- Alison KM Hadwin added an answer:1Any advice on why there is no band on gel for sample, but band for +ve control and poison PCR?
I have crushed up woodlice, serial diluted and spread plated and have growth of fungus on SDA+antibiotics.
I have also done a DNA extraction and run PCR with fungal primers, bacterial primers and have no band for the woodlouse DNA, I ran a poison tube with woodlouse DNA and the positive control and I got a band, meaning there is no inhibitor in the woodlouse DNA.
For the DNA extraction I used a kit that said to elute into DES water, I found that the samples degraded in the fridge so eluted into TE buffer, would this cause a problem, some has mentioned TE buffer being a chelator and may bind to mg and prevent the extraction from working?
I have no idea what to do now, anyone know why it may not be binding to the primers?
TE buffer contains EDTA, so yes, it can bind to the Mg in your PCR reaction and inhibit the reaction. Did you add the same volume of extracted DNA to your "poison" reaction as to your sample reaction? If so, it doesn't sound like there is enough EDTA to inhibit the reaction.
If you still think the TE is a problem, you could try adding extra Mg to the PCR reaction to make up for the Mg binding to the EDTA. Or, instead of using TE for elution, try using Tris-HCl (5mM, pH 8.5) to avoid the EDTA all together.
If you are having problems with DNA degradation you might also want to store your samples in the freezer and double check that all your plastics are DNAse free.
Have you tried amplifying the colonies you get on your plates using your fungal primers? This would help you confirm that the primers are actually able to amplify the fungi present in your sample. I would also extract using the same technique that you use to extract the DNA from the woodlouse. Fungal DNA can be very difficult to extract and I find most kits don't do a very good job even on a pure culture, so your woodlouse extraction may simply not contain any fungal DNA. The technique I usually use to extract fungal DNA is to grind the sample under liquid nitrogen and then use the CTAB extraction. I've never tried it in a situation like this, but I don't see why it wouldn't work.
Is there any way you can reduce the amount of woodlouse DNA relative to the fungal DNA in your sample? If you are interested in fungi in the gut, for instance, maybe you could dissect out the gut and only extract DNA from that, rather than the whole organism. This would allow you to have a better idea of how much fungal DNA is present in your extract, rather than just looking at the concentration of total DNA.
- Rais Ahmed added an answer:4Problem in real time PCR optimization; probes are not binding.
I am facing a problem in real time PCR (CFX96 Bio Rad) optimization. I have to find out the melting temp Tm of primers that is 54 C, and program setting is 95c for 10 min, 95c for 15 sec and 54 C for 30 sec, but probe is not binding. When I run real time PCR, no peaks appear on the screen. I run amplicon on gel, but bands were seen. Where is the problem? I think the probe is not binding. What do you suggest? Please give your fruitful suggestion to solve this problem.
Dear Jan Lennart Korner, i am using CFX96 bio rad system. i am using TaqMan probe. yes i have first run gradient PCR and find Tm of primers 54C and on gel i got clear band. then i had run real time PCR at annealing temp of 54C, but no peaks are seen on screen but i got clear band on agarose gel..i think amplification is going on but probe is not binding? what you suggest?Following
- Lieven Waeyenberge added an answer:5Any advice on a 2nd PCR Problem- Smear?
I did a 2nd PCR to amplify the PCR products from first PCR I did. The first PCR products are in lanes 3 and 4. The product bands are quite light but still there. There are two bands underneath the DNA band- we think those are the forward and reverse primers (why did they show up?). Despite this, I did a second PCR with the products from the first PCR and the results are in lanes 5 and 6. There seems to be a high-molecular weight problem and a smear underneath. I really don't know what the problem is. Could the problem have something to do with the fact that I did a second PCR with the first PCR products in spite of the extra primer bands? And what should I do next? Should I go all the way back to doing PCR with the original template or can I modify my first PCR or second PCR products somehow?
I see your PCR-product is very weak. So I would not recommend to dilute it or to purify it for the 2nd PCR as you will end up with even less PCR-product. Using nested primers is possible but than you need to design or find somewhere nested primers and it will make your PCR-product (a bit) smaller. I would do the following: try the "band-stab-method". How? First prepare master mix like you did for the first PCR and divide it in tubes (number of tubes equals number of samples, in your case 2 tubes because you have 2 samples). Put most of your PCR-product (from 1e PCR) on agarosegel and do a normal electrophoresis. Make your band visible (DNA-dye in gel or color gel afterwards) on the illuminator (like you did for the photo). Use a pipettip to stab the gel on the position of the PCR-band, so you need to stab the PCR-product on the gel and transfer your tip into to the tube with mastermix. Repeat this with another tip for the second sample. Shake the tip gently in the master mix, remove it and repeat your PCR (= 2nd PCR). If all goes well, this will re-amplify your PCR-product without creating smear as all unwanted PCR-products, primerdimers and other things are left behind on the gel which you can throw away. Only DNA from the band you touched/stabbed will be present in the mastermix. So you can see it as a kind of purification without using a purification kit. This method works for me, even on weak products.
