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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Hello,
I am studying DNA barcode (COI) on amphipods but so far, I can´t achieve product PCR+. I have used three primer (Folmer/HCO2198 LCO1490 – Zplank/ZplankF1 ZplankR1- JG/jgHCO2198 jgLCO1490), parameter different of annealing temperature, +/- DNA concentration, + Mg+2, Trehalosa (potent PCR enhancer) to 5%. I used two extraction method DNA: 1) Hot-shot and 2) Wizard SV Genomic DNA. The amphipods are marine (Stenothoidae, podoceridae, maeridae and caprellidae). The concentration of DNA is between 5 ug/uL to 50 ug/uL for Method Hot-shot, and 1 ug/uL to 10 ug/uL for Method Wizard SV Genomic DNA.
Something idea about How I can achieve product PCR+?.
I would be very grateful for his assistance.
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The prblem could be with your dna. Check the od260/280 ration which should be close to 1.8. Only use 50ng dna per reaction. In case your primers are degraded get some dna from another researcher that amplifies in their hands to act as a pcr control. Start with an annealing temperature 8c below the lowest annealing temperature of your primers. Once you have a dirty amplification you can raise this temperature to get a cleaner product
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I am following this paper (DOI: 10.1128/AEM.02895-10) to run a PCR that could differentiate between live and dead bacteria.
The method is based on propidium monoazide (PMA), which enters the dead cells, binds to the DNA, and then polymerizes, impairing the subsequent PCR step. The s s done with UV irradiation. The authors report this step as: "10 min of incubation in the dark, samples were exposed for 5 min to a 600-W halogen light source at a distance of 15 to 20 cm from the light source".
I have available a cross-linker with an emission at 254 nm but with only 5 W, not 600 W.
I have killed the bacteria by incubating them at 99 degrees for 15 mins. I added PMA at the suggested concentration of 100 μM in the dark for 10 mins to both the treated cells and a control. To compensate, I left the samples for 30 mins under my lamp in a tissue culture plate (to be sure that light reached the samples).
However, the PCR showed only a decrease of one circle in the treated samples compared to the control.
Am I correct in thinking that 5 W is too little? Or is there another fundamental step I am overlooking?
A 600 W lamp is not readily available. I have a 40 W lamp in the cabinet. Would that be better? and how long should the samples be irradiated?
Would the normal 1.5 mL tubes be transparent to the 254 nm light to are there special tubes?
Or is the fully chemical approach to differentiate live from dead bacteria by PCR?
Thank you
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ADDENDUM: it turned out the problem here was with the extraction. After repeating, there is a 10 qPCR cycles difference between dead and alive bacteria, which is what I was looking for. Case Closed. Thank you for the support
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Hi everyone and good morning!
In our lab we want to make a sanger sequence opf the RPGR gene (ORF1/ NM_001034853.2) but is technically very dificult to make any PCR within that zones (it´s a mutational hotspot so, it has to be made)
¿anyone has any idea? we have arrived at the point to have very few not specific bands (so we cut the band of interest and so on) but I wanted to learn from others experience working on STR´s zones.
thank you very much... it´s technically challenging and fun!
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we send for other mutations both forward and reverse primers and I observed this phenomena on frameshift mutations (wich make them easy to identify on the SeqMan) I'ill make what you say and maybe send also some internal primers.
thank you Paul!!!
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We are currently having problems with PCR amplifations of a ~200bp gene fragment. We cannot see any product after a 35 cycles amplification round and usually use a simple reamplification protocol: 1ul of our first PCR to a 20ul reaction. Resulting positive bands are not so clear so I wonder if you could kindly share your experiences with reamplification protocols. Thank you!
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re amplification has a problem both with overamplification and the production of incorrect bands. 10 cycles of pcr is 1000 times more pcr product so often weak error bands become strong and also the greta over amplification of the right product means that product molecules anneal and you get a long smear of amplimer rather than a single band
I would dilute the first pcr product 1/1000 in water and set up identical pcr re amplifications but remove tubes at 14,16,18 etc cycles and you should get clean strong bands at a few different cycle numbers then the product will smear and become messy at too many cycles but it is necessary to tes how many cycles is the right number
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I am attempting to amplify nirS and nirK in some salt marsh sediment samples for my Masters thesis studying denitrification. My nirS gene is amplifying great in both PCR and qPCR. In the literature, nirK seems to amplify well at an annealing temperature of 57-58 C. However, when performing PCR, the target sequence amplifies well in my bacterial strain control at 50 C annealing temperature using HotStart Polymerase. I switched over to qPCR using 2X SYBR Green polymerase and increased the annealing temperature to 55 C because that was the standardized protocol in the lab. It worked incredibly well for nirK except upon inspection of a gel I have a beautiful band at the wrong bp length (700bp and my target sequence is 160)! No other bands show up at the correct length. I could lower the annealing temperature down to 50 C but that is incredibly low considering that I believe no specific amplification is occurring already at 55 C. I could also increase to 58 C but amplification doesn't occur at that temperature either. What should I do?
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The melting temperature of primers depends on the buffer that you are using for the PCR reaction. This website calculates melting temperature for primers for common PCR buffers: https://tmcalculator.neb.com/
If you are unsure about the melting temperature, then I would recommend looking up touchdown PCR (TD-PCR). This is a normal PCR except the melting temperature is lowered by one degree after each cycle, which allows you to get specific amplification of your product. If you still see a band of the incorrect size after doing TD-PCR, then either the primers are poorly designed or there is contamination of some other DNA that is being used as the template.
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I am trying to extract DNA from a single insect from different sites. After extraction nanodrop readings show around 12-15 nanogram per microlitre concentration which is normal for the insect. However PCR amplification from two specific sites are failing everytime inspite of DNA being present. I have checked for presence of inhibitors by extracting DNA from four sites simultaneously which included the two sites mentioned above. DNA concentration is sufficient. However PCR amplification failed again. What could be the reasons for the failing PCRs?
I am attaching the gel image of the samples.
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The PCR result is influenced and inhibited by substances used in DNA isolation. You should pay attention to the concentration of the substances you are using and bring them to the appropriate concentration.
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I am currently optimizing a RPA-based protocol. As everyone knows, this nucleic acid amplification technique is based on isothermal amplification (around 37-42˚C) in combination with recombinases and single-stranded DNA binding (SSB) proteins.
