Here is something that is super simple, but may work: How about put a His-tag on one of them but not the other. Run the tagged domain over a Nickel column to bind it. Then flow the second domain over the column, with some washing. Then elute everything with imidazole and see if you have both domains, since the second one should have only remained via interactions with the first domain. It isn't quantitative, but it will give you information on whether the two domains bind each other.

You can try FRET measurement after tagging your proteins with CFP/YFP FRET pair which is a bit difficult but can be good to measure transient interactions between proteins.

Interesting problem!

I agree with Muhammad's suggestion that FRET would be a very good approach in this case. It might be a lot easier to just label the proteins with fluorescent molecules (e.g. fluorescein) in the first instance, as re-cloning with the tags could be a lot of work.

Another approach, less refined but perhaps a good first step, would be to try chemical cross-linking. This is fast, and might give you the confidence to go on and try some more involved methods.

Since you already cloned your proteins of interest, can't you subclone them into other vectors to perform a yeast-2-hybrid study? The initial investment may be a bit high, but you could then rapidly perform site directed mutagenesis etc. to test which residues are important for the interaction...

I'd go with Nicholas's suggestion. Using a xrosslinker such as Glutaraldehyde will let you know if there is an interaction. You can then decide if you want to go down the yeast 2 hydrid system.

Check the activity in the presence of 2-mercaptoethanol

How do you know that the activity of this enzyme needs its two domains to interact with each other? Is it possible that the two domains work independently?

Here is something that is super simple, but may work: How about put a His-tag on one of them but not the other. Run the tagged domain over a Nickel column to bind it. Then flow the second domain over the column, with some washing. Then elute everything with imidazole and see if you have both domains, since the second one should have only remained via interactions with the first domain. It isn't quantitative, but it will give you information on whether the two domains bind each other.

If your enzyme requires some co-factors you can add the required co-factors while performing your interaction assays. Also you can try in-vitro pull-down assay.

you can just try Yeast two hybrid easy and low cost ! or BiFC

FRET and Y2H both require additional tags that may interfere with the interaction if added in the region of contact. It is certainly not unheard of for an enzyme or protein to refold and regain activity in trans as you describe. Stable proteins such as GFP can refold from fragments and this is used in assays such as BiFC (bimolecular fluorescence complementation). A cross-linker is a good idea, but it will not tell you whether the two regions are associated in a functional manner (as opposed to non-functional aggregation of 'sticky' exposed regions). How much enzymatic activity are you detecting relative to intact enzyme? What is known of the domains in terms of folding and function?

Simple and elegant assay is GST-pull down. You need to synthesize labeled one protein and interact it in vitro with the other one tagged with GST. Use Glutathione-beads to pull-down the labeled protein and demonstrate the interaction using appropriate controls. Analyze the interaction by SDS-PAGE and autoradiography.

I would go with Patricia's approach. It is simple to do, albeit it requires subcloning to incorporate the his-tag and won't give you any quantitative information. Alternatively, you could label both proteins with small fluorophores (one each protein) that make a good FRET pair (such as Rhodamine and Alexa488). It would give you quantitative information such as Kd. However, labeling both proteins would involve a prior preparation of C-less constructs, and Cysteine substitution at some region of the protein (which will be a lot of work and could affect the folding of your protein if some Cysteine residues are important for it).

The method suggested above require special instrumentation to demonstrate the color changes in vitro. You can do that using gels but it is much more cumbersome than radioactive probes which is very sensitive.

Personally I like to chemically cross-link the proteins, then run them out and look at them via western. You can guestimate the size of the associated-protein(s) based on the shift of the molecular weight of your protein of interest and then you may just be able to strip the blot and reprobe for a candidate protein based on molecular size. This technique may also produce a variety of different sizes as different proteins may be interacting with your protein of interest. Furthermore, you can test different conditions and see how the factors that interact with your protein change.

You can try to measure kinetics of binding of those two proteins by surface plasmon resonance (SPR) if you know someone who can measure it. You can get KD value and Kon and Koff of the kinetics and it is sometimes quite interesting to know it.

