Amplified fluorescence sensing of protease activity with conjugated polyelectrolytes

Abstract

Fluorescent conjugated polyelectrolytes with pendant ionic sulfonate and carboxylate groups are used to sense protease activity. Inclusion of the fluorescent conjugated polyelectrolyte into the assay scheme leads to amplification of the sensory response. The sensing mechanism relies on an electrostatic interaction between the conjugated polyelectrolyte and a peptide substrate that is labeled with a fluorescence quencher. Enzyme activity and hydrolysis kinetics are measured in real time by using fluorescence spectroscopy. Two approaches are presented. In the first approach, a fluorescence turn-on sensor was developed that is based on the use of p-nitroanilide-labeled peptide substrates. In this system enzyme-catalyzed peptide hydrolysis is signaled by an increase in the fluorescence from the conjugated polyelectrolyte. The turn-on system was used to sense peptidase and thrombin activity when the concentrations of the enzyme and substrate are in the nanomolar regime. Kinetic parameters were recovered from real-time assays. In the second approach, a fluorescence turn-off sensor was developed that relies on a peptide-derivatized rhodamine substrate. In the turn-off system enzyme-catalyzed peptide hydrolysis is signaled by a decrease in the fluorescence intensity of the conjugated polyelectrolyte.

Water-soluble, fluorescent π-conjugated polyelectrolytes (CPEs) afford a unique platform for the development of highly sensitive fluorescence-based sensors for biological targets (1–4). Signal amplification in CPEs results from a combination of factors, including delocalization and rapid transport of the singlet exciton along the π-conjugated backbone, along with the propensity of oppositely charged excited-state quenchers to ion-pair with the ionic groups attached to the CPE chains (1, 5–7). In effect, the CPE acts as a highly effective “antenna” that channels excitation energy to a quencher trap site that is ion-paired to the CPE. Chemo-optical signal amplification as high as 1,000-fold has been achieved with CPEs, as assessed by the ability to detect molecular fluorescence quenchers (1, 5).

By taking advantage of the intrinsic fluorescence signal-amplification properties of CPEs, several groups have recently developed sensitive assays for biologically relevant targets including proteins (1, 8, 9), DNA (3, 10–12), glycopeptides (13), and carbohydrates (14). Typical CPE-based fluorescence assays allow detection of the target analytes in the nanomolar concentration range (1, 3, 10), and, in a few cases, detection limits in the picomolar range have been reported (11). The CPE-based assays share the common features of being relatively easy to implement and giving a rapid response. Moreover, because the response is detected by using fluorescence, the systems can be adopted to a format that is compatible with f luorescence-based high-throughput screening (HTS) assays (15).

In most cases, CPE-based bioassays take advantage of a competition between a nonspecific ion-pairing of an ionic quencher moiety to the CPE chain and the highly specific recognition and binding of a “ligand” that is covalently bound to the quencher unit to a receptor target. For example, in one approach, a cationic viologen quencher was covalently linked to biotin, and this quencher-tether-ligand assembly was used to detect biotin-avidin or biotin-strepavidin binding (1, 13). In another approach, a quencher-labeled peptide nucleic acid sequence was used as a quencher-ligand to detect a complementary single-stranded DNA sequence (3).

Enzymes present another important target for bioanalytical chemistry because of their importance in a variety of biochemical processes. For example, proteases play critical roles in physiological and pathological processes, such as protein catabolism, blood coagulation, cell growth and migration, protein activation, cell regulation and signaling, tissue development, inflammation, tumor growth, metastasis, and pathogenesis (16, 17). Rapid, sensitive assays are needed for HTS of enzyme inhibitors as potential therapeutic drugs. Dependable HTS assays must have low detection limits and ideally should provide real-time readout of enzyme kinetics. Colorimetric and fluorescence-based homogeneous assays have been validated for a variety of proteases (18). These assays rely on the development of color or fluorescence in solution as a result of substrate hydrolysis. Although these methods are being widely used, in general, their sensitivity is limited to the micromolar or submicromolar range; consequently, measurement of initial rates at ultralow substrate concentrations or at low enzyme activity can be problematic. Thus, improvement of sensitivity and shorter readout times of HTS assays is an important goal.

