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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Does anyone have an updated list of the spa types (with repeats) in Staphylococcus pseudintermedius? There is the Ridom spa server for aureus, but the ones for pseud do not appear to be online.
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Dear Laura,
The website http://www.pse-spa.org is out. Could you help us?
We have some data of spa sequences to analyze.
Thanks.
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Hi,
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!
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The highest concentration of actual RNA I've ever gotten (confirmed by dilution series and Qubit) was around 2000ng/uL. This sample was viscous and sticky, it looked and felt very different from 'normal' RNA samples. If your samples still look and feel like water, the high reading is probably contamination. If they are very viscous, try diluting 1:10 and repeating the measurement. If you did just extract a REALLY large amount of RNA, diluting it will dilute any impurities too, and it might be fine for reverse transcription.
If you can, I highly recommend you try to get access to a BioAnalyzer or a TapeStation, which can tell you enormously more about the quality of your RNA sample. For RT-PCR, it matters that your RNA samples are at least reasonably intact, and that the amount of degradation is similar between samples that will be compared. Nanodrop can only tell you approximately how much RNA is in the sample, and approximately what impurities may be present. BioAnalyzer can tell you whether your RNA molecules are intact or degraded.
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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.
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Dear Papri,
Yes, you are right about the efficiency of the kit, even I found it in several protein engineering publishings. As I am naive to this area of work, I need to standardize it first, however, I can say I am in the process :). As you mention about the digestion of the parental plasmids, I think in this case it happens after paternal plasmids enter the host cell by MrcBC endonuclease. Nevertheless, I shall request you to go through the manual once. I will make good use of your insights.
Cheers!!
Diptesh
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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.
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I have stored the reaction mix at -20. Whenever I take it out, I keep it in a cooler prechilled at -20. There is least chance for the mixture to be at a temperature >-10 for few seconds.
Thanks for suggesting the alternative kit.
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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.
My Program
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.
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Have you checked the Q5 protocol from NEB, the Denaturation is 98oc, not 95oc.
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Hello All,
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.
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You have no product because you have a huge primer dimer problem. The 2 primers are complementary at some position and are binding to each other giving a very small product which only contains the 2 primers. This melts easily and has perfect homology to both primers so amplifies very well and removes all of the primer from the reaction mix so large primer dimer intensity and zero specific product. This happens either when the primers are poorly designed or when you use far too much of each primer so that there is a lot of primer to start the dimerisation process. Try the reaction with 1/4, 1/8th, 1/16th and 1/32 the amount of each primer to try to get a product. Also run more cycles so that you get visible amounts of product while stting up the assay
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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.
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Nucleated cells. Mature mammalian RBC are not nucleated.
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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?
Warm regards,
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The enzyme will be fine and still active and all of the reagents will be stable at the annealing temperature.I would expect a slightly higher level of non specific amplifications but if the pcr always amplifies clean then if you reset to the proper parameters I would expect the pcr to work
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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
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If you know the sequence of your template then align the primers against that sequence. Then add the next 5 bases to the 3' ends of each primer and check the annealing temperatures are well above 60c and also check that there is only a low probability of the primers forming primer dimers
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Hi all!
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!
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Hi Thomas Riedinger,
What platform/software did you use to design your primers or did you do it manually? Firstly, I would suggest that you align all your target COI sequences through a multiple alignment tool (I used MEGA X) instead of relying on pre-designed sequences (I assume designed individually per species). This will help you narrow down conserved areas that are similar in all your target species, making your universal primer more effective.
Next, I would suggest you design your primers using a software design tool (I personally love PrimerExplorer). Using a software makes primer design so much easier as it helps to suggest primers that will potentially work, mitigating those outside the accepted parameters so you don't have to. I would say it is easier and smarter to let the software do the brunt of the hard work when it comes to the design, and you then validating the primer. (Saves you the headache as well).
Did you also check your universal primer through BLAST? Did it amplify your target species? Is it possible that other non-target species were amplified in your universal primer? You would want your universal LAMP to only identify your intended target species, so please ensure how specific your primer is.
