Fluorescence Polarization FP

FP is an alternative readout principle for endopeptidase activity assays. FP or anisotropy measurements allow the detection of changes in the rotational correlation time of particles. These differences in the rotational correlation (or relaxation) time are related to different masses of particles. The experimental determination of steady-state fluorescence anisotropy requires the linear polarization of the light used for the excitation of the probe as well as linear polarization of the emitted fluorescence. Based on data of an appropriate experiment, the fluorescence anisotropy can be calculated as:

where Ivv and Ivh are the vertically and horizontally polarized fluorescence emission intensities measured with respect to vertically polarized excitation light (Lakowicz, 2006). The factor g is used to correct the usually imperfect optical system. The g factor is given by:

For horizontally polarized excitation light, Ihv and Ihh are the intensities of the vertically and horizontally polarized fluorescence emissions, respectively. It is strongly recommended to use the background-corrected intensity values for the calculations. The background correction is determined by subtracting the intensity value obtained for the buffer with a certain polarizer setting from the intensity value for the sample obtained with identical settings.

Practically, the g factor can be determined using a free fluorophore with a known anisotropy value as a standard. Free fluorophores like TAMRA (5,6-carboxytetramethylrhodamine) and fluorescein with rtrue values of rtrue = 0.0202 (ptrue = 30 mPU, see below) and rtrue = 0.0134 (ptrue = 20 mPU, see below), respectively, can be used (some sources report 27 mPU for fluorescein in solution):

The anisotropy measurements yield only the relative amounts of the different species in solutions. The fraction of the product can be calculated as:

where the factor Q is necessary to correct for differences in the fluorescence intensities (FIs) of the fluorophore in the parent substrate (FIsub) and in the product (FIprod) (Marks et al., 2005):

Q = FIprodlFIsub

It should be noted that all relations given in the formulas above are restricted to fluorescence anisot-ropy. However, it is popular in the screening community to use fluorescence polarization over fluorescence anisotropy. Polarization and anisotropy are related by:

Fluorescence polarization values are usually shown in millipolarization units (mPU); mPU = APU = 1000 mPU. A prerequisite for fluorescence anisotropy-based assays is that the enzymatic cleavage event leads to a significant change in the mass of the molecular moiety carrying the fluorophore. When short peptides are used as protease substrates in an FP assay, the difference in size between substrate and the fluorophore containing product is in many cases too small to generate changes in the polarization value needed to develop assays robust enough for high-throughput compound testing.

However, the development of robust FP-based protease assays is feasible as described by Tirat et al. (2005). The authors developed FP assays for UCH-L3 and USP2 based on both undecapeptides and on ubiquitin constructs as substrates, C terminally labeled with TAMRA. Dynamic ranges of >100 mPU at 100% substrate conversion sufficient for automated compound testing under initial velocity conditions were observed only with large substrates based on ubiq-uitin. In general, our experience is that robust FP assays for proteases can be developed if the molecular weight (MW) of the proteolytic product carrying the fluorophore is smaller than 10 kDa and the MW of the intact, uncleaved substrate is at least 10 times larger than the cleavage product when fluorophores with short lifetimes of 1 to 4 ns (such as fluorescein or rhodamine) are used.

The concept of MW enhancers was developed to increase polarization value changes caused by large size differences between substrate and product when small peptide substrates were used. The most frequently used approach is based on the attachment of a biotin molecule to the terminus of the peptide on the opposite site of the scissile bond to the fluorophore. After the incubation of the substrate with the protease, streptavidin is added to the assay. Streptavidin binds to biotin and enhances the masses of the substrate and the unlabeled product, thus causing an increase of the polarization

FIGURE 2.6 Fluorescence polarization readout principle. In the intact peptidic substrate (amino acids symbolized by X, Y and Z) labeled with a fluorophore (dye) and a molecular weight enhancer (MWE) at the opposite sites of the scissile bond, a high polarization is observed (long black arrow). After cleavage of the substrate between amino acids X and Y by a protease, the MWE and dye are decoupled and a low polarization of the labeled cleavage product is generated (short black arrows). A decrease of the polarization dependent on the enzymatic velocity is recorded.

value of the substrate (Turek-Etienne et al., 2003; Flotow et al., 2002; Leissring et al., 2003; Levine et al., 1997; see Figure 2.6). Examples FP-1 and FP-2 in Table 2.2 indicate the dynamic ranges that can be achieved with FP assays with MW enhancers.

Other MW enhancers have been described. Peptides with oligonucleotides as enhancers were attached to substrates for a polarization assay with caspase-3 (Lopez-Calle et al., 2002). As strepta-vidin has four binding sites, fluorescence self-quenching effects between the dye molecules can occur with increasing binding degree resulting in depolarization (Pope et al., 1999). The alternative MW enhancer approaches may overcome this limitation.

In contrast to the other fluorescence-based methods described, FP offers the opportunity to measure the binding affinities (KD) of inhibitors in addition to the inhibition of the catalytic activity. The application of this approach is straightforward if binding of a peptide with a MW below 10 kDa to the protease is known. The peptide can be labeled with a fluorophore distant from the binding site without influencing the binding properties. Such binding studies can help to explain the conformational changes of proteases (Nomura et al., 2006), but can also be used to determine binding affinities of protease inhibitors by competition experiments. The approach is much more difficult where a small molecule inhibitor must be labeled, because significant efforts in terms of structural analysis and synthetic chemistry are required to avoid or minimize the influence of the fluorophore on the binding characteristics. Moreover, FP is a ratiometric readout and is thus less sensitive to autofluorescence and quenching artifacts by test samples (Levine et al., 1997; Owicki, 2000). Fluorescent compound interference with a readout can be minimized by using red-shifted dyes such as Cy3B (excitation and emission at 530 and 580 nm respectively) and Cy5 (excitation and emission at 640 and 680 nm respectively) (Turek-Etienne et al., 2003). One major disadvantage of the FP method is that the FP value is basically calculated from two separate intensity measurements. Hence, small changes in FP occur as small changes on already high signals. The resulting sensitivity of this readout is usually sufficient for applications like HTS in which a graduation of the readout in 10% steps is adequate. Other applications like the compound profiling require smaller graduations. In these cases, the resolution of an FP measurement conducted with a standard microtiter plate (MTP) reader may be too inaccurate.

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