Daniel:Notebook/HiResChrPaint/2013-7-11: Difference between revisions
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Notes: *denotes beginning of a new dye tube; X denotes the end | Notes: *denotes beginning of a new dye tube; X denotes the end | ||
==AmpR11 and AmpR12== | |||
Trying again to make sense of this dye procedure. The above table suggests that the last few drops of | |||
the dye per reaction tube always seem to be better than the first. It might just be a problem of mixing. | |||
So I'm going to try again but this time mix it really well (vortex and pipette) before I add the dye. | |||
1. Add 5 uL DMSO to vial of dye; vortex vigorously and pipette thoroughly. | |||
2. Add 1 uL DNA in 19 uL labeling buffer (Component C) | |||
3. Denature DNA for 5 minutes at 95C; snap cool on ice | |||
4. Add 1 uL dye to sample; add 4 uL labeling buffer | |||
5. Incubate for 15 minutes at 80C, snap cool on ice |
Revision as of 22:39, 11 July 2013
S2 Probe Prep (Started 5/29/2013)
ULYSIS Dye Coupling (Test 6)
Should be the last test. I'm going to test at 15 minutes and 20 minutes incubation time, 1 uL each. If the results go well this morning I will be labeling the V6S2 set this afternoon.
- Following ULYSIS protocol
1. Add 1 uL DNA in 19 uL labeling buffer (Component C) 2. Denature DNA for 5 minutes at 95C; snap cool on ice 3. Add 1 uL dye to sample; add 4 uL labeling buffer 4. Incubate for 15 minutes (AmpR9) or 20 minutes (AmpR10) at 80C, snap cool on ice 5. Purify using Centri-Sep Column
Results
ng/uL DNA | pmol/uL dye | ug DNA in 20 uL | pmol/uL DNA | dye/probe | dye/100 bp | Base:Dye Ratio (:1) | |
AmpR9 | 64.6 | 0.6 | 1.3 | 3.9 | 0.2 | 0.3 | 326 |
AmpR10 | 71.1 | 0.4 | 1.4 | 4.3 | 0.1 | 0.2 | 539 |
It would seem that I can always tell beforehand whether a yield will be good or bad, by the color of the solution post incubation. If the solution has a faint green color it will be a good yield. If the solution remains clear it will be a bad yield. Therefore I'm going to ignore columns for future trials (yes, it looks like I need more troubleshooting). That will save on time and $$, and hopefully I'll get to the bottom of this before long.
Review of AmpR Experiments so Far
Sample | uL dye Added | Incubation Time (min) | pmol/uL DNA | pmol/uL dye | dye/probe | base:dye ratio | |
AmpR1 | 1 | 15 | 3.5 | 1.2 | 0.341 | 147 | |
AmpR2 | 1 | 15 | 3.1 | 1.2 | 0.384 | 130 | |
X | AmpR3 | 2 | 15 | 3.4 | 1.9 | 0.552 | 91 |
*X | AmpR4 | 5 | 15 | 1.9 | 1.4 | 0.7 | 69 |
* | AmpR5 | 2 | 30 | 3.3 | 0.5 | 0.2 | 330 |
AmpR6 | 2 | 60 | 3.8 | 0.4 | 0.1 | 474 | |
X | AmpR7 | <1 | 20 | 2.6 | 2.3 | 0.9 | 56 |
* | AmpR8 | 1 | 20 | 3.6 | 0.7 | 0.2 | 259 |
AmpR9 | 1 | 15 | 3.9 | 0.6 | 0.2 | 326 | |
AmpR10 | 1 | 20 | 4.3 | 0.4 | 0.1 | 539 | |
BLANK | 0 | 15 | 4 | 0.2 | 0 | 1009 |
Notes: *denotes beginning of a new dye tube; X denotes the end
AmpR11 and AmpR12
Trying again to make sense of this dye procedure. The above table suggests that the last few drops of the dye per reaction tube always seem to be better than the first. It might just be a problem of mixing. So I'm going to try again but this time mix it really well (vortex and pipette) before I add the dye.
1. Add 5 uL DMSO to vial of dye; vortex vigorously and pipette thoroughly. 2. Add 1 uL DNA in 19 uL labeling buffer (Component C) 3. Denature DNA for 5 minutes at 95C; snap cool on ice 4. Add 1 uL dye to sample; add 4 uL labeling buffer 5. Incubate for 15 minutes at 80C, snap cool on ice