In terms of option #3… it’s common that after you separate complexes you want to find out what’s in those complexes. To do this you can separate the proteins within the complexes. So basically you cut a lane out of your native PAGE gel (which, remember contains the complexes stuck in place). And you denature those proteins so that the complexes will come apart and the proteins will be able to travel independently from one another, just like in SDS-PAGE. But unlike when you normally run SDS-PAGE, you don’t have liquid samples you can load into wells. Instead, you take that gel slice & And you rotate it 90 degrees and put it on top of an SDS-PAGE gel. (or actually pour the gel kinda around it so it gets stuck in there during the polymerization).
Instead of the horizontal position being determined by what order you loaded wells in, the horizontal position here corresponds to where the protein (in its complexed form) ran on the native PAGE. So, when you run that SDS-PAGE gel, the horizontal position will correspond to the complex it was in (i.e. proteins in the same “column” will have come from the same complex but there won’t be nice clear separations between lanes because you don’t have lanes!).
The vertical position (how far the protein travels down the gel) will correspond to the size (length here and no longer shape as well) just like you’re used to with regular SDS-PAGE.
Now you can stain the gel for proteins like normal. But, depending on what your starting sample was, you might see something pretty weird looking with a bunch of spots and smears (you won’t get nice bands because you don’t have lanes! So how clear it looks will depend in large part on how the native run went.
If you were to draw a line vertically through the gel, the proteins along the line (likely) came from the same complex, so you can then either know what they are based on their size because you started with a pure mix of known proteins or you can probe for them with antibodies specific for proteins you suspect they are. Or you can do things like cut out the spots and send them for mass spectrometry (mass spec) analysis to see their identity (even if you have no idea what they are).
In terms of option #1 (staining directly after native PAGE)… you can “fix” the gel to precipitate the proteins in place so they won’t diffuse out (wander randomly from where they stopped) before staining or you can stain without fixing and then cut out the bands and get the protein out to identify (with methods such as electrocution or diffusion, none of which I have ever done but there are thorough instructions in the Nature Protocols article I link to at the end).
*Note: some proteins form multimers through disulfide bridges (cystine cross-links). These are covalent bonds as opposed to just attractions. So they can’t be broken up with just SDS & heat. Instead, you need to add a reducing agent such as BME (beta mercaptoethanol) to split them up. This will reduce the cystines to cysteines, going from |-S-S-| to |-SH HS-|. If you suspect you have multimerization through disulfide bridges, you can run SDS-PAGE under nonreducing (no BME, DTT, etc.) and reducing conditions to see if you see a difference. (and you can do this without having to do the native PAGE step as well).
note: BN-PAGE is commonly used for membrane proteins, so if you look up protocols they often talk about solubilizing membranes and stuff and include a mild detergent cuz those proteins are used to being surrounded by lipids.
Blue Native PAGE (BN-PAGE) - here are the notes I took for my lab book, but there are better protocols linked below. There are lots of different protocols and you can use different gel types and stuff but this is how I (just) learned. I’m planning to post a video showing me doing it too. Because it’s so cool looking!
I used the Biorad 4-20% Tris-Glycine TGX gels
Prepare your samples and instead of SDS loading buffer, use glycerol to final concentration of 20% (I started with an 80% glycerol stock because it’s much easier to pipet than pure glycerol! So, 80%’s my 4X loading buffer in a way… and the glycerol keeps the samples from running away.
DO NOT BOIL! Boil in SDS-PAGE to denature, but here you do not want to denature
Set up your cassette in the running module
pour no-dye buffer (1X Tris-Glycine) into inner chamber (cathode buffer)
load wells
load 20% glycerol in empty lanes to get consistent flow
add dye to inner chamber - pipet 2mL of 100X concentrate into the bottom of the chamber, then pipet up and down with a large pipet to mix
run for ~20 min at 100V - proteins should enter gel
remove inner dye-containing buffer & replace with fresh no-dye buffer
continue running until dye front comes out - can also increase V to 200
some good posts/articles:
Original article: Hermann Schägger, Gebhard von Jagow, Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form, Analytical Biochemistry, 1991, doi.org/10.1016/0003-2697(91)90094-A Detailed protocol: Wittig, I., Braun, HP. & Schägger, H. Blue native PAGE. Nat Protoc 1, 418-428 (2006). doi.org/10.1038/nprot.2006.62 Practical look: Bench Tips: Blue Native-PAGE by Eric Torres: blog.benchsci.com/bench-tips-blue-native-page
more on Coomassie staining here: bit.ly/cbbgelstaining
Here’s more about isoelectric points: bit.ly/isoelectricpoint & ua-cam.com/video/CLgzYBm_ymk/v-deo.html
Here’s more about native & blue-native PAGE: see bit.ly/nativepageoverview ; UA-cam: ua-cam.com/video/0w-Y33FmuWs/v-deo.html & ua-cam.com/video/ID4rCtcR8TE/v-deo.html
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
Thank you so much madam you deserve more subscribers than you would ever imagine. Ma'am I am a student who just finished a levels in Sri Lanka.I work in a lab now for volunteer purposes.My hand shakes while I am doing pippeting.is it a normal thing or should I be worried. Thank you so much madam
Thank you but I'm not in it for subscribers & I don't monetize my work, I just really want to help people so I'm glad I am! It's totally normal to be shaky starting out, but over time you'll get much steadier. It just takes practice and building up those weird muscles!
I come from your instagram post. I learned a lot from your videos. Thank you so so much
So happy I could help!
