As for the "how to prepare samples and run the gel?" part of your question: I have run some gels for in-gel fluorescence, my protein was sfGFP-tagged, with a good result. Sample preparation is essentially the same as for denaturing gels, but as you pointed out you should not boil the samples - it kills GFP fluorescence completely. You may try heating it up for 5 min at 50-60 oC - in my case this sample preparation gave a good result, but it wasn't a membrane protein. After running the gel as usual, at room temperature, you don't stain it, only wash with water, and it is ready for in-gel fluorescence detection.

For the second part of your question, I have not tried it with this particular machine, but since the excitation wavelengths are the same I would give it a try, with an appropriate filter.

Good luck!

Generally for PAGE to give nice, crisp bands you have to run it under de-naturing conditions. If you run the gel under conditions where the proteins are still folded they tend to aggregate in the wells. It is recommended to run native gels on ice.

If you're set on trying it, I'd recommend using a 3-12% gradient gel to give the protein the best chance of making it out of the wells. Just expect to get a smear rather than a crisp band, and the MW markers will be pretty much useless.

Hi Samuel,

As Chris pointed out, it is tricky to run a gel without denaturing conditions. Even more so if you have a membrane protein. If you are using the Biorad system, send their customerservice this question, I think they can help.

For the 395 fluorescence: it is the right wave length for your GFP, but you also have to check that you have the richt filter for the emitted light (not sure what that is for uvGFP).

Maybe you should consider to run the gel under denaturing conditions and rather use an antibody for your GFP tag? That saves you a lot of hassel.

Good luck!

As for the "how to prepare samples and run the gel?" part of your question: I have run some gels for in-gel fluorescence, my protein was sfGFP-tagged, with a good result. Sample preparation is essentially the same as for denaturing gels, but as you pointed out you should not boil the samples - it kills GFP fluorescence completely. You may try heating it up for 5 min at 50-60 oC - in my case this sample preparation gave a good result, but it wasn't a membrane protein. After running the gel as usual, at room temperature, you don't stain it, only wash with water, and it is ready for in-gel fluorescence detection.

For the second part of your question, I have not tried it with this particular machine, but since the excitation wavelengths are the same I would give it a try, with an appropriate filter.

Good luck!

Thanks for your answers everyone. Just to clarify, I will be denaturing the samples as SDS will be in my sample buffer like normal, I just won't be boiling them or will be at the most heating them at a low temperature like Dominika suggests. The idea is to use the same gel to hopefully get a total protein (analogous to commassie) from the stain free detection feature of the gels, the in-gel GFP fluorescence and also a WB against the twin-Strep tag.

I am also going to be trying to run these on blue native PAGE with in-gel GFP fluorescence to verify the quaternary structure and homogenicity using the "light blue" version as it's termed with Thermo's native gels for the cathode buffer. In theory it is workable according the the instructions as it gives enough charge to let the samples enter the lanes, but is light enough to allow fluorescence to be detected. I'm having to do this as unfortunately we don't have access to fluorescence size exclusion chromatography, so I'm exploring alternatives like using Native PAGE fluorescence.

I have done lots of work with GFP-tagged membrane proteins. One thing I learned was that certain gels work better for this. In particular Tris-glycine gels worked really well, as I think GFP seems to unfold/lose fluorescence in other gels (possibly to do with the pH). If you are trying to load whole cells you may have some issues with gloopy samples that refuse to load on the gel. What I have done to overcome this is solubilise samples in DDM or another mild detergent, then ultracentrifuge the sample and then load the supernatant on the gel (in loading dye with no heating at all).

Thanks David, I'm going to be extracting the membranes by ultra centrifugation first. I'll likely at first just be running the total membrane fraction to confirm expression etc, but I will then move on to solubilising with detergents to see the soluble and insoluble fractions.

Hi, in case anyone wonders how this evntually panned out, I can tell you that I was able to get in-gel GFP fluoresence to work eventually. Essentially you just need to make samples up as normal in loading buffer, but then add protease inhibitors and just leave them at room temp for 20 to 30 mins with occasional light vortexing. Pre-chill the SDS-PAGE running buffer, but other than that run the gel as normal (I used 200V for 40 mins rather than our usual 300V for 17 mins though to limit gel heating). I was able to occasionally see a faint signal with Bio-Rad Geldoc EZ and the blue plate, but eventually found we had an Amersham Typhoon imager elsewhere in the building, which was able to show a robust signal with its 488 nm laser line.

One word of warning, initially I thought I could see expression of my protein on the in-gel fluorescence, but mass-spec protein ID could only find the GFPuv fusion tag. It transpired that the GFPuv was dimerising under the high concentration it was at, so I would recommend putting a monomising mutation (such as A206K for GFP) in the GFP tag to prevent this.

Hi,

I am trying to do the in-gel fluoescence for my fusion protein a small membrane protein and EYFP tag in the C-terminus. My general protocol is:

1, add 4X SDS- sample buffer with protein

2, heat at 50C for 20 min

3, add to sds-page gel 5-15% gradient gel and run at 120 V for 1 hr

I wonder whether I can still see the YFP fluorescence in the gel? Thank you.