By the way: it is quite normal to see left-overs of primers on the bottom of your gel. The other band probably are primer-dimers. Don't worry about these too much when it is possible to create sufficient amounts of PCR-product. However, if the PCR is not performing well resulting in a very weak PCR-product, the band of primers is stronger (most primers were not incorporated in the PCR-product) or a lot of primer dimers are visible. This means that the primers are not suited very well for making the PCR-product or the design of the primerset is not optimal. In this case it would be better to design other primers. If this is not an option (on short-term), use the "band-stab-method" in stead.Following
- Shefali Desai added an answer:3How do I set the parameters and assess the results in primer-blast of NCBI
Recently, I begin to use PCR for my research,the first step is to design the primer.Obviously,primer-blast might be the most conveniently tool for this job.But,I got a little confused about setting parameters that may lead to a primer which could balances the specificity and productivity and avoids forming some secondary structure or hairpins.As I have tried before,when putting my target gene in it,I got some primers,and different results gotten with different parameters,it is really a confound thing for me to assess the results,although i have read the article1 related to this tool.So I am wondering that is there anybody who use it often could give me some advices or some articles to help me!Thanks a lot!
(1.Ye, J., Coulouris, G., Zaretskaya, I., Cutcutache, I., Rozen, S., & Madden, T. L. (2012). Primer-BLAST: a tool to design target-specific primers for polymerase chain reaction. BMC Bioinformatics, 13(1), 134. http://doi.org/10.1186/1471-2105-13-134)
Try to check on GC % which should be ~50 % which would help you in lower Tm and good annealing temperature.Following
- Joe Baxter added an answer:3I have to design primers for RNA sequence: How can I make sure that my primers do not amplify repeated regions or contaminant DNA?
The are no exons or introns in the sequence that can be used for Exon junction span, what to do??Following
- Tyler Square added an answer:1Does it make any difference if a target site in my CRISPR design is on the reverse strand or does it have to be in the forward strand?
I am designing CRISPR sites in zebrafish and I am using T7 promoter for guide RNA synthesis. Does it matter if the target site I get is on the reverse strand?
Not if you are using the standard Cas9. This means you can essentially target either GG(18N)NGG, or CCN(18N)CC. We do both in the publication I have linked to this post. Cas9 makes a double stranded break, so as long as you direct it to the right place, it should work.
However, the dCas9 ("dead" Cas9, which simply "sits" on the target site and blocks transcription) apparently works better if you target the non-template strand (e.g. http://www.ncbi.nlm.nih.gov/pubmed/26168398).Following
- Can Kiessling added an answer:3What positive controls can I use for PCR?
I am doing PCR for cDNA obtained from various tissues, to check which ones express my gene of interest. Aside from using primers for a gene that I know is expressed in all tissues, what other positive controls can I use?
Your positive control should be a sample that gives you consisent amplification as you have shown that tissue expresses that gene.
When you are running your experiment and include this positive control and it amplifies, you know that your primers work. If there is no amplification in tissue of interest then it is likely not expressed in that tissue.Following
- Muhammad Tayyib Naseem added an answer:8I designed primers based on a cDNA template, but I got a stripe from the genome DNA template. When I ran PCR, I got nothing from the cDNA. Why?
I designed primers based on a cDNA template, but I got a stripe from the genome DNA template. When I ran PCR, I got nothing from the cDNA. Why?
You can try with different dilutions of cDNAFollowing
- Lucia Taja added an answer:14I have a PCR purified fragment for sequencing but when I sequenced it, I suspect there are two amplicons. What cloning strategy can I use to fix this?
A fragment of DNA from a genomic prep was amplified by PCR using specific primers. This fragment was PCR purified and sent for sequencing. The sequencing results however indicated that possibly more than one amplicon is present in the sample (mixed sample). What cloning strategy that can be used to resolve this issue so that the nature of the PCR products can be accurately deduced?
If you cannot separete both bands by gel electrophoresis, you can clone the PCR producto using the TOPO-TA cloning kit and sequence several clones in order to identify both productsFollowing
- Yu Tao added an answer:2How can I make a stock solution of (cDNA,primer, syber green, dd.w) for real time PCR?