I have designed several candidate primers to optimize my RPA. All of them were searched in literature and designed considering the criteria to be used in an RPA reaction.
These same primers were verified by conventional PCR (At different Tm: 55-65ºC), obtaining a successful result.
The problem started when I used the RPA master mix (lyo version from Twistdx) and performed the RPA reaction with the same primers and samples used in the conventional PCR. ALL THE RESULTS BECAME NEGATIVE?!
Primer concentrations used in RPA reaction was 400nm (recommended by twistdx) and the reaction time was 30 minutes at 39 °C (conditions recommended for the set of primers tested). Visualization of the amplification was done on agarose gel (1.5%) and doing a posterior melting curve assay. In all cases, no amplification was detected.
Does any one have a clue on what is happening???
I don´t know which other variables I can change to obtain good results
Suggestions?
Thank you
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Primer concentration is important in this experiment (usually in all amplification experiments). In addition to it,my question is whether the recombinases enzyme and sample in your experiment are working properly?
All the best
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Two days ago, I carried out the Agarose gel electrophoresis. And also, I performed the PCR prior to the electrophoresis (bt_3294 [KCTC 5015]). (The process that I did is on the picture.)
But I got some problem with the electrophoesis. I got a normal band at 'a' and 'b', however I cannot get the band at 'c'.
At first, I thought the primer did not function effectively during the PCR. However, if that was the case, there would likely be a primer dimer band present at 150-200bp.
So here is the question.
1. Can the band not appear even if the primer dimer occurs? 2. If there were any other causes, what could have caused it?
I will conclude this writing by wishing you a good day today.
Best regards,
Joonseo_Cha
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I recommend all the above answers in addition check the voltage used in the electriphoresis process if your gene size is very small it could run out of the gell if the voltage is not suitable
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I want to make a mastermix for PCR. I already know the primers and stuff I want to use.
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1.Non-specific band at 400 bp, even though by gene size is 650 bp
2.No primer dimer
3.PCR Conditions using right now:
Denaturation - 98 degrees (45 sec), ext.- 98 degrees (30 sec)
Annealing - done for 55 degrees to 61 degrees (Tm for primers was- 58 degrees)- 45 seconds, 35 cycles
Extension- 72 degrees (45 sec.)
Final extension- 72 degrees (15 seconds)
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  1. Primer Design: Check the design of your forward and reverse primers. Ensure that they are specific to the target DNA region and do not have any significant mismatches or secondary structures that could lead to non-specific amplification. Confirm that you are using the correct primers designed for the 650 bp region and not for a different, shorter fragment.
  2. Template DNA: Verify the quality and integrity of your template DNA. Degraded or contaminated DNA can result in non-specific amplification or shorter products. Ensure that you are using the correct template
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Recently carried out PCR in order to amplify MMP9 Product from A431 Cells in order to prove its presence in the cancer cell. However, when Electrophoresis was carried out post-PCR here was the result:
*Image attached below*
(Left to Right: 100bp DNA Ladder, Non-Template Control, Wild Type A431, Positive Control, Negative Control, MMP9(1) & MMP9(2))
Bands were formed at ~500bp instead of the expected 106bp. In fact, ZERO product was formed at 106bp. The ideal was for bands to form at ONLY 106bp for each well apart from the NTC, yet there was only one single incorrect band. (-ve Control yielded no result from this PCR likely due to human error) This result is unacceptable as the primers used were specific for MMP9 product and was ordered from a locally renowned Biotech Company, with reference to research papers and was tested again Primer-Blast.
Does anyone have experience with PCR to identify this band?
(Can MMP9 specific primers possibly amplify non-targeted DNA during PCR?)
Is it possibly somehow MMP9 related?
(Does DNA form Pentamers of sorts in excess?)
This PCR-Electrophoresis was conducted several times and this was one of the better results. I've also attached the Temperature Gradient PCR for reference.
Thanks in advance!
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Depends if the gene has repetitive regions and can form concatemers or your primers can anneal to other parts of the gene/genome. Have you tried different extension times? If in doubt send a band for sequencing to find out what you are amplifying.
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I performed electroporation to introduce a plasmid into Agrobacterium and then plated the transformed bacteria onto media containing antibiotics to select for the presence of the plasmid, but subsequent PCR reactions for the target gene are negative
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Dear Mohamed Samir Youssef did you try to pate your Agro without transformation or a mock transformation to see if there is some contamination that is not Agro or an Agro resistant to your antibiotic of choice or that antibiotics are ok? After purifying plasmid from Agro, do you run it on gel before doing PCR? normally the concentration is very low, but you could still try to see if you got any plasmid at all or nothing.
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I have run PCR and digestion (Not1-hf & Nde1) that gave good bands. The plasmid was PET-28-make123 and the PCR product was franken-flag-dsred.
Then I performed ligation and ran a gel with ligation reaction, plasmid only, and PCR product only control and no bands were present.
How do I find out what caused this?
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Barely. You can visualize a single band that has maybe 5-10ng in it, although faintly. So you should be able to see that amount of DNA if in a single band. But after ligation you might have a number of various ligation products so that each band could be faint.
I never bothered running ligation products on a gel, I just ensured that I ran the proper controls and then transformed.
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I'm encountering a PCR issue in our experiments. We employ a master mix as control and utilize 3 or 4 primers in a single reaction. Since our samples are typically heterozygous (containing both wild-type and mutant alleles), it's expected to observe two bands as results. In one particular PCR, I'm consistently observing a faint wild-type band in the master mix. I've repeated the experiment using all-new reagents, but the outcome remains consistent. This light band consistently aligns with the length of the wild-type. Is it common to observe such bands depending on the choice of primers? What are the places I can improve.
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Run a PCR with 2 vials each of no primer, no-template, and master mix alone. And in case you are using strips, better do it in vials so that contamination can be avoided.
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Is there anyone who has done TaqMan assays using average regular use PCR mastermix (not the TaqMan assay specific mastermixe) using cDNA as template for the qPCR test? I wanted to know the ins and outs of the procedure and the optimization you did to get accurate results.
Thanks in advance.
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No, you cannot perform TaqMan assay using average regular use PCR master mix.
