advances-in-protein-chemistry

Protein unfolding kinetics has been observed by monitoring changes in emission maximum or fluorescence intensity
due to perturbations in protein structure in presence of destabilizing agents [94-96]. Extrinsic fluorophores also have been
used in studying protein stability or protein structure and microenvironment. These extrinsic fluorophores either bind
covalently to primary amino groups, or specific group likes thiols or non-covalently on basis of charges or hydrophobicity.
Ammonium 8-anilino-1-naphthalenesulfonate or ANS is one such probe that binds to hydrophobic regions of a protein
which is accompanied by blue-shift of the fluorescence maxima from 545nm for free ANS to 470nm for bound [97]. Once
bound to protein it can be used as probe to monitor protein conformational or structural changes due to altered solution
conditions.
Ligand binding to proteins has been studied extensively using fluorescence spectroscopy by studying changes in intrinsic
tryptophan fluorescence, changes in extrinsic bound fluorophores or by studying changes in the fluorescence properties of
the incoming ligand as a result of protein-ligand interaction. Protein-ligand interaction using fluorescence spectroscopy
is performed by titration experiments and the extent to which fluorescence is quenched or enhanced is proportional to
formation of the protein-ligand complex. Change in fluorescence is measured as a function of ligand concentration and
is expressed in form of what is called ‘binding isotherm’. This approach can be used to determine the extent of maximal
fluorescence change when protein is fully bound to ligand (i.e. at saturation) and consequently fraction of bound and free
protein at any concentration of ligand can be determined. Ligand binding at equilibrium in this fashion can be explained by:
[ P][ L]
Kd
=
(19)
[ PL . ]
where K d
is the apparent dissociation constant, [P] is the concentration of the protein, [P.L] is the concentration of the
protein-ligand complex and [L] is the concentration of unbound ligand. The K d
values can be determined from non-linear
least squares (NLLS) regression analysis of titration data using different variants of the above equation. There are many
commercially available software packages like GraphPad Prism, GraFit, KaleidaGraph that can be used for treatment of
the binding data obtained in fluorescence experiments. These programs have preloaded equations and models that take
into account different possibilities that can exist during and after ligand binding to protein; the two important factors
being the number of potential binding sites and cooperativity of binding. These software packages also provide options
to incorporate equations for models that are not already present and then use them for data fitting and analysis. Few
examples of application in studying protein ligand interactions can be found in references [89-92] and [98-100]. Another
approach for studying protein-ligand interactions using fluorescence is Macromolecular Competition Titration (MCT)
method which is also a thermodynamically rigorous method. MCT is also useful in cases where formation of the complexes
is not accompanied by any adequate spectroscopic signal change [101]. It involves quantitative titrations of the reference
fluorescent ligand into a protein solution in presence of a fixed concentration (final experiment needs to be conducted
over range of concentrations) of a non-fluorescent ligand whose interaction parameters are to be determined. MCT allows
determination of total average degree of binding and the free ligand concentrations, over a large degree of binding range,
and construction of a model-independent, thermodynamic binding isotherm.
Fluorescence anisotropy (FA) can be employed in such cases in which intensity changes neither in intrinsic fluorescence
nor the fluorophore fluorescence are good enough as a parameter of binding data [99,102-105]. To have a certain value of
initial anisotropy the fluorophore needs to be attached to a molecule larger is size than itself and for this reason anisotropy
based experiments are mostly used in studying protein-DNA interactions. A certain disadvantage of fluorescence anisotropy
is that it can be used to measure binding constants if molecular size of protein-ligand complex is significantly different
from free fluorescing component. Another important consideration while using fluorescence anisotropy is that it relies on
an extrinsic fluorophore. This dependence on extrinsic fluorophore is for multiple reasons. First to maximize difference
between bound and free states the fluorophore needs to be attached to smaller partner. Second extrinsic fluorophores like
ones based on fluorescein or rhodamine derivatives and attached at multiple places to ligand (like on both ends of DNA)
have a large extinction coefficient which makes possible conducting experiments at nanomolar range concentrations [99].
In addition to anisotropy, fluorescence polarization can also be used to study protein-ligand interactions [99,105].
Fluorescence resonance energy transfer (FRET) is a fluorescence based phenomenon used to study biological interactions
at short distances (1-10 nm). Both interacting species need to have a fluorophore and emission spectrum of donor molecule
should overlap with excitation spectrum of acceptor molecule. The extent of overlap between two spectra is referred to as
spectral overlap integral (J). Transfer of energy happens through non-radiative dipole-dipole coupling hence donor and
acceptor transition dipole orientations must be approximately parallel. Forster has demonstrated that the efficiency of
process (E) depends on inverse sixth-distance between donor and acceptor:
E R
= R r
6
0
6 6
0
+
where R 0
is the Forster distance at which energy transfer is 50% efficient and r is the actual distance between donor and
acceptor. For studies involving biological systems distances between 2-9 nm are useful. FRET has been employed in
biological systems to usually study protein-DNA and protein-protein interactions. In studying protein-DNA interactions
the two fluorophores are on DNA molecule at distances where in unbound form they don’t involve in energy transfer
through FRET. Protein binding to DNA induces changes in the structure of DNA like unwinding or bending that brings
the two fluorophores close increasing the FRET efficiency. Increase in FRET efficiency creates changes in the fluorophore
of the acceptor molecule which can be monitored as a function of the binding protein molecule that induces this change or
can be monitored over time [99,106-108]. FRET in protein-protein interactions has been utilized for detecting interactions
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