It is also possible that during the experimentation, cross contamination occurred causing the smearing and low bands (non intended ones). LAMP is highly sensitive to cross contamination.You might have carried over DNA from your previous assay onto this one. Ensure that your DNA extraction workspace differs from where you carry out the LAMP assay and make sure your workspace is always sterilized, and you should make sure to open the reaction tubes at a different place compared to the incubation area to prevent aerosol contamination.
I assume that the optimization done on the various reactions times, primer concentration & temperatures didn't change the outcome. Did you try adjusting the other chemical concentrations as well?
To summarize: 1) Realign all your COI sequences and take the conserved region and run it through a primer design software. Validate your primers through BLAST and also ensure your workspace is sanitized to prevent cross contamination.
Hope this helps Thomas!
Do keep me posted.
Lavan x.
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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!
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I highly recommend the primer cocktail (cocktail 2) proposed by Clare et al. 2007, which is a modification of the cocktail proposed by Ivanova et al. 2006 to amplify the DNA barcoding region of the mitochondrial COI gene. This primer cocktail works pretty well for taxonomically distant species, which I believe is what you are looking for.
My colleagues and I used this cocktail 2 very successfully in Pinto et al 2018. We did not have to troubleshoot the PCR conditions to obtain bright amplifications that yielded high quality sequences.
References:
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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.
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Hi Samitha,
Some thoughts on your sequences.
DA1TRNHPSBAfp
Starts with strong (>1000) well spaced base peaks dropping slowly from base 105 to about 330 bases with a signal/noise of 200. Quite good sequencing but this kit is capable of 900 bases of strong intensity signal. This is usually cause by insufficient primer or template. If this sample has primer dimer ( PD) then this will measure as DSdna so less template will go into the sequencing mix and the signal will die early and the remaining template will give a weak but readable signal
DA2TRNHPSBAFP
The signal strength is only 28 and s/n ratio is 4 so no readable sequence at all.The reaction has failed completely so possibly primer and template do not match
DI5MATKKIMM13FP
Clean strong base peaks up to base 53 ( strength 1600)then signal drops to zero.
This can happen with some secondary structure problems in the template dna but can also happen if you run out of primer. In this case I suggest that the sequencing primer has been used up sequencing PD
DI7MATKKIMM13FP
Good clean strong ( 3700) sequence up to 46 bases then dropping to baseline intensity. There is no huge unincorporated dye peak so the sequencing reaction is working but clearly no on the template dna so I think that we are sequencing a strong PD
De1TRNHPSBAFP
Very large dye peak ( mixed bases) at start of sequence (>4500) then no peaks. The sequencing reaction failed completely. Either no primer, no template or the wrong primer used in sequencing.
D13MATKKIMM13FP2
160 strong bases all Ns. We have 2 primers annealing in different places and producing 2 sequences. As it is unlikely that you supplied 2 templates so something is acting as a second primer or one primer is annealing in 2 places on the template dna ( unlikely), The sequence stops much too early. I do not know the cause for this one but if you had not told me that there was no PD I would say the pd has dissociated and caused F and R primers to anneal to the template and has used up the sequencing mix too early.
RecA C92f
Huge unincorporated dye peaks (17000 and 4500) at ends of sequence with low intensity clean sequence in between. I think that you have light PD which has sequences and used much of the sequencing primer and the rest has produced a weak but accurate sequence that has not been basecalled. The reason for not calling the bases is that the unincorporated base peaks are so high that the base calls are below the set cut off value of 5% of the highest peaks so they are ignored.
RECAC92F
Huge (>12000) unincorporated dye peak at start dropping very rapidly after 35 bases. Dye peaks at 85 and 120 bases indicate that the sequencing has failed. I think the primer was all used on the PD sequence and the unincorporate dbases are from the unused sequencing mix
Overall I feel that your problems are caused by PD using up all the sequencing primer not leaving enough to get a good signal from the template dna. Additionally pd is double stranded so will measure as DS dna when calculating the amount of dna to sequence so you are actually sequencing too little dna.