If you wish to perform PCR using the regular PCR mix, the assay is no longer called TaqMan Assay because the defining feature of a TaqMan Assay is the probe. This small piece of DNA matched to the DNA template being measured has two special molecules attached, a fluorescent reporter dye (R) and a quencher (Q). While both molecules are attached to the probe, the fluorescence of the dye is suppressed by the quencher. These probes bind to the template DNA after it has been denatured into single strands but before it has begun duplicating, making sure that all duplications of the template interact with the probe.
During the PCR reaction, Taq DNA polymerase extends the primer through the polymerase activity, as it approaches the probe it displaces the probe and cleaves it through the 5′ to 3′ exonuclease activity. This separates the reporter dye and the quencher dye from the probe, which results in increased fluorescence of the reporter. Accumulation of PCR products is detected in “real-time” directly by monitoring the increase in fluorescence of the reporter dye with an automated PCR system.
The assay which you would wish to perform is called two-step reverse transcription-polymerase chain reaction. In this assay, two enzymes are used namely, reverse transcriptase to produce single-stranded cDNA copies, which are then used as templates in an amplification reaction catalyzed by a thermostable DNA polymerase. This assay is the traditional method of RT-PCR in which the two synthetic reactions are performed separately and sequentially.
The TaqMan Assay is a real-Time PCR assay which detects the accumulation of amplicon during the reaction. The data is then measured at the exponential phase of the PCR reaction. The assay which you may plan to perform using average regular use PCR master mix is a type of conventional PCR using agarose gel which is not as precise as qPCR. By using the regular use PCR master mix, you cannot perform qPCR because for qPCR one requires the fluorescent reporter molecule such as fluorescent dye, a labeled oligonucleotide primer or probe such as (TaqMan Probe) for fluorescent detection which is monitored by the automated PCR system. Real-Time PCR makes quantitation of DNA and RNA easier and more precise than conventional PCR.
So, if you wish to use the average regular use PCR master mix, you need to perform the two-step reverse transcription-polymerase chain reaction and not qPCR.
Best.
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Actually, I'm using the PCR purification kit of Qiagen, I am amazed why the solution is yellow, and as they indicated if pH >7.5 then add 3M sodium acetate, the solution turns again yellow from orange or purple. what is the chemical mechanism behind that?
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I think that the pH indicator is the buffers is Thymophthalein, pH shift from colorless to blue at pH 9.3
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We are interested to analyze the expression of a test gene by Real Time-PCR. Primers and PCR conditions were the same as in a previous paper with the same organism but in our hands the product is not obtained. Primers of housekeeping genes work well. What do you recommend to overcome this?
Regards
Raúl
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I have been through this too, and sometimes they just don't work, just because a primer is published doesn't mean it will work. But before you discard them you can try some things. I agree with Paul, first check in PrimerBlast if the primer sequences are specific to your target, if they do, then try different melting temperatures, you said the PCR conditions were the same and even if the sequences are the same the Tm can vary between laboratories. Also try to standardize them in a sample where the gene is highly expressed, sometimes you can't see it just because it isn't present in the sample. Hope this can help, if doesn't go for Oscar's advice.
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I have been trying to insert a 1.2kb fragment in pGEMT vector. I have used taq polymerase for the PCR amplification and did use gel extraction process to purify the PCR product. After ligation and transformation into the vector I got 7 white colonies in blue white screening but none of them has the insert within. Can anybody please tell me what can be the possible reason and how to troubleshoot?
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Sreya Ghosal, did you add X-gal and IPTG to your agar plates? If you did, then the problem might be with the vector or, more specifically, with lacZ gene (possible mutation?). Has the vector worked before for blue/white screening for anyone?
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Hi, this is about what is polymerase chain reaction what is its scope, and Application in the biology field, and how it amplifies genes. How Is this process used in gene cloning techniques
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Anyone know of a buffer that will work adequately for PCR with Takara's Primestar GXL? I have tons of enzyme but am out of buffer. Takara will sell a pack of extra buffer tubes, but it won't arrive for a month!
I have old tubes of Pfu, Taq and Vent buffers or could make up a generic buffer if I knew the pH, ionic strength and Mg2+ concentrations that GXL likes, but Takara won't say...
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you can attempt to create a generic PCR buffer with common components and optimize your PCR conditions. Here's a basic recipe for a generic PCR buffer. Generic PCR Buffer:
10x Tris-HCl buffer (pH 8.3 to 9.0)
50 mM KCl
1.5 mM MgCl2 (you may need to optimize this concentration)
You can adjust the pH using concentrated Tris base or HCl. The pH range is broad because different enzymes have varying pH optima. For Taq polymerase, a pH around 8.3 is often used, but some enzymes may work better at a slightly higher pH.
Regarding Mg2+ concentration, you'll need to optimize it. Different polymerases have varying requirements for Mg2+. For a start, you can prepare a range of concentrations (e.g., 1.5 mM, 2.0 mM, 2.5 mM) and test them in your PCR reactions to see which gives you the best results.
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I need a suggestion. I am working on molecular characterization by using an SSR marker. I have recorded data in 1, 0 in format. What will be the input file for association mapping?
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Thank you sir
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I have run multiple agarose gels in which I have at least two combs.
I continually see the lower part of the gel visualising much fainter than the top portion.
The last band of my ladder (250 bp) often disappears. It is quite a problem as I often don't see faint bands in this area if I have loaded my PCR products in the lower portion of the gel.
See images attached - look especially at the ladders in the top vs bottom
In the one image you can see the gel itself after a run in the tank, where the loading dye is significantly lighter in the bottom portion than the top - so it's not a problem with the visualising equipment.
I have run the gel for different times (30-60 min) as well as at different voltages (100V vs 120V) and see the same.
The TAE buffer has been changed
I have observed the same with another gel tank.
The intercalating dye, SBYR Safe, has been replaced with a new aliquot.
Other individuals have also experienced the same issue
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I use ethidium bromure (BET) to visualize PCR products. This dye migrate in opposite direction of DNA. If my gel run for a too long time the dye risk to be over of small sizes. I supose it is your case.
What you can try: increase agarose concentration at 2% or 3%.
Reduce the time for the migation : 15 min should be enough. You can adjust the voltage.
You can increase your SBYR amount.