To address the need for improved sensitivity in proteolytic enzyme assays, we implemented a general approach to chemooptical signal amplification that uses CPEs. In the present report, we describe two different approaches to this problem. Both approaches use anionic CPEs as the signal-transduction element along with readily available quencher-labeled peptides. The assays are carried out in solution, they provide a real-time signal, and they can also allow determination of enzyme kinetic parameters at very low substrate and/or enzyme concentrations. The assays are carried out by using a conventional bench-top fluorescence spectrometer, but the format is such that it can be easily adapted to a standard 96-well microliter plate for implementation in HTS assays.

Materials and Methods

Materials. PPESO3 and PPECO2 were synthesized according to methods described in ref. 7. All solutions were prepared by using water that was distilled and then purified by using a Millipore purification system (Millipore Simplicity Ultrapure Water Systems). Buffer solutions were prepared with reagent-grade materials (Sigma-Aldrich). Concentrated aqueous solutions of the polymers were diluted with appropriate buffers to a final concentration of 1 μM. Stock solutions of the enzymes in the appropriate buffer were prepared immediately before their use in the fluorescence assays. The enzyme solutions were maintained at 2–4°C before use, and they were always used within 4 h of preparation. (The concentration of all enzyme stock solutions was adjusted to 0.5 mg·ml^-1.) Papain (lyophilized powder, 14 units·mg^-1, Sigma-Aldrich) was dissolved in activating buffer (1.0 mM sodium phosphate/5.0 mM EDTA/5.0 mM cysteine, pH 6.0); the assays were conducted in 1.0 mM phosphate buffer containing 0.01 wt% of mercaptoethanol (pH 7.1). Peptidase (porcine intestinal mucosa, lyophilized powder, 50–100 units·g^-1, Sigma-Aldrich) was dissolved in 1.0 mM sodium phosphate (pH 7.1); the assays were conducted in the same buffer solution. Thrombin (bovine plasma, Factor IIa, lyophilized powder, 1050 NIH units·mg^-1) was diluted in 1.0 mM Tris buffer and 0.01 wt% of PEG6000 (pH 8.0); the assays were conducted in the same buffer. The enzyme substrates N,N′-bis(carboxybenzyloxy-l-arginine amide)rhodamine-110 dihydrochloride (Rho-Arg-2, Molecular Probes), N-benzoyl-Phe-Val-Arg-p-nitroanilide hydrochloride hydrate (Bz-FVR-pNA, Sigma-Aldrich) and l-Lys-p-nitroanilide dihydrobromide (K-pNA, Sigma-Aldrich) were obtained from commercial sources and were used as received. Concentrated stock solutions of the enzyme substrates were prepared in DMSO solution and were diluted in pure water before enzyme assays to a final concentration of 100 nM.

Fluorescence Assays. All enzyme assays were conducted at 37°C. A 3.0-ml aliquot of polymer solution was placed in a poly(methyl methacrylate) cuvette with a 10-mm path length, and the initial fluorescence intensity was measured after the sample was allowed to thermally equilibrate. The substrate was then added, the solution was incubated for 10 min, and the fluorescence intensity was again recorded. An aliquot of the enzyme solution was then added, and the fluorescence intensity was recorded at 10-s intervals. For PPESO3-based assays, the fluorescence intensity was acquired with excitation and emission wavelengths of 420 and 520 nm, respectively; and for PPECO2-based assays excitation and emission wavelengths of 400 and 460 nm, respectively. When applicable, fluorescence intensities were converted to substrate concentration as a function of time by using Eq. 1 which is derived from the Stern–Volmer equation (19).

[1]

In Eq. 1 [Q]_t is the quencher concentration at time t, [Q]₀ is the initial quencher concentration, I₀ is the initial fluorescence intensity of the polymer solution, I_Q is the fluorescence intensity after addition of quencher and before the addition of enzyme, and I_t is the fluorescence intensity at time t after enzyme addition. The initial reaction rate was calculated by the tangent method by using the first five acquisition points.