You may need to clean up your pcr ( use less primer for a start) or gel separate the template and clean it up before sequencing.
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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.
Mary
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Surcharged wells with DNA. You could load less volume (3-5 ul) or even dilute your PCR products.
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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?
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Some restriction enzymes are very fussy about cutting in pcr buffers which often contain many different salts. Bfa1 is a poor cutter in pcr buffers generally. What taq are you using to make the pcr product and what volume of amplimer are you cutting and is it purified product or just amplimer in taq reaction mix? Are you adding additional Bfa digestion buffer if you are cutting unpurified pcr product?
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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..
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For pKD46 mediated gene deletion, we use the following protocol: -
1. Subculture the cells (Grown at 30 degrees) in 10 mL LB and add 1mM arabinose for induction. Grow up to 0.6 OD again at 30 degrees.
2. Wash the cells thrice with ice-chilled water in a falcon tube.
3. Transfer the cells to a microfuge tube and spin for 5 min at 5K.
4. Resuspend the pellet in 100 microliters chilled MQ and take 50 microliters for electroporation.
5. Add 300 - 500 nanograms of your PCR DNA and give the pulse.
6. Immediately add 1 mL of warm ( kept at 37 degrees) 2YT media (You can do it outside also) and keep it at 37 degrees for 4 hours.
7. Plate the cells in selective plates.
Hope it will work.
Good Luck
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Hi everyone,
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?
Thanks!
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As mentioned in the instructions provided by the supplier, add 1 volume of 6X loading buffer for 5 vol. of sample. In your case, 5 ul for 25 ul of sample. Just make sure that the well in the gel is large enough to accommodate 30 ul, if need be by taping together two or more teeth of the well-forming comb.
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Hi everyone,
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.
Best
Ruojin Tian
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I have seen these positive results before from colony PCRs when you use the colony directly from the transformation plate. If you are using primers inside your insert and a good DNA polymerase, you can amplify your insert from the fragments of DNA that are spread over the plate, not inside the cells. This DNA comes from the ligation you used to transform the cells and the colonies that you obtain contain a empty vector.
To solve the problem with the colony PCR, you should use primers located in the vector that produce amplification of your fragment. If the fragment is really long, you can use a primer inside the insert and other in the vector.
Concerning the toxic expression of your insert, do you know if the expression is due to your vector or is your insert which is been recognized by the transcription machinery?
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Hi!
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!
Thanks
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Hi
I think you can use the following for bacterial and viral business
In silico PCR amplification
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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?
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I agree with John Hardy Lockhardt, but if you wish to avoid spurious mutations, use a high fidelity DNA polymerase, like Phusion or Q5, rather than standard Taq.
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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!
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Hi Chandra Somasundaram , thank you for your reply. I did not have any antibiotics or antifungal agents in the media. The media I submitted for testing was the same batch I'd been using to culture the cells.
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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?
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Ahmed H. Shatta There are optimization procedures for finding the lowest volume which efficiently amplifies your DNA, which can be based on orthogonal experimentation. I would be happy to give you an elaborate explanation in the messages if required.
Explanation - Efficiency of PCR reaction can be better and can be worked with reduced volume also if your proportions of input DNA and primer/probe concentrations are proportionate even in low volumes so that the plateau phase of PCR doesn't occur before the average cycle count you are providing.
Suggestion based on explanation - So you can provide reduced primer/probe concentration and also reduce input DNA proportionally so that at least a limited amount of volume can be reduced.
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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.
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Hi Sebastián, after digesting with restriction enzymes 5' end will have a P. No worries, it will work.
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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.
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Arunabha Khara It is good that you have sequenceable amounts of pcr product with the extra cycles but for the future you may have to ask yourself why 37 cycles are necessary. I think that Katie A Burnette is correct and that your dna has pcr inhibitors reducing the efficiency of the pcr so that it needs extra cycles so if this continues to happen then amplifying less template dna will probably be the answer
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I am trying to design primers for some miRNAs but I keep getting errors pertaining to my minimum and maximum PCR product size. Any help?
