Native-PAGE. Could anyone have a protocol or some advice to give me?

I want to analyze proteins in transfected HeLa cells using native gel electrophoresis.

Since the samples has to be prepared in a non-denaturing condition, could someone suggest a good recipe for cell lysis? I also heard that one must dialyze the lysates with a suitable buffer before sample preparation. How effective is it?

I've recently performed a western blot for my Native PAGE. My protein of interest is around 120kDA however it formed oligomers with size up to 250kDA. I used Precast Gel Any KD when running my NATIVE PAGE (I already performed a native page then coomasie stained it and I indeed observed tha band. For my western blot protocol, I searched what method to follow and this were the steps that I followed. I prepared cold Protein Transfer Buffer (1X PTB: 25mM Tris Base, 195mM glycine, 0.05% (w/v) SDS). I did not add methanol to my transfer buffer . I soaked the following the following in their respective solutions for 30 mins: PAGE gel in cold PTB filter paper in cold PTB and scrubbing pads in cold ddh2O. The membrane that I used is 0.2um PVDF and I pre-wet it with the following solutions in order: 100% methanol for 15 seconds, ddH2O for 5 minutes and cold PTB for 10 minutes. After assembling the cassete, I place it in the western blot tank and was slowly filled with cold PTB since a magnetic stirrer is placed at the bottom. I placed cooling pack inside and along the tank. I run the apparatus at 100V for 1hr and 30 minutes.

After the run, I considered my blot successful since the dye from the ladder that i used, i.e dual color precision plus, successfully transfered to my membrane. I rinsed the membrane with sddH2O and store it overnight at blocking solution (5% (w/v) whey) at 4degC. I wash the membrane with TBST (0.1% v/v Triton-X in TBS) for 10 minutes with shaking at room temperature and repeated this three times. I transferred my membrane to my primary antibody solution (1:2000, prepared with blocking solution) and incubated it at 4oC overnight. I then washed the membrane with TBST the same manner as mentioned above. I then transferred my membrane to secondary antibody (1:5000) and incubated it at 1hr, RT with rocking. I then washed it again with TBST 3 times then incubated it in Alkalin Phosphatase Buffer for 10 mins. After which i detected the proteins in the membrane with NBT/BCIP.

My problem was that I was able to detect bands however not in the correct position. To be specific, the bands I detected were found in the area when you load your proteins in your gel. I was wondering if the method I used is really used when blotting proteins in their native state.

I am setting up experiments to look at redox interactions between Ref-1 protein and transcription factors upon different treatment conditions. I am looking for western blot protocol for native gel condition.

Please correct me if you have tried any other method or reagents.

1) 12% separating gel (5ml): 30% acrylamide: 4ml, 0.375 M Tris-HCl (pH=8.8): 5.89 ml, 10% APS = 100 ul, TEMED= 10ul

2) Stacking gel: 0.375 M Tris HCl (pH=8.8)= 4.275 ml, 30% acrylamide = 0.67 ml, 10% APS= 0.05 ml, TEMED= 5ul

3) Running buffer (1X)= 25mM Tris, 192mM glycine, pH=8.3

4) 2X sample buffer: 62.5mM Tris-HCl (pH=6.8), 25% glycerol, 1% bromophenol blue

5) Not going to heat the sample

6) Running gel at 120 V

I'm using total proteins treated with desired drug to see the expressions of Trx. But I also read that with acidic and basic proteins due to different charges, they do have different running conditions. I'm not sure if I need to add any other buffer to running buffer to ensure movement of charged proteins. And also we need a different marker than SDS/PAGE??

I'll appreciate you if you can suggest me and correct me for this experiment.

Few things about blue native page I couldn't find at all. First, why do we have separate cathode and anode buffers (in traditional SDS PAGE tris-glycine is the only buffer) in BN PAGE?. Moreover, there's plenty of coomassie g250 in the loading buffer, then why do we need 0.02% coomassie in the cathode running buffer? Can the gel be run just with tris glycine buffer on traditional tris-HCl gel (stacking gel pH 6.8, resolving gel pH 8.8, running buffer pH 8.3) if we add coomassie g 250 in the loading buffer (the whole system in this case will be devoid of any SDS).