Results and Discussion

Overview of Approaches. The structures of the two CPEs used in the present work are illustrated in Fig. 1a. PPESO3 is an anionic poly(phenylene ethynylene) (PPE) that features sulfonate side groups. In aqueous solution the polymer absorbs with λ_max = 445 nm and it exhibits an intense yellow-green fluorescence (λ_max = 530 nm, ϕ_fl = 0.10). PPECO2 is also an anionic PPE-type polymer which absorbs with λ_max = 425 nm and it features strong, blue fluorescence (λ_max = 470 nm, ϕ_fl = 0.20). PPESO3 and PPECO2 are quenched strongly by a variety of cationic small-molecule quenchers (e.g., dimethyl viologen and cyanine dyes); values of K_SV = 10⁷ M^-1 are typically observed (7, 20).

Fig. 1.

(a) Structures and acronyms for polymers and quencher substrates. (b) Mechanism of the “turn-on” and “turn-off” CPE-based sensors.

In the course of this investigation two different approaches were developed to monitor proteolytic enzyme activity (Fig. 1b). In the first approach (fluorescence turn-on), a cationic peptide (the enzyme substrate) labeled with a p-nitroanilide (p-NA) unit is used in conjunction with the anionic CPE PPESO3. The p-NA moiety is a strong quencher of PPESO3 fluorescence; and, because the cationic peptide ion-pairs with the polyelectrolyte, the p-NA-labeled peptide quenches the CPE fluorescence even at concentrations in the nanomolar range. Introduction of a proteolytic enzyme to a mixture of the p-NA peptide and PPESO3 induces hydrolysis of the p-NA group; and, because this moiety is not charged, its ability to quench the PPESO3 fluorescence is eliminated. This outcome results in an increase in fluorescence from PPESO3 concomitant with peptide hydrolysis. By using Eq. 1 it is possible to determine the substrate concentration during hydrolysis, thereby allowing one to measure enzyme-catalyzed reaction kinetics. (Although not specifically shown in Fig. 1b, it is likely that the enzyme-catalyzed hydrolysis reaction requires that the substrate-quencher is free in solution, i.e., it is not ion-paired with the CPE. However, this requirement does not interfere with the enzyme-catalyzed reaction because the substrate-quencher exists in equilibrium between free and ion-paired forms.)

The second approach (fluorescence turn-off) is demonstrated by using PPECO2 combined with a “caged” peptide substrate-quencher. In particular, the bis-Arg derivative of Rhodamine-110 (Rho-Arg-2, Fig. 1a) is colorless and nonfluorescent; it does not quench the fluorescence of PPECO2. However, the monoamide derivative (Rho-Arg) produced by hydrolysis of one amide linkage in Rho-Arg-2 is an “uncaged” quencher. Rho-Arg absorbs strongly between 450 and 530 nm; and, because it is positively charged and has good spectral overlap with PPECO2 fluorescence, it strongly quenches the PPECO2 fluorescence by resonance energy transfer. Introduction of a protease into a mixture of PPECO2 and Rho-Arg-2 induces amide hydrolysis, and in the first step of this process the Rho110-Arg quencher is formed, resulting in a decrease in the polymer's fluorescence. Because Rho110-Arg-2 can be used directly as a fluorescent assay for enzyme hydrolysis (i.e., the hydrolysis product, Rho-Arg, is fluorescent), it is possible to demonstrate the signal amplification afforded by inclusion of PPECO2 into the assay.

