I am surprised about many of the answers, well intended but they are only partially addressing your problem (by the way feedback inhibition-a term from Monod- refers to endproduct inhibition i.e., when the product of the last enzyme reaction inhibits the first enzyme reaction of the pathway), The main problem of your question is, that you do not described whether you checked for time and enzyme linearity in the first place. Do you have a one substrate one product reaction or a two substrate reaction (thus Lineweaver Burk will not give you the Km and Vmax because you need to do replots (see Enzymology books: MM-kinetics are only valuable for 1S-product reactions). The observed inhibition at higher substrate concentrations can have many reasons some of them are addressed above by the other researchers.It can be that your assay is not time linear, the substrate interferes,with the fluorescens measurement (so get the product and check for linearity, i.e., making a standard curve for product and fluuorescence) that will tell you whether the substrate may interact with the fluorescence measurement. You can have true substrate inhibition (frequently when at high S conc more than 1 S binds at the catalytic side. you can have at high product conc reverse reaction, or competitive inhibition at the S biding site (read up on this. Doing enzyme kinetics with partial knowledge of what is going on is a waste of time, i.e., taking vmax =v=o.5 as the true Vmax. S0.5 values (half velocity in S-saturation curve as a true Km etc.,etc.

Generally to determine KI values you have to do inhibition studies at various I conc. at Km or the apparent constants (see above) and at saturation (If you only do it at S-saturation and lower I conc and high KI (I 0.5) you will not see an effect if you have competitive inhibition) It depends what the final goal of your studies is how much work you must include. But anyway you have to do your measurements at low and close to saturation of your substrate(s) depending on your S-saturation curve after you insure that time and enzyme linearity is secured andyou are not in the Substrate inhibition range. Although, if your inhibitor is a substrate analogue you will get both inhibition effects. Keep in mind that time linearity depends for an endpoint assays also on your S-conc. you should not have under any circumstances converted more than 20% (most say 10%) of your substrate to reduce your error.

Finally and most importantly before you waste your time and money by measuring things you have to do all over or getting false results, get an intermediate enzyme kinetic book and read it from cover to cover (or take a class!). Don't relay on internet info only some of the descriptions are misleading.

You are right. Addition of inhibitors should also result in changes in the fluorescence and therefore from the determination of the binding constant values you could make the right choice of the inhibitors. Sometimes, just fluorescence experiments may not rightly help you to determine these binding parameters. If you have an access to Isothermal titration calorimetry, you could make the choice of the inhibitor better. Therefore, the best practice to use multiple approaches. So in your case enzymatic assay (in the presence and absence of inhibitor), fluorescence measurements (which you could perform if you have multiple inhibitors) and ITC is the best choice.

Be careful, sometimes the excess of substrate or product decreases fluorescence by quenching phenomena. But if you have actually inhibition by excess of substrate, you have to use a substrate concentration lower than the point where the inhibition is clear. According toy your data, I do not know if that concentration is higher than Km. If so, Km is OK.

It may be very simple also... Saturated detector? ;)

The concentration so substrate in a screen should be used such a way that one can monitor changes on the both the sides, i. e. increase or decrease in its value. But in your case since you are looking for inhibitor(s) that is going to reduce your readout as your reaction readout is fluorescence from the product. So you can use the maximum concentration of substrate which is having linear correlation with the fluorescence from the product. If you are using more than that then your readout will be in saturation zone and you may not see the effect of inhibitor. Hope this help.

I'm not sure I am understanding the answers here, but normally we found that the highest sensitivity to the inhibitor, in terms of decrease in absorbance in our cases, should be at 1-2 x the Km. I would think it should be the same for fluorescence, as you have suggested. I do not see how the substrate inhibition that you mentioned (quite normal for high concentrations of many substrates in many enzymes) would affect this, but as long as the substrate inhibition is not significant around the Km (it will obviously affect your apparent Km, but you could fit your data to the substrate inhibition equation if you need to account for this, I doubt it is necessary for simply screening for inhibitors). As noted by Abdalla, there are other ways to screen for binding, if that is what you want, but I suppose you need the inhibition data, too.

Do you have access to a known inhibitor for your enzyme? If so, try a dose titration of this inhibitor and see if the IC50 value in your titration comes close to the literature value. If not, I would just try a non-fluorescent version of the substrate and see if that inhibits the processing of the fluorescent substrate. Then your assay will be validated for testing new compounds.

I think that, you can determine the optimum concentration by making a serial concentration of your substrate and detect the concentration that gives you the highest fluorescence. the decrease of your readings may be due to feedback inhibition or the conversion of your product to another product (you can check the final product), any way you can select your optimum concentration via series of substrate concentration.

