Sample Prep in SDS-PAGE

[pseizure]

So I've attempted a myoglobin protein extraction on some muscle tissue following a protocol. Now I am moving on to the SDS-PAGE part of the experiment, but I am getting confused with protein concentration and how much to load.

According to some protocols I have read Myoglobin (mM) = [A525/(7.6 mM-1cm-1x1cm]

So I took my absorbance at 525 and got 0.060

Using the formula above I calculated 0.00789 mM

So how do I know if I need to concentrate or dilute my sample for SDS-PAGE? I think I read somewhere that 0.5 ug of protein in 5 ul was good but I am very confused now.

[SciB]

Hi,

When we run proteins on SDS-PAGE for western blotting we always use about 100 ug, but that is total protein from a cell extract, whereas your sample is purified protein. The molecular mass of bovine myoglobin is about 17,000 Da, so your 0.00789 mM solution would contain 0.00789 x 17,000 = 134 mg/L or 134 ug/mL or 134 ng/uL.

The minimum amount of protein detectable in the gel will depend on how you stain it. Silver staining is very sensitive and can detect protein bands in the picogram range, while Coomassie staining works well in the nanogram range.

So, your estimate of 0.5 ug or 500 ng is well within the range for Coomassie staining. I would try running 1, 4 , 10 and 20 uL of your protein extract and see how strong the bands are.

Remember to run a prestained protein molecular mass marker and bromophenol blue tracking dye. What percent acrylamide were you going to use? I would use 10-12% for a 17 kDa protein like myoglobin.

Good luck and do let us know how it comes out.

Best wishes,

Sybee

[pseizure]

Thank you. I am using an 18% gel to ensure I get myoglobin separated clearly. I am planning to have the band (protein) sequenced at a protein mass spec lab.

I just noticed my protocol doesn't have a fixative step before staining with Coomassie Biosafe. Do I need to add one? I think I spoke with someone before who said that might screw things up but I'm not sure and nervous about what to do.

Thanks.

[deleted-140482]

I've never fixed a gel before Coomassie staining it, but I also wasn't familiar with this "Biosafe Coomassie." I looked it up, and found a product I assume is similar made by BioRad.

The instructions on their website do not mention a fixing step, so I think you should be fine without it. (see below)

Instructions for Staining Polyacrylamide Gels
SDS-PAGE Mini Gels

Wash the gels 3 times for 5 min each in 200 ml of ddH2O.
Remove all the water form the staining container and add 50 ml of Bio-Safe™ Coomassie stain or enough to completely cover the gel.

Gently shake for 1 hr. Protein bands will be visible within 20 min and reach maximum intensity within 1 hr.

Longer incubations will not increase the background.

Rinse the gels in 200 ml of ddH2O for at least 30 min. Stained gels can be stored in water.
Note: Residual SDS in the gel may cause background staining and interfere with band intensity while the gel is in the stain. Rinsing the gel extensively in water after staining will remove background and allow proper visualization of the bands.

SDS-PAGE Gels
For large gels (16–20") increase the volumes in the above method twofold.

Good luck!
JMP

[SciB]

Hi,

Just for your information. It IS Ok to fix your gels in acetic acid for later mass spec analysis, because that is how we have been doing it. But, as with many things in research, there are more ways than one to accomplish the same end and JMP's method is just as valid as ours.

Here's the protocol we use: http://skirball.med.nyu.edu/resources/f ... l-staining

From JMP's instructions for the Biosafe Coomassie stain, it is apparent that acid fixing is not necessary, so just follow their protocol and you should be fine. Be sure to read the protocol in my link because it has some additional tips for lowering background and improving sensitivity of the mass spec analysis.

Good luck and do let us know how it turns out.

Sybee