In terms of option #3… it’s common that after you separate complexes you want to find out what’s in those complexes. To do this you can separate the proteins within the complexes. So basically you cut a lane out of your native PAGE gel (which, remember contains the complexes stuck in place). And you denature those proteins so that the complexes will come apart and the proteins will be able to travel independently from one another, just like in SDS-PAGE. But unlike when you normally run SDS-PAGE, you don’t have liquid samples you can load into wells. Instead, you take that gel slice & And you rotate it 90 degrees and put it on top of an SDS-PAGE gel. (or actually pour the gel kinda around it so it gets stuck in there during the polymerization).
Instead of the horizontal position being determined by what order you loaded wells in, the horizontal position here corresponds to where the protein (in its complexed form) ran on the native PAGE. So, when you run that SDS-PAGE gel, the horizontal position will correspond to the complex it was in (i.e. proteins in the same “column” will have come from the same complex but there won’t be nice clear separations between lanes because you don’t have lanes!).
The vertical position (how far the protein travels down the gel) will correspond to the size (length here and no longer shape as well) just like you’re used to with regular SDS-PAGE.
Now you can stain the gel for proteins like normal. But, depending on what your starting sample was, you might see something pretty weird looking with a bunch of spots and smears (you won’t get nice bands because you don’t have lanes! So how clear it looks will depend in large part on how the native run went.
If you were to draw a line vertically through the gel, the proteins along the line (likely) came from the same complex, so you can then either know what they are based on their size because you started with a pure mix of known proteins or you can probe for them with antibodies specific for proteins you suspect they are. Or you can do things like cut out the spots and send them for mass spectrometry (mass spec) analysis to see their identity (even if you have no idea what they are).
In terms of option #1 (staining directly after native PAGE)… you can “fix” the gel to precipitate the proteins in place so they won’t diffuse out (wander randomly from where they stopped) before staining or you can stain without fixing and then cut out the bands and get the protein out to identify (with methods such as electrocution or diffusion, none of which I have ever done but there are thorough instructions in the Nature Protocols article I link to at the end).
*Note: some proteins form multimers through disulfide bridges (cystine cross-links). These are covalent bonds as opposed to just attractions. So they can’t be broken up with just SDS & heat. Instead, you need to add a reducing agent such as BME (beta mercaptoethanol) to split them up. This will reduce the cystines to cysteines, going from |-S-S-| to |-SH HS-|. If you suspect you have multimerization through disulfide bridges, you can run SDS-PAGE under nonreducing (no BME, DTT, etc.) and reducing conditions to see if you see a difference. (and you can do this without having to do the native PAGE step as well).
note: BN-PAGE is commonly used for membrane proteins, so if you look up protocols they often talk about solubilizing membranes and stuff and include a mild detergent cuz those proteins are used to being surrounded by lipids.
Blue Native PAGE (BN-PAGE) - here are the notes I took for my lab book, but there are better protocols linked below. There are lots of different protocols and you can use different gel types and stuff but this is how I (just) learned. I’m planning to post a video showing me doing it too. Because it’s so cool looking!
I used the Biorad 4-20% Tris-Glycine TGX gels
Prepare your samples and instead of SDS loading buffer, use glycerol to final concentration of 20% (I started with an 80% glycerol stock because it’s much easier to pipet than pure glycerol! So, 80%’s my 4X loading buffer in a way… and the glycerol keeps the samples from running away.
DO NOT BOIL! Boil in SDS-PAGE to denature, but here you do not want to denature
Set up your cassette in the running module
pour no-dye buffer (1X Tris-Glycine) into inner chamber (cathode buffer)
load wells
load 20% glycerol in empty lanes to get consistent flow
add dye to inner chamber - pipet 2mL of 100X concentrate into the bottom of the chamber, then pipet up and down with a large pipet to mix
run for ~20 min at 100V - proteins should enter gel
remove inner dye-containing buffer & replace with fresh no-dye buffer
continue running until dye front comes out - can also increase V to 200
some good posts/articles:
Original article: Hermann Schägger, Gebhard von Jagow, Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form, Analytical Biochemistry, 1991, doi.org/10.1016/0003-2697(91)90094-A
Detailed protocol: Wittig, I., Braun, HP. & Schägger, H. Blue native PAGE. Nat Protoc 1, 418-428 (2006). doi.org/10.1038/nprot.2006.62
Practical look: Bench Tips: Blue Native-PAGE by Eric Torres: blog.benchsci.com/bench-tips-blue-native-page
more on Coomassie staining here: bit.ly/cbbgelstaining
Here’s more about isoelectric points: bit.ly/isoelectricpoint & ua-cam.com/video/CLgzYBm_ymk/v-deo.html
Here’s more about native & blue-native PAGE: see bit.ly/nativepageoverview ; UA-cam: ua-cam.com/video/0w-Y33FmuWs/v-deo.html & ua-cam.com/video/ID4rCtcR8TE/v-deo.html
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
#scicomm #biochemistry #molecularbiology #biology #sciencelife #science #realtimechem
Thank you so much madam you deserve more subscribers than you would ever imagine. Ma'am I am a student who just finished a levels in Sri Lanka.I work in a lab now for volunteer purposes.My hand shakes while I am doing pippeting.is it a normal thing or should I be worried.
Thank you so much madam
Thank you but I'm not in it for subscribers & I don't monetize my work, I just really want to help people so I'm glad I am! It's totally normal to be shaky starting out, but over time you'll get much steadier. It just takes practice and building up those weird muscles!
Nice slides. Do you create the figures yourself?
Thanks! Yes I do
Thank you!! ❣