You may have substrate inhibition - which occurs more often than people think. If so, the v against [S] curve should go up to a peak and then decline, smoothly. Of course, there are assay-related possibilities which others have suggested above. Product inhibition should be avoided if you are measuring inital rates. (Another possibility is that you are missing the initial rate....if the reaction is very fast this is possible and then you actually measure a later part of the rate where the substrate has been used up...check to see if the inital readings are approximately the same....if these rise for your higher values of [S] then you are probably missing the initial rate. This can be addressed by reducing [E]...if you have to use different values of [E] then you can correct for this by dividing v by [E] and plotting v/[E] on the y-axis.)

Determination of inhibition parameters at S=Km is not a right idea. At this concentration, reaction velocity is most sensitive to any slight variation of effective [S]. If your inhibitor affect Km, as well as Vmax, then in the presence of inhibitor effective [S] will be lower than you expect, and you'll get decrease in velocity due to 2 parameters simultaneously. Usually inhibition is measured at saturating [S], or peak [S] in your case.

Other approach may be useful too, to measure v([S]) at different fixed [I], not v([I]) at fixed [S]. Then you can determine Ki by Dixon equation.

Use a substrate concentration in the range of saturation concentration, where you will obtain a lower variation of the reaction rate. Even more, you will have a higher sensitivity when you assay the inhibitors. In a first step, you can assay thee inhibition capacity comparing the reaction rate with and without inhibitor. As a second step you can determine the type of inhibition in the classic way: at a constant inhibitor concentration for different substrate concentration.

Is this enzyme one well described in the literature? If not, it should be interesting to study its whole kinetic behaviour. Another question is if you are working with a purified preparation, to avoid possible interferences. The next thing I would do is to determine if self substrate inhibition is real or an artifact, as it was suggested above. If substrate inhibition is real and you are considering to study this enzyme kinetics, I would sweep the kinetics (S curve) at different I concentrations to determine if there are modulatory effects only at the active site or also at possible inhibitory sites for the substrate if present (i.e. as it occurs with acetylcholinesterase). One I concentration may give you some information, but you cannot be sure if you chose the proper or correctly calculate Ki

It's not so easy for me to comment on a fluorescent assay since I don't know the structure neither of substrate nor of product. My opinion (as stated also by other comments) is that you must check for inner quenching, that is to say, if the substrate contains a chromophore which absorbs light (=energy) in the emission band of the fluorescent product, you cannot increase the substrate concentration above a value corresponding to ~0,01 AU. Fluorescence measurements are highly sensitive to environmental influences, and this is particularly the case if you have got a substrate which contains an intramolecularly-quenched fluorophore which becomes fluorescent when converted to product(s).

Any other suggestion on how to perform kinetic measurements (although excellent as many of the comments above) must await the resolution of this concern about the fluorescence quenching.

I wish all the best to you and a New Year of happyness and successful work.

I am surprised about many of the answers, well intended but they are only partially addressing your problem (by the way feedback inhibition-a term from Monod- refers to endproduct inhibition i.e., when the product of the last enzyme reaction inhibits the first enzyme reaction of the pathway), The main problem of your question is, that you do not described whether you checked for time and enzyme linearity in the first place. Do you have a one substrate one product reaction or a two substrate reaction (thus Lineweaver Burk will not give you the Km and Vmax because you need to do replots (see Enzymology books: MM-kinetics are only valuable for 1S-product reactions). The observed inhibition at higher substrate concentrations can have many reasons some of them are addressed above by the other researchers.It can be that your assay is not time linear, the substrate interferes,with the fluorescens measurement (so get the product and check for linearity, i.e., making a standard curve for product and fluuorescence) that will tell you whether the substrate may interact with the fluorescence measurement. You can have true substrate inhibition (frequently when at high S conc more than 1 S binds at the catalytic side. you can have at high product conc reverse reaction, or competitive inhibition at the S biding site (read up on this. Doing enzyme kinetics with partial knowledge of what is going on is a waste of time, i.e., taking vmax =v=o.5 as the true Vmax. S0.5 values (half velocity in S-saturation curve as a true Km etc.,etc.

Generally to determine KI values you have to do inhibition studies at various I conc. at Km or the apparent constants (see above) and at saturation (If you only do it at S-saturation and lower I conc and high KI (I 0.5) you will not see an effect if you have competitive inhibition) It depends what the final goal of your studies is how much work you must include. But anyway you have to do your measurements at low and close to saturation of your substrate(s) depending on your S-saturation curve after you insure that time and enzyme linearity is secured andyou are not in the Substrate inhibition range. Although, if your inhibitor is a substrate analogue you will get both inhibition effects. Keep in mind that time linearity depends for an endpoint assays also on your S-conc. you should not have under any circumstances converted more than 20% (most say 10%) of your substrate to reduce your error.

