Have you thought of fluorescence spectroscopy. You would have to label either the peptide or the protein.

Note: I have never done it myself and even forgotten the name of the specific method.

Dear Subhash

if your peptides are short and soluble I think that you can try:

1) Differential scanning fluorimetry ( https://www.nature.com/articles/nprot.2007.321) which is able to provide you the melting temperature of your protein. Is probable that the peptide binding will stabilize the protein and in presence of the peptides, if the binding happen, you will se an increase of melting temperature. Sincerelly i never performed it with peptides, but many times with other cofactors as metals ( Article Identification of a Novel Zinc Metalloprotease through a Glo...

Article The structure of Clostridium difficile toxin A glucosyltrans...

It require protein concentrations in the order of 10-30uM depending on the protein MW and small volumes (40ul/well)

2) Otherwise you can try to perform a sort of ELISA by immobilizing the peptides on a Covalink plate, blocking the non specific binding site, incubate with the protein, wash away the non buonded protein and reveal the bindng using an antibody able to recognize the protein with a protocoll similar to the ones used for peptide array ( Article Identifying Protein-protein Interaction Sites Using Peptide Arrays

Personally i prefer the first approach but not all the proteins show good profile at DSF, you need a RT-PCR machine and i never used it with peptides therefore is good to have some backup plans in case it not work.

Ciao

Manuele

You can use fluorescence polarization (or anisotropy) to detect interaction of a small ligand with a peptide and need only a few micrograms to do this. Label the peptide with an appropriate fluorescent dye and records its anisotropy. Add incremental amounts of protein to the cuvette and record the anisotropy each time. when the small fluorescent-containing ligand (peptide) binds a larger molecule (protein) the change in anisotropy will be very noticeable. You can plot the change in anisotropy versus amount of protein added and can calculate various binding parameters. Good luck.

DSF could work well as mentioned above (it works very well for small molecule ligands) - this will give you a yes/no answer but wont' give you any binding affinities which might be ok depending on what you want. Alpha screen might also work or you could try SPR (either on an SPR machine or on an Octet multi-channel machine.)

You could do a pull down against your protein if you can immobilize your peptide onto beads? That might be the easiest - Especially if you can add a biotin tag fairly easily onto the end of your peptide or something like that.

Alternatively if you have an antibody against your protein (or a tag that your protein has) you could use a peptide scanning dot blot where you print small amounts of peptide onto a membrane (changing the region of the peptide that you're looking at, at each point) and then wash the membrane with your protein which will hopefully stick to the dots that contain the fragments of peptide that it binds to. You can then use an antibody that sticks to your protein to identify these spots. It uses a very low amount of material but you do need someone to print the membrane for you.

Importantly, if you want to determine accurate binding constants, you need to be working at a concentration higher than the Kd of binding. A lot of these techniques use very small volumes of ligand but need high concentration which is something to bear in mind as to whether your limitation is solubility of peptide or just how much material you can afford.

I don't know if your peptides or proteins have tryptophan or other aromatic amino acids, but intrinsic tryptophan (or other amino acid) fluorescence is one possibility. No labeling is required. Using a fluorimeter, one monitors the natural fluorescence emission spectra (~320-400 nm scans) of tryptophan in the protein in a cuvette after excitation at about 290 nm. Ligands (peptides) that bind to the protein may alter the fluorescence intensity or shift the wavelength of maximum emission. You can monitor these changes while titrating in the ligands.

Kinetic studies can be performed using SPR - a plasmon-enhanced reflectivity measurement over gold chip using microfluidic setup. It is extremely sensitive, no labeling is needed, and both association and dissociation phases are tracked. For a commercial state of the art, take a look at Biacore. If you need even more sensitivity, ellipsometry can be incorporated, resulting in the TIRE technique, see e.g.

Article Treatment of SPR background in Total internal reflection ell...

Hey Subhash,

There are obviously a lot of choices out there for what you're asking, but a colleague of mine uses LigandTracer to monitor the interaction between proteins and small peptides, so long as the small peptides in question are either radio or fluorescently labeled. The assays are simple to run and all you have to do is attach your proteins to a Petri-dish. Of course all of this is dependent upon the properties of your particular protein

Of course, there are many alternatives to ITC and MST.

However, Subhash Narasimhan, I would not say that "ITC requires milli-molar concentrations... and MST also requires low milli molar concentration". In particular, ITC requires a larger amount of sample compared to other techniques (e.g., fluorescence polarization), but the concentrations are usually in the submillimolar range (unless there are many binding sites and/or the binding affinity is low). The standard set-up is: 10-20 microM in cell (300-1500 microL) and 100-200 microM in syringe (40-300 microL). The indicated volumes depend on the calorimeter model. I have been able to perform ITC experiments with 2 microM in the cell and 30 microM in the syringe. At the end, the observed signal depends on the binding enthalpy and the binding affinity.