I am working on environmental DNA (eDNA) in aquatic environments. As mentioned by Barnes et al. (2014), environmental conditions influence eDNA stability in aquatic systems.
Recently, the eDNA that I collected from a small and relatively acidic pond has relatively low yield when extracted, hence it effects the downstream procedures, such as PCR. I am troubleshooting with the PCR step rather than the phenol/chloroform extraction step.
Is there any PCR protocol that would essentially work on this particular issue (acidity causing degradation of eDNA), ie adding some particular reagents, and or changing temperature settings? Thus far, I could not find any particular solution in the literature.
If I understood correctly, then the template DNA of your samples is already degraded. So you start the PCR with degraded material.
There is no way to "undo" degradation that already happened.
Now consider the PCR with a pair of primers to amplify a target sequence. The stronger the degradation, the more of the original intact DNA modelules containing the target sequence will be broken somewhere between the the two primer binding sites. Those molecules that have at least one break inbetween won't be amplified (exponentially).
This probability depends, as said, on the degree of degradation (what you can not influence) and on the distance between the primer binding sites (the larger the distance, the more likely will be a break inbetween). So the only way to improve your chance of a successful amplification is to chose primers with a short distance inbetween (producing short amplification products). The products may be as short as approximately 80bp (shorter products will not be easily distinguishalbe from primer-dimers than may also be produced).
Have you tried Betaine and DMSO enhancers for PCR? i have used them for my dscDNA, they are distinctively effective compared to water....
Dear Hediche Zunnun, I have not yet tried them out. Upon seeing your answer, I've searched for the effect of these agents on PCR, and found this:
http://www.promega.com/~/media/files/resources/promega%20notes/65/betaine%20and%20dmso-%20enhancing%20agents%20for%20pcr.pdf
Their results indicate that when only DMSO was added, there is the expected product, but also some nonspecific bands. When only Betaine was added, the yield seems to be less than in reactions with DMSO. Combination of these two does not have any specific improvement on the brightness of the band as compared to the PCR reaction including only Betaine as an additive enhancing agent.
I am planning to add DMSO in my next PCR reaction. Any suggestions on the concentration?
Well, I added 1.25 ul to total volum of 25ul Taq PCR reaction, but betaine was 6.25ul to 25ul of total reaction. you can do little more research on their quantitiy online, because i did not have record of the concentration of both Betaine and DMSO(sorry).
for one gene sequence DMSO was quite effective, but when i used it to another Gene amplification, there was not product of interest. Length of genes are pretty much the same. Wish you all the best !
Dear Jochen Wilhelm, thanks for your detailed explanation. As for the latter issue you raise, I think I would not be able to change the length of the amplicon, since I am targeting the short fragment of the mitochondrial COI region (313 bp) for metabarcoding. Amplifying a shorter region would not help my purpose, unless there are other shorter and variable enough COI regions used for NGS, which I am not yet aware of. I will hence work around the enhancing agents for PCR, rather than changing the primers.
What DNA extraction protocol did you use? What pH was the water?
Do you use barcoding primers? I prepared some samples few months ago that amplified well with the standard primers but when I used the barcoding primers used for MiSeq... well, I had serious problems with them
I am doing phenol/chloroform extraction. The pH is around 5 in this pond, I've been told by people carrying out physicochemical analysis in this water. And I am using short (313 bp) metazoa COI primers.
I think you can first try different kits or procedures for extracting DNA, because once you get a good quality DNA then you can easily do PCR. you can check DNA qty and purity by nanodrop.
If you pelleted your sample and resuspended in a buffer or even in saline, the pH shouldn't be a real problem as the waters are poorly buffered, usually. So with the equilibrated phenol won't be an issue.
Have you measured the [DNA] after extraction? Do you have enough? You can also check for inhibitors in your sample by spiking with a good DNA you know it amplifies well. DO you add BSA? My environmental samples don't usually work if I don't add it even after using kits with inhibitor removal steps
I usually resuspend my pellets in distilled water. I did measure the DNA after extracting, and there seems to be DNA, with concentrations ranging between 20-40 ng/ul. However I am getting this really weird curve, which I am currently trying to figure out why occurs.
Also for BSA, I used to add it for my previous PCR reactions, but not on the environmental DNA yet. It did not make much of a difference for my previous PCR settings, I was usually getting similar bands with/without BSA.
Like mentioned earlier, I do not see any problem with the pH as long as you use a buffer. Eventually you can just neutralize your sample before approaching the extraction. Or better, if it is just a water sample, try to filter and work from the filter as starting material. But according to your spectrum, you do have DNA (absorbance at 260) so I think the problem is that you are co-extracting other things that absorb at lower wavelengths. Not sure what type of sample is that, but you could be co-extracting humic acids and/or most probably metals (the solubility of many metals increases at acid pH). If that is the case, I would definitely filter and work from the filter. Have you checked your extracted DNA on a gel? What does it look like? If the integrity is ok, you can try to purify from the ge or even PCR from the gel. Keep us updated!
Dear Antonio García-Moyano, these are great insights! For my purpose, I do not prefer filtering the water samples, since it would potentially cause the loss of floating eDNA, which I want to capture as well.
I did use Qiagen kit on phenol/chloroform extracted DNA, so it was an additional purification step. It indeed helped, after which I was able to get rid of that peak at 230 nm. I agree with you that there might have been other substances that were co-extracted with DNA, and an additional purification step took care of this. My only worry is that it might have caused the loss of some of the DNA as well. The bands were relatively faint.
When I measured the concentration using Qubit it was fairly low, ca 5 ng/ul. But overall what matters is the amount that I have for the latter NGS step, so I think I am safe for now.
1) I hope you agree that the lambda to use is 260 instead of 230.
2) Did you actually quantify how much bacteria are in the sample before going in for extraction.
3) Going on a hunch here, can you perform a spectral scan to see if you have a phenol carryover, this can really kill the PCR. If so a couple of ethanol extractions might help.
4)How does your positive control for the PCR look ?
Dear Jay Siddharth, yes I definitely agree. However my point was that I ideally should see a higher peak at 260 nm as compared to other wavelengths, since I extracted DNA with the phenol/chloroform method. The problem is that (apart from the fact that we are not living in the ideal world) I extracted other things along with DNA, which give me that peak at 230 nm.
I am not really interested in bacteria in my samples, and am targeting the metazoan taxa. Just out of curiosity, how would you quantify their amount, i.e. what tool would you use?
For detecting phenol contamination, I think that one needs to check if there is a peak 230 nm as well, here is some details on how phenol's UV spectrum looks like: http://seqanswers.com/forums/showthread.php?t=21280
I do get a shoulder at 230 nm, which could be due to some humic acids as well as phenol. So, I am not sure how to dig into this information.
Sadly, I have not had any positive control for my PCR reactions.
Also dear @Siddharth, you can take a look at my other question, which explains the issue in a bit more detail: https://www.researchgate.net/post/How_can_I_troubleshoot_with_high_absorbance_at_230_nm_260230_is_12_for_a_DNA_extract
Given the spectrum you linked, wouldn't it be better to use the OD260/270 ratio to check for phenole contamination?
OD230 will be influenced a lot be Tris, acetate, DTT and Sodium that may also be present in relatively high concentrations, obscuring the phenole absorption at this wavelength.
Yes it is true. In this case, I do not think the contamination in my samples is due to phenol, since there is no such a peak like the one in the link. Also 260/270 ratio seems to be more than 1 for my nanodrop measurement.
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