@karim. It may be difficult to see an initial rate at high substrate concentrations, if there is product inhibition. The onset for inhibition could be very quick,. The shape of the progress curve and what are the kinetic parameters from the fit will tell you what the mechanism is. But I would switch the assay if I could to a spectrophotometric one. I think there are ways to couple GDP to an enzyme that utilizes NADP/NADPH etc. or use ADAM's idea by measuring inorganic phosphate, but with the continuous assay.

I am not sure in understand. At 250 and 100 uM you end up with curves, or hyperbolic plots? So I don't see the problem. At higher concentrations, the reaction will take longer, and at 10 mM GTP, the concentration is very high so you will need to do a time course to get an endpoint or Vmax.

What do you mean by a linear curve?

Hi Charlotte,

First, there is something I am not sure to understand: when you say you determine an initial velocity at a given time point, does it mean you only have one point (plus the zero time ) to determine your rate? I hope not, otherwise how can you know that you are really in the initial conditions? (i.e. linear part of time-dependent plot and substrate consumption inferior to ~20% over the kinetic duration).

Second, the linearity of your substrate-dependent velocity plot would for sure show that the km constant is probably >10 mM, but does not tell a lot about the kcat/km and does not seem to indicate a lack of co-factor (would be limiting for enzyme reaction).

Third, is your assay sensitivity limited by the initial quantity of substrate or by the released product? If it is only the latter you should dilute your enzyme for kinetic assay at higher substrate concentrations, if it is the former a sample dilution before hplc analysis should solve the problem

Hope it will help

We get a liner curve in some case but when we minus the values of sample blank/substrate blank we get start getting hyperbolic curve. Try to minus blanks values . If it do not work then you recheck you calculations, rather then increasing the substrate amount better to dilute the enzyme extract and also run a positive control

Using a lower enzyme concentration would solve the assay problem, and/or take your samples within a shorter time from when you start, or simply dilute your samples before analysis.

Why do you assume that Vmax should be reached before 10 mM? This asumption implies a low KM, which might not be the case.

...sometimes a high enzyme conc will result in prolonged non-steady state conditions. I have no idea what the situation is here, but your mixed results may point to that. I would redo all measurements with e.g. ten times lower [E] to see if the result is the same.

Again, I can't see why the possibility of a high KM is not plausible. Are similar enzymes typically displaying low KM:s?

In this situation, Km values can be useful. From previous studies to find these values (for example enzyme database- Brenda), you can scan an area 100 fold.

The high KM could be because you don't have enough Mg++.

Do you have enough of that?

Lower enzyme may exacerbate your problem, as you already don't reach Vmax in some cases. But 1.5uM does seem like a lot. I think you may need to try to get the assay conditions right for one concentration of GTP. You may need to add things to stabilize the enzyme. I don't know much about GTPases, but cofactor addition IMO would be essential. You can try adding detergents, ie at low concentrations to aid in the stability, or BSA, or reducing agents, etc etc.

also, if at higher concentrations you don't reach the same Vmax, it could also be because of product inhibition and not enzyme stability. I don't know how you can check that because you measure the ratio of GDP/GTP. Typically, you can add the product back at varying concentrations and plot 1/ v vs. 1/P. I would think there is a way to do this. Maybe someone else can comment. Or if you have an expert kineticist working with you, you can fit the progress curves in Dynafit to get the info you need. I would not necessarily expect your system to follow simple kinetic models.

If you measure initial rates, which you should, you will not see any product inhibition. Substrate inhibition will be visible as a decline in rate at higher concentrations, which you haven't seen (yet). If you want to add more coenzyme, a metal ion in this case, I would do that at the expression level (in the cultivation broth). After this, if you use the same enzyme batch for the measurements for the same curve on the same day, stability issues shouldn't be a big problem, usually...

1. Check your GTP substrate to make sure it is not contaminated with a significant amount of GDP, which could act as an inhibitor and also add background to your assay.

2. Watch out when using such a high enzyme concentration that you are not getting side reactions from contaminants in the protein prep. You could have more than one GTPase.

3. A simple approach to measuring the Km and determining a suitable enzyme concentration to use at the same time is to run a matrix of enzyme and substrate concentrations and take time points. This is easiest to do with a plate reader-friendly continuous assay, but can also be done with a bit more effort using a discontinuous assay.

Since inorganic phosphate is one of the products, you can measure it continuously as an absorbance change using nucleoside phosphorylase + methylthioguanosine, or discontinuously using Malachite Green/molybdate reagent. All the materials for these assays are available commercially. It would be important to make sure the phosphate contamination of your GTP was low.

First, I would say I am not so familiar with this protein, so I just comment on general rules for kinetics.

There are two ways to determine initial rates:

1> Get the full time course , determine the linear phase by eye and the slope gives the initial rate;

2> Fit the full time course to a polynomial equation (a + bx + cx^2....) and b is the initial rate;

So you probably should not determine the specific time point for one concentration and apply that to all concentrations.

And if for some reason, you could not saturate the reaction, so you could not get an accurate measurement of Km, you still get useful information for kcat/Km from the slope, which probably is a more important parameter for the protein.

