I use the traditional pull down assay

Usually, GTPrS and GDP loading is used as positive and negative controls respectively. Depending on your experiment, I would do it in its natural state and compare it with the GTPrS and GDP loaded ones. You also might have to look into the time required to get maximum RhoGTPase activity (I usually see higher level activity around 5 mins (after stimulation) and then it starts going down) 

Thank you Avinash! I'm also checking for the natural state of the RhoGTPase, and do the GTPrS and GDP loading as controls like you do. Just that the signals were kind of weak ( I did time points at 3' and 10' too). I feel like that a small disturbance on the background might jepodize the results when analysing. I saw paper published that people tested the activated state (by loading with GTPrS), it looks like a magnification of the signals. Just that I'm not sure whether it is acceptable.

I usually use the tagged protein (like PAK - GST or Rhotekin - GST) in excess. I find that with more of that, the pulldown shows better results. Just an idea that might be worth trying for you

Hi,

I'm also using regularly GST-PAK and GST Rhotekin and I used to agree with the comment above that results look better when more beads are added, but I could also see in later control tests pulling down dominant negative GTPases that the increase in beads also increases the amount of unspecific binding of GDP bound inactive protein, so watch out. The good thing of using non-hydrolyzable GTP as control is that it also gives you info about the capacity of your beads, meaning whether they are saturated or not. In a "linear" pull down the amount of free binding sites in the beads exceed the amount of natural-occurring GTP-bound protein so you should always see more protein in a pull down in presence of GTPgamma that in your problem samples. On the other hand, if you have similar amounst of protein pulled down from your sample and positive control you might be in saturating the beads.

good luck

Thank you Sebastian, your answer is very helpful! I think I'm already over the question whether to check for natural, activated or inactivated state of RhoGTPase based on my observation comparing the three states with the same treatment on cells. As shown on the slide I attached, samples loaded with GTPrS or GDP didn't differ among different time points of stimulation. I believe this is what you said about the saturated effect on GTPrS loading. I have two more question based on your comment: 1) what's that unspecific binding of GDP bound inactive protein, is that extra bands or the darkening of the target band, how do you tell? 2) What do you do if you're sturating the beads? Is that a problem of the beads or something else? Thanks Sebastian!

Hi Zhimin,

Time ago when I was making the same question than you representatives from companies explained me that the unspecific binding of Rho-GDP (or even other GTPases) occurs through -weak-binding of the GTPases with RBD domain, that's why washing is an important step here. So when increasing the amount of beads the amount of available sites also increases and so the chances of unspecific binding. The problem is that at the end what you see in the blot is just RhoA, you cannot distinguish between GDP and GTP. What I did was to transfect cell lines with dominant negative-GFP tagged RhoA and then, after the pulldown, I test the membranes against GFP and could see that when the amount of beads was drastically increased above 40 ug (in general I used no more than 20) GFP signal started to show up.

I regularly use hippocampal primary neuronal cultures so the amount of total protein I handle is  quite low compared to what  from what one may get from cell lines. I did some saturation curves with GTP gamma to set up my conditions.

You can take the amount of beads you use and incubate them with increasing amounts of protein in presence of GTPgamma, when you reach the point in which the amount of pulled down RHO is not changing with the increase of protein your beads are saturated. Here you can reduce de amount of starting protein in your assay or increase the amount of beads. From what we discuss before I would go for the first option.

Cheers!