Turn-On Approach. This approach to enzyme activity sensing is demonstrated by using an array consisting of two commercially available p-NA-derivatized peptide substrates, K-pNA and Bz-FVR-pNA, and two proteolytic enzymes, peptidase and thrombin. Peptidase is a nonspecific aminopeptidase that cleaves terminal lysine peptide bonds (21, 22). In a preliminary experiment carried out with UV-visible absorption to assay for the release of free p-nitoaniline, we demonstrated that K-pNA is cleaved by peptidase. Thus, as shown below, experiments were carried out with the system comprising K-pNA/PPESO3 and peptidase. Thrombin paired with the tripeptide substrate Bz-FVR-pNA was chosen because the kinetic parameters have been reported for this system (23), allowing us to evaluate the accuracy of the kinetic data that are recovered from the CPE-based assay. Thrombin is a serine endopeptidase that selectively cleaves Arg-Gly bonds in fibrinogen to form fibrin and release fibrinopeptides A and B (24). Thrombin also cleaves Arg-aromatic amine peptide bonds in small synthetic substrates (25), and it has been shown that Bz-FVR-pNA is a good substrate for this protease (23).

The sensitivity of the PPESO3/p-NA peptide/enzyme system is determined, in part, by the efficiency by which the p-NA peptide quenches the fluorescence of PPESO3. To determine the sensitivity, Stern–Volmer quenching experiments were carried out by using the peptide substrates in buffer solutions of varying concentration. Table 1 compiles the Stern–Volmer quenching constants (K_SV) obtained for quenching of PPESO3 by the two p-NA substrates used in the assays. Two trends are clear from these data: (i) K-pNA is a more efficient quencher than Bz-FVR-pNA, and (ii) the quenching efficiency for both peptides decreases with increasing buffer concentration.

Table 1. Stern–Volmer quenching of PPESO3 by p-NA peptides.

	Ksv, M-1
[Buffer], mM	K-pNA*	Bz-FVR-pNA†
1.0	1.7 × 107	2.3 × 106
5.0	4.4 × 106	1.2 × 106
10.0	4.3 × 106	1.1 × 106

PPESO3 concentration is 1.0 μM.

^*Phosphate buffer

^†Tris buffer

These effects are understandable because the primary mechanism for fluorescence quenching is static quenching due to an ion-pair complex formed between the cationic peptide and the anionic polyelectrolyte.† At pH 7, K-pNA is a dication whereas Bz-FVR-pNA is a monocation; therefore, it is to be expected that the association constant between K-pNA and PPESO3 will be larger. The effect of increased ionic strength on K_SV arises due to an electrostatic screening effect. This effect is important because many enzymes assays are carried out in the presence of buffer. To minimize the effect of the added salt on the sensitivity of the PPESO3 fluorescence to quenching by the p-NA peptides, the assays were carried out with buffer concentrations of 1.0 mM.

Fig. 2a shows the fluorescence spectral changes observed in a typical PPESO3-based fluorescence turn-on protease assay. The initial fluorescence from PPESO3 (c = 1 μM, solid line in Fig. 2a) is quenched ≈75% by addition of K-pNA (c = 167 nM, dotted line). After addition of peptidase to the solution (c = 3.3 μg·ml^-1) hydrolysis of K-pNA is signaled by an increase in the PPESO3 fluorescence intensity with increasing incubation time. A subtle blue shift in the PPESO3 fluorescence that is observed after addition of peptidase to the PPESO3/K-pNA solution may arise due to nonspecific interactions between the polymer and the protein (26, 27).

Fig. 2.

(a) Fluorescence spectroscopic changes observed during turn-on assay. (—) Initial fluorescence, [PPESO3] = 1.0 μM, phosphate buffer solution, pH = 7.1; (•••) fluorescence after addition of 167 nM K-pNA. Fluorescence intensity as a function of time after addition of porcine intestinal peptidase (c = 3.3 μg·ml^-1): 5, (– – –) 20, (–·–) 50, (– ·· –)70,(- - - - -) 100, (– – –) 140, (......) and 200 (–·–) min. (b) Fluorescence spectroscopic changes observed during turn-off assay. (—) Initial fluorescence, [PPECO2] = 1.0 μM, [Rho-Arg-2] = 500 nM, phosphate buffer solution, pH = 7.1, 0.01% mercaptoethanol. Fluorescence intensity as a function of time after addition of 3.5 nM papain: 1, (– – –) 3, (- - - - -) 6, (–·–) 18, (– ·· –) and 28 (......) min.