If you can immobilize your enzyme substrate on a resin or use an immobilized substrate analog you could pull down both domains in the right stoichiometry if they are interacting.

for a quick estimation of protein interaction changes, i would use any spectrometric analysis and look for change in spectra. it would help if your protein has any helicity change, and you can observe in CD; or change in functional group shielding - peaks in NMR NOESY or change in stokes radius - spin the complex in analytical ultracentrifiguation, or observe complex formation in in real time under dynamic light scattering.

SPR is a fine method, but it is not really easy to perform. Immobilisation and regeneration steps alone need so much fine tuning and will take up a lot of time. Try the other methods first.

Thank you all.... I had followed Patricia Liwang's technique and yes, there IS interaction between the two domains when SDS PAGE of eluate is done (Actually, i had been trying that since a long time, but I had no hopes is getting result. But Liwang's comment showed that I m not alone with this kind of thoughts.... :P). Thank you, Liwang. And special thanks to Dr. Petri Kursula.... Ur analysis on the subject was extra-ordinary. I would definitely try to employ your suggestions in my work. Once again, Thank you all.....

SPR and CD are fairly good techniques for evaluating protein-protein interactions. Why not to use isothermal microcalorimetry? By such a technique, quantitative data can be obtained with small amounts of material. Another useful approach could be to attach a molecule onto a stationary phase (e.g. nitrocellulose or plastic well), and then measure the binding of the other molecule (previously labeled by 125-I, biotin or terminal NH2-derivatizing fluorophores/chromophores). These suggestions are for quantitative analysis.

Hi Siva,

Two approaches you might consider are 1) crosslinking with suberimidate (this links lysines in adjacent proteins) or 1) yeast two-hybrid (this is good if you have clones for the 2 domains) - this can even be done as a commercial service.

Best wishes

Prof David Sheehan

I'm not an expert on that, but I know that protein interactions can be measured with NMR. It can also inform you of the residues involved in the contacts, but that requires assignment of the residues which takes some time.

If these proteins have been crystallized and you can get the PDB, you can also try to predict their interactions computationally and see if the results make sense.

I do not know which native gel you tried, so I will mention the one that has worked a lot for my work in the past. Depending upon the pI of the two domains, you can hand-pour different pH native gels, and see for interaction as described in McLellan T, Anal Biochem, 126, 94–99 (1982) OR A concise version in Bio-RAd Mini-Protean Tetra cell manual pg 18.

As an alternative to using protein, a two-hybrid system (either yeast or bacterial) would be a relatively straightforward (and rapid) method to examine potential interactions (as mentioned above). However, these systems are quite prone to false positives, so an additional and more robust method (e.g. co-immunoprecipitation, surface plasmon resonance, etc) would be needed to confirm this putative interaction.

AlphaScreen: simple and fast.

Size exclusion (gel filtration) chromatography might be worth a shot. Very simple if you have the columns/instruments. In a non-denaturing, isotonic buffer (eg PBS) look at each individually and then together. Interacting complexes will elute at higher MW (earlier). You might get 1, 2 or 3 peaks. The equilibriums may change with dilution in the column but as these species will have migrated fatser already, they will still elute earlier. Will need to choose the column such that it separates appropriately. If lucky, and planned well, these kind of experiments can also be used to look at stoichiometry, equilibriums etc

Here is something simple. Run UV/Vis of each of them in the region 250-300 nm and also 200-250 nm. Run each separately then mix and rerun the spectrum. I suggest mixing at 1:1 stochiometric ratio. Take difference between them, taking into account concentration differences to generate a difference spectrum. You should be able to generate a binding curve this way if you want to vary the ratios. CD difference should be even more sensitive. This is as easy as the light scattering method but both methods should work.

do you know how strong the interaction is? I have been thinking about doing the same experiment as suggested by Patricia, but I don't know if the proteins will dissociate from each other during washing due to a change in equilibrium?

