No.  1)  A crystallographic hexamer does not mean that your protein is a hexamer in solution.  You'd need some other evidence for that, such as SEC, SAXS I(o) analysis, or MALS, or sedimentation analytical ultracentrifugation approaches.  2) Fitting a stoichiometry to ITC data can be tricky, but possible in some cases.  You need to be working at a fairly high (>250) c value, and in that case the 'breakpoint' or inflection point in the titration is the molar ratio at which your protein binds the inhibitor.  If your protein is indeed a hexamer and all 6 of the monomers bind the inhibitor, this would be 1.  It can be dangerous, not knowing these details, to fit the titration to a model with many sites, and it certainly by itself is not evidence for stoichiometry.

As far my understanding in ITC data analysis, Kd1 and Kd2 values are same  it means your those 2 binding sites are having same affinity with inhibitors or ligands. Kd3 and Kd4 values are increased, it is indicating that after certain range, protein affinity is decreased. Still I am not understanding that last two Kd values are decreasing, means again affinity increased.  The final conclusion is like first two sites have same affinity, then started decreasing and then why sudden it have started increasing, that is bit strange for me also.

It will be grateful for me also if somebody will discuss that data here.

No.  1)  A crystallographic hexamer does not mean that your protein is a hexamer in solution.  You'd need some other evidence for that, such as SEC, SAXS I(o) analysis, or MALS, or sedimentation analytical ultracentrifugation approaches.  2) Fitting a stoichiometry to ITC data can be tricky, but possible in some cases.  You need to be working at a fairly high (>250) c value, and in that case the 'breakpoint' or inflection point in the titration is the molar ratio at which your protein binds the inhibitor.  If your protein is indeed a hexamer and all 6 of the monomers bind the inhibitor, this would be 1.  It can be dangerous, not knowing these details, to fit the titration to a model with many sites, and it certainly by itself is not evidence for stoichiometry.

Hi Joe,

When you fit your data to 6-site sequential model you are basically fitting you data to 6*3 i.e 18 parameters. With that many parameters you can fit any data nicely.

You need other experiments (as suggested above) to confirm that your protein is indeed hexameric in solution. With ITC, a realistic model would be where all sites are equivalent i.e same Kd. Depending on how you define your protein concentration (for monomer or hexamer) you should get a stoichiometry of 1 or 6.

Good luck!

Hi Joe, yes, let me just back up & complement Soumaya's answer. Already fitting 2 binding events for a single ITC titration is a very unstable operation and 6 binding sites would certainly be too many parameters per data. You probably will get several quite different results if you do repeat fits of the data where you initialise the parameters differently.  I would start with a 1-site binding model and see what your N is, and how good the fit is, at least visually. For example if you get a good fit with N=6, then you could conclude that all your 6 interactions might be pretty similar. If you get poor fit with N different from 6 and significant slopes before and after the main transition, then you may have a mix of high and low affinity interactions. All the best for your analysis!

Could you, please, show an image of the thermogram and the fitting analysis of the isotherm?

Anyway, you must consider that the Ki's (step-wise association constants) you get from the Sequential Binding Model (very misleading name, by the way) are not the site-specific binding constants, but they are combinations of the site-specific binding constants and cooperativity constants. For example, for two binding sites K1 is not the association constant for a specific binding site, but it is equal to k1+k2 (that is, it reflects the formation of the singly-ligated complex); and K2 is equal to k*k1*k2/(k1+k2), where k1 and k2 are the site-specific association constants and k is the cooperativity constant (thus reflecting the formation of the doubly-ligated complex from the singly-ligated one). And, very importantly, the values of K1 and K2 cannot be used directly to assess the potential cooperativity; cooperativity and independency of binding sites must be discussed considering the expression 4*K2/K1 (if it is equal, larger or smaller than 1). Therefore, there is a straightforward, but not trivial, mapping between the two sets of constants. The larger the number of binding sites, the mapping equations get more complicated, but they can be generalized.

Many people still misinterpret the Sequential Binding Model and draw erroneous conclusions from their analysis. The Sequential Binding Model is just the General Binding Model defined in terms of the step-wise association constants. Therefore, there is no need for a specific name.

Regarding your question, there are some concerns with your analysis (e.g. too many parameters, the Kd's are not site-specific binding constants, values of enthalpies?), and showing the data and all the fitting parameters could help in deciding if the analysis is good or not, and figuring out what the most appropriate model could be (or what could not be).

Thank you all for the answers/conversation!! All very helpful. I've attached a fitted data. Please let me know what you think. Also please note that the curve starts exothermic and end endothermic, perhaps that could be added to the conversations as well. This data do not fit in one or two site binding model, and fit poorly in 1 and 2 seq binding modes... thank you all! 

You might also show the thermogram (thermal power vs. time) in order to better judge the titration.

Which are the concentrations of protein in cell and ligand in syringe? Have you performed a ligand dilution control experiment? Where protein and ligand in (exactly) the same buffer?

The heats observed along the titration seem to be very small, because the enthalpies are quite small. In this case, it is important to assess the dilution of ligand into buffer.

ligand/protein is in same buffer. ligand 2mM protein 25uM.

Ligand concentration is quite high (appropriate for the titration, but high). Therefore, you must perform a control experiment injecting the ligand at the same concentration into buffer. This control experiment should be performed routinely in ITC.

done that. baseline is clear..

Well, then it would be interesting to see:

1) Thermogram corresponding to that experiment.

2) Same experiment performed using a buffer with different ionization enthalpy.

3) Same experiment performed at a different pH.

One of the main concerns is the small value of the enthalpies. Those three points are indicated regarding that point.

The other main concern is that some of the potential binding sites might be considered as a subset of identical binding sites, but the analysis software does not allow you to do that. And that would reduce the number of adjustable parameters.

"Structural" binding sites do not necessarily correspond to functional binding sites.

Try to plot your enthalpy data versus the number of "expected" binding sites/protein per liter and assume a 1:1 ligand/receptor ratio. See what happens: the saturation level should suggest the actual concentration of "functional" binding sites.