PCR - Science method
PCR is an in vitro method for producing large amounts of specific DNA or RNA fragments of defined length and sequence from small amounts of short oligonucleotide flanking sequences (primers). The essential steps include thermal denaturation of the double-stranded target molecules, annealing of the primers to their complementary sequences, and extension of the annealed primers by enzymatic synthesis with DNA polymerase. The reaction is efficient, specific, and extremely sensitive. Uses for the reaction include disease diagnosis, detection of difficult-to-isolate pathogens, mutation analysis, genetic testing, DNA sequencing, and analyzing evolutionary relationships.
Questions related to PCR
I am doing a trizol based RNA extraction on a knee joint sample. My 260/28o ratio is anywhere from 1.85 -1.95 for most of my samples, but my 260/230 ratio seems quite low (1.08-1.2). My RNA concentration also reads anywhere from 1800 ng/uL - 4000 ng/uL... I dont even know if that is possible. Does that mean my sample is super contaminated? What can I do to improve this? I need to run RTPCR on these samples eventually. Thanks for your help!
I have been trying to perform a single amino acid mutation using a the GENEART site-directed mutagenesis system. I isolated the plasmid, fixed its concentration at 50ng/ml for the mutagenesis PCR (received multiple faint bands on 0.8% agarose) then performed recombination reaction followed by its immediate transformation in "already provided chemically competent DH5a cells .Firstly i tried the as per the instructions given in the kit's manual ( no colonies), later I tweaked the transformation protocol ( made fine adjustments such as: increased ice incubation for after transferring recomination reaction by 8 mins increased heat shock duration by 60 sec, etc.) I tried transforming the DH5a cells with the wildtype plasmid and it did give colonies. But no colonies with the recomination reaction whatsoever. Any suggestion might save reactions in the kit for my further experiments. Thanks for your valuable time.
I have tried In-fusion cloning few times and it used to work in my hand. Suddenly, after few months when I again wanted to do some more cloning, it is not working as in I am not getting any clones in the test plates.
I have used gel purified vector (fully double digested as observed in gel) and gel purified PCR products. The homology overlap of the PCR products were 15 bp.
I have used 50ng of vector and the Vector : Insert molar ratio was 1:4. The In-Fusion reaction incubation was done in a water bath set at 50C for 1 hr. It would be of help if I can get any advice regarding this.
Hi, I am linearizing a 8kb lenti puro vector with PCR prior to Gibson assembly. I have precious few reactions in my Gibson kit so I don't want to use something until I'm sure what it is. Currently I am preoccupied with the mystery of why the PCR seems to work very well (get expected band) only when I use the tubes that melt in the thermo cycler, but when I run the same reaction with tube strips that never melt, I barely see any band (mostly a smear).
Reaction - 51.5uL total
Template Plasmid Dna 43ng/uL - 1uL
mol. bio. H2O - 35uL
DMSO (has been freeze-thawed) - 2.5uL
Q5 5X reaction buffer - 10uL
Q5 Polymerase - 1uL
10uM Primers (F/R mixed) - 1uL
According to eurofins genomic and snapgene my primer Tm are 65-66 C.
95C - 5min
95C - 20 sec
61C - 20 sec
72C - 3 min
25X cycles from steps 2-4
72C - 5 min
4C - infinity
I am currently running an experiment to see if splitting the reaction volume in 2 parts or increasing the Ta to 67C will help.
TCC CCA TCA TCC CAC TCT CCA (56.3) 57%
This is my Forward primer, Tm calculated using OligoCalc,
Temperature tried are 50.5, 52, 53, 54, 55, 57, 61 and 68C
Thermo Tm calculator suggests Annealing temp as 65C , have tried that also still no PCR observed.
Reaction has LA Taq polymerase, undiluted /diluted (1:10) primers, template 30-50ng.
95C 5mins denaturation
Varying annealing temperature
68C extension for 8mins (vector 6Kb)
cycle of 18
10mins 72C final extension.
Please suggest a way out.
I need to extract RNA from white blood cells (buffy coat layer) in human blood samples to do further RT-PCR studies. I am using Monarch's Total RNA Miniprep kit; however, the protocol card gives two a Mammalian Whole Blood option and a Tissue/ Leukocytes option. If I extract RNA from whole blood, what cells is the RNA coming from? Leukocytes, immature RBCs, etc?
If the RNA is from the WBCs, we would be able to do extraction directly on the whole blood sample.
I put a PCR reaction on (Used MyFi buffer and polymerase) and the annealing temperature was 54 degrees C. We realised after one cycle that the time was incorrect and reset the parameters. Anyone got any insights?
Here are the things :
4kb tagged DNA
Extension time provided is of 4 minutes
DNA concentration is about 1995.6ng/ul
I used 1:10 and 1: 100 dilutions as well but still not getting the band.
Please provide insights
I am trying to develop a universal LAMP assay for the CO1 mitochondrial gene. The plan is to amplify the gene of multiple different species using LAMP based on pre-designed sequences attached to the target DNA fragments via PCR.