I did real time PCR but time standard curve of b-actin is not quantified so I think there is problem in my stock solution.
Maybe there was something wrong with your standard plasmid DNA.Following
- Sa Roth asked a question:NewWhich are good RT-PCR primers for Mcl-1?
I am looking for Mcl-1 RT-PCR primers (I am using POWER SYBR from Thermo Fisher).
Have you worked with Mcl-1 and found good primer pairs?Following
- António Maximiano Fernandes added an answer:3Does any of you have experience in using primer BLAST for primer design?
I have to design species specific primers and to check the specificity of some primers. I have not much experience about it so I was using primer BLAST. Do you think it is a good tool? Do you have any suggestions on its use? Thank you a lot
From NCBI ----- http://www.ncbi.nlm.nih.gov/tools/primer-blast/
- Add acess number: eg (M84709)
- Change parameters (if necessary) as Asif said
- Press get primers
See paramers and protocol in atach
Sigma also have online design wonderfulllll.....
- Marcos De Donato added an answer:3Why do I get a weird background on Southern blot and cannot detect ladder?
Got problems about Southern blotting...
There are two problems for my Southern blot using 32-dCTP labeling.
One is that there’s no ladder.
The other is the huge weird background. (very clear black in the upper part and white in bottom part)
Here is my experimental procedure...
I tried to genotyped the recombinant yeast strains, so I treat the genomic DNA with XbalI for 37C 4hr. Then I loaded my samples on the 0.8% agarose gel and run at 60volt O/N.
Then I check the gel and found that the DNA was cut appropriately.
After that I start the Southern electro-transferring for 2 hr. Before the hybridization, I soaked the membrane with 0.8N NaOH (250ml) for 15 min and then change the sol to 10X SSC buffer for another 15 min.
Next, I washed the membrane with 0.5M phosphate buffer in the Southern glass tube. And then I pour 25ml Southern hybridization buffer (65C) into the tube and put it in the oven (65C) for 1 hr. [Pre-hybridization]
At the same time I prepared my mixed probes which contain: 100ng of my PCR probe, 0.1ng lambda DNA BstEii and some MQ water.
First I keep the mixed probes at 100C for 3 min and then put it on ice.
Next, add 5ul High primer mix and 4ul 32P-dCTP. Incubate the mixture at 37C for 20min.
Add the mixture into a Mini Quick Spin DNA column, and centrifuge the mixture at 3400rpm for 4 min.
Check the radioactivity of the mixture using gager counter
Incubate the mixture at 100C for 3 min, then put it on ice
Add the mixture in hybridization buffer and pour it in the glass tube (with membrane inside)
Rotate the glass tube at 65C overnight
Wash the membrane four times (15min each with wash buffer at 65C)
Expose the membrane.
Sorry about the lengthy experiment procedure typed above…
Any comment is welcome!!
If you have access to a real time thermal cycler with HRM capabilities, you can also do high resolution melting analysis (HRM), which has advantages for discovering new SNPs, although it does not work in all SNPs.
Check this paper (sorry I do not have any published of my own):
- FİLİZ RZ added an answer:5Any suggestions about a specific primer in RT-PCR which gave very clear and specific band several times but stopped working?
I have been using one primer in my RT- PCR experiment. I got very good specific bands several times in several weeks However, suddenly, It stopped working. I ordered new stock, but new stock is also not working. But interesting point is, control actin band is as normal as were before. I did check gradient temprature again but negative. Now I am confused on this problem. Can anyone help me figure out?
Maybe RNAs can be degradation for any reason...Really, RNAs is very sensitive.
You should check your RNAs on the gel and any programme(just like nano-drop)..
Shortly,RNA isolation is very important for RT-PCR.Following
- Diana Duarte-Delgado added an answer:3What are some reasons that could contribute to some bands of a PCR being dim while others noticeably brighter?
This an image of one of the PCR results received with the named problem of inconsistent band brightens.
Or perhaps, some DNA samples are of not good quality, are degraded and present troubles in the amplificationFollowing
- Simeon Rossmann added an answer:2How does the CLC (Main Workbench) TaqMan primer designer compare to other tools?
