Previous qPCR was done incorrectly (wrong primers), so am repeating with the correct primers. Performed qPCR using q18s primers on DNased RNA (1:100 dilution to match final concentration of template after making cDNA). qPCR set up and plate layout are here.
gDNA dilutions were used as positive controls. gDNA = BB11 (0.49ug/uL) from 20090519. Used 5uL of 1:10, 1:100 and 1:000 dilutions.
Results: Still had 5 samples that came up positive. Will re-DNase treat these samples and then re-qPCR them.
Performed qPCR using q18s primers on DNased RNA (1:100 dilution to match final concentration of template after making cDNA). qPCR set up and plate layout are here.
gDNA dilutions were used as positive controls. gDNA = BB11 (0.49ug/uL) from 20090519. Used 5uL of 1:10, 1:100 and 1:000 dilutions.
Results: All samples, including positive controls, came up negative! Realized that I accidentally used the EF1 primers which will NOT amplify gDNA. And, to top it off, this was BEFORE going to seminar (i.e. before having any beers).
Performed qPCR using q18s primers on DNased RNA (1:100 dilution to match final concentration of template after making cDNA). qPCR set up and plate layout are here.
gDNA dilutions were used as positive controls. gDNA = BB11 (0.49ug/uL) from 20090519. Used 5uL of 1:10, 1:100 and 1:000 dilutions.
Results: gDNA dilutions look good. However, some samples are definitely coming up before the 40 cycle mark. Will re-DNase treat these.
Master mix containing Gigas gDNA will be used to verify that the recalibration did work. qPCR setup/plate layout is here. I’ve made a master mix using Cg_HSP70_F/R primers designed by Mac. gDNA is BB #12 (0.445ug/uL) from 20090519. The gDNA will be added to the master mix.
Results: The results are a bit disconcerting, as this run shows virtually the exact same pattern in fluorescence detection as that on 20090722, despite using a different set of gigas gDNA. Below is a set of graphs comparing Column 1 Ct values of the two tests from 20090722 and today:
Clearly, both runs exhibit virtually the same pattern of relative Ct values to each other in each respective run. Not cool.
According to the Bio-Rad rep that analyzed the calibration test on 20090824, recalibration might be a good idea. As such, we’ll do that. Old calibration file was found and backed up prior to proceeding, as generation of a new calibration file would replace the old one.
Results: The Bio-Rad rep (Carl Fisher) has responded and said that the range of fluorescence is within the expected 2-fold difference and looks fine! Finally!
Based on recs from Bio-Rad rep (Carl Fisher), will repeat Opticon 2 calibration (see 20090813) test according to Opticon manual. Then, will rotate plate 180 degrees and repeat test and upload data to Bio-Rad server for analysis and evaluation.
Results: According to the Bio-Rad rep, he thinks recalibration is in order. He pointed out that we’re seeing a 3-fold spread in fluorescence, which is outside of the “tolerable” range (which is 2-fold). Will recalibrate.
Ran a qPCR on Rick’s Lake Trout DNA from 4/28/2009 using primers in Carita’s CMA01 Primer Plate (Excel file). DNA was pooled (2uL from each sample), spec’d and diluted to 10ng/uL. qPCR set up is here. Plate layout matches Carita’s primer plate layout. Since we’re just looking for positive/negative samples, I ran this on the Opticon 2 despite the recent “problems” we’ve been having with it. Cycling params used with the 2x HRM M.M. are as follows:
95C – 10mins
40 cycles of:
95C – 10s
60C – 15s
72C – 25s
Results: Overall, looks good. Negative control is clean. Based on signal strength and clean, tight melting curves, the following primer sets (location in CMA01 primer/qPCR plates) will be used for HRM analysis tomorrow: C11, D3, E12, D3. Actually, never mind. Steven’s sent me contig info for the lake trout so we’ll just order and test primers that are more likely to produce good data instead of worrying about the salmonid primers.
Received new FAM calibration reagent. It comes pre-prepared in a 1x PCR buffer (0.3uM), however there is only enough for a single plate (use 50uL/well). Will run plate in Opticon 2. Run according to the calibration protocol in the Opticon 2 manual (p. 10-4).
Results:
Well, there’s actually a signal this time as opposed to the run on 20090806. However, it’s pretty clear that the signals aren’t even close to being uniform. Or, as the manual says “tightly clustered lines”. I’m also not sure why the fluorescence decreases over time, although it could simply be degradation of the fluorophore after being hit with light. I’ve sent the results to Bio-Rad for help interpreting them.
Due to some weird anomolies seen during my previous qPCRs with the H.crach RNA/cDNA samples (positive controls produced good fluorescence when tested in Column 1 of the Opticon, but consistently failed to produce virtually any fluorescence when in Column 6 of the Opticon), I’ve decided to check the Opticon’s fluorescence detection.
qPCR setup/plate layout is here. I’ve made a master mix using Cg_HSP70_F/R primers designed by Mac. gDNA is BB #11 (0.49ug/uL) from 20090519. The gDNA will be added to the master mix. All wells will be tested.
Results: Unfortunately, this does not look as tight as it should. The Cts range from 23.9 – 26.6 (Cts from Opticon 2 with default threshold settings). This is nearly a 3 cycle difference which represents a nearly 10-fold difference in “expression.”
Now that we have a solid positive control, I’ll use the H.crach_h-1fg_intron primers to check the existing cDNA and DNased RNA. qPCR plate layout/set up is here. Anneal temp 50C.
Results: Looks like all the cDNA and DNased RNA are negative ! Finally! Will make cDNA from the DNased RNA.