Therefore, I designed PCR primers with a large overhang (100bp) that contain all primer sequences needed for a LAMP assay (F1/B1, F2/B2, loopF/loopB and F3/B3). So far, I was successful in amplifiying the product with the overhang primers but the LAMP reactions have not been as successful yet. My best results on agarose gels show smears and very low bands (100bp). Troubleshooting ideas I have tested so far:
- Various reactions times & temperatures
- Increased primer concentrations
- EtOH/SodAct purification and initial denaturation of target PCR product
Some other issues I thought of are:
- DNA target too long: ca 550 (from 3’ F2 to 5’ B2c): the target I am trying to amplify is longer than suggested for LAMP reactions (200bp)
- LAMP primers designed incorrectly: I was most unsure about the F1c/B1c sequences. From my understanding of the reaction they should be reverse complementary to the F1/B1 sequences so they fit when they fold over, is that correct?
- There are potential hairpins in my primers. I have checked LAMP primers of other studies and they are potential hairpin regions as well, therefore I was unsure how big their inhibition effect is
As this is my first attempt in designing a LAMP assay and maybe the approach is a bit unconventional, I am sure there are several factors that can be optimized.
I appreciate any further insights on this. Thanks all!
The goal of the project is to collect around 200-250 samples (primarily muscle tissue) from a bushmeat market and transport to laboratory for species barcoding. DNA extraction will then be performed, followed by PCR using universal mammalian primers (and if no product is yielded, universal vertebrate primers), confirmation of product using agarose gel electrophoresis, DNA purification, Sanger Sequencing, then alignment to the lowest possible taxonomic unit. The results will be compared to reported species by the seller (to assess misidentification) as well as used to identify protected species.
Current plan is to first use the following universal mammalian primers (majority of samples expected to be mammals):
MTCB-F (Size~1420) 5'-CCHCCATAAATAGGNGAAGG-3', targets cyt b (Naidu et al 2012)
MTCH-R (Size~1420) 5'-WAGAAYTTCAGCTTTGG-3', targets cyt b (Naidu et al 2012)
Then if those don't yield products use the following universal vertebrate primers:
L1085 (Size 215) 5'-CCCAAACTGGGATTAGATACCC-3', targets 12S rRNA (Kitano et al 2007)
H1259 (Size 215) 5'-GTTTGCTGAAGATGGCGGTA-3' targets 12S rRNA (Kitano et al 2007)
Question: This is my first time performing species barcoding of any kind (and also only have minimal bench experience, background is clinical); are these appropriate primers? Any critique/advice on methodology?
Thank you so much!
I am currently conducting a study about molecular characterization of Dioscorea spp. in Sri Lanka. I have prepared PCR products using both matK kim m13 and Trnh-psba primers.Clear bands were obtained through agarose gel electrophoresis but when sequencing only few base pairs were sequenced.
I used 2X Phusion Master Mix during my PCR reaction and after that I run the PCR product (10ul).
I can observe a faint band but I do not why it looks like degradation. I attached a picture to ask what could be the reason of this big smear on the gel?
Thanks a lot for your help.
I am trying restriction digestion of my PCR product by using BfaI. After restriction digestion, I could see only PCR products of size 425bp, but not the restriction products of size 240bp+185bp. Interestingly, the PCR product contained the restriction site (CTAG) but it does not digest the PCR products (Lane 5). Initially, I doubted the restriction enzyme, but it was working for genomic DNA digestion (Lane 1 and 3 are genomic DNA whereas 2 and 4 are respective restriction digested genomic DNA). 'L' represents the DNA ladder. I did not face any such issues with other restriction enzymes. Can anyone suggest how to troubleshoot the issue?
I m working on gene deletion in Salmonella Gallinarum by using lambda red technology . I have transformed pKD46 into Salmonella Gallinarum by electroporation. But I m stuck in transformation of PCR product .. Kindly guide me what could be the possible reasons for the failure..
I have a question that might sound silly. I'm performing a PCR with Q5 polymerase, in a reaction volume of 25mL. Now, I would need to run all of the volume of the reaction to excise a band. The question is, how much loading dye (I'm using a 6X TriTrack) should I add to visualize the bands in the gel?
I have a very troublesome time when I trying to build the construct of my gene. Since the gene sequence of my target insert is toxic due to the leaky expression, I used the competent cell that ordered from NEB company: NEB® 5-alpha F' Iq Competent E. coli (High Efficiency) to reduce this problem, and I encountered trouble when I tried to extract the DNA from the starters.
I used traditional digestion to get my insert and the vector, the I did ligation. After the colonies grew on the ampicillin plats, I did colony PCR by the use of my insert primers, it shows the very good result that my insert is inside of the gene of the colonies' cell.
Then I used the colonies to grow the starters (O.N. in liquid LB+amp.). I also did the starter PCR, but in the comparison of the positive control (the plasmid where I get my insert from), which has strong right size band; the samples shows nothing.
