Analyzing Multivalent Binding Events

Publications focusing on assessing multivalent interactions often highlight the difficulties associated with assessing binding kinetics and thermodynamics for such processes [11,15,39]. Determining the true equilibrium constant for a multivalent binding event is complicated because, as described above, multiple types of binding interaction may contribute to the interaction under investigation. Thus, an inability to dissect the contributions of monovalent binding, multiple site binding, subsite binding, rebinding of multiple epitopes within one or more binding site, and other effects such as steric stabilization renders the analyses of these systems difficult.

The evaluation of protein-carbohydrate interactions is complicated by the tendency of these substances to participate in multivalent binding and the low affinities of their monovalent interactions. These features dictate that the assay used to measure ligand activities be chosen carefully. Assays of several different types may be required to properly evaluate ligand potency. Moreover, generating data relevant to ligand function under physiological conditions can be a difficult and onerous task. Because throughput can be limited in assays that closely mimic physiological conditions, the best assay for screening potential inhibitors might necessarily be artificial, and also may not identify the best ligand (see Sections VI.4 and VII.B.2). Some of the more effective methods that have been employed to evaluate inhibitors of pro-tein-saccharide interactions are described in the subsequent sections.

A. Fluorescence Anisotropy

Fluorescence anisotropy is a technique that can be used to monitor low-affinity monovalent lectin-ligand interactions [52-54]. An advantage of this technique is that it can be used directly to determine ligand binding constants and also to confirm that ligands are competing for a specific saccharide binding site. The basis of the method is the detection of changes in the fluorescence anisotropy of a ligand-bound fluorophore. When a protein binds the labeled ligand, there is an increase in the rotational correlation time of the fluorophore because the resulting complex is large and tumbles more slowly than the free ligand. An increase in fluorescence anisotropy results (Fig. 6), and these changes can be analyzed to afford a binding constant. Moreover, binding affinities of unlabeled ligands can also be obtained accurately and

Figure 6 Schematic depiction of a fluorescence anisotropy assay. Fluorescently tagged molecules are excited by plane-polarized light; only molecules in the proper orientation are excited. Emitted light is detected in the original plane and in the perpendicular plane. The quantity of fluorescence observed in the two orientations is determined by the rate of tumbling, which depends on particle size and relates to binding.

Figure 6 Schematic depiction of a fluorescence anisotropy assay. Fluorescently tagged molecules are excited by plane-polarized light; only molecules in the proper orientation are excited. Emitted light is detected in the original plane and in the perpendicular plane. The quantity of fluorescence observed in the two orientations is determined by the rate of tumbling, which depends on particle size and relates to binding.

efficiently by monitoring their ability to compete with a fluorescently labeled control ligand. The disadvantage of this method is that high concentrations of protein are needed to accurately determine the weak association constants typically observed for protein-carbohydrate interactions. Even when the protein can be readily produced or isolated, the high concentrations required can result in protein aggregation and/or precipitation. Overall, the method has many advantages, but its applicability depends on the system.

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