Another question comes to me is about the molecular weight ladders. If we coat the normal standards (commercially available, meant to be run on SDS PAGE) with coomassie G250, they should run across the gel. In principle, these standards should give approximate estimates of molecular weights in BN PAGE. Kindly correct my thoughts on this issue.

Hi Folks,

I purchased the Novex Native PAGE, which relies on Coomassie G250, and had a successful electrophoresis where I've seen the ladder separate.

However, for some reason the transfer continues to be unsuccessful. I am using PVDF membrane with the BioRad Transblot SD Semi-Dry transfer cell.  I have tried the following conditions

1.) Transfer in Tris Glycine, 1% SDS, 20% methanol, ph8, at15V- 30 mins to 2 hours

2.) Transfer in Tris Glycine, 1% SDS, 20% methanol, ph8, at 25V- 30 mins to2 hours

3.)Transfer in Tris glycine, 1% SDS, 10% methanol, ph 9.4, at 25V- 45 mins to1 hour- and have a membrane which blue stained but the ladder has not migrated onto the membrane

4.) Transfer in Tris glycine, 1% SDS, 10% methanol, ph 9.4, onto pvdf membrane (on both sides of the gel) at 25V- 1 hour

I am not even getting the ladder to shift.  I wonder whether there is something in the chemistry of the Novex gels which requires a special type of transfer buffer?  If anyone has any experience with Western Blot following Novex Native Page, I'd appreciate your advice.  

Thanks!

Hi,

I have native plasma membrane extracts from mouse brain which I am running on BN-PAGE gels.

We have tested several concentrations of DDM and Digitonin, and 1% DDM + 1% Digitonin in 1X Native PAGE sample buffer gave best results.

I am using 4-16% Novex Bis-Tris NativePAGE gels. They are run at 150V and I replace the dark blue cathode buffer with light blue cathode buffer 1/3 through the run.

I add 1uL of Coomassie G-250 5% sample additive to each sample.

My protein of interest is a connexin (which forms homohexamers around 500kDa and gap junctions around 1000kDa) with a high pI (~9). Heterohexamers can be of varying sizes.

I am using an aquaporin as a "control" (pI 7).

I have attached an image of my problem (membrane after blotting, cut and developed with Cx and AQP antibody).

As you see in wells 5-7:

- I have strong staining for my connexin, especially in the wells.

- I have bands in the right range (for hexamer and gap junction), however, they are completely streaked (could be explained by heterohexamers which do occur).

- The aquaporin runs fine and shows expected bands at right size.

Things I have tried:

- Running at 120V for 10 min before increasing voltage to 150V for the remainder of the run.

- Adding Triton X-100 to the sample (0.01%)

- Adding double amount of loading dye (With Coomassie).

- We have tested running 1.25 to 20ug of protein - samples still get stuck in well.

- I have also tried TGX gels with Tris-Glycine buffer (no SDS) and "inverting the electrodes" which did not seem to work (no protein on gel or membrane).

My biggest problem is that most of the connexin signal is stuck in the well.

Does anyone have an tips for improving the migration of connexin in the gel?

Best,

Nadia Skauli

I want to get isozymes from pea leaf extract. I need a complete protocol for native- PAGE as well as a protocol for catalases, peroxidases and SOD.

I want to run a native PAGE gel of my purified protein as we suspect that it may be forming oligomers. The protein I am working with is a 50kDa soluble protein with an acidic pI.  I recently ran a clear native page by essentially just leaving SDS and DTT out of the usual buffers. This approach seemed to work quite well (in so far as the protein formed a band in the gel), but I am now concerned that as this approach relies on the intrinsic charge of the protein, it is not an accurate way of estimating the molecular weight. I was thinking of trying a blue native PAGE but it seems that this was developed for membrane proteins and some of the protocols I have found are quite complicated. Can anyone point me towards a good protocol for blue native page? Another thing I was wondering is that most of the native PAGE protocols seem to use gradient gels rather than uniform gels, is this necessary?