I've been trying to extract proteins from microalgae, and I'm getting around 0.7 mg/mL from a 20 mg pellet, but when I load it on my SDS-PAGE I'm not getting any bands, according to my calculation I'm loading around 0.3 mg/mL of protein per well which should be more than enough as I've read that coomassie blue R-250 staining can detect as little as 0.5 ug of total protein per band.
Do you have any suggestions that explains why I'm not getting any bands?
...and finally a classic sensitve silver staining potocol by Bassam et al. (1991)!
Hope this helps!
Roughly 100ng (give or take, depending on your protocol), so at .3ug/ul you should be detecting it.
If you posted your protocol and a picture of a gel after mild de-staining, it would be helpful.
What method are you using to measure protein concentration?
How thick are your gels?
Your ladder/positive control should take Coomassie and be very visible after destaining. Is this the case?
You could try G-250, which has better sensitivity (~30ng).
Hello Grant,
Thank you for you reply, I'm measuring my proteins using lowry modified assay and BSA as standardt,
I'm using coomassie blue R-250 at 0.5%, with 50% methanol and 10 % acetic acid for staining. For destaining I'm using 40% methanol and 10% acetic acid O/N.
I'm using a 10 % poliacrilamide Gel using the miniprot system from biorad (0.75 mm). I dont have any pics at the moment but there is always a very strong blue line in the middle of the gel and nothing else.
I load 10 uL of my sample which have been previously prepared with 2X laemmly solution with DTT for 10 min at 95ºC.
I dont have G-250, I'm stuck with R-250 =P (Lab budget).
I think there is a problem with my staining because when I run a gel using a protein lysate my bands are very faint, but I have tried different concentrations of R-250 and is always the same (0.05%, 0.25% and now 0.5%)
I always clean my gel with dH2O before several times to remove the excess of SDS and then I start the staining which I also let O/N.
Thanks!
Just as a word of caution, BSA is a poor standard. A search of ResearchGate should provide you with a number of reasons why.
Getting into your problem, you may want to modify your staining/destaining protocol. I use .25% CBB in the same solution you use. To stain, I cover the gel in staining solution, microwave it for 30 seconds and gently shake it for ~30 min. I then destain per the usual way with ethanol/acetic acid, with the following recipe for 1L:
785ml ddH2O
165ml EtOH
50ml Glacial Acetic Acid
You may be destaining too harshly with methanol. I have attached an image of what one of my gels looks like after a few destains.
What is the strong blue line in the middle of your gel? Is this your ladder?
At .3ug/ul sample with 2X Lamelli, load volume of 10ul (6.5ul Sample, 3.5ul Lamelli), you are loading a maximum of 1.95ug protein. This is very very low, but should be detectable by Coomassie. Can you load more sample? Can you run a standard at a higher concentration to verify your staining is effective?
The concentration of the Coomassie shouldn't matter too much. Only so much of the dye in your solution can be bound by the proteins (which you don't have a lot of) anyway.
Hope this helps.
Hello Grant, thank you for the nice tips.
I'll try and load more protein or try to concentrate it first using 10 % TCA, I'm also gonna try a new protein lysate from my microalgae to see if I can improve the yield ( I should be getting a lot more, but they're very hard to break). Then I'm gonna try with the protocol you gave me to check, hopefully it will work.
The blue line is going across my gel through every lane and is about in the 55 Kda mark, I've read it could be contamination but I used gloves all the time and clean glassware so I dont know what it can be.
One question, is ok if I use Ethanol instead of Methanol for the staining solution as well?, I dont like working with methanol so much cause of the fumes and all.
BTW how did u get your gel to look green?, using UV? it looks cool.
I've never stained in MeOH but I can't imagine it would matter much. Isn't it the acetic acid that smells strongly?
I scan my gels using the LiCor system. Coomassie dye fluoresces under the 700nm channel.
You should have plenty of sample.
I've always carried out the staining step for 30-35min - make sure you're doing this part long enough.
Hi there,
If nothing seems to work, you can always try silver stain which has 100X greater sensitivity than Coomasie staining (~2-5 ng protein).
The staining solution is simple and should not be a problem. The only real disadvantage of the method is the interference of SDS in the protein band visualization and thus several techniques to overcome this have been developed (see attached material).
Silver staining is indeed more complicated than Coomasie, but it's an alternative should all protocol optimizations fail.
Here is the material...
(Sorry, I couldn't attach it at first!)
More for silver stain protocol (Goodlett LabUniversity of Washington)...
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