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Compare with your DNA Ladder/ Marker
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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. 
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@Maria, have you solved the problem with false positives? I have been getting similar false positives only in Hex probe tagged gene in a cheaper enzyme only at Ct between 29 and 34 which is typical for all the false positive cases. In other standard enzymes it does not occur though the primer mix used is the same. This is a multiplex reaction and the other probes are FAM and Rox which are working good. The shapes of the curves for false positive are mostly sigmoidal and some are non-sigmoidal. Please help.
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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?
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Hello everyone,
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
Ege
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the major problem with 1 step pcr would be non-specific amplifications.
But if you want to perform one step pcr to know the accurate annealing temperature of your primers then you could go for gradient pcr
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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?
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Taylor Buckley ExoSAP-IT, is used to purify the PCR product, could not be the problem because you got band in the half of the samples. Try to do electrophoresis of the PCR product without ExoSAP-IT and if you dont get band you may have a problem in your PCR.
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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!
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What's your isolation procedure Trizol? Often times simply diluting your DNA extraction product will eliminate the off effects of contaminants when you PCR. Have you checked the 260/280 ratio on the spectrophotometer (Nanodrop) prior to amplification? Hope this helps. Best of luck.
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Hello everyone,
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!
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https://asm.org › Articles › October
SARS-CoV-2 Sequencing Data: The Devil Is in the Genomic Detail
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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.
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Dear Abhay,
Import your run file (.eds) format into the QS Design and Analysis software then click on analyse. Your results will appear. Then select the Standard curve from the drop-down list and click on the eye icon. Your plot will appear. Select the target and all the wells. The slope, R2 value, PCR efficiency and the error will be displayed below the plot.
Good Luck!!
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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
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aggree with Dr Paul.
and if you concern about the inhibit factor from the blood, you could try neb Hemo KlenTaq. it works even with 10% blood.
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Hi,
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?
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Perhaps the RNAse protocol has a high temperature incubation period that has degraded or fragmented the RNA a bit and generates shorter copies of the transcript, thus showing the smear. Some RNA structures are more resistant than others. Try another RNAse protocol that inactivation of the enzyme is not carried out by an increase in temperature.
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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!
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Yes, you can definitely store them. -20 -80 and dry should all work. Dry samples can be a bit tricky to isolate from but are very easy to store. You just want to avoid stuff thawing or getting wet.
On a storage note, plastic bags get very brittle at -80, it is easy to have bags break and spill, you may want to switch to tubes
If you are collecting these samples again I would suggest having duplicates (or more) of each plant and to store them in different locations (like different freezers, or one dried and one frozen). That way you will have a backup!
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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?
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Hi Jonas Boateng I would like to know which species-specific gene did you choose to target Proteus vulgaris.
Mohamed Mahrous Amer I would like to read the paper you mention: ''Zh Mikrobiol Epidemiol Immunobiol. 2005 May-Jun;(3):33-9.
[Species-specific detection of Proteus vulgaris and Proteus mirabilis by the polymerase chain reaction].'' Could you send to me the full article?
Thanks in advanced, best regards.
María
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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.
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It is probably because a previous reaction has not been aborted. Just go to the status tab ( should be the second last in the menu) abort the already running program and then try again. It should work.
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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?
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Try the La Taq DNA polymerase with GC buffer in combination with the deaza dGTP
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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?
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Are you sure that the product at 55c is not just primer dimer?
Can we see a picture of the gel image. I would run a temperature gradient from 55 to 65 using a dna sample known to amplify with any other primer set to find a temperature that works as expected
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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.
TBE buffer
0.7-1% agarose gel
70V
~1hr
And, by the way, it's happening for two of my different PCR samples, which makes it even more confusing for me.
Any suggestions?
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Xiping Zhang Any idea how to get rid of it?
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I want to set up PCR for 16s rDNA in bacteria. At least how many ng/ul should the nanodrop measurement be as a result of DNA isolation?