Another tip : don't use detergent to clean your gel tank
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If my plasmid doesn't have Ecor1 and sal1 but i need my plasmid to have those, I am aware that i can make primers with Ecor 1 and sal1 overhangs but how is that done from the beginning?
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Dear Yukti
you can certanilly do it trough mutagenesis. However i suggest to you to evaluate enzime free cloning approaches as the PIPE cloning (that could be also used for plasmid mutagenesis) that let you free to the presence of restriction enzimes.
if you are interested to know more details about it, read the following papers
or look to the following videos available on my blog, ProteoCool
best regards
Manuele
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Hi,
1. Does any one know what is the maximum length of amplicons we can amplify using PCR? I need to amplify fragments for assembling a big vector (approximately 100kb).
2. Do you think it's achievable if I amplify 10 genes of approximately 10kB and stitch them together using GIBSON assembly?
Any recommendations/suggestions are appreciated.
Thanks
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Harculase II claims it can do fragments up to 23kb.
If Gibson assembly does not work you could always use sequential recombineering or SIRA to assemble your final construct.
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The Quantstudio 3 qPCR machine works really well with low-ROX SYBR premix PCR, I just concern about whether it can work in high-ROX premix or no-ROX premix? In the software, the reference dye can be chose as ROX or none, etc, but not having options like low-ROX or high-ROX?
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Thanks for your recommend. I tried by myself and it looks like not good idea for using high-ROX premix with Quanstudio 3 machine :))))
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I have 2 protocols for PCR sample purification. One uses Sodium for salt and other is using Potassium for salt. What is the reason for using different chemicals for salt in purification protocols. Which one should I use?
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The principle behind both of them is the same; however, Potassium is more prone to induce precipitations than Sodium, which may be problematic for some analytical techniques. If these consideration do not affect your analysis, you may use Sodium salt for easier handling.
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Who uses Tetra-Arms PCR method ? Who can help us in using the method specific in using primers(outer & Inner Primers) ??
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Dear you can use this tool
but finally you should try the primer because primer designs by bioinformatic tool for ARMS_PCR some time fail in real discrimination of the SNPs
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I have carried out a PCR reaction, resulting in a product of ~227bp. I quantified the raw product using PicoGreen, while this isn't entirely accurate with other components potentially interfering, it resulted in concentrations of ~ 200ng/ul. Gels also showed good bands.
However, after PCR purification with the PureLink kit (using Buffer 2, not Buffer 3) I have been left with 7 ng/ul (Nanodrop figure) and a negative PicoGreen value. I used 23uL of PCR product for purification.
Can anyone help with this drastic loss of product? I tried to re-run the elute through the column a second time, with no difference being seen!
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are you sure that the binding buffer had isoprpanol added and that the wash buffer had ethanol added either of which would result in low binding of dna to the column. you could try eluting the purified dna with hot (70c) elution buffer left on the column for twice as long as recommended which does increase yield but the likely explanation is poor binding in the first place
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I 'll design the large sequences (approximately about 1000 bp) as the homology arms.
so for transformation, which ones are better? the linear integrative plasmids or linear PCR cassettes?
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Hi again, just for info you don't need large sequences of homology to promote recombination. 20 to 40 bases are enough...
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I am amplifying gene (1488 bp) from RNA genome using specific primers and run in reverse transcription PCR(RT-PCR). The result, I am getting 2 bands my target band (1488bp) and extra band with size ~ 400bp)
To remove the extra band, I cut my target band from gel and purified my product by using Purification from gel -Agarose Gel DNA Extraction Kit. Then amplified PCR product, but I got the same result my target band (1488bp) and extra band with same size(400bp) .
I repeated this several time and every time found the same thing.
Tried using a different annealing Temp(high and low) , but the problem persisted.
i attached the pic of gel (lane 1 ladder, 2 and 3 lane PCR product )
I would really appreciate any advice on how to troubleshoot this issue.
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Thanks all for offering your valuable advice.
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Gene around 5000bp
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Why not design primers to make 2-3 pieces and clone them using Gibson Assembly? I assume that your final goal is to clone into an expression vector. Besides, my answers might be more helpful if you are more specific about the gene (GC-content...).
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I am doing overlap PCR to join 5 fragents to form expression cassette and product is always trapped in the wells and is not migrating down, I did purification by PCR buffer A but still the result shows bright light up and down and and a less intense band if anyone can guide me I will be thankful..
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Are you trying to join all 5 fragments at once? You didn't mention the size of any. I assume the final product is quite big. Try using a range of different percentages for the agarose gel and run your product. You may find the optimum percentage of gel that will show the migration.
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Actually I am having issue with double purified PCR bands for gel electrophoresis..
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You cannot send the mixed pcr product for sequencing. It will look like a mess of peaks. Your main options are
1 cut the bansd out of the gel and column purify each band separately and sequence. This is best if you think that the 2 bands are related like normal and deleted template amplimers
or 2 add dmso up to about 8% maximum and hope that the increased stringency removes the wrong band
or 3 increase the annealing temperature until the wrong band disappears,
2 and 3 deal with the problem of the 2 bands not being rellated and one band just being caused by inappropriate primer binding.
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Qiagen PCR purification kit is the DNA purification kit., after following the manufacturers protocol I am unable to get the yield.
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What is your starting input? How are you measuring the yield? Is all the ethanol gone after the wash steps?
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Can anyone help me understand what do different bands mean in gel electrophoresis after performing PCR?
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It probably means that there is a small amount of non specific priming in your pcr. To minimise it you can use less dna, less primer and make your annealing and extension times shorter in your pcr reaction, If these do not work then adding 5% DMSO to the reaction mix often works
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There is no change in the experimental conditions or the experimental procedure, but the electrophoretic bands become very shallow in the latter one, and this has been happening many times lately.
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The problem could be the pcr. Order new primers, run the pcr at 2c lower annealing temperature , run 3 more cycles of pcr and do not run the sample on the gel for so long and you should get a stronger amplification. The suggestion of using both a positive and a no template control is a good one
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I would like to know how I can find out the size of the amplicon from my primers. I have these primers: Forward: GCTCCGCAGCCTGCATGGA Reverse: ACGATGCCGCCATCCTCCT. I saw the suggestion of putting the sequence of the forward primer followed by four N's and then the sequence of the reverse primer, but it doesn't work, and I also can't find that sequence in BLAST.