After that, I redid everything, and miniprep (i.e., the DNA extraction). In addition, I digest the extracted plasmid. The intact plasmid looks 2kb smaller compared to what it should be, also the digested bands.
What could be the problem during the process? If the idea that using low-copy number e. coli is not really working, is there any other way that I can use to build the construct with this toxic gene? My aim is to transfer the gene into tobacco and produce protein.
Thank you so much and looking forward to hearing any constructive suggestions.
Anyone knows of a software that can generate band patterns from whole genomes and custom primers? I have seen it for digestion patterns, but could not find one for PCR. My primers have some degenerate nucleotide, as I'd need to do fingerprinting PCR, so the software must also accept a few mismatches. Ideally, the software would be either free or low cost...or can be rented by the month.
I have done most of the project with real PCRs, but a few samples were lost in shipment, so I am looking for a work-around to complete my data.
Any suggestions are welcome!
I have gel purified a specific gene from pcr, using 2% gel run at low voltage for longer time to get better separation. I used the Qiagen PCR clean up kit.
The concentration is pretty good (~50ng/ul) considering how much dna can be lost with clean up kits.
I haven’t digested the product at all, so the primer sites/ cut sites should still be intact, and the purified product shows only one band, corresponding to my control.
Can I run another PCR, if I end up wanting to get a higher concentration of the gene?
I know the salts and residual oligos can cause issues, but post-purification, is there anything I would need to look out for in order to do a standard Taq PCR?
Our in-house mycoplasma testing flagged both my media types as positive for mycoplasma, but the cell line media samples I provided all tested negative. The tests were performed on different days and are going to be repeated, but I was wondering if anyone had any insight as to why this might have happened? Could it be a problem with the FBS source for each media type, or are the cells likely to be a false negative? Thanks!
I am working on the molecular characterization of several genes, using standard PCR. Is there is a specific approach that could be used for achieving this reduction?
Hi, i am an undergraduate student and i'm trying to clone a gene fragment in a plasmid vector for the fist time for me. I have some questions, one of them is that i was reading that in some protocols gives the advice that if you want to insert a PCR product it most have be phosforilated before the ligation step, so you have to use a kinase to achieve that, but i also understand that restriction enzymes (at least some of them) leave a 5P end. So when i digest my PCR product it will be enough to leave 5P ends? Thanks in advance for your answers.
I am trying to amplify 16S regions from insect gDNA samples. I am getting the target bands with low intensity. I am following standard NEB protocol for 25ul reaction tube. 7ul of PCR product was used for gel electrophoresis.
I ran samples (looking for c.bovis in swabs and tumors) using the same primes/probes, Taqman Master Mix, DEPC H2O that was used last week on samples that worked fine. These new samples that should had been negative tested positive for c. bovis. I replaced all reagents and reran the samples. Still positive. Some samples are negative and the NTC is negative so I do not thing contamination is the cause. I can not think of any other cause for the false positives. Can anyone offer any assistance? Thank You.
I have also included an image of the results for better visual.
I have been using NEB Hifi Gibson assembly for a couple years now and I've been quite happy with it. I regularly make plasmid constructs with 4-8 fragments, and always >1/4 of the colonies are "perfect," while the remaining ones may have some SNPs at the joining sites, or be misassembled due to a repetitive region.
Some colleagues said that Golden Gate has even higher efficiency. That even with 8 fragments, it is normal for 50-100% of colonies to be perfect. Is that true? Is Golden Gate that perfect?
We are trying to set up a one-step PCR for COVID19. We are trying to implement the Midnight 1200bp, by using a one-step RT-PCR protocol instead of the recommended two-step alternative. We are using Takara's PrimeScript. We are able to get a product with ARTIC primers (standard protocol) ~400 bp PCR product, regardless of sample quality, however in Midnight Protocol when we use better samples, the results seem to be more reliable however there is still a large amount of variability compared to one another. Would anyone have an idea on how I can get rid of this problem? Should I try and do the PCR two-step?
Thanks for any help
We first ran our samples through PCR and diluted them, then added ExoSAP. After running a gel, we got bands for the undiluted and diluted samples, but only half of the samples showed a band after using ExoSAP. Has anyone else run into a similar problem or could perhaps offer a solution?
Hello- I am a graduate student studying avian malaria from birds in Louisiana, USA. I am currently trying to amplify DNA from Haemosporidian parasites with PCR, following the protocols in the attached paper (Hellgren et al. 2004).
Specifically, I have not been able to amplify DNA during the first PCR, which uses primer pair HaemNFI - HaemNR and should target Leucocytozoon, Haemoproteus, and Plasmodium species.
I have tried various optimization trials and have not been successful. Although I have tried increasing the annealing temperature in the PCR thermocycler, I have not seen changes or any bands appearing in the gel. Thus far we have used AmpliTaq Gold 360 Master Mix and Hot FirePol Blend Master Mix.
Possibly it could be many things that are going wrong, so I wanted to ask for any advice.