To demonstrate the suitability of the fluorescence turn-on method for acquisition of kinetic data, a series of assays was carried out in which enzyme and substrate concentrations were varied. Similar results were obtained from assays carried out with peptidase and thrombin; therefore, only the thrombin assay data are presented herein. In these experiments the PPESO3/Bz-FVR-pNA/thrombin system was used. Kinetics experiments were carried out by monitoring PPESO3 fluorescence intensity, and the time-dependent intensity data were subsequently used to determine Bz-FVR-pNA concentration by using Eq. 1. First, the dependence of the hydrolysis rate on thrombin concentration was determined while maintaining the initial Bz-FVR-pNA substrate concentration constant at 490 nM. Fig. 3a illustrates plots of log ([Bz-FVR-pNA]) vs. time for five thrombin concentrations ranging from 2.6 to 13.0 nM; V_max/K_M ratios were calculated from the slope of the plots for each enzyme concentration. As expected, the plot of V_max/K_M vs. [thrombin] is linear (Fig. 3a Inset) (28), and the slope of the plot affords a value of k_cat/K_M = 0.86 liter·μmol^-1·s^-1. This value is in good agreement with a value obtained by using the same substrate, but by monitoring the hydrolysis kinetics by absorption spectroscopy (0.53 liter·μmol^-1·s^-1) (23). This experiment demonstrates that by using the PPESO3 fluorescence-based assay it is possible to detect the proteolytic activity of thrombin within 100 s for enzyme concentrations ≥100 ng·ml^-1 (2.7 nM or 0.1 NIH units·ml^-1). This response is considerably more rapid and is nearly 100-fold more sensitive than a previously reported fluorometric assay for thrombin activity (29). Extending the incubation time to 50 min reduces the detection limit for thrombin activity to 1.8 ng·ml^-1 (50 pM).

Fig. 3.

Enzyme kinetics measured by using PPESO3/Bz-FVR-pNA/thrombin assay system. (a) Concentration of Bz-FVR-pNA substrate as a function of incubation time plotted for various thrombin concentrations. Conditions: [PPESO3] = 1.0 μM, Tris buffer solution, pH = 8.0, 0.01% PEG 6000, [Bz-FVR-pNA] = 490 nM. [Thrombin]: •,0; ▴, 2.6; ▪, 4.3; ▾, 6.9; ♦, 8.7; and ○, 13.0 nM. (Inset) Initial hydrolysis rate as a function of [thrombin]. (b) Concentration of hydrolyzed peptide substrate as a function of time at various initial Bz-FVR-pNA concentrations. [Thrombin] = 8.7 nM; [Bz-FVR-pNA]: •,0; ▴, 147; ▪, 392; ▾, 490; ♦, 980; and ○, 1,470 nM.

In a second series of experiments the thrombin-catalyzed hydrolysis of Bz-FVR-pNA substrate was monitored as a function of the initial substrate concentration (Fig. 3b). In these experiments the initial substrate concentration ranged from 0.1 to 1.5 μM, and the thrombin concentration was 8.7 nM. Despite the low and relatively narrow substrate concentration range used in this set of experiments, it is clearly possible to resolve the differences in the signal for the various samples within a 100-s incubation time. In addition, it is clear that the initial rate increases with substrate concentration, as expected because, in all cases, the substrate concentration is less than K_M (K_M = 72 μM; ref. 23).

p-Nitroaniline-labeled peptides are often used in absorption-based assays for enzyme activity. These assays rely on the use of absorption spectroscopy to detect p-nitroaniline that is produced by enzyme-catalyzed hydrolysis. Given that p-nitroaniline has ε = 8,800 liter·mol^-1·cm^-1 at 410 nm, (25) and assuming a p-NA substrate concentration of 1.5 μM (the maximum concentration used in Fig. 2b), the maximum absorbance change possible for the assays described here would be 0.013 AU (for a 1-cm path length). This absorption change is close to the detection limit of a typical bench-top absorption spectrometer. Although this comparison is qualitative, it is evident that the use of the PPESO3-based fluorescence assay affords an increase in sensitivity by 2 or more orders of magnitude relative to the p-NA-based colorimetric method, allowing assays to be carried out at significantly lower initial p-NA substrate concentration, but still affording reliable real-time monitoring of the hydrolysis reaction.