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Dear Berfin, nanodrop can safely measure the lowest possible concentration of 0.4 ng / ul. But for PCR I would highly recommend to use at least 10 ng / ul in total starting template provided that your samples OD 260/280 and 260/230 ratios both are within their normal ranges to confirm the purity and quality of your extraction. I usually perform conventional PCRs and qPCR both with at least
25 ng / ul of sample, and if ever I faced any problem it is mainly related to primers not the template (DNA / cDNA) itself.
Regards....
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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.
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Some prevalent errors might cause the PCR product to get bogged down below:
  • Misread gel: If the gel is not poured correctly, it will not polymerize or harden uniformly so that the molecules smear.
  • Well overloading: If the wells are too much packed, or the sample isn't diluted appropriately, the surplus sample might be smeared through the gel. Moreover, when the gel is shifted after placing the selection well, the model might break down. This might be spread as well.
  • Improperly Prepared Sampling. Divides into an enzyme that cuts the molecule in particular areas to prepare the protein or nucleic acid sample (restriction digestion/). If not correctly performed, the enzyme might uncontrollably break down the molecule too much and cause a smear. Moreover, the wrong buffer or temperature choice might lead to an inappropriate enzyme function, producing flushing, and not most minor.
  • Pollution: if a sample of DNA is polluted by a protein, which can also produce smear.
This is a relatively common problem for agarose gel electrophoresis, and there can be plenty of information elsewhere.
Good luck!
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Hello everyone,
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.
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Hi Kateryna,
The fluorescent signal seems to be very similar between the different kinetic curves, so there may be the possibility that the assay is not working as expected.
Of course you can also try to optimize the acquisition parameter on your plate reader. Could I ask what are the current parameter that you are currently using, specifically:
- What are the wavelengths in excitation and emission?
- What number of flashes are you using?
- What gain setting (if Manual, optimal, or calculated from well)
Best wishes,
Nicola
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Hello!
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?
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AS it appears these are Multiplexing Read Sequencing Primers, have a look at this(Illumina Adapter Sequences) and search your primers seq inside the paper, you will find something which may help you:
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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.
Thanks
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quick question, are you sure these are paralogs and not alleles of the same gene? Lots of plants have heterozygous allele sequences (depends on the species, cultivar, how much it was inbred etc.)
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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?
Thank you!
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Hello
I am not having ant CTs after performing the qPCR. Anyone can provide some help?
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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 -
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Alexander, check your vector. It usually has T7/Sp6 promoter regions flanking the MCS. After cloning your sequences into it, it's easy to amplify them with a pair of these primers.
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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).
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Thank you Dejan Vidanovic I am not doing official diagnostic testing. I am doing a validation study to identify and quantify wildlife diets using eDNA in their feces. Given the low abundance of DNA in the feces, I want to add confidence in my negatives -- a negative detection means the target food item is not in the animal's feces and therefore the animal likely did not consume it.
In testing the IPC with spikes, I am getting the 'right' pos/neg end point calls with my positive and negative controls, so I think it's working OK. I am just concerned about that decrease in end point RFU values, but maybe the absolute magnitude of fluorescence isn't important in my situation.
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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?
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I would highly suggest sequencing your final cloned product, that way you can be sure that you have cloned the correct fragment. If there is a useful internal restriction site in your fragment you could do a diagnostic digest prior to sequencing, that way you can pre-screen for clones that should be correct.
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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.
Thanks!
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Congrats on your success in designing a stable primer!
Actually primer designing is crucial and pivotal! There are actually some unique factors to declare the best primer for a particular sequence. Adjusted Tm value, GC content (40%-60%, 50% is more preferable), product length (100 -800), primer sequence length ( 20-25), and the hairpin loop can be considerable. But it depends on the way of your research.
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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?
Thanks!
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After SDS and prot k you need to further purify with trizol and chloroform followed by isopropanol ppt and wash of pellet in 70% ethanol or at the very least the ppt and wash step
Without that your lysate will be packed with salt which will inhibit downstream emzymes (reactions)
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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!
