see the cytokine signals in situ - and worked perfectly (but expensive). So, I would suggest first trying your LiCl-'purified' RNA samples in a one-step reverse transcription quantitative PCR (RT-qPCR) approach first, since you have a better chance of detection that way than trying to visualize what will end up being very faint to non-visible cytokine bands on your gels. A one-step qRT-PCR kit from Quanta would work nicely for this. Your LiCl-purified RNA might not have to be diluted 1:350 beforehand... and you will want to prove that by performing a 20-point serial 1:2 sample mixture dilution study up front to see where your valid dynamic dilution range is for the RT-qPCR for each of your targets.  The small amount of mucosa able to be extracted for total RNA in these studies is always the bottle-neck.
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Hi everyone ! would like to know  about exact  difference between negative   and positive control in PCR
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A positive control is one that you expect to work under the conditions given.  The positive control will test your master mix, MgCl2 amounts, primer annealing temperature, and extension times.  If your positive control does not work, those results indicate that something is wrong with your annealing or extension times or temperatures, or something is wrong with your MgCl2 or master mix set up. If your positive control does work and your test samples do not, then there could be something else going on such as not enough or too much template.  I will often use a plasmid with the desired sequence I want to amplify for my positive control (typically around 500 pg as an amount).
A negative control for PCR is one which should not give you amplicons, typically the negative control will contain no template or will have one or the other primer.  Setting up two negative controls, each containing only the forward or reverse primer, should not provide visible amplicons.  Therefore, any visible bands might be a result of contamination or multiple opposing binding sites for the designed primers. 
Short and sweet of it: A negative control is one you expect not to work under the conditions.  A positive control is one you expect to work and to provide you with the expected result.
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Does anyone know the primer for Chrysaora sp. DNA amplification?
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Sir,
I am mistaken in any step of isolation. Because few samples of Pelagiidae species DNA isolated earlier with good result. I have extracted DNA from the tentacles, bell margin and gonad using (Orgin) Marine DNA kit. Please give me some suggestion to get a good result.
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Did anyone used LED Transilluminator (instead of UV) for visualizing DNA or PCR ... Amplification of bacterial DNA,using 16Sprimer and Midori Green Advance instead of ethidium bromide?
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I use the small blue light transilluminator from Clare Chemical. Very happy with it. We don't use ethidium (not allowed it), I have tried sybrsafe, gelred, gelgreen and gelview - all work fine with the blue light. Blue light is fantastic for cloning and allows you to take your time during gel extractions without UV causing DNA damage, giving you more colonies (especially when cloning larger fragments).
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Hi there,
I would like to do a tumor burden follow-up by measuring circulating cell-free DNA (ccfDNA) at different time points in a mouse experiment (by treating or not mice after tumor initiation).
Does someone have a protocol to amplify DNA as I can draw only 100-200µl of blood/week in mice.
I have a protocol to isolate DNA (QIAmp DNA blood mini kit) from plasma.
Thanks in advance!
Christophe
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Thanks a lot Paul, I'll come back to you when I would have time to retry it!
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I wish to know the size of PCR amplified beta-actin cDNA (from Hela cell line) on an agarose gel (0.8%).
I read differences in band size for a beta-actin gene, In my case, it is showing near 100bp. 
Kindly tell me the correct band size (on an agarose gel ) for PCR amplified beta-actin cDNA?
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This will depend upon the primers that you have used to amplify the template.You need to use your primer sequences (both forward and reverse on the beta actin reference sequence  using NCBI Blast . This will tell you the exact product size. Moreover to 0.8% agarose gel is used for running products more than 1kb in size . For a shorter product size ranging between 100bp to 600bp genrally 1.5% to 2% agarose gel is recommended.
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I am trying to check the sequences of some Actinomyces species we are growing. I extracted DNA from them and checked the purity; almost all looked good to me. 
I then put them through a PCR with universal 16s primers using DNA I had already validated as a positive control and water and my negative. 
From the gel you can see that some of the DNA may be highly sheared, but there are no bands anywhere except for the control. 
What might be the problem? The IDs are being checked in case there is any contamination, but the samples I took DNA from were struck out to isolation on a plate so they should all be pure cultures. I also highly doubt that all 14 samples were contaminated.