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hello, I dont know any thing about this matter. sorry
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There is no amplification. I don't know what happened. If anyone can help me with some advise.
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Hello, as for me, I conducted a research on Herbivore(elephant) diet from their fecal sample source.
I used Uniplant primers and noticed no amplicons on the first run. I then decided to just try out doing a second PCR on the amplicons using the same primer and check...(more like a nested PCR) and saw bands in the gel and a third one just to be sure, since the BP are almost close to primer dimers, I did a third PCR same primers on the 2nd amplicons, the bands were brighter...and when purified and sequenced, I found out the diet contents.
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I did gel electrophoresis for my PCR product. The size for the amplification was inconsistent but still in the targeted range. Then, I proceeded with purification but the end result obtained was increased in size (100+ bp) than the targeted and previous result. why?
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If your purification method involved a high salt step and salt was carried through with the purified dna then it would run slowly (larger)
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Hello dear scientists.
Can the specificity of primers change over time or depending on the sample? It looks like the primers I used in my recent PCR to target exon13 of CFTR gene on chromosome 7, actually amplified a different location on chromosome 15. The gel looks like ladder with two sharp bands.
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The company probably put the primer sequence on the vial. It might be worth Blasting the primer sequences just to check if they have any relevance to cftr in spite of their name might clarify the problem
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My lab is having trouble with mycoplasma contamination, so we're trying to figure out the source of contamination. We've already verified that our DMEM and FBS are clean, next we want to test our trypsin (0.05%). But since it can digest proteins, we're worried that it might render our PCR reaction useless. Has anyone had experience with using trypsin in a mycoplasma PCR test?
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the most likely source is your inoculum cells
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For diagnostics of Avian infectious bronchitis viruses, I use an end-point RT PCR reaction, and before the RT srage I do a primer annealing stage with the target reverse primer (2uM). My queation is: If I have 2 targets, is it right to use a 1:1 mixtue of their reverse primers (2uM) instead of using rendom hexamers, in order to get better sensitivity for these two targets amplified from the same RT product?
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Assuming the sequence heterogeneity is low at your primer binding sites you could replace the random hexamers with your reverse primers.
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Hello Everyone,
the description of NEBuilder HiFi DNA Assembly mix says: "No PCR clean-up step required".
I always used this mix after DNA extraction from agarose gel , but now we have some problems with this step... Does anyone use this kit without PCR clean up, directly on the PCR product? Does it work? How many of the product is necessary? How could I measure or estimate the concentration of the DNA in this case?
Thank you very much,
Magdi
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Hi Magdolna Keszthelyi , I 've nerver used the NEb HIFI without the purification on agarose before but if you want to try I think you can estimate your concentration of DNA on agarose gel with the DNA ladder you're using. You can load a few microliters and estimate (approximately) the concentration with the intensity of the band you obtain (you can read about it here https://www.researchgate.net/post/How-do-I-quantify-the-DNA-accurately-on-agarose-gel);
May I ask what problem you're having with the purification on gel ?
Have a nice day
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How to calculate the optimum annealing temperature for PCR?
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Here's a general guideline to calculate the optimum annealing temperature for PCR:
Initial Estimate: Start with a rough estimate of the annealing temperature. This is typically done by considering the melting temperatures (Tm) of the forward and reverse primers. A commonly used equation is:
Tm = 2°C(A+T) + 4°C(G+C)
Calculate Tm for both primers and take an average. This initial estimate is a starting point for optimization.
Also, this formula is valid for primers up to 20 nucleotides length. For longer sequences, especially those with lengths greater than 20 nucleotides, using more advanced formulas to calculate the Tm!
Gradient PCR: Perform a gradient PCR, where you set up multiple PCR reactions with varying annealing temperatures in small increments.
Amplification Optimization: Run the gradient PCR and then analyze the results. Look for the reaction that produces the strongest and most specific band on an agarose gel or another suitable method for visualizing DNA. A single specific band indicates successful amplification of the target sequence. Bands at higher or lower temperatures could indicate non-specific products.
There are various software tools and online calculators available that can perform calculations. Some commonly used ones include: OligoAnalyzer (IDT), Primer3, UNAFold, MeltCalc
When working with longer DNA sequences, using these specialized tools can provide more accurate Tm values and improve the success of your PCR experiments.
''The most appropriate Tm temperature is specified in the synthesis reports coming from the companies that you have made the primer synthesis.''
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This is my PCR 1.2% agarose gel result. I used a 25uL reaction system (0.3uM primers, 12.5uL Taq master mix, 1uL genome template and 5.5uL H2O). The primer Tm is 62℃, and reaction procedure is 94℃ 3min--(94℃ 30sec--57℃ 30sec--72℃1min)25cycles--16℃. What is the possible reason for the existence of the bottom band around 100-250bp? How to optimize my PCR procedure? Thanks!
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Hi Paul, Hi Anita
it seems that PCR amplicons are not at same size among samples. is it true?
in addition to degraded DNA, you should also see smears above PCR bands, you can have excess of primers and also multimerizations of primers.
primers are always in excess, and overall PCR must be standardized with right conditions, right amount of each goods...you could try using less amount of primers, just to see.
all the best
fred
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I am currently struggling with genotyping a set of my mice. We obtain genomic DNA from tail lysis and my positive control I am using is from a mouse I have previously genotyped and is homozygous for the gene I am looking for. My controls are working however, I do not get any banding for the samples from the litter I am currently trying to genotype. I have tried re-diluting my working primers in milli-Q water, changing the TAE buffer, re-isolating the DNA, modifying the amount of template, and optimizing the annealing temperature for my primer. This PCR was working perfectly fine before but now it is not does anyone have any suggestions?
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I agree with Katie A S Burnette about the pcr reagents and about pcr inhibitors especially about using less dna.
One other possibility is the quality of the reagents used to lyse the tail sample and prepare the dna. For instance if using a hot alkali method then NaOH and KOH both absorb CO2 from the air and become carbonates which will be bad for the dna preparation. It may be worth preparing new dna isolation reagents and using these in the dna preparation
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Polymerase Chain Reaction (PCR) testing is a molecular biology technique used to amplify and analyze DNA or RNA samples. So, To perform PCR testing, which list of laboratory apparatus and equipment is required?