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Just to check- did you make sure you knew the copy number the genes you were attempting to knock out and were you able to confirm you knocked out all copies? And do you now whether the genes you're knocking out are essential?
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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?
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Thanks for sending me the GEO accession number. You cannot find CEL or CHP files, because you have Illumina chip data here. I have downloaded the file and have sent you the zipped file some minutes ago. Unzip it and you will find the expression values (Signals). I hope you will find your gene.
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I am trying to design primers for some miRNAs but I keep getting errors pertaining to my minimum and maximum PCR product size. Any help?
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Hi Emmanuel
Amplicon size for real good RTPCR need to not exceed 100-150Bp when respecting the delta delta CT protocol.
for miRNA since the targets are really smaller, amplicon size turns around 50Bp.
see this old paper, it can help:
all the best
fred
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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?
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Shouldn't be a problem since your PCR was successful. What are you using/doing for PCR purification?
There are kits that you can use to quickly clean up PCR products, such as the Column-Pure Gel and PCR Clean-Up Kit from abm. cat number: D516, or the QIAquick PCR Purification Kit Cat. No. / ID: 28104. Both works well for me, but abm is cheaper and does the job.
Alternatively, if you're sending your sample to a third party for sequencing, you can also ask them to perform the purification for a small additional cost.
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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?
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Something may have mixed with the sample umber 4. What ladder/marker have you used in your gel?
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Hello all-
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?
Thanks!
Andrew
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Andrew Kleinhenz, I also did nested PCR but I did not do purification! However, as the PCR products was in higher amount for the first run, I had to dilute it. I got good results! So, it is not always necessary to do purification.
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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?
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The Benzyl chloride method for DNA extraction aslo can solve your problem.
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Is it ok to use water for injection (WFI) instead of ddh2o for molecular biology work, such as pcr or dna purification?
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It is certainly OK, but probably a waste of money. Water for injection is guaranteed pyrogen (endotoxins)-free, which is not a worry for most molecular biology applications. Distilled or reverse osmosis water is fine; prior autoclaving takes care of inactivating nucleases.
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Is it necessary to gel extract either the vector or the PCR insert prior to the in-fusion HD cloning reaction? (as suggested by Takara)
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I am having students do this who have limited time so I want to make it as straightforward as possible yet successful. Spin column purification would remove enzymes after insert PCR and after vector linearization . My only concern is incomplete cutting but it would be worth it to screen more colonies if need be than have them try to gel extract the vector.......
Thank you for your input!
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Hello,
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 ?
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"All DNA concentrations were measured by nanodrop and reduced to one concentration" -do you mean RNA?
Because you really shouldn't spec cDNA: this is largely meaningless (unincorporated nucleotides and primers all contribute to the absorbance).
If all else is equal, my suspicion is that cDNA synthesis was just...really bad for those specific samples. Have you looked at any other genes? If the cDNA synthesis was low efficiency, then everything will be low.
If this is the case, then my suspicion would be "dirty RNA", generally due to poor 260/230 ratio. Did you determine this ratio for all your RNAs?
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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?
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Yes I expect it to work. The primers were not active and the enzyme activity will not be effected by the short heating step and all reagents are very heat stable. I would add the primers and restart the cycling
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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?
Thanks!
Eli
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Dear Eli Perr
I was just wondering about the fluctuation and inconsistent result you’ve faced. Hope you can find the way out.
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Hi,
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,
Kind regards,
Laurens
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Hi Laurens!
What do you want to do in the end? Perhaps your task can be solved through a gels analysis software?
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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?
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Have you tried touch down PCR with your nested primers
This is a common problem with standard round of PCR
Touch down starts from above the melting temp of your primers and during the course of cycles goes down in 1C intervals to Tm-2 where it then stays for the remaining (10-15 cycles)
This high stringency tends to result in better bands in your situation
I can send you my TD protocol if you wish
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Hello,
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 ?
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Hello
No, it is not culture contamination.
What is the passage number of the mesenchymal stem cells?
It is known that under extended cultivation there is a marked decrease in the content of proteins involved in cytoskeleton structure dynamics.