Is it possible that the ratio of PCR ingredients is incorrect?
Any advice or helpful tips are much appreciated. Thank you!
I'm currently trying to sequence SARS-CoV-2 genomes at the medical lab I am working at. I am using the improved ARTIC protocol (Eco PCR tiling) and an ONT MinION.
After some initial difficulties, I was able to perform 2 sequencing runs which yielded a coverage of >95% for almost 50% of the samples.
However, the past two runs were much worse and I don't really know what the problem is. I can think of two things that might be the problem: a) the extraction method I am using (maybe there are too many contaminants present?) and b) I so far haven't diluted samples samples with lower ct values.
Does anyone have any suggestions on where to troubleshoot?
Thanks in advance!
I have performed a qPCR test to check my primer efficiency. I got the rough data now which I have opened with Design and Analysis software 2.5.1. It is quite confusing and I am relatively new in operating this software post qPCR run. I would be grateful if you someone can provide me instructions on this software for beginners, especially related to the primer efficiency test.
Thank you in advance.
I've been trying to run PCR on blood infected with Plasmodium which has low levels of the parasite. The PCR target is a gene of Plasmodium.
Has anyone had experience with this low parasitaemia causing non-amplification? I have tried different DNA extraction kits as well as a range of thermalcycling conditions, gradient PCRs, etc.
I have tested the primers in silico and they work, they are also from published literature.
Any advice would be appreciated
I am having some problems with my PCR. I have performed a RNA extraction with RNeasy Mini Kit, and after that, DNase treatment (RQ1 RNase free Dnase, Promega) on 2ug of the RNA obtained from the extraction. Then I perfromed actin amplification to evaluate the yield of my experiments. I've loaded the products on a 2% agarose gel, and then I saw this:
in the second lane there's the pcr product obtained without DNase treatment,
the third lane is the pcr product after DNase treatment,
my question is: What is that smear? Can it be DNA?
Hey everyone. I collected some leaf tissue samples from the plant Phragmites australis from which I am hoping to extra DNA for sequencing. I will be extracting DNA using Qiagen DNeasy Plant Mini Kits. I was in a rush when storing them, however, and I just placed them in ziploc bags in the freezer at -20oC. They had been kept in the same bags in a cooler while transporting them from the field to the lab.
Is this going to be okay? They've been in there for a few weeks at this point and it may be another few weeks or even a month or two before we will be able to begin lab work. Would it be possible to move them to -70oC now or is it too late? Can they be thawed and dried at room temp in silica gel? Just wondering what my options are here and what I need to take into consideration. At this point, it is too late in the season to collect new tissue samples so this is all that I have to work with.
Thanks in advance!
I am designing a PCR to detect the presence of Proteus vulgaris. Unfortunately, all the literature I have come across use either the 16S rRNA gene or the 16S-23S ITS. Does anyone know of a virulent or structural gene specific only to Proteus vulgaris?
I am unable to run PCR reactions in the Eppendorf Mastercycler nexus gsx1 model. Every time I try to run a reaction, the screen says "no cycler available". I have looked online for solutions and read manuals but to no avail.
I tried amplifying the CGG at 5'UTR of the FMRP1 gene by TP-PCR but it was not successful. I ended up with no amplification but when I used the primers flanking this repeat (short PCR) it worked with accuprime GC rich taq pol. So any suggestions from experts who have worked on this gene?
I am doing an allele-specific PCR to detect a point mutation in the DNA of Aedes aegypti, when I used a tm temperature of 55 °C, I observed some PCR products in negative control, and some nonspecific bands in the samples and positive controls. When the tm temperature was increased to 65 °C, the negative control was clean, positive control without unspecified bands, but, no amplicons were observed in any sample, what should I do?
I cut an amplicon band from agarose gel and purified it with a silica column to get rid of a non-specific PCR product. But after I ran my purified DNA on the gel again, the non-specific band didn't disappear!
I've performed the whole procedure several times without any luck.
0.7-1% agarose gel
And, by the way, it's happening for two of my different PCR samples, which makes it even more confusing for me.
Hello, I am doing PCR on fungal DNA samples and use EF1 and EF2 primers and the New England Bio Labs Phusion Taq Polymerase. I have had successful PCR reactions but over the past month I have been getting a uniform smear in each well including my negative control. I still have nice crisp bands, but the smear is a problem because I am sequencing these products. I use filter tips and my prep areas for pre-PCR and post-PCR are separate. If anyone has experience with this or any advice, it would be much appreciated. I have attached an image of a recent gel, the first well beside the ladder (farthest to the left) is my negative control (PCR mixture with no DNA, ultra pure water replaces the DNA). I have changed out my reagents/water and the smearing has not stopped.
I finally got my primers and was able to perform PCR to obtain my linear DNA. In order to check if they are correct I am using the CFPS and my PCR product.
The protein is FtsZ-YFP, so I am checking the YFP fluorescence. But my graphs are looking "strange", blank (containing only CFPS components) is decreasing and is similar to my samples (the graph attached is an example of how it looks like).