Finally and most importantly before you waste your time and money by measuring things you have to do all over or getting false results, get an intermediate enzyme kinetic book and read it from cover to cover (or take a class!). Don't relay on internet info only some of the descriptions are misleading.

I am agree with Juergen Wiegel

I feel Juergen answer does directly address the problem, and it is hard to add much more, without information in the nature of the enzyme or the reaction it catalyses.

Linearity of the assay (i.e. initial velocity) is a key aspect to explore enzyme inhibition. Check if, under the condition of measurement, the reaction is reversible, because if your reaction may be reaching equilibrium in some, but not all of the assay conditions (i.e. the net conversion is exhausted at some point along the assay), this may give you the erratic behavior you observe, without substrate inhibition or other kinetic complexities.

The concentration of substrate required to detect inhibition depends on the inhibition kinetic mechanism. You should check a standard enzyme kinetic textbook to see the details. For a detailed study of enzyme inhibition by a ligand, you actually need the enzyme response to variable amounts of inhibitor, at several fixed concentrations of substrate.

HAPPY NEW YEAR

Rogelio

You take subtrate concentration at which your protein show maximum activity. At higher concentration of subtrat also inhibit activity or end product may inhibit activity

The comments dealing with substrate inhibition are full and to the point but the problem may be a relatively simple one in that it is quite possible that the substrate quenches the fluoresence of the product. I recommend that before trying anything else, the effect of increasing substrate concentration on the fluoresence of the product should be tested.

Dear Venkat,

In my opinion, Dr. Juergen Wiegel has provided a path of your further exploration. Further, in case of fluorescence, there are two reactions, excitation and emission. so you need to coordinate with time factor while assaying your enzyme. Besides, quenching phenomenon also interferes with factual reading of fluorescence. I hope now you will understand how to process for.

Wishing you all the best in year 2013,

Sanjay Mishra

What you are observing might be correct. Substrate at higher concentration could inhibit enzyme activity. But you need not bother with the kinetics. Your basic interest is to characterize inhibitor. Many workers assay enzyme inhibition at sub optimum substrate level. You assay enzyme inhibition at various sub optimum levels of substrate concentration ,each time with various doses of inhibitor. Draw dixon plot for each doses of inhibitor. Your kinetics would tell you what type of inhibitor you have obtained. Now you could fix your assay method depending on competitative, non competitive or uncompetitative mode of inhibition of your enzyme.

@ Subhabrata Sengupta: Your statement is relevant and thus quite convincing to the researchers who are on the way to working in the area of 'Enzymology and Protein Engineering'.

Wishing you a very happy and joyful year 2013;

With regards,

Sanjay Mishra

In my opinion, Dr. Juergen Wiegel's view and suggestion over the problem is most appropriate. I only want to add up a very simple suggestion. Be sure whether above a certain substrate concentration, the activity remain constant or decreases. If decreased than what is the rate and what is its relation to the initial increase. Be sure that in any experimental setup for studying enzyme behavior, there should be only one variable in one setup. The ultimate result will come out with correlated interpretation of the set of experimental findings with different variables introduced as prob parameter.

In my opinion, the problem is simple and should be solved methodically with a clear and workable concept over the basics of enzyme kinetics.

Hi Venkat Pulla, what fluorescent substrate are you using? If it is MUB-substrate I suppose you first dilute your MUB-substrate in DMSO and then in sterile distilled water. At high MUB-substrate concentrations there can be inhibition effect of DMSO. However from my experience there are huge differences between different MUB-substrates and samples (i.e., water, soils, etc.). There are many and different inhibitors in different samples, so I recommend to measure your enzyme activity in range of 0-400uM of MUB-substrate (dense conc. at the low range). From this you will see where you are in 0-order (i.e. saturation) for each enzyme and sample.

I would use the substrate concentr. at 0-order range and NOT at close to Km value, since this is 1st order kinetic range and your enzyme activity depends on substrate conc. From my experience 100-300uM range of MUB-substrates are fine for most soil samples.

Anyway, you should measure every time at least 3 different MUB-substrate conc. to chose the right noninhibiting substrate concentration.

Best,

Jiri

You are probably facing a case called substrate inhibition, which probably results in binding of subsrate to another site of enzyme, changing the conformation leading to lower activity. Stick to a substrate concentration 10Xkm

Most likely substrate inhibition, which occurs due to non productive binding. Something like one leg each in two boats.