Thank you, Dr. Edelmann. Far western blotting is indeed easy and elegant method. I would try that also. Thank you once again.

You could do that by FRET.

Siva Ram Uppalapati,

Do you think this indicates a salt-dependent association? Nickel columns are typically run with high-salt buffers to reduce electrostatic interactions, but electrophoresis is generally done under low-salt conditions to minimize conductivity. It seems that this might help explain your initial negative results.

If you have purified proteins, I recommend Biocore and Far WB. Gel filtration with WB analysis of various fractions antibodies for both proteins of interest provides complementary info. In addition, confocal microscopy helps you obtain supportive evidence for in situ colocalization.

I'll second the yeast 2-hybrid recommendation; that'll also set you up for figuring out the basis of the interaction by mutational analysis.

If you can express either of them as fluorescent fusions (or fluorescinate them after purification) you can use fluorescence polarization assays to get the Kd of the interaction.

If you'd like a quantitative method have a look at, "Isothermal titration calorimetry of protein-protein interactions.", by Pierce et al. (1999).

http://www.ncbi.nlm.nih.gov/pubmed/10527727

PLA - proximity ligation assay can be used with endogenous proteins inside the cells, and it's quite good also to quantitatively measure the interactions.

http://en.wikipedia.org/wiki/Proximity_ligation_assay

You can try co-immunolocalization with 2 or even 4 different proteins. It will not give you interaction directly, but at least you will now precise protein co-localization. Good luck!

Try BioID is a method to screen for interacting proteins in living cells.

If you suspect that a protein , say protein S is interacting with the protein under investigation, say protein I. Use a rabbit polyclonal to immunoprecipitate protein I. Run a blue native PAGE of this IP in the first dimension, and SDS PAGE of the spot in the second dimension. Stain the second dimension gel. Having known the details of protein I, a silver/coomassie staining of the second dimension gel will give details of protein S. in addition, other interacting proteins also will be unraveled., that can be lidentified by mass-spec and/or confirmed by immunoblotting with a mouse monoclonal antibody.

I simply did native PAGE, It worth trying because of simplicity !!!!!!!!!!!!! More detail ; Pandey et al., PNAS 2006.

NIMR, New Delhi

Well, you could try to determine the structure of your domains/enzyme. However if you have trouble obtaining crystals you may have to try other methods.

Since the adduct is able to exert enzyme activity it should be stable enough in solution so, why don’t you simply try with a gel filtration chromatographic approach ? If needed you may add one substrate to further stabilize the enzyme (some time it works).

When you say native PAGE did not show interaction, did you do an in-gel activity assay? This would show whether there is activity when the two domains associate, and provide built-in control for no association. However, it could be that you will see no stable association by any interaction assay, but when the concentration of each is high enough in the "cocktail" there is sufficient contact that activity is possible.

Personally I use tagged proteins and as already suggested here FRET or Fluorescence polarization.

Most simply: Run size exclusion chromatography of individual domains and of the mixture and compare retention times.

Assuming that the reaction is performed by only one of the two domains (and not two individual enzymatic steps by each of the domains), it appears that the other domain is needed for activation. Since you are unable to measure interaction with CIP or native gel electrophoresis, the interaction most likely is weak. Hence techniques that include washing steps (e.g. size exclusion or trapping a His-tagged version) will probably fail. lf you can lay your hands on a label-free biosensor technology such as SPR (as suggested by Gunnar) you have a better chance to be successful, since these techniques are able to measure even low affinities. One domain has to be immobilized on the surface of the sensorchip by chemistry or antibody, the other can be injected to measure interaction. If you fail there, you have to look at co-factors that are needed for the reaction and that may have gone lost in the purification of the domains.

If your familiar with yeast and you are wanting to see if two known proteins interact. Look into doing a yeast two hybrid analysis. Briefly it involves tagging one protein with the activation domain of a transcriptional activator and the other with the binding domain of the same transcriptional activator then expressing them in yeast. If they interact the activator will promote expression of a reporter gene. Of course this would only work if your proteins are able to be expressed in yeast.