Of course the first thought is that primers are not correct.
Otherwise I was wondering if it makes sense to control the solution under the fluorescence microscope (confocal) to check and maybe adjust settings in Tecan.
I need to identify some microorganisms we isolate from nature. The company we have an agreement says that I need to design a primer as
- fw primer: 5′ ACACTCTTTCCCTACACGACGCTCTTCCGATCT – [your gene-specific primer]
- re primer: 5′ GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT – [your gene-specific primer]
and do the first PCR with these primers before sending the DNA samples.
I will use this for yeast-fungi identification but I am confused. Should I add ITS1 / ITS4 at the end of their primers? Also, are there any primers that are better for fungi than ITS1 / ITS4?
I need to design primer for two paralog genes, one of the gene doesn't have any unique region, Can anyone suggest how design the primer for real time PCR for this gene.
I am doing ATAC-Seq with OMNI-ATAC-Seq protocol (Kaestner Lab 2019). I did PCR amplification for 5 cycles of my library. Now I use 5ul of the PCR product to do a qPCR to determine how many additional cycles I need.
I have 4 questions about the detail of how to do this qPCR.
1. What method should I choose on the qPCR program, absolute quantification, relative quantification or delta delta CT or any other method? Should I add melt curve steps after the PCR program?
2. How much ROX did you add in your reaction system? None of the ATAC-Seq protocol (either omni-atac seq or Buenrostro's protocol) mentioned how much Rox should be added in the reaction system. We have a AppliedBiosystem QuantStudio 3 real time PCR machine which requires low rox. I usually purchase commercial SYBR GReen mastermix which includes rox, sybr green and everything in it. But in every ATAC-Seq protocol, it just said to add 0.09ul Sybr green I, it didn't mention how much Rox, or how to set up the setting of the reaction.
3. What is the amplification curve of your qPCR for ATAC-SEQ? I added low rox dye and everything else following OMNI-atac-seq protocol, but my amplification curve looks very weird. It's not a smooth curve, it's disconnected.
4. Some people mentioned to use KAPA Library Quantification kit to do this qPCR and determine number of additional cycles needed. We have the kit. Could anyone advise how to do this in detail?
I am planning on ordering a number of DNA sequences that range in size from about 50-200bp, to be cloned into a vector. I think the best/ most economical way is to order the sequences as ssDNA, that are flanked by a pair of 'universal' primer binding sequences, so that I can amplify all of them with a single primer pair. I could use a random pair of primers that have used previously, but I was wondering if there is a more commonly used pair of primers for this purpose?
The alternative is ordering it as two ssDNA sequences and annealing them, however this would be considerably more expensive. It would also give me a fairly finite amount of DNA.
Below is a workflow schematic for what I am referring to.
1. Synthesise ssDNA commercially:
primersite1 - RE site - DNA sequence - RE site - primersite2
2. PCR with primer1 and primer2 (Phusion)
3. Purify dsDNA product
primersite1 - RE - DNA sequence - RE - primersite2
primersite1 - RE - DNA sequence - RE - primersite2
4. Digest with REs to remove primer binding sites
- RE - DNA sequence - RE -
- RE - DNA sequence - RE -
I would like to use an exogenous internal positive control in my qPCR reactions (probe-based with gDNA). The IPC protocol states a 60C anneal/extend step, but my target assays require this step to be 63.3C.
I tried running the IPC at 63.3C (singleplex, not with my assays) to see how it performs at this higher temp. I had previously tried running one of my target assays (FAM) with the IPC (VIC) at 60C using the manufacturer's protocol.
The average RFU at 63.3C (IPC alone) was 765, and the average RFU at 60C (IPC + FAM assay) was 6700 and 9400 (tried it twice on separate days).
The call type was as expected (positive call for all reactions except for those with the IPC blocker), but the RFU values are very different.
Does it matter that the RFU values are so different? Is this difference because the reactions were run in singleplex vs duplex (lower fluorescent signal?) or because the temperature is higher? I just want to confirm the IPC's performance and determine whether or not I can reliably include this control in my reactions.
Product used: TaqMan™ Exogenous Internal Positive Control Reagents (thermofisher.com)
Purpose: to test for inhibition (sample is avian feces).
I did a pcr and got a non-specific band after gel electrophoresis. So, I cut the correct band from the gel and did a column purification. Then, I digest the amplicon with two restriction enzymes to use it for ligation.
After I ran my products on the gel, the digestion product gave me a clean, specific band. But, the product from my gel purification still showed the non-specific band!
Could anyone explain why this happened?
Can I still use my digestion products for ligation and cloning?
I've been reading about the length of the primers and I found that is around 18-30 bases and the shorter the primers are, the more efficiently they will bind or anneal to the target. However, I added like 8 bp to mine and they became more stable, it increased the concentration and the Tm changed. Please help me to understand what happend because I haven't found information about that, and if you have a paper or link to support your answer, that would be great.