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First and foremost, you will need a separate laboratory space/room maintained at the required working temperature to conduct the PCR test. PCR should be conducted in a PCR hood which is a workspace enclosed on three sides that provides a space for performing the amplification of DNA.
For the PCR technique, the most important requirement is the thermal cycler also called PCR machine which is an instrument that amplifies target nucleic acid sequences into millions of copies via PCR. Besides the thermal cycler, you will require a -20-degree freezer to store the samples and PCR reagents.
The main PCR reagents include:
1. The PCR primers which are short strands of nucleotides of approximately15–30 bases. PCR primers are complementary to the template DNA and serve as a DNA synthesis starting point for DNA polymerase.
2. DNA Taq polymerase is a type of DNA polymerase that is active at high temperatures and since PCR is carried at high temperatures, this enzyme is the best choice. This is the molecule that helps in amplifying the sample.
3. Deoxynucleotide triphosphates (dNTPs) are the building blocks of nucleic acids. They consist of four basic nucleotides - dATP, dCTP, dGTP, and dTTP, and they are needed to synthesize DNA by DNA Taq polymerase.
4. PCR buffer which provides a suitable chemical environment for DNA polymerase to perform efficiently. They help maintain a stable pH during PCR and ensure that the reaction is conducted under optimal conditions.
5. Lastly, the DNA template which is the specific sequence of DNA that is the gene of interest which needs to be amplified. It is the starting material for the PCR reaction.
Additionally, you will require a tabletop microcentrifuge to centrifuge the sample/reagents, a vortex mixer to mix small vials of liquid, and a water bath set at 37 degree C (optional).
Among the consumables, you will need 0.2ml PCR tubes, filter pipette tips that prevents PCR contamination, and micropipettes {P2 (0.2 - 2ul), P10(0.5 – 10ul), P20 (2 – 20ul), P50 (5 – 50ul), P100 (10 – 100ul) and P200 (20 – 200ul)}.
Please note that if you wish to analyze RNA samples, Taq polymerase will not work on RNA samples. The incorporation of the enzyme reverse transcriptase (RT), however, can be combined with traditional PCR to allow for the amplification of RNA molecules.
Best.
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I'm having trouble obtaining a well-defined band of the COI gene in mollusk (Aulacomya atra) adductor muscle samples, despite using 35 cycles and TBT-par. Other mitochondrial and nuclear genes amplify smoothly using the same samples, such as the mitochondrial 16S gene and the nuclear 18S and 28S genes. What recommendations would you provide to optimize the PCR and achieve a clearer band?
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You can use another type of DNA polymerase for example Phusion polymerase which is highly specific and has a low elongation time with the presence of ssO7D, or even KOD polymerase which has proofreading ability, that will decrease the error rate that you can face
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Hi all,
I need support in calculating the analytical sensitivity of molecular assay ( detection of DNA) for detection of an bacterial infection from oral swab samples. Usually we spike various dilutions of pure culture suspension to the oral swabs collected form healthy volunteers and perform the assay.
For example, if we spike 10ul of 105 CFU/ml of Pure culture suspension to the oral swab, perform the DNA extraction and elute the DNA in 200ul of buffer, use 5ul of template DNA for 20ul PCR reaction, What will be the CFU equivalents in the 20ul reaction?
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Thankyou Michael.
Now I am confident that my calculations are right.
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Hello
Good time everyone
I ran the cloning product on a 1% gel and observed the following band. Can anyone tell me what is the reason why my band has widened?
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I disagree, the marker looks fine, the image just does not have a high resolution and therefore looks grainy. Yes, there are some smudges on the gel, but this - as we all know - sometimes happens. Just load less PCR product and you will see a sharper band.
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I am trying to optimize a SYBR assay and one primer set is giving me dimers every time. I've increased the annealing temperature to close to the calculated value for that primer set, and the other set is running great so I don't want to mess with the thermal profile any more. I've already decreased the primer concentration to 50 nM (25 ul reaction) and there are still dimers amplifying around Ct 30-35. I know the next step is back to primer design, but we are hoping to compare this work with another published paper that used the same primers. Maybe I just need someone to tell me the hard truth that it's time to give up and redesign. Thanks!
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I suppose it really depends on _when_ you are getting primer dimers.
Given that PCR is an exponential process once there's even a single template there, if you run a reaction for enough cycles, you'll almost inevitably amplify something.
I run my qPCRs for 40 cycles, and I will occasionally see signals (and melt peaks) consistent with primer dimers in my negative controls, but these usually only emerge after 36+ cycles. Given that my actual positive signals emerge at much earlier cycles (20-25), I can assume that even if primer dimers are contributing to the fluorescence in my actual samples, those primer dimers are contributing very, very late in the process, long after the 'quantitative' bit of the qPCR has been determined.
I mean, the ideal world is 'no primer dimers', but still. If you see primer dimers emerging in the late 20s/early 30s, it's perhaps cause for concern, but anything after cycle 35 is readily identifiable as non-specific amplification.
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Is it possible that the amplification failure products can be visualized in electrophoresis? Due to the failed amplification results it shows bands in my electrophoresis with bands that are quite clear. My amplification curve clearly shows amplification failure, but when I look back at it with electrophoresis there are some obvious bands, how is that possible?
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I think this band is because of the primer bind with itself we call it dimer
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I have set up a PCR using primers that amplify only the exons of my gene of interest. The template I am using is cDNA (mouse samples). I set up this PCR a year ago with an annealing temperature of 57 C, using Kapa Master Mix (the product size is about 2.8 kb). I repeated this PCR 3 weeks ago and I could get sharp bands for some of the samples (Fig. 1), but when I repeated the PCR again (2 days after), I only got smears (Fig. 2) (In the next repeatations changing extention and cycle numbers, sometimes for some samples bands appeared (Fig 3) and in the next round for the same samples I only got smears!) I changed each component and used filtered tips. I also tried different extension times and reduced cycles. I have used diluted templates (1/2, 1/4 and 1/8). I cannot understand why the PCR that worked is not working anymore! I have changed the stock of primers and I still get smear in my samples and water control (Fig 4). When I use other primers targeting different gene, it works (Fig 2). Regarding the water control, using the same master mix but different primers resulted in a clean water control compared to the water control for my gene of interest.
