newmanreb

Still confused about this if anyone is able to help me out! Thanks.

Hi there, I'm having trouble with a restriction digest of a 5 kbp PCR product. The product was purified using a Qiagen PCR product clean up spin column kit. This product is too big to travel much on the gel, so I confirmed that it is the sequence I think it is by performing a nested PCR on the 5 kbp product to amplify a 300 bp product. This was successful. I am using a double digest of brand new NEB enzymes MseI and MluCI. These enzymes are TimeSaver certified (should digest in 5-15 minutes at 37 degrees) and both have 100% efficiency in the rCutSmart buffer. I set up half reactions of what NEB suggests with 0.50 ug DNA. I also tried doubling the amount of enzymes and doubling the time from 15 mins to 30 mins. Still, nothing I've tried results in the band on the gel changing. The main band remains relatively unchanged and there is a faint smear on the rest of the lane. I am stumped. I'm concerned that maybe the digest is working but I simply can't see any bands because these enzymes cut so many times? They should generate 49 fragments and the largest one is around 500 bp. I tried cutting this product with BamHI though as well and it didn't change anything on the gel even though it should generate fragments of 3456, 1088, and 364 for this sequence. Any advice on how to troubleshoot this would be greatly appreciated. Thanks in advance!

Thank you CharonY! This helps me out a lot. As for the probe that binds to the target region, how long in bp would that tend to be? I suppose I assumed that the probes would be about the same length as primers, and although primers specifically bind to the target, they are clearly not enough to determine the identity sequence. However, they may be thinking of universal primers used to amplify the 16S region, for example, which are not designed to be species-specific and therefore not intended to identify the sequence.

Hi all, I'm doing some research at my company about implementing qPCR, because we are still in the dark ages with conventional PCR (cPCR). I came across this information in a publication, and I haven't been able to find more help online to clarify what the authors are saying. I was hoping someone here could illuminate things for me. This is a quote from a 2002 paper, citation below. It says, "A major reason why classical PCR has not been adopted by most plant disease regulatory and diagnostic laboratories is the time and labor required to confirm the identification of the PCR amplification product. The simple presence of a particular molecular weight DNA fragment in an agarose gel does not prove the identity of the resulting band, and verification of the amplified product must be done by Southern blot hybridization. Another major factor is that the technique is not much more sensitive than ELISA, and it is much less sensitive than isolation of the organism on semiselective agar media (Wang et al. 1999). However, these concerns do not apply to qPCR." Later in the article, the authors include this statement: "Real-time PCR has many important advantages over classical PCR: (i) it eliminates the need to do a Southern blot to confirm identification of PCR product...." I have a few questions about these statements: 1. Has cPCR advanced since 2002 to the point where confirmation of the resulting sequence is not always needed? I'm fairly certain cPCR is more sensitive than ELISA, potentially by a lot. We do some plant diagnostic tests where a strong band is considered a positive, but usually we will use Sanger sequencing for identification. (That could just be lazy assay design on our part - we're working on that.) Back in 2002 Sanger sequencing was definitely around, but maybe it was significantly more expensive than Southern blotting? 2. My main question is this - why is it that qPCR products do not need sequence identification? My understanding of qPCR is that it's almost the same chemistry as a cPCR reaction, it is simply monitored with fluorescence. Is it because of the increased sensitivity? How would that be explained? Thank you so much for any thoughts & discussion. Rebekah Schaad, Norman W., and Reid D. Frederick. "Real-time PCR and its application for rapid plant disease diagnostics." Canadian journal of plant pathology 24.3 (2002): 250-258.

Hi all, we are thinking of upgrading our gDNA quantification method from the agarose gel method to the NanoDrop. I've been trying to research online what quantification methods are best for DNA extracted from plant tissues, including wood and decayed wood, because I know these extracts can introduce unique contaminants that would not be present in something like blood, for example. I haven't found much information online about this topic specifically, just lots of discussions about NanoDrop vs. Qubit vs. qPCR quantification, etc. I think the NanoDrop will be a good move for us but I was wondering if anyone out there with expertise in plant DNA extracts has a favorite quantification method. Thank you!

Hi there! I am with a company who is lucky enough to be expanding our operations into a new building. Our PCR laboratory will feature 3 rooms: A reagent prep room, a sample preparation room (DNA extraction) and a post-PCR laboratory. Since we are designing the building, we get to decide what air flow we need throughout this unidirectional workflow. Now. It's clear to me that the reagent prep room, the "cleanest room", needs positive pressure to keep contaminants out. Also clear - the post-PCR room needs negative pressure to keep amplicon contaminants IN. What is not clear to me is how to treat the extraction room. In both of my previous labs, we didn't have the luxury of having a separate room for DNA extraction. DNA extraction was considered a "pre-PCR" activity and done under positive pressure in the pre-PCR area. HOWEVER, upon further research, I've found multiple sources which say the DNA extraction room, being "dirtier" than the reagent prep room, should have negative pressure like the post-PCR room for the reason of keeping DNA inside the room and nowhere else. This resource, on page 13, shows that the nucleic acid prep room should have negative pressure, and then on page 16 shows an example floor plan with POSITIVE pressure in the extraction room! https://www.aphl.org/programs/newborn_screening/Documents/2015_Molecular-Workshop/Molecular-Laboratory-Design-QAQC-Considerations.pdf What pressurization would you use in a DNA extraction room: positive or negative? I have found conflicting sources online about what to do about this. I would really appreciate any insight. For context, our rooms are separated by a shared hallway. My concern with putting negative pressure in the DNA extraction room is that we will pull in amplicon contamination. This is the room in which we will be opening our reaction tubes to add DNA template - I wouldn't want to introduce amplicon contamination at that point. Thank you for any input!