I'm trying to isolate DNA from fibroblast cultured cells without using a kit (with lysis buffer and prot K), but after PCR amplification, no products is shown on the agarose gel. However, the same DNA isolation method works in HEK293T cells.
Does anyone have experience with DNA isolation of fibroblast with lysis buffer or tips?
I am trying to knock out a gene involved in virulence from a genomic region of roughly 150kb. So far I have been using CRISPR to target individual candidate genes, but I have recently exhausted my list of candidates without identifying the actual gene, so I have decided to take a different approach.
I plan on using split marker PCR to eliminate chunks of my genome in a stepwise fashion, with the hopes that one of these knockouts will induce a change in virulence, allowing me to focus in on the specific region that was knocked out to find my gene of interest. I understand the process of split-marker PCR, though most of the literature I've read involves using it to knock out specific genes, whereas I will be using it to systematically knock out genomic regions underlying my QTL of interest.
To minimize the number of transformations needed, I'd like to knock out as much of the region as possible in each transformation. The selectable marker I'm using is roughly 3kb, and I've designed my first primers to knockout roughly 5kb, as I've heard of previous successful knockouts of this size. I've also heard that the transformation efficiency decreases proportional to the increase in size disparity between the genomic region knocked out and the selectable marker that replaces it.
I'm currently planning on increasing the size of my knock outs by 2 or 3kb until I fail to get a positive transformant, but I was wondering if anyone has insights or experience with this sort of thing that they could provide me. Or perhaps some molecular theory as to the upper limit of genomic knockouts. I really don't want to do ~30 individual successful transformations to eliminate my full 150kb region. Thanks!
I m a first year student so this question might seem a little dense. but currently I ran PCR analysis of some gene. I got the potential interesting genes from the microarray data from the geo data. first time i run the PCR, the Ct value was high. when i ask my supervisor, he ask me to check the raw expression level that if the raw expression value is low, it could be a possible problem. but there is only comparative expression level(log2FC). could it be that i couldn't find or is there a case that the researcher didn't upload the raw expression level on the geo data?
We forgot a washing step during DNA extraction with 70% ethanol but washed it with 100% ethanol which was the next step, however PCR gave good bands at the expected size range. Will this salt remain during sequencing after purification of the PCR product? what problems can it cause?
This is a general bacterial PCR using 8f and 1541R primers. Only the positive control showed but what is the double band and at what bp?
I am trying to only detect (meaning no relative expression yet) a certain marker via taqman probe in DNA from old FFPE samples. I have tried to "increase my signal" by performing a nested PCR by running the profile without the probe and then purifying the PCR product (column based-Qiagen). I then went on to the secondary PCR using the Taqman probe. My detection was clean but low (high Ct values). The nested workflow resulted in pretty much the same Ct values as a single run (non-nested). I fear I am losing anything I am gaining during the purification process.
Is it absolutely required to purify between primary and secondary PCR's when doing a nested process?
I extracted DNA from some kombucha products with a manual protocol, with one final step using phenol, for 16S and ITS sequencing.
However, there is a problem for the majority of my samples in PCR step. Apparently, there is/are some inhibitors in my samples that impede PCR. My samples pH is around 3 and they contain polyphenols.
Did anyone have any similar experience or any suggestion?
I have a problem with the PCR results from my mesenchymel stem cells lysates. I use beta-actin as a reference gene and for some time now the Ct value is at the level of 30-35 cycle, a few months ago I did not have this problem, Ct was at the level of 15-16.
These results are passage 2, passage 6 and passage 12, everywhere a bad result which therefore does not depend on the passage.
All DNA concentrations were measured by nanodrop and reduced to one concentration, reverse transcription was performed under the same conditions. All probes were analysed on the same PCR plate at the same time. It is not the fault of the starters, because they were frozen in -20, used in the same concentration, on one plate. In new samples they showed the level of 30-35 (bad result), and in the old ones they are fine 15-16 (good result).
So what is the cause of such discrepancy in actin values?
What could this change result from? Is cell culture contamination possible resulting like this ?
I started a PCR process and recall I didn't include the Primers of the gene of interest in the cocktail preparation process. The pcr was at the initial denaturing step (@1:00), Will my reaction be successful if I stop, and include the primers in the pcr mixture? To restart he pcr process?
I am at my wits end with this PCR so I am hoping someone in the world might have some ideas. I am consistently getting inconsistent results.
I am following the PCR protocols adapted from " Joung, J., Konermann, S., Gootenberg, J. et al. Genome-scale CRISPR-Cas9 knockout and transcriptional activation screening. Nat Protoc 12, 828–863 (2017). https://doi.org/10.1038/nprot.2017.016"
I need to use these 2 PCRs to amplify and subsequently analyze DNA from a CRISPR screen. I am using the above PCR protocol and primers that have already been optimized for this purpose and have worked successfully previously for our lab. Importantly, I am using herculase which is a high fidelity (expensive) polymerase, which I also heard can be unstable.