Could you please share your thoughts on this? I appreciate any comments.
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try another mastermix company type ,it may help
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I did a transformation of the BL 21 cells with my plasmid DNA. Then I ran the dna plasmid pcr to check the inserted dna and both samples showed bright, clear bands. However, when PCR with a larger volume was available for sequencing, one sample was very faint (with increased plasmid volume) and one sample had many bands. Is the first result a false positive?
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Huyen Ngo Please, repeat both PCR in small and large volume, and if small is OK, just amplify DNA in several tubes for sequencing. Large volume has different kinetics of the PCR.
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Hello, we've been trying to replace a ~100bp region in E. coli with a ~2.5 kb DNA using Red recombination. The insert is a linear PCR product containing the chloramphenicol resistance marker. After recombination, we selected singled colonies on chloramphenicol plate. The genomic DNA of the colonies were isolated and confirmed by PCR using different sets of primers (pls see the attached figure). All picked colonies gave similar results. Using primer sets of (1,3) or (1,4) or (2,4), we got PCR products with expected length. Using primer sets of (2,6) or (5,6), we got no PCR products or products with wrong length. We thought it may be caused by bad design of the primer 6, however, changing the sequence of primer 6 resulted in similar outcomes. And weirdly, we also did PCR using primer (1,6), the product has a length around ~100bp suggesting no insertion. However, based on the PCR results using primer (1,3)/(1,4) and also the phenotype, clearly the insertion happened.
Such results suggest that the insert is linked to the upstream region correctly, but something is wrong at the downstream region. What happened? Any advice would be much appreciated! Thank you.
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Robert Adolf Brinzer We transfected using PCR product. We did PCR on the untransfected host strain using different primer pairs. Primer (1,6) gave the expected result. Other primer pairs didn't result in clear bands.
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Hello
I am running a PCR to quantify bacteria (namely Escherichia coli). I would like to count only live cells and I have seen there are several protocols that employe chemical dying of the death cell's DNA followed by DNA purification and then qPCR.
To avoid the costs of purification, is there a protocol for differentiating live from dead cells that includes simple boiling of the cells followed by PCR?
Thank you
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Assuming the dead cells are porous or at least prone to pore forming agents you could do a DNAse digest first.
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Good day everyone,
I had a problem with my RT PCR machine, while it was performing it stopped suddenly because of another program, and I didn't get a chance to resume it (Although there is no option like this), and it reached cycle number 18 out of 40 cycles. while I was searching in the settings I found an option called quick start and I started my run over again. In this case, should I expect any right results?
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Hi there,
In my opinion the problem is that you cannot trust the results you get from this experiment.
As long as you still have cDNAs and reagents, just repeat the experiment to be on the safe side.
Good luck,
Sebastian
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(Text in bold was the error conducted)
Mutation: WPD to FPD
Restriction site: BspEI
Template size: 7.16 kbp
The following reagents were added to make a total volume of 50uL:
5x PCR buffer: 10uL
dNTPs: 2uL
Primer mix: 2uL
dH2O: 25uL
Phusion polymerase: 1uL
template 60ng: 10uL
The experiment method wanted 5ng/uL plasmid DNA solution to add 50ng of template DNA to the reaction. However I made an error in dilution and resulted in 6ng/uL, but still added 10uL as indicated above.
Would or How would this affect PCR results? Thank you in advance for your help.
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Using more template just means there is a slightly higher chance of transfecting your starting plasmid into your competent cells and not your PCR mutagenized product.
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I have a problem with real time PCR, please help me if possible
I was extracted total RNA of human peripheral blood and OD260/280 was in range of 1.6-2. cDNA was specifically synthesized with a stem loop primer specific for miRNA. I did real-time PCR for miRNA with the SYBR Green method with following conditions in 40 cycles: 94 degrees for 10 minutes of activation. The cycling temperature was 95 C° for 15 seconds, 58 C° for 30 seconds, 72 C° for 45 seconds.
I have done real time PCR several times, and every time the same problems are repeated: 1- Some samples have CT value and some samples do not, or in two duplicate reactions, one sample has CT value and the other does not. ,
2- the melting temperature curve does not show the existence of a specific product. I changed the annealing temperature from 57 to 60, used 1/10 and 1/20 dilutions of the cDNA, but the results did not change. Please help me to solve this problem
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Look at the scale of the Y-axis on your graphs. You are getting no amplification. This is just background noise.
Add in a good positive control and try again.
Good luck!
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I want to do rolling circle amplification with phi 29 polymerase enzyme . Previous article show they use 10x reaction buffer. Can I use 10X reaction buffer from thermofisher for the PCR.
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Hi there,
Use the 10x stock of the buffer specific to the polymerase you intend to use.
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Hello,
I am currently working on making mutations in my plasmid conducting Quikchange. The plasmid size is 3800bp and I am using primers with several mutations. However, both my parental plasmid and the mutated product have two bands (see attached gel)
The gel shows L1) GeneRuler 1kb ladder, L2) my parental plasmid from a mini prep, L3) the plasmid product (mini product) after conducting PCR -> DpnI -> PCR cleanup -> Transformation -> Mini, L4) the plasmid product (mini product) using the mini product from L3 as template for a new round of PCR -> DpnI -> PCR cleanup -> Transformation -> Mini.
It looks like I get less of my desired plasmid for each round (~3800 bp) and more secondary product. The loss of primary DNA is not feasible, as I need to conduct three consecutive rounds of mutation PCR. Can anyone help me explain what the secondary product may be? I have never seen it before.
The lab works with M13 phages containing ~6400 bp ssDNA. Could the upper band of L2 be the plasmid integrated into phage genome (~10,000 bp total) from previous experiments? And could L3 and L4 be the product of more plasmid integrated into the phage genome? Or is this genomic DNA contamination?
I hope that you can help.
Kind regards Emil
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If you think you may have M13 contamination, then transform your plasmids into a non-male host strain (F-) which won't be infected by M13 and then make a new DNA prep.
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Hello dear scientists
Where can I find the Liz 1200 size standard for Gene marker software ?