Our group used a yeast two hybrid system as well. You can really chop up proteins and target very small domains to verify the interaction. Using intact proteins, differential tags might work over a column using resins for FLAG or His. Size exclusion chromatography is a great first step for this analysis as well since it is a diluting technique and you'd be using affinity techniques in the second step. Both techniques also allow native elution conditions. Good luck.

Along with all of the other nice suggestions, how about something like a TAP-tagging approach? That has been used to identify weak complexes in the past. Also, if your interaction is transient, you could try non-specific crosslinking. It is also possible that they only interact when substrate is present, so you might want a huge excess of substrate in your assay.

But at the same time, what is your reasoning for identifying interaction between the two domains? If I understand correctly, the two protein domains are normally part of a single protein and would be covalently attached. So, what are you trying to determine by showing whether or not they interact separately?

- Are you trying to demonstrate that they can interact enough to form the active protein?

- Are you trying to identify how they interact? If so, you would need specific crosslinking or mutagenesis studies, not a simple bind/not bind assay.

- Something else?

The exact question you're asking should determine what experiment would be the most helpful. Good luck!

As suggested before I would try Analytical Ultracentrifugation. Because both proteins are in solution at the same time under centrifugal force you wouldn't have to worry about interaction with the matrix or washing steps.

If you know the interface of the interaction (and if you know that it is different than the active site) you could try mutate amino acids at the interface and see if direct interaction is needed for the reaction to occur or if it is only two half-steps.

Good luck!

There are many methods based on surface plasmon resonance, e.g.:

Article Comparative study of Surface Plasmon Resonance, electrochemi...

Simply I would use a glutaraldehyde or some other cross linking agent (depending on the residues on the stipulated dimer interface) and do an sds-page..

Since the adduct is able to exert enzyme activity it should be stable enough in solution so, why don’t you simply try with a gel filtration chromatographic approach ? If needed you may add one substrate to further stabilize the enzyme (some time it works).

There are also 2 techniques whcih allow you to study protein intreactions in situ and withr Co-Ip. ,Here is a link: http://www.nature.com/nmeth/journal/v8/n11/full/nmeth.f.351.html

Another technique to analyze protein/protein interaction is Biacore, although you would need to find a lab to work with that is quite experienced with this technique (https://www.biacore.com/lifesciences/research/proteins/introduction/index.html).

I do suggest to go ahead with Yeast two hybrid system, which is relatively very simpler technique. --> to proceed with it, you need to have your protein domain genes in two different yeast expression plasmids Or shuttle vectors one having DNA binding domain and other with activation domain.

--> As you have already come to know they both have activity together, you can definitely confirm it with Y2H system.

--> You can even do Surface plasmon resonance (SPR) for protein-protein interaction by Biacore/Biorad or GE instruments. In this case, you need the proteins to be in pure form and the availability of instruments.

If you want to understand more of the protein-protein interaction, and can express and purify enough of the 2 different proteins, Isothermal titration calorimetry is the gold standard. Will give you a Kd/Ka, stoichiometry and the thermodynamics (enthalpy, entropy and head capacity) of the binding between the 2 proteins.

Hi Siva,

Yes, there are many techniques by which you can identify the interaction between two proteins. First it depends where you want to do it, within the same system or you have two proteins expressed and purified separately. As earlier Patricia has mentioned using a His-tag on one protein and mobilizing it on the column and add the other protein and let it bind/if there is some interaction then elute both proteins together with Imidazole.

Secondly, you can use SPR.

Thirdly, you can use NMR, to do a protein-protein interaction study. It will be even better if you know the structure of one the the protein, you can tell where the other protein binds to your protein.

Hi Siva,

Yes, there are many techniques by which you can identify the interaction between two proteins. First it depends where you want to do it, within the same system or you have two proteins expressed and purified separately. As earlier Patricia has mentioned using a His-tag on one protein and mobilizing it on the column and add the other protein and let it bind/if there is some interaction then elute both proteins together with Imidazole.