Increase in passage number leads to a senescent stage characterized by proteome alteration at the level of cytoskeleton organization, actin dynamics and decrease actin turnover.
This could be a possible reason. I suggest you use cells of low passage and re-check.
Best Wishes.
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Hello!
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!
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Is it possible to use inverse pcr? ie cut the inserted sample genome with a common cutter that does not exist within the insert. religate under self annealing (low concentration) conditions and then amplify the unknown sequence with primers internal to the insert but facing outwards from the insert ( therefore inwards to the circularised sequence thus generating the flanking sequence. Alternatively NGS
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Hello Researchers,
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?
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Sounds like the issue is you are looking for a low-abundance template. You'll need more DNA.
What DNA extraction protocol did you use? You might need to add in grinding in liquid nitrogen, freeze-thaw cycles, or a proteinase K digestion.
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Hello everyone,
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?
Thank you!
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Dear Dr. Oscar
When I just gave a random human gene accession ID and designed primers for 50 returns (50 pairs result), before the result appeared, I got a page which said that there are some matches with another gene in the database.
.. Finding primers specific to your PCR template (using Primer3 and BLAST)
Input PCR templateNM_000184.3 Homo sapiens hemoglobin subunit gamma 2 (HBG2), mRNA Range1 - 586
Your PCR template is highly similar to the following sequence(s) from the search database. To increase the chance of finding specific primers, please review the list below and select all sequences (within the given sequence ranges) that are intended or allowed targets...
If I tick mark the extra similar gene, I got two primer pair results, each for the two genes.
If I did not tick mark the extra suggested gene and continued, I got 50 pairs result. Except the first one which was my target gene, other appeared as:
Products on potentially unintended templates
> NM_000559.3 Homo sapiens hemoglobin subunit gamma 1 (HBG1), mRNA
I later found that, if you uncheck
... Primer Pair Specificity Checking Parameters
Specificity check
Enable search for primer pairs specific to the intended PCR template ......
in the first page where we enter all the required parameters for primer design, the results are specific to the gene of interest only.
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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.
Thank-you
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Dear Sara
The simple explanation for that is that if you use degenerate primers your amplified mixture might not be completely monoclonal and thus your sequenced ladder will be confusing by virtue of overlapping sequences
Cloning solves this problem as does secondary amplification with internal nested specific primers (but that might not work in your application)
In addition cloning out will culminate in better sequencing efficiency so with these two parameters in mind better signal to noise: in other words a clean unequivocal sequencing read rather than potentially a confusing ambiguous polyclonal read out
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Hello everyone!
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.
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Your nanodrop is probably measuring free d NTPs as well as the pcr product. Exo sap only phosphatases the NTPs it does not remove them.
You can precipitate then redissolve the dna or pass the samples through very small desalting spin columns prior to nano measurement.. If using column purification then heating the elution buffer to 70c and leaving it on the column for twice as long as the recommended time will increase the eluted yield.
You could also run a couple of known quantified pcr product as control samples on an agarose gel along with a known volume of exo sapped pcr sample then visually compare the intensity of the bands to assess how much sample to sequence. If your dept has a Qubit then it is capable of good DNA quantification but sequencing is quite robust and lower technology methods do work well especially for short sequencing templates
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Hi,
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?
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The use of the geometric mean for averaging multiple reference gene Ct-values is often not recommended (see Jochen Wilhelm's contributions in this thread: https://www.researchgate.net/post/How_to_analyze_a_qPCR_with_2_or_3_housekeeping_genes).
Thus, alternatively, you may use the normal arithmetric mean Ct-value of multiple reference genes and/or stick with the "Livak-method"...
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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 ?
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The type of modeling algorithm to be used is specific to the type of data you have and depends on the linearity or not, but for linear modelings you can apply the PLSR that can better fit this case, but also why not trying some machine learning tools (ANN, random forest...) as some experiences have proved that they led to better models. Another point is that care should be taken to the data preprocessing step.
Good luck
Issam
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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?
