A couple final notes: It’s good to keep an eye on the dye front from the start (not just when you think it should be close to done) - this lets you see if there are any problems. What you want to see is that the dye front is moving evenly (all the lanes at the same rate). If you see one side running higher than the other, this is sometimes because the buffer level is too low (is your cassette leaking? If so, once your samples are in the gel (cleared past the wells) you can take it out and fix it) or you might see a smiling effect where the edges run slower than the center due to uneven heating. If you see that, turn down the voltage. The sooner you can detect a problem, the sooner you can try to fix it. more here: blog: bit.ly/gel_smile ; CZcams: czcams.com/video/a8geIlRuwrc/video.html In terms of the dyes in nucleic acid sample loading buffers, you often see 2 or 3. This allows you to estimate where various size fragments are in the gel during the run. The dyes run differently in different gel types and percentages, so check a dye migration chart to see what size fragments the dyes will run alongside in the gel you’re using. If they overlap with yours they could make it hard to see your bands so you may want to use another dye. You may need to make one yourself but it’s super pretty to do so! Here are a few I use: SDS-PAGE (denaturing protein): The recipe we use in our lab for 100mL of 6X is: - 5.91 g Tris-HCl pH 6.8 - 6 g SDS - 48 mL glycerol - 30 mg bromophenol blue You can store this at room temperature (RT). Then, before using, add 90 μL of β-mercaptoethanol to 910 μL of this mix - you can freeze this aliquot and it will be good for a while. Note: many protocols use higher [BME] than this (5% is common, but our recipe uses 1.5…) working concentrations: - 80 mM Tris-HCl pH 6.8 - 1% SDS (w/v) - 8% glycerol (v/v) - 0.0005% bromophenol blue (w/v) non-denaturing nucleic acid: * 6X Non-denaturing sample loading buffer (6X NSLB): 10mM Tris pH 8; 1mM EDTA; 15% w/v Ficoll 400; 0.25% w/v Bromophenol Blue denaturing nucleic acid: * 2X Denaturing Sample Loading Buffer (2X DSLB): 98% v/v formamide; 10mM EDTA; 300μg/mL bromophenol blue * 2X Xylene cyanol formamide loading dye: 95% formamide, 5mM EDTA (pH 8.0), 0.05% (w/v) xylene cyanol for more on SDS-PAGE loading buffers: blog: bit.ly/sds_loading_dyes ; CZcams: czcams.com/video/IDENwOYpGRs/video.html more on denaturing vs. reducing bit.ly/denaturing_vs_reducing ; CZcams: czcams.com/video/YoMH7CmYHwA/video.html more on SDS-PAGE: bit.ly/sdspageruler & czcams.com/video/FgXlPAKVSGY/video.html more on native protein-PAGE: bit.ly/nativepageoverview & czcams.com/video/QRL5cfJNwm0/video.html more on nucleic acid PAGE: bit.ly/ureapage & czcams.com/video/MHJqnur6yqk/video.html more about agarose gel electrophoresis: bit.ly/agarosegelelectrophoresis & bit.ly/agarosegelcompare & czcams.com/video/vbuxf3rcMxg/video.html more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
A couple final notes:
It’s good to keep an eye on the dye front from the start (not just when you think it should be close to done) - this lets you see if there are any problems. What you want to see is that the dye front is moving evenly (all the lanes at the same rate). If you see one side running higher than the other, this is sometimes because the buffer level is too low (is your cassette leaking? If so, once your samples are in the gel (cleared past the wells) you can take it out and fix it) or you might see a smiling effect where the edges run slower than the center due to uneven heating. If you see that, turn down the voltage. The sooner you can detect a problem, the sooner you can try to fix it. more here: blog: bit.ly/gel_smile ; CZcams: czcams.com/video/a8geIlRuwrc/video.html
In terms of the dyes in nucleic acid sample loading buffers, you often see 2 or 3. This allows you to estimate where various size fragments are in the gel during the run. The dyes run differently in different gel types and percentages, so check a dye migration chart to see what size fragments the dyes will run alongside in the gel you’re using. If they overlap with yours they could make it hard to see your bands so you may want to use another dye. You may need to make one yourself but it’s super pretty to do so!
Here are a few I use:
SDS-PAGE (denaturing protein):
The recipe we use in our lab for 100mL of 6X is:
- 5.91 g Tris-HCl pH 6.8
- 6 g SDS
- 48 mL glycerol
- 30 mg bromophenol blue
You can store this at room temperature (RT). Then, before using, add 90 μL of β-mercaptoethanol to 910 μL of this mix - you can freeze this aliquot and it will be good for a while. Note: many protocols use higher [BME] than this (5% is common, but our recipe uses 1.5…)
working concentrations:
- 80 mM Tris-HCl pH 6.8
- 1% SDS (w/v)
- 8% glycerol (v/v)
- 0.0005% bromophenol blue (w/v)
non-denaturing nucleic acid:
* 6X Non-denaturing sample loading buffer (6X NSLB): 10mM Tris pH 8; 1mM EDTA; 15% w/v Ficoll 400; 0.25% w/v Bromophenol Blue
denaturing nucleic acid:
* 2X Denaturing Sample Loading Buffer (2X DSLB): 98% v/v formamide; 10mM EDTA; 300μg/mL bromophenol blue
* 2X Xylene cyanol formamide loading dye: 95% formamide, 5mM EDTA (pH 8.0), 0.05% (w/v) xylene cyanol
for more on SDS-PAGE loading buffers: blog: bit.ly/sds_loading_dyes ; CZcams: czcams.com/video/IDENwOYpGRs/video.html
more on denaturing vs. reducing bit.ly/denaturing_vs_reducing ; CZcams: czcams.com/video/YoMH7CmYHwA/video.html
more on SDS-PAGE: bit.ly/sdspageruler & czcams.com/video/FgXlPAKVSGY/video.html
more on native protein-PAGE: bit.ly/nativepageoverview & czcams.com/video/QRL5cfJNwm0/video.html
more on nucleic acid PAGE: bit.ly/ureapage & czcams.com/video/MHJqnur6yqk/video.html
more about agarose gel electrophoresis: bit.ly/agarosegelelectrophoresis & bit.ly/agarosegelcompare & czcams.com/video/vbuxf3rcMxg/video.html
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com