Secondly, you can use SPR.

Thirdly, you can use NMR, to do a protein-protein interaction study. It will be even better if you know the structure of one the the protein, you can tell where the other protein binds to your protein.

You might try BiFC, a technique in which you make fusion proteins with half of a fluorescent protein attached to each of your domains. Neither fusion protein is fluorescent on its own, but the dimer is. See http://www.ncbi.nlm.nih.gov/pubmed/18573091

If you have the cDNA clones of the proteins of study, clone the cDNAs into the two hybrid vectors. The tethering and interactions can be measured by the induced Luciferase activity. If protein A interacts with Protein B and you can immobilize Protein A or B as GST-fusion protein, interaction can be determined by GST-pull-down and SDS-PAGE. For this experiment, you need to generate one of the proteins labelled with 35S-methionine by transcription & translation in vitro.

Best of luck

There are many good suggestions. Assuming you want a qualitative method, you could try a native gel.... which you did. The drawback is that if the interaction is weak you may miss it. This also means that gel filtration likely will not work. You can use cross linking to overcome this and then run an SDS gel. This is a simple experiment and since you have the proteins already does not require anything new other than the x-linking agent.

Another alternative is a diffusion measurements (via NMR) that can give you a good indication even if the interaction is weak. (preferably the proteins have different sizes and one of them is relatively small)

Here is a question! why would anyone down vote Patricia Liwang's comment? It is a reasonable suggestion that may or may not work depending on the affinity of the interaction. So why the down vote ?

Anyhow, as Markus indicated, weak interactions are hard (some would say impossible?) to detect by co-IP, native gels, or gel filtration. And failure to co-IP your two separate domains strongly suggests that your interactions are not super-strong, which would also rule out a pull-down approach. Some of the in vitro approaches suggested -calorimetry, NMR- require purified proteins in substantial quantities, and will not answer the question of whether your two separate domains interact with one another in live cells.

On the other hand, mixing both domains (I assume mixing means purified recombinant protein domains in a test tube) gives you some kind of enzymatic activity. What other explanation of that finding is there than an interaction between the domains?

If you need independent confirmation of the formation of some kind of complex in a test tube, Markus suggestion to use cross-linkers is very sound. If you need confirmation of the interaction in live cells, I would encourage you to use a fluorescence assay. The BiFC approach I suggested above works just fine. We use it in the lab to detect G protein beta-gamma dimers.

I would opt for the His-tag + cross-linking idea. I suggest using cleavable cross-linker so you can cross-link, co-purify on Ni-NTA column, test activity and then cleave the cross-linker. Then you can either run the mixture on SDS-PAGE and get relative quantity or re-purify the cleaved complex and demonstrate the activity is lost again. For cross-linkers I had good experience with Pierce (no stocks)

If protein's have cysteine's you could label them with FRET pairs and do a FRET assay. But your fluorophore-labeled cysteine's will have to be within 50 Ang of each other to show a FRET signal. Sometimes you can also see fluorescence quenching upon association or Anisotropy increase upon association.

I do not know what Native PAGE gels you have tried, but I had good experience with using Native gels as described in the paper [McLellan T, Anal Biochem, 126, 94–99 (1982)] (a concise version is in pg 18 mini-protean bio-rad tetra cell manual (no stocks)]. None of the commerical native gels worked in my case. Depending on pI of the two proteins you can choose which would work, and what direction electrodes should be in, a couple of tries will give you the best gel. It is always best to do native gels in cold room, hand pour them after degassing the acrylamide mixture for fine bands.

In my case, his-tag pull down technique did not work as good, primarily as my non-tagged protein also stuck to the column non-specifically and all tricks to avoid this gave little effect.

Good luck and keep us posted to what works.

Hi, Siva, actually, Co-IP and other in gel assays will tell you only about the probability that 2 proteins can potentially interact with each other. To get real evidence for interaction you have to use either FRET (as mentin by Bihal) , or, simple and more precise, PLA assay. So far PLA is one of most suitable and simple technique to prove protein-protein interactions in situ. With this method you can easy overcome all possible problems with data interpretations form in gel assays, including Co-IP and will have a direct evidence for the interactions in cells. Good luck!.