I want to design primers/probes for a TaqMan application. Since I am used to do primer design in the CLC Main Workbench that would obviously be my first choice. It offers convienient options to design primers from alignments that are able to discriminate between rather homolog sequences. However, I didn't use it for TaqMan probes yet. While checking the software options I saw that most people recommend Primer3 in this context. I didn't really get any insight into the algorithm that CLC uses to determine primers in the manual. However, since you are quite free to set a multitude parameters, I guess that the hits are scored mainly based on fitting these as well as the alignment with the homolog sequences. (Edit: From the help menu: "CLC Main Workbench employs a proprietary algorithm to rank primer and probe solutions. The algorithm considers both the parameters pertaining to single oligos, such as e.g. the secondary structure score and parameters pertaining to oligo-pairs such as e.g. the oligo pair-annealing score." Not very helpful in my view.)
Does anyone have experience with using both, CLC Main Workbench and other available TaqMan primer/probe design software (especially Primer3)? Would you prefer one over the other (why)? Does anyone have any details on the algorithm that the Taqman primer/probe prediction in CLC is based on (in other words, is it related to Primer3)?
I guess it won't make much of a difference and I plan to counter-check both methods for my targets and use basic additional checks (BLAST of primers, order and test multiple options). It would still be nice to gather some input from people with some experience beforehand.
Edit: I am currently running Main Workbench 6.9.1, however I didn't see any changes in the Primer designer in the changelogs to the current version, so that should not really affect the results.
It's a helpful and simple article about the basics of TaqMan primer/probe design. Does not really answer my question but might still be useful for other people stumbling upon the question :)Following
- António Maximiano Fernandes added an answer:2Dose any one explain this picture from UV electrophoresis after running in PCR?
Knowing the first line from right side it is negative control and the second line for internal control and third line for positive control?
and we have also in my kits
* HPV-267-325 bP
* Internal control-723 bP
I need to know these is negative result or primer- dimer?
See the picture attached please,
I agree with Paul . It may be related also to high concentration of primers. Check the concentration in work solution.
- Sabine Strehl added an answer:1Can anybody please provide me neomycin resistance gene sequence or primers for neo gene incorporated in HEK293T cells?
I am trying to develop stable cell lines for my desired protein in HEK293 cells through G418 selection. However, HEK293 cells are not getting killed by G418 even at high conc (1000ug/ul). Thus, I would like to look for whether my cells already possess a neomycin resistance gene (neo phosphotransferase) or not? So please help me by providing the sequence or primers for neomycin resistance gene?
Read this thread and you can spare yourself the PCR; but you have to clone another resistance cassette into your vector...
- Touraj Farzani added an answer:4Is it possible to design primers (R and F) that are different in size?
I am designing a set of primers that have His tag in N side. My primers length are 45 and 25 and the specific regions for my interest gene are 20 for the both primers. Is there any chance of success?
I deeply appreciate you.Following
- Laurence Stuart Hall added an answer:9How to achieve better amplification in qPCR?
I am currently comparing ULBP-4 expression in cancer cells. The method I used for qPCR as follow:
a) cDNA synthesis = 1ug of RNA in 20ul
b) I diluted the cDNA 10X prior to qPCR
c) I used 4ul of diluted cDNA, the final concentration of primers (F/R) is 100nM, total reaction volume is 20ul
a) Average Ct value: 29
b) Average melting point (from melt curve): 75C (I believe this is the melting point of primer dimers)
How can I improve my technique to produce reliable results?
IDT primer design tool is excellent
This will screen for primer dimers (hetero dimers) as well as intra primer annealing via hairpin loopsFollowing
- Richard Campbell added an answer:1Do various DNA templates for PCR affect annealing temperature for the same primer pair?
I have done PCR for a primer with genomic DNA templates which are isolated from two different bacteria. At 55 degree C, only the first template generated target amplicon (sharp, bold, specific) whether the second one didn't (the amplicon was seen at Ta = 46 degree C, light band). Can anybody help me to explain this phenomenon?
When using different templates you may need to adjust the annealing temperature to compensate for potential off targets. I would check the genomes of the different bacteria for your primers to ensure you do not have off targets. Also make sure the sequences that your primers are targeting are the same between the 2 templates being used.
Are the different bacteria different strains or genomic preps from different colonies?
If it is different preps of the same strain, then it may just be a DNA quality issue.Following
- Vivica Grotelueschen added an answer:2How to prepare an equimolar mixture of primers?
I want to study the AOB oxidizing community in the soil samples.For that I am selecting the primers CTO 189F / CTO 654R.(Kowalchuk et.al 1997).
As said in the article [attached] , PCRs were conducted with an equimolar mixture of three forward primers (CTO189fA-GC, CTO189fB GC, and CTO189fC-GC) and a reverse primer containing a single ambiguous base.The forward primers CTO189f-A and CTO189f-B (GGA GRA AAG CAG GGG ATC G) and CTO189fC-GC (GGA GGA AAG TAG GGG ATC G) were synthesized separately and collectively referred to as CTO189f-GC.