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Thermo fisher or their local stockist
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Hey All,
How i can avoid or prevent Primers dimer during PCR amplification? I Know about the solutions that creating optimals primers and using proper primers concentration, also to change or adjust the annealing temperature.
But Iam looking for another solutions can use it to Helps us .
Second, what the best application can helps me to to check PCR primers ( Free app).
Thanks,
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Hi, so I was just wondering what would be a good Ct value to aim for in real time PCR. My lab typically uses about 150ng of cDNA per well for a 10uL real time qPCR reaction. But in a recent time course treatment experiment, not a lot of RNA was collected and my PI wants to run 13 genes. I didn't have enough cDNA to make triplicates of that many genes and I ended up running a second time course to get more RNA (but that isn't ideal obviously). I read that real time qPCR can use as little as 100 pg of cDNA. The highest Ct that I have gotten with the old procedure for a gene has been around 26. Can I dilute it so that the Ct is around 35 or is that too close to 40 for comfort? I realize that a 2^9 fold dilution is a bit extreme, but this is just for future reference. Thanks.
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150ng per well is masses.
You are probably getting high Cqs because you're massively overloading the well, and moreover adding too much cDNA synthesis buffer (which is inhibitory to PCR).
Dilute your cDNA. Always dilute your cDNA.
This seems to be one of those things that "everyone knows but never says", so people keep having to discover this the hard way.
For reference, I use ~8ng per well, and I work on some pretty low abundance stuff.
Make your cDNA, dilute it ~1/10 (I dilute 1/20 and then use 2ul per well because I find that easier, but 1/10 and 1ul per well is the same thing), then run your PCRs again.
You may well find that you have much better Cq values. Also, your cDNA will last a lot longer because you have a much larger volume now.
As a ballpark reference, Cq ~35 means you started with probably only one target molecule in that well. You will usually find that Cqs from ~29-35 are pretty variable between wells, because when you're dealing with (and quantifying) countable numbers of molecules, you can easily get uneven partitioning between replicate wells.
If you really want to quantify targets of such low abundance, you can simply include more replicates: the average behaviour of six replicate wells is much more representative than that of three replicate wells.
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According to IDT oligo design, Ta Optimum = 0.3 x (Tm of primer) + 0.7 x (Tm of product) – 14.9; where Tm of primer is the melting temperature of the less stable primer-template pair, and Tm of product is the melting temperature of the PCR product. How should I calculate the Tm of my pcr product for the above mentioned formula?
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You should perform a melting curve to know the Tm of your PCR product. You can also estimate the Tm using this formula but it assumes certain conditions: https://www.rosalind.bio/en/knowledge/what-formula-is-used-to-calculate-tm
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There are many companies that offer primers for pathogens found in industry animals (cattle, equine, etc.), but I cannot really find anything related to pets (cats, dogs, etc.). Is there something I am missing and maybe it is just easier to try to design the primers myself?
Thanks!
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Hi,
Did you consider talking with your mentor as well as trying to use PUBmed?
Otherwise, a more specific question will help you find a specialist.
Regards
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I am trying to design a primer for a specific gene and the Tm is coming near to 60 degrees when I select a minimum of 30-35 bp, for forward as well as for reverse primer. As the template is too long I think it might lead to non-specific binding, what are the possible solutions for this problem? I am using Snapgene to design primers for cloning
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1. Adjust primer length: Increase or decrease primer length slightly.
2. Evaluate Tm prediction methods: Use different algorithms or tools for Tm calculation.
3. Adjust primer sequence: Make small modifications to fine-tune Tm.
4. Check for secondary structures: Identify and minimize hairpins or self-complementarity.
5. Consider primer placement: Avoid repetitive or variable regions.
6. Follow primer design guidelines: Adhere to established recommendations.
7. Validate and test primers experimentally.
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For sanger sequencing reaction with BigDye terminator mix, I will need to purify the excess unincorporated terminators, primers, dNTPs from my PCR product. But in times when I won't be able to purify immediately, how do I store the unpurified PCR product? Is it acceptable to simply store them in -20C until I am ready to purify? Will the terminators and other components of the BigDye reaction mix interfere with the PCR product?
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Read the protocol that came with the BigDye kit. They recommend keeping the plate on ice or storing at 4C, but not indefinitely.
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I am doing absolute quantification RT-qPCR for the beta actin gene. I start with RNA, turn this into cDNA and use that for qPCR. I am going to get a custom oligo of the PCR amplification product to use for making the standard required for absolute quantification.
This is how I got the sequence for the custom oligo. First, I input my primers into this website: http://www.bioinformatics.org/sms2/pcr_products.html along with the mRNA sequence for beta actin. The website gave me the amplified product, which is 61 bp as expected.
My question is should I use this result sequence as my oligo, or should I reverse complement it? I am wondering this because I input the mRNA sequence into the website to get the amplicon, but I am using cDNA which would be the complementary sequence. Please let me know. Thank you
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Can you get the synthetic oligo as a double-stranded molecule? Single-stranded molecules will not work as a standard.
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Hi everyone,
Why could the band appear smudged?
I have amplified a 300 bp DNA G-block using Q5 2X mastermix (NEB) for 34 cycles, using 10-uM primers and 1 ng of template G-block as per protocol (M0492S). The primers generate overhangs and are as follows: fwd: Tm=47'C, 20mer, 35% GC, rev: Tm=59'C, 8%, 148mer, 8% GC. My colleague ran gradient PCR to determine best annealing temp of 57'C.
Gel electrophoresis: 1% TAE agarose gel, ran at 120V for 30 min and 90V for 15 min at 400mA. Ladder: 1Kb Plus DNA ladder (NEB)
PCR regime: 98'C:30 sec initial denat., 34X[98'C:10sec, 57'C:30 sec, 72'C: 20 sec], 72'C: 2 min final extension, 4'C hold.
Lane1: ladder
Lane2: 20 uL of PCR rxn+3 uL loading dye,
Lane3: 20 uL -ve control (no template) + 3uL loading dye.
I am wondering if:
1) I added too much PCR rxn per gel?
2) Gel concentration is too low?
3) Rev primer is way too large and formed dimers
4) Ladder did not separate too well so it is unclear what the MW of the smudge is
5) Annealing temp is not suitable?
6) Extension time too long?
7) Running buffer needs to be changed?
8) Voltage is too high?
9) Too ma