When I first tried to amplify this set of DNA as smaller scale PCR test for for PCR 1, I saw no bands except for the genomic DNA extracted, so no amplification.
I tried this again, this time comparing the amplification of a DNA sample that had worked pretty well before (enough DNA extracted from gel to mantain coverage)--> still no bands except genomic DNA.
(I had tried this before but had not maintained enough DNA after PCR2 for enough coverage of the screen and had to repeat again with some new DNA I extracted.)
I tried once again but using a different herculase (we have 4 separate vials) for each reaction to see if any had lost their activity. The DNA sample that used to work and a DNA sample from a colleague that he claimed amplified.
Now, I am either receiving very bright bands or no bands at all (at the expected ~300bp size) but there doesn't seem to be a pattern as to why. The herculase that didn't work before for my one DNA sample, now works but not for my colleague's sample. Some of the herculases work some of the time but not others.
I repeated again, this time I had someone else in the lab (AG) try it as well alongside me (EP), using two different DNA samples that worked well enough before (enough DNA extracted from gel to maintain coverage).
We both observed the same inconsistent results. This time instead of observing all or nothing I am seeing some wells with faint bands, some with bright bands and some with no bands.
I am mixing the components, mastermix and PCR tubes well before amplification and before loading onto an agarose gel. I see the same results when loading some of the sample into multiple gels so no problem with that.
I don't understand why this can be occurring.
If some of a reagent went bad it should be well mixed so that all samples get about the same amount of it, not leading to inconsistent results.
I have also previously checked (with 192 PCR reactions for a PCR reaction that always works with hotstart taq pol) that the PCR machine works well and there are no problems with pipeting.
If I cannot fix the problem and ensure I am always observing good, consistent amplification, I will be at high risk of not getting high enough coverage for my screen.
Let me know if you have any ideas or you would need more details.
Anyone have experience with Herculase and notice this before?
I'm going to do microsatellite genotyping.
Therefore, after PCR I will be doing gelelectrophoresis.
I will take pictures of this gel, but I will eventually need output in a .fsa file.
Is there free software available to make this conversion?
For example uploading pictures -> peak conversion -> peak scanner -> .fsa file?
I have been searching a lot, but I have trouble finding a solution.
Thanks a lot in advance,
I normally use a nested PCR on 16S-23SrRNA region, for diagnosis of a plant pathogenic bacterium. It is a well-established and almost “trivial” protocol, shared in many laboratories.
Recently when the nested PCR products are loaded on 1% agarose gel, smirs appeared instead of usual bands, even in blanks. This happens with samples previously checked by qPCR, for pathogen presence and concentration.
On the contrary, the products of the first PCR are better defined.
We tried to solve the problem by changing the different variables of the system:
primers working aliquotes, primers pair (different pairs can be used for the same genomic region and with the same amplification protocol), Taq Polymerase, dNPTS; thermocycler; TBE buffer, electrophoresis cell and power supply, agarose, DNA staining (new Gel red and Red Safe aliquotes); we also tried by lowing the template concentration, but nothing worked.
Other types of nested PCR on the same DNA samples, but on different genes, didn’t have any problems.
Could someone please suggest me an explanation for those bad results?
I have a problem with the PCR results from my mesenchymel stem cells lysates. I use beta-actin as a reference gene and for some time now the delta Ct value is at the level of 30-35 cycle, a few months ago I did not have this problem, delta Ct was at the level of 15-16.
I use the same methods, the same isolation procedure and reverse transcription assay of RNA, the same primer concentrations and so on.
What could this change result from? Is cell culture contamination possible resulting like this ?
I have a mouse line with an inserted experimental gene.
I know the sequence of the inserted gene, but I do not know where it is inserted in the mouse genome (very new mouse line with little documentation). I want to know where it is inserted, which I assume would require two primers: one forward primer in the inserted gene (known sequence) and one reverse primer outside the inserted gene in the mouse genome (unknown sequence). How do I create the reverse primer without knowing the sequence?
I was thinking of just using a one-primer PCR reaction (forward primer), of course I would get much less product. Could I still sequence what little product I have and discover what's on the other side of my inserted gene? Then design the reverse primer? I just don't know if it is even possible to amplify then sequence using only one primer.
Any advice will help so thank you in advance!
I am performing leaf endophyte 16S amplicon from Arabidopsis plants grown in natural soil in growth chamber. I have collected the leaves (5 weeks), surface sterilised and lyophilised the leaves before DNA extraction. After DNA extraction, I've performed 799F-1391R amplification as per protocol by Chen et al. (Chen, T., Nomura, K., Wang, X. et al. A plant genetic network for preventing dysbiosis in the phyllosphere. Nature 580, 653–657 (2020).)
However, I see no bands for microbiome after PCR in agarose gel, although a band for mitochondrial DNA is there. Can anyone guide me what could be the reason?