I have a suggestion... But not sure if this is help... The binding proteins (interactive domains) are less sensitive (can resist) to the presence of proteiases (protein lyses enzymes )...try...some ...if available ...

Only about 10% of scientist here suggest the size-exclusion in the case of this problem. Selected by me 3 of them are advising this simple experiment:

- Prof. Umberto Mura (who's score is higher than 95% of ResearchGate members)

- Prof. Guillermo Romero(his score is higher than 97.5% of ResearchGate members)

- Prof. Fatih Uckun (his score is higher than 97.5% of ResearchGate members)

This led me to the conclusion: then more experience has the researcher, then more simple solutions is finding.

Interestingly people suggesting glutardehyde are not warning that this chemical added in excess can cross-link not-interacting proteins, especially if they are concentrated. When my students were having fun with set-up of this method I had feeling they can cross-link everything, even two proteins kept in separate tubes, but close to each other;)

I don't understand fully the enthusiasts of the two-hybrid. If I would decide to edit an novel entitled "False positives in two-hybrid system", I will easily find ten volunteers in my Institute to write a chapter for me.

Please when advising in this essential question for the young scientist to keep in mind "are there any easier techniques" was pointed out there.

Hi all,

At the very outset, thank you for all of your valuable suggestions.

Finally, I m here to tell you all that my experiment is over and a manuscript is under preparation. Dealing with the protein-protein interaction issue, I had done invitro pull down assay, invitro domain complementation assay, Far western blot assay and Dynamic light scattering assay. All these four experiments unanimously proved that there is a definite interaction between the two domains. Also I had used bioinformatic tools to identify the possible interacting amino-acids between the two domains.

Once again thank you all.....

You might want to employ MicroScale Thermophoresis! This technique allows you to measure interactions also in cell lysate if you express one of the interacting proteins as a fluorescent fusion protein. Please visit our homepage for further information:

http://www.nanotemper-technologies.com//home/

Hi, Heide, this is amazing technique, however, MicroScale Thermophoresis give you only a probability that interaction may happens. Thereafter one need prove interaction directly in situ.

I just wanted to mention www.nanotemper.DE as perhaps a potential possibility do further address the question - and then, the company already replied and mentioned that (two messages before)... So, I am little too late.

Please note: I am not affiliated with that company, nor do I even know the members personally, nor am I anyhow expert in their technology - I never used it (so, I may be wrong using it for your problem). But from what I think of testing, their technology may be useful in protein-protein bindings.

Unfortunately (for me), I registered a very similar website (which I am not mentioning here) before they started their business and sometimes people erroneously keep sending me emails instead of them.

It is an amazing technique! You are right, measuring in cell lysate, you can not exclude that other factors are interacting with your fusion protein as well. But it will give you some first insights and in a next step you can perform some more MST experiments with purified proteins. You will not only explore if they are binding to each other, but you will be also able to determine the affinity of this interaction!

I'd like to add that FRET doesn't prove direct interaction and only shows that the fluorophores connected to the enzymes are close to each other. You can definitely get FRET from two non-interacting proteins that are merely in close proximity to each other so be aware of that.

Is that really true about the fret assays? I would love if you could give a reference. I am interested in this because that for FRET substrates, which I use, the acceptor and donor pair have to be on the same molecule. If the fluors are released, there is an increase in fluorescence. I know we used a dansyl cyclosprin A derivative, and the change in fluorescence upon direct binding was monitored. One can also use tryptophan fluorescence. On another note, if one is an enzyme, one can use an activity assay. This is usually direct and easy. But there would have to be some kind of change in activity in order to assess if binding occurs.

I like the His tag idea, and this has been done extensively by others.

I meant to say if one protein is an enzyme, then an activity assay can be used. Sorry for my typing.