@karim. It may be difficult to see an initial rate at high substrate concentrations, if there is product inhibition. The onset for inhibition could be very quick,. The shape of the progress curve and what are the kinetic parameters from the fit will tell you what the mechanism is. But I would switch the assay if I could to a spectrophotometric one. I think there are ways to couple GDP to an enzyme that utilizes NADP/NADPH etc. or use ADAM's idea by measuring inorganic phosphate, but with the continuous assay.

I agree withmany of the suggestions made above. For a spectrophotometric assay you can couple GDP formation to pyruvate kinase and lactate dehydrogenase monitoring the oxidation of NADH at 340 nm. This would be more sensitive and more rapid than the HPLC assay. It will also remove the issue of product inhibition as GDP is converted back to GTP and has the added benefit that substrate concentration is held constant.

Mg2+ is highly likely to be a co-factor and using this assay you can also quickly test for other activators / inhibitors, optimise for pH and ionic strength.

You don't mention the purity of your enzyme. An E.coli contaminant (e.g. HSP) could be responsible for the GTPase activity?

It may be that an interacting partner protein (or other activator) is missing from the assay so that the enzyme active site is not in an active conformation for GTP binding.

Or it may be that the enzyme is not saturable, in which case you can determine Kcat/Km from the slope of the rate versus enzyme concentration. This may give you a clue as to whether the measured rates are likely to be physiologically relevant.

When you say that you plan to stop the reaction earlier, what do you mean more exactly? I understood it as you are running HPLC and taking samples at chosen time points, correct? If this is what you are doing you can take your samples within a short time from when the reaction started. Then when you plot the conversion vs time you only include the points which are linear. Did I misunderstand you? Are you measuring the conversion, one point, after a certain time?

Alan's answer inspired me. Actually, you probably can try another coupled assay that will make your experiments easier. When GTP is hydrolyzed to GDP, inorganic phosphate (Pi) will be released. Do you know a protein called "phosphate binding protein"? This protein captures released Pi and gives a fluorescent signal if you attach a fluorophore (MDCC) to it. But if your reaction is too slow this assay might not be suitable for you.

Here is the reference:

http://www.sciencedirect.com/science/article/pii/S0003269707005143

1. Why do you need to add so much enzyme? Let us assume your protein has a 25 kDa molecular mass, and you are using a 100 µl assay volume. 1 µM = 1µmol/l = 1nmol/ml = 0.1 nmol /(100 µl). Then 0.1 nmol = 0.1 E-9 mol (25000 g/mol) = 2.5 µg of protein in 100 µl equals a final protein concentration of 25 mg/ml. That is a high protein concentration, for crystallization trials you may have enough with 10 mg/ml. I wonder how concentrated is your stock?

At such high protein concentrations, the product formation rate can be extremely fast. Then at the time you stop your assay almost all substrate was long before converted into product, and the more your add, the more product you get. In this case your rate estimate is wrong, because you ignore the true time interval that the enzyme required to catalyze the transformation.

I suggest you do a calculation to estimate the absolute amount of product your are forming and the absolute mass of substrate present in your reaction tube at the start. To remain in the steady state linear phase of the reaction, no more than 5-10% of substrate should have become product at the stopping time.

Sorry to point this out Rogelio, but there is an error in your calculations. 2.5 µg per 100 µl is 25 µg/ml not 25 mg/ml. If you dilute your stock 10-fold (i.e. a 10 µl to a 90 µl reaction mixture) then the stock solution of the enzyme is 250 µg/ml (0.25 mg/ml) which is perfectly reasonable.

@ Charlotte, if you go up in concentration of GTP, ie 1mM or higher, you will need to increase the Magnesium concentration due to mole: mole ratios.Also spectrophotometric continuous assays are super easy, less labor intensive, and provide much more useful info, because you get the complete progress curve in a single experiment.

Sorry Alan,

You are right, I was sloppy, but it is still high for an enzyme assay if your protein is pure, unless the enzyme kcat is low. Roughly, from the integrated MM equation, an enzyme with a kcat of 10 s^1 and a Km of 500 µM (a low specificity constant of 1E4 M^1 s^1), 1 µM of Active sites will consume 10% of 1 mM initial substrate in ~63 s, and will reach ~65% after 10 min. With 100 µM you will get 70% conversion in 10 min. and with 10 mM you would get 37% conversion in 10 min. That is assuming no product inhibition. If you take a two point slope, your rates would be 7 µM/min with 100 µM initial substrate, 65 µM/min with 1 mM initial substrate, and 3170 µM/min with 10 mM initial substrate. It looks like no saturation, but it really is a consequence of taking each estimate from data obtained at very different points in the progress curve.

Your assay is pretty rough. You assume the product is 0 at time 0, and your measure GDP formation at another time point. Using two data points to determine initial rates. It is rough but I don't see any problem with it. I think your problem has two possibilities.

1. did your quench work? Imagine you start your reaction and quench at certain time. Then you load your sample to HPLC to do the measurement. What if your quenching doesn't work? You reaction was still going on until all the substrate hydrolyzed. Check your quenching.

2. You said your essay is out of range when substrate concentration is larger than 10mM. It means your quantification probably has problem. You should quantify the product GDP according to relative integrated peak area of both GTP and GDP, not the absolute area. In this way, you can load less sample when the substrate concentration get very high to saturate UV detector. Also you need do GTP and GDP control very well to avoid measuring a wrong peak.

One more less likely possibility is simply your enzyme has extremely weak Km., not kcat/Km. This happens when GTP is not natural substrate of your enzyme.