Regarding this I have two questions.Please help.
Q1.This means that I will order three forward primers where in primer named A and B , the R will be replaced by A/C and primer named C will be as it is.
Is my understanding correct?
Q2.How can I prepare equimolar mixture of three forward primers?
Normally,I am diluting my stock(100picomole) to 10picomole and then I am using 0.4ul of each primer (F and R) for 25ul PCR reaction mixture.On this case,if I dilute each forward primers to 10 picomole and then add equal volume of each to make it 0.4ul for 25ul PCR mixture will it be correct?Please help.
Q1: As far as I know, the "R" means A or G (not A or C). Besides this, I agree with you that one of the primers is ordered with an "A" at the position of the "R", the other with a "G" instead or "R".
Q2: In the end you will have to find the best concentration of primers empirically. There are two ways of interpretation, resulting in different total primer concentrations in which the molar relationship of the thre forward primers remains the same, but the total primer concentration in the PCR mixture and the relationship with the reverse primer differs:
(1) If you first mix the working concentration of your primers and use 0.4µl, the total concentration of forward primers will be the same but the concentration of each individual primer is only one third. This way you might get less primer-dimers and I think normally there is more than enough primer in a PCR, so it want matter if you have less of the individual ones.
(2) Dilute the stock solutions of the primers with the other primers, meaning, that for your 1:10 dilution instead of using 10µl of the primer and adding 90µl of buffer/water for each primer individually, you take 10µl of primer A stock, 10µl of primer B stock and 10µl of primer C stock and fill up with 70µl of buffer/water. Now you have a equal molar working concentration of the three primers (10picomole each) and can work with your usual 0.4µl/25µl. Of course you have a higher total concentration of primers (or your dilute the mix by factor 3 again). Easier with same result: Add 0.4µl of each primer to you PCR mix and adjust the water/buffer volume.
I guess, what you want for your experiment is the first thing, otherwise you would have an untypical missmatch between forward and reverse primer concentrations and an unnormally high total primer concentration which might disturb your PCR since the primers might prefer playing with themselves instead of with your template.Following
- Kaja Nilsen added an answer:4Why are my amplification curves non-exponential?
Did a specificity test of newly designed primers. They have been working well, no abnormalities, until tested on human DNA. 2 samples were negative, but 3 were positive, with non-exponential amplification curves.
Does anyone know what might cause this?
Also these positive samples had double-peaked melting curves.
The products have not been analyzed further by gel electrophoresis, as the experiment did not require this. But I am eager to learn, and really wonder what might cause these weird curves..
Ok, I see.
Thank you so much for taking the time to answer! This really helped me a lot!Following
- Zhiwei Chen added an answer:8Any advice on a wrong PCR product size?
I amplified a gene(np gene of TSWV) form total RNA.
total RNA was extracted from leaf of tobacco.
I had used these primer pair:
F : ATGTCTAAGGTTAAGCTCACTA
R : TTAAGCAAGTTCTGTGAGTT
these primer pair amplify 777 bp pcr product.
I had checked these primers in "Primer Blast". primers was specific for TSWV.
but my pcr product is about 500 bp.
what is there problem, why pcr product is not 777 bp?
the pcr cycle was:
94 --- 94 51 70 X35 --- 70
3:00 ---0:30 0:30 1:00 ---- 10:00
I had used Red Master Mix from Ampilicon.
Make sure your RNA extraction for virus. Did you use cDNA as the templet ? And is there any other bands in your amplification? Then you can sequence your product to see if it is a virus sequence or just a sequence from plants.Following
- Antônio Augusto Fonseca Jr. added an answer:3How can I troubleshoot my standard curve in real time RT-PCR?
Hi! I did some primer design for my real time RT-PCR. I did the conventional PCR to check whether they amplify the right region in the right condition. I designed the IL-1b primer so the amplicon to be around 100 bp, not more than 130 bp so I can use it for real time.They did amplify the region. Next, I ran my samples in real time with the same primer. However, the standard did not make any linear line. In another time, it makes a linear line but otherwise with the normal one, so the usual slope of -2 or -3 was not found. I thought the sample was the problem so I did run the samples with the same procdure,reagent, and standard samples for GAPDH and NS-1 primers, but they amplify perfectly with slope of -3.
I wonder if I have to re-design my primer. Any thoughts?
Thanks in advance
Did you check primer Tm and primer concentration? Is it sybr or probe? did you add MgCl2?Following