I will have to read about the nano temper technology. Sounds cool.

I know with some enzymes, if they are heterodimers for expression, each subunit is placed in the vector, in opposite directions, so that when expression occurs, each subunit is expressed in equimolar concentrations. There are lots of reasons why the enzymes would not have activity when expressed separately. Catalysis requires both subunits.

One could have the binding site for a cofactors, for example, and the other, for the substrate. Or more likely, the active site is formed by interactions between the heterodimers, or possibly even tetramer. One subunit could be an activator.

If you want to know what is going on, simple kinetic studies with the substrate for the enzyme could shed light on things. If the dimer or multimeric enzyme is necessary, catalysis should be concentration dependent. So if a concentrated solution of active enzyme, is diluted enough, the enzyme activity should fall off. This would indicate a direct interaction between the two subunits, and you can get a binding constant. So one can look for concentration effects on your kinetics.

Another explanation, is that each protein is not properly folded without the other, and then when added together, refolding takes place. This might be able to be followed by circular dichroism.

Please feel free to contact me if you have any questions regarding the technology and possible applications!

Thank you Heide

Obviously it will depend on the affinity. Anyway, since you have recombinant proteins... 1. you could try calorimetry but you will need a good amount of protein. 2. You may try NMR. You will need one protein labeled with 15N, then do a titration course with several concentrations. You don't even need to assign the protein, all you need to see is changes in the 2D spectrum if the proteins are interacting. And if you do assign the amino acid residues... you will also know the interacting site. Again, you will need a good amount of protein. Both techniques can give you an estimated Kd as well (the ITC obviously will be much more accurate). An ITC should be easy to find in any institute.

You might think about photo-activated cross linkers which might be able to capture the two proteins in their native interaction state but will, when 'flashed', form a permanent bond between them.

well, there are multiple ways: affinity -MS or Affinity - PAGE (practically you immobilize one covalently on a sepharose column and see if the other one is binding), biosensor and biosensor -MS , ELISA (if you have a specific antibody against one of them), ESI-MS in native conditions, ion mobility- MS.

Please consider my following idea just a a brainstorming: I am not really expert in what you did, but it sounds to me, it could be exciting, if true. As I suggested before you could try to contact that international (young) company to ask, if they could help you in any way - but without the need for you to really buy their instrument. Maybe, anyone in your area already has such an instrument, or there is a way the company could just do the experiment for you (you probably have to pay their service or negotiate something out). My experience in general (I do not have experience with that new company) is that new technologies are kind of happy to be mentioned in a first-class publication, or any publications with new findings to have their exclusive method recognized. But maybe I am wrong and their technique is already so established that they can not really offer much for your specific questions. Or, my assumption could be wrong that this technique could really bring you forward. Again, I am not affiliated with that young new tech company.

You may consider the proximity-ligation assay. It is a little expensive, but gives you very robust results. And great pics!!

Since you have a functional assay, and what you really want is active enzyme, I am not sure why everything cannot be done with the assay or assays if there are two different enzyme activities. Many proteinases have the same issues, ie they have to be dimers in order to be active. So I would be concerned with what expression and reconstitution conditions that give you the best specific activity. You can thEn do the assay as a function of concentration of protein. Please read the papers on cysteine proteinases or HIV proteinase. They will be helpful. If you show, for example, one protein allosterically activates the other protein, varying the subtrate and activator concentration, in a cheap and simple experiment will give you the info you need. But you do need fairly purified components.

The data can be fit using equations from Siegel's Enzyme Kinetics using PRISM software. PRISM also has some built in enzyme kinetics equations.

To made funcctional assay is an excellent idea. In that case one can prove interaction in the level of proteins functions. This assay can be the last step after 4 previous:

1. Probablity that proteins can interact (Co-IP).

2. Years 2-hybrid system.

3. Interaction in the specifci cells and specific compartments (PLA - proximity ligation assay)-.

However, I am not sure how easy one can perform functional assay in the case of organs. Prehaps, it will be easy in the cell cultire.