Details of PCR: Template used 10 ng. Run for 35 cycles. Negative control is fine and positive control (Synthetic community) giving desired band. Is it due to very low biomass of microbiome in samples, How to troubleshoot?
I am designing some primers with Primer-BLAST and this problem keeps happening to me:
I specify the characteristics of the primers, the amplicon, the number of pairs, the organisms in which I want it to test for specificity...
But I am not able to prevent that those pairs with more than one unintended templates from appearing.
I say "more than one" because the first one is always the correct one (I don't know why it appears there but doesn't bother me). Then I have to filter "manually" the pairs that have only this one, (as the one in the attached image), and then choose between them. The real problem is that these pairs could be, maybe one or two in fifty (if there is any).
So, how can I prevent pairs with those "Products on potentially unintended templates" from appearing? Is there any option I am missing?
Hi, I am investigating ZIP proteins in yeasts and I used degenerate primers. I want to isolate my PCR amplicons, but I was told I need insert them into a PGEM-T vector and then sequence it using the vector primers for the blue-white screening. I would like to know the reason why they can't be isolated directly.
I've been using ExoSAP-IT to purify my PCR products before sending them to sequencing (Sanger). The problem is that I need to measure it before sending, so I can send the correct amount and when I try to use the nanodrop for doing it, the 260/280 absorbance gets extremely high (from 9 to 20).
What can I do for decreasing it?
Also, the quantity of DNA is getting very high, something like 3000. If I dilute it, do I have a chance of having better results of quality?
Before this step, I was purifying my PCR products from agarose gel with the GeneJET Gel Extraction Kit (Thermo Fisher Scientific) but the quantity was never enough for the Sanger Sequencing, although the quality was better.
I have a question on RT qPCR:
until now I used Livak method to analyze my qPCR data. I normally use 8 different reference genes and determine the three most stable ones. Then I use the geometric mean of Cq of these three reference genes for 2^-ddCq method.
Now I want to include PCR efficiency values via Pfaffl method. I am able to do the calculations referring to each single reference gene. But I don´t know how to combine more reference genes in the analysis.
The formula contains the efficiencies for target and reference genes. What is the efficiency for the control gene if I use 3 different ones?
Comparing PLSR, PCR, SVMR and MLR, which one is better for the correlation between linear data (MIR and NIR spectroscopies) and chemical data (pH, peroxide value ...)? There is(or are) other tools better than the four ones cited former ?
I have encountered a potential problem with filter sterilization of supernatants from Enterococcus strains, which according to PCR data, are hosts of several multiple bacteriocin genes, but show very little activity against Listera. Does anyone have practical experience with absorption of bacteriocins, and especially enterocins, on filter materials like cellulose acetate, PVDF, PES etc?
I want to use 3 ug of RNA, although the recommended amount is up to 1 ug. If it is possible, should change the amount of the other componenets respectively?
I need the cDNA for qPCR.
Thank you very much,
I have a few lines of a particular gene of which the earlier (T2 ratio) are not known. I want to do antibiotic selection of these seeds and I want to know which plants of next generation (T3) are going to be homozygous. Is there an easy way to confirm number of T-DNA insertion of my gene present in genome of individual plants? Like some PCR strategy.
I'm wondering about primer dimer formation in qPCR despite I've run them on PCR then on gel and they didn’t form a primer dimer on that so where’s the problem here ? otherwise, i ran 40 cycles on qPCR and 35 on PCR so could this have an affect?
Most of the literature used partial cytochrome b gene sequences for animal phylogenetic studies instead of the complete sequence. What is the reason behind this where full or complete cytochrome b gene sequence could explain more details?
Thank you very much!
Maybe someone knows why in the first cycles the amount of dsDNA increases and at some point it begins to decrease.
I have used PCR-RFLP for the genotyping of fowl adenoviruses previously but now I am interested in the genotyping of FAdVs by real time PCR. Kindly share your experience if someone has already used this way. Thank you
I used a kit to clean up a PCR and purify my DNA. The DNA Needs to be at a higher concentration for me to use in electroporation experiments. How do I concentrate my DNA? Should I be worried about concentrating the salts in the buffer? I would assume it is not possible to run again in the kit. In the future, should I elute in water instead of the buffer provided with the kit?
I used EcoR1 to digest pGEM®-T Easy Vector and there are bands that correspond to the released genes that were inserted but no bands at all for the cut plasmid. The lanes which have arrows only have the genes released and no bands at all for the cut plasmid. 20ng of DNA was used to calculate the prepare prepare the master mix for restriction. Would restraining the gel show the bands or increasing the amount of DNA? what might have caused this to happen?
I extracted DNA from Cocoa leaves and amplified some fragments (SSRs markers), the thing is I get those bands! (attached image), and don't have a defined band. I'm using a new TBE 1X with the gel, running with TBE 1X buffer, 3% Agarose gel, and running it 60V/150 min. But I can't get defined bands!
Maybe it's because of the thermal PCR program? should I use a TouchDown PCR or something? any idea to get defined bands would be Helpful! Thank you!