Ran conventional PCR to amplify the full length abalone withering syndrome 16s product for subsequent cloning. Used four different DNA plasmid preps as template, for comparison purposes. Plasmid preps used were:
Ran 20uL (of 50uL reaction) of each sample on 0.8% TBE gel.
Lanes (left ro right):
1 – Hyperladder I (Bioline)
2 – p16RK7 (from 20120718)
3 – p16RK7 (from 20120718)
4 – p18RK7 (from 20120718)
5 – p18RK7 (from 20120718)
6 – pWC8 (from 20120718)
7 – pWC8 (from 20120718)
8 – p16RK7 A (from 20131106)
9 – p16RK7 A (from 20131106)
10 – NTC
11 – NTC
Expected a band of ~1500bp. Bands of all samples (except pWC8) run at ~1500bp. pWC8 runs a bit higher and will not be used for cloning, as part of our troubleshooting the failure of our plasmid qPCR standard curve for withering syndrome.
Essentially the same results as the previous run. No template controls do amplify, but EXTREMELY weak and late. Melt curve analysis shows that the signals for the no template controls don’t cross the threshold set by the software.
However, I just looked back at the qPCR results from 20120208 where I used these V. tubiashii 16s primers and realized I got the same results from the cDNA (double-peaks in melt curves and amplification in the no template controls)!! So, I suspect that this primer set isn’t that useful. Will have to examine other sets of V. tubiashii 16s primers to use. Will discuss with Steven.
Ran qPCR on the Taylor water filter DNA extracts from yesterday using V.tubiashii 16s primers (SR IDs: 455, 456). Used RE22 DNA as a positive control, provided by Elene. Master mix calcs are here. All samples were run in duplicate. Plate layout, cycling params, etc can be found in the qPCR Report (see Results).
All samples amplified, including the negative controls. Negative controls exhibited very weak, late amplification. Additionally, many of the samples have a “shoulder” or apparent double-peak present in the melt curves. Will repeat to see if I can eliminate amplification in negative control samples.
Performed qPCR on all 12 samples. Used Cg_EF1aF/R2 (SR IDs: 1410 & 1412) for one set of qPCRs and Vtub_16s_F/R (SR IDs: 455 & 456) for the other set of qPCRs. Used pooled C.gigas cDNA (from 20110311) and RE22 DNA (provided by Elene) as positive controls for C.gigas and V.tubiashii, respectively. C.gigas gDNA (7ng of BB16 from 20110201) was used as a negative control for EF1a. Master mix calcs are here. Plate layout, cycling params, etc can be found in the qPCR Report (see Results). All samples were run in duplicate.
C.gigas EF1a – Positive control amplified. Negative control and no template control were all clean (i.e. no amplification detected). The majority of samples had amplification, however two samples had no amplification at all (samples 132 & 136).
V.tubiashii 16s – Positive control amplified. No template controls exhibited amplification in both replicates. All samples exhibited amplifcation, however nearly all of the melt curves have multiple peaks present, suggesting that more than one target is being amplified. I suspect this is due to residual gDNA, but this fails to explain the amplification in the no template controls which also exhibited dual peaks in the melt curves.
Spoke with Steven and he suggested to skip troubleshooting the V. tubiashii 16s for now and proceed with trying to qPCR some additional V.tubiashii genes. Will talk with Elene to see if/which additional genes she has primers for.
This is a repeat of the previous qPCR from earlier today, BUT I think I might have used the wrong primers in the earlier qPCR (see below). Set up qPCR with the correct (I’m 100% sure of this) primers. Plate layout/workup is here.
Results: Well, in retrospect it looks like I DID use the correct primers earlier! However, the problem is the same. But, the melting curves in the H2O-only samples don’t seem to be the same as what is being seen in the RNA samples, suggesting that the signal in the H2O-only samples are likely primer dimers (melting curve peaks are shifted to the left and are VERY low signals; barely above background).
So, what to do now? Mac has a mad ea suggestion to spike some water with gDNA and DNase treat the sample to assess whether or not the Dase treatment is actually working or not. I think I’ll do this.
Results: Looks like gDNA contamination is still present!! This is insane! However, the two water-only samples produced a signal suggesting that something else is contaminated. Will try just qPCR-ing water to see if I can get a clean signal. Will use “store-bought” PCR water instead of NanoPure water.
*UPDATE**: Possibly used 16s universal bacterial primers instead of H.crach 16s primers! Doh! Will re-qPCR using the correct primers.
This is a repeat of yesterday’s PCR due to the presence of bands in the water-only samples. Will use reagents and universal 16s bacterial primers (27F & 1492R) provide by the Horner-Devine lab in hopes of: 1) getting this two work and, 2) figuring out the source of the contamination.
All rxns were prepared sterily and all instruments, racks, tubes, tips and water were UV-sterilized for ~45mins in the biological hood. Rxns were prepared in the biological hood. PCR setups are here. Anneal 60C. Cycling params same as yesterday.
Lane 1 – 100bp ladder
Lane 2 – DNA (HD Rxn 1)
Lane 2 – H2O (HD Rxn 1)
Lane 3 – H2O (HD Rxn 1)
Lane 4 – DNA (HD Rxn 2)
Lane 5 – H2O (HD Rxn 2)
Lane 6 – H2O (HD Rxn 2)
Lane 7 – DNA (SR Rxn)
Lane 8 – H2O (SR Rxn)
Lane 9 – H2O (SR Rxn)
Lane 10 – 100bp ladder
Results: Well, we got our band and NO contamination in any H2O lanes. The super-bright, 1500bp band will be excised and purified using Millipore spin columns and submitted for sequencing. However, this gel is interesting because the primers provided by Mike (used in HD Rxn 1 and SR Rxn) did not amplify anything…