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Thank you all for your answers. I am aware about the pH dependent absorption of bacteriocin to producer cells according to the articel from Yang (2000). However I have been researching a little more and I see that Enterocins A and B are actually more active at pH 6.5 rather than at lower pH. Still no idea though how they behave against filtering material as well as plasticware and glassware.
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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,
Sapir
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No, you cannot. optimum primer concentration will be very less in the reaction. Therefore primers will be insufficient for the cDNA synthesis.
You need to add extra random hexamer or oligo dT (whichever method if you are using). And you need to increase the synthesis time to increase synthesis turnover number for each RT enzyme.
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Hi everyone!
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.
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I used a 3 primer method for identifying the insertion of my gene in Brachypodium T-DNA mutants , the protocol was given on the T-DNA mutant website.
They must have a protocol for Arabidopsis on the TAIR or SALK website, did you check it?
Briefly it mentions using your Gene specific primer with the forward or reverse T-DNA primer. In one reaction you add both T-DNA forward+ reverse+ your Gene specific primer. I have attached the protocol for your reference, it worked perfectly after the PCR on the gel you will see one band for homozygous insertion and two bands for heterozygous insertion.
Hope it answers the question, feel free to ask any further questions.
Best of luck Avinash Sharma
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When we do not have DMSO, can we be satisfied only with the thermal denaturation of the double strands of RNA (e.g. 5 min at 95 °)?
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Hello
At higher temperatures partial denaturation of dsRNA may occur, while at moderate temperatures denaturation of dsRNA is not guaranteed.
On the other hand, DMSO brings about effective denaturation of dsRNA into corresponding ssRNA preventing reannealing.
Use of DMSO becomes essential especially when your end application is sequencing. DMSO helps to improve reverse transcriptase PCR and Sanger sequencing.
I would suggest you use DMSO for good results.
Best Wishes.
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Hi there!
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?
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generally, qPCR is high precise than conventional PCR. Occurrence of primer dimer will be there if you would run qPCR for 40 or 37 or 35 cycle it doesn't matter.
if you might think something wrong about your sample. 1st thing, it may be sampling error or handling error. 2nd is after preparing premix don't keep it for long time as soon as possible feed it and load it.
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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!
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Hi!
You got the answer.
Simply to show/prove populations are phylogenetically distant from other species three regions of mtDNA viz. cytochrome b, 16S RNA and and D-loop regions will be used.
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Maybe someone knows why in the first cycles the amount of dsDNA increases and at some point it begins to decrease.
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Theoretically, pairs of primers are required to amplify double stranded DNA in every PCR cycle. In case of asymmetric PCR using single primer, in early cycle of amplification the primer can anneal to template and extend to form double stranded DNA. At early PCR cycle, we might obtained certain amount of dsDNA that can be detected by fluorescence dye but longer amplification will produce more ssDNA compared to dsDNA. Certain fluorescence dyes are not efficiently bind to ssDNA thus produce faint band during agarose gel electrophoresis. You might use fluorescence dye that specifically design to intercalate ssDNA (example SYBR Green II, Diamond Nucleic Acid Stain).
Hope can help. Thank you.
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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
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This is a good source of primer/probe for detection of FAdVs by rRT-PCR:
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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?
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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?
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I think you're having a few different issues here.
Firstly, you should probably run more DNA in each lane to better visualize the bands. For example, your uncut plasmids should produce 2 distinct bands (circular and supercoiled).
Secondly, the size of the extracted genes does not match the expected size. pGEM-T Easy is a 3kb plasmid and your undigested bands run at >20kb, implying a 17kb insert. However, the bands in the digested lanes are all <1.5kb, far smaller than expected.
I would strongly suggest that you verify the plasmids you are using by digest screening or sequencing.
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Hi researchers!
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!
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This kind of result is often caused by removing the comb from the gel before full setting of the gel has occurred but in your case the size standard (ladder) is running cleanly so the misshaped bands may be caused by either too much salt or too much glycerol in the loading buffer. Try running less dna ( about half) and use less loading dye and make sure the sample and dye are well mixed before loading the samples