I am doing a 12% SDS PAGE gel after total plant protein extraction from cotton leaf by TCA/ACETONE precipitation method. I am loading the crude protein without dilution after keeping it for denaturation at 92.7 degree C for 5 minutes in a PCR block. Unfortunately I am getting only the higher MW bands and not the Lower MW bands. Why? Also note that my loading buffer contains both SDS and DTT for denaturation. But why not getting denaturation? Also note that when I am keeping gradient from 85-95 degree C for 5 mins I am getting denaturation in all but when keeping singly at one temperature not getting denaturation? Very surprising!
Can you attach images of the gels. Are the bands spreading? I am having a hard time understanding your questions. Protein denaturation by temperature follows a first order kinetic, what matters is the time and temperature.
Why are you using TCA/acetone precipitation? How does the band pattern compare to that of a non-precipitated extract?
When presenting a gel it is necessary to provide size markers, otherwise it is difficult to understand which "size part" you are refering to.
Very small peptides may be lost because of incomplete precipitation. Also, you may not have resolved the precipitate completely. See e.g. http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0124723
It seems that you have a lot of proteins and thus running out of sds. Sds must be always in excess for proper denaturation and charge.
I diluted my proteins in various concentrations and then heated in PCR block at 92.7 ' C for 6 minutes. Still not getting denaturation. Should I prepare my loading buffer again? It has got a bit old. Should I change the composition of my loading buffer. How long can a loading buffer be used?
My loading buffer composition is:
Tris-Cl 100mM, pH 6.8
DTT 200mM- 100 ul
Glycerol 20% - 400 ul
BPB- 4mg
SDS 4%- 800 ul
Deionized water- up to 2 ml
The protein is dissolved in a buffer of Tris-Cl pH 8.1.
While I add the sample I take 20 ul of diluted protein sample along with 25 ul of extra DTT plus 75 ul of the above loading buffer.
Are you heating the sample after adding the loading buffer or before?
Original SDS-PAGE is not suitable for separating proteins smaller than about 20 kDA. Use tricine gel or tricine with urea to see small proteins on SDS-PAGE.
Protocol: http://www.nature.com/nprot/journal/v1/n1/full/nprot.2006.4.html
Can you explain details of sample you have loaded in each well (ie. protein conc., loading volume).
What abt the sample loaded in the lane next to molecular wt. marker, since some small mol. wt. protein has been resolved in that.
As Valentina said, we can not resolve small mol. wt. proteins/peptides in usual SDS PAGE gel with tris-glycine buffer system.
I prefer using this buffer:
Buffer: 12% SDS (wt/vol), 6% mercaptoethanol (vol/vol), 30% glycerol
(wt/vol), 0.05% Coomassie blue G-250 (Serva), 150 mM Tris/HCl (pH 7.0)
Mix 15 ul of sample with 5 ul of buffer. Heat the sample to 37 C for 15 min (up to 60 min). Some proteins irreversibly aggregate with SDS if you boil the samples. I never boil the samples. Give it a try and see what happens, if a solid still exists even with diluted samples you must have something else in your sample that are not proteins.
Hi,
Lower MW proteins missed so I suggest to use fixation buffer before staining( acetic acid,methanol and water)
I have a similar problem. My gel is 4% stacking and 10% resolving and I used the comercial standard marker of Promega. The proteins are separated good until the 25KDa band, from this band to the smallest band (10KDa), my marker is not separated. I prepared fresh buffers and I have checked the pH. My electrophoresis buffer has 4% of SDS but the problem cotinues.
Dear Anirudha Sahu,
why do you do the TCA/ACETONE precipitation step? For leaf extracts the protein concentration is typically high enough to directly use them for SDS-PAGE.
Then you need to be aware that Rubisco is very dominant, comprising up to 50% of the total protein. Are you assuming that the low molecular weight proteins have been lost? Or could it be there are only few of them anyways?
If you want to resolve these proteins, you need to use higher % PAA gels or gradient gels. It looks like you cast the gels yourself, you can easily go up to 15%.
To get sharp bands you may have to either permanently block the Cysteines or ensure the remain reduced, i.e. use a negatively charged anti-oxidant in your buffer. 2-ME and DTT are not charged and will not run into the gel, you may get oxidation and cross linking during separation. The result is that bands are not as sharp, making it difficult to spot low intensity proteins.
Best, Markus
12% gels are not much use for anything less than 20kDa. Additionally, you are not really resolving anything below your HMW plant bands, meaning the proteins are messed up / insoluble. DON'T do the precipitation step; in my experience, proteins don't like TCA/acetone. There is a simple ethanol precipitation step you might try, but first do as Markus says (hi, Markus!) and just run straight extract.
Laura Martínez Rachadell did you find the solution for your problem. i recently , for the first time got similar problem. we use 6% stacking and 18% resolving gel. Usually all the proteins are nicely resolved. but for the first time it happened that proteins below 35Kda are not resolved at all, as evident by ladder, while higher mol wt proteins are very nicely resolved. Under usual conditions lower mol weight proteins resolve better because they migrate more distance as compared to higher ones.
To find out the possible reason, I was going through the basics of SDS PAGE...the whole ph difference thing and chlorine , glycinate ions.
i suspect it has something to do with ph. I suspect either there was some problem with running buffer ph or by mistake I used ph 6.8 buffer in resolving gel instead of 8.8.
can anyone help?
Heena Pahwa : you sure there isn't a problem with your running gel acrylamide concentration? If the gel was 8% instrad of 18%, that could explain things. OR you're right, and the pH / composition of the buffer is wrong. In which case, you just have to make new...B-)
Ed Rybicki if running gel acrylamide concentration was less, then smaller molecular weight protein would have run out of the gel. Isn't it? Ok I forgot to mention one thing. This time my gel ran longer than usual. Usually we wait for dye front to move to the bottom before we switch it off and it takes about 90-100 minutes but this time it took 156 minutes (60 V for stacking gel and 95 V for resolving gel). While it was running, I did not pay attention to the ladder, I was only waiting for dye front to go down. While transferring, I happened to look at the ladder, that it had not resolved below 35 kDA.
So if only acrylamide concentration was less, then wouldn't it run normally, as in smaller molecular weight protein will migrate faster and leave the gel instead of waiting there for larger molecular to resolve?
Yes I just have to run the gel again with right ph and acrylamide concentration and everything will be normal again, but because this thing was very strange so was just curious to know the phenomena behind and the root cause of the mistake. :)
I found the solution to my problem and there was the Tris reactive with I did my buffers to do the gels. Try to change it
Laura Martínez Rachadell i didn' t get it....
My problem is also solved. there is no problem with resolving buffer or acrylamide concentration. I changed two things. Running buffer I made fresh (that we generally use twice) and I used a different power pack. Either that running buffer was destroyed (cant say how) or power pack is the problem. because I noticed that current that it was giving at 100V was 6mA, which is very low. usually current is 19-25mA.
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