Turn-Off Approach. This approach to a CPE-based enzyme assay was demonstrated by using a system consisting of PPECO2, Rho-Arg-2, and the proteolytic enzyme papain. PPECO2 is used rather than PPESO3 in this assay, because its fluorescence is blue-shifted relative to that of PPESO3. Because of the blue-shift, PPECO2 has improved spectral overlap with the absorption of the rhodamine chromophore (30, 31), which leads to an overall improvement in the sensory response for the polymer/Rho-Arg-2 system. (The improved sensitivity is believed to result from the fact that the quenching phenomenon responsible for the sensory response is singlet–singlet energy transfer from PPECO2 to Rho-Arg, see below.)

Fig. 2b illustrates the changes that occur in the fluorescence spectrum of the PPECO2/Rho-Arg-2/papain turn-off assay before and during incubation with the enzyme. At the outset of the experiment, a solution of PPECO2 (c = 1 μM) and Rho-Arg-2 (c = 500 nM) when excited at 400 nm exhibits a strong fluorescence with λ_max ≈470 nm. Immediately after addition of papain (83 ng·ml^-1, c = 3.5 nM), a pronounced decrease in the PPECO2 fluorescence intensity is observed. The intensity continues to decrease for ≈30 min whereupon further changes in the intensity are not observed. Note that in the spectrum obtained after 28 min of incubation the fluorescence exhibits an additional band at λ_max ≈515 nm; this fluorescence arises from Rho-Arg, which is produced by the hydrolysis reaction (30, 31). Because Rho-Arg does not absorb at the excitation wavelength (400 nm), the fluorescence that is observed from this species is sensitized by PPECO2, confirming that the polymer fluorescence-quenching mechanism is singlet–singlet energy transfer from PPECO2 to Rho-Arg.

To demonstrate the application of the turn-off sensor to real-time monitoring of papain activity, a series of assays was carried out by using the PPECO2/Rho-Arg-2/papain system at various concentrations of the Rho-Arg-2 substrate. Fig. 4a illustrates the PPECO2 fluorescence intensity as a function of incubation time for assays in which the papain concentration was fixed (166 ng·ml^-1, c = 7 nM) and the Rho-Arg-2 substrate concentration was varied over the range 17–333 nM. Note that the reactions are essentially complete within a 15-min incubation period. It is evident that even for [Rho-Arg-2] < 20 nM, it is possible to detect papain-catalytic activity within a 2-min incubation period, demonstrating the high sensitivity and rapid response of the turn-off sensor system.

Fig. 4.

PPECO2/Rho-Arg-2/papain assay. (a) Fluorescence intensity as a function of time for various initial Rho-Arg-2 substrate concentrations. Conditions: [papain] = 7.0 nM, [PPECO2] = 1.0 μM, phosphate buffer solution, pH = 7.1, 0.01% mercaptoethanol; initial [Rho-Arg-2]: •,0; ▴, 16.7; ▪, 66.7; ▾, 167.7; and ♦, 333.3 nM. (b) Comparison of change in fluorescence intensity (ΔI_flr) for Rho-Arg-2/papain (□) and PPECO2/Rho-Arg-2/papain () assays; initial [Rho-Arg-2] = 333.3 nM.

With the Rho-Arg-2 substrate it is also possible to use fluorescence to directly monitor its hydrolysis (30, 31). This is because the hydrolysis product Rho-Arg is fluorescent, whereas the substrate Rho-Arg-2 is only weakly fluorescent (30, 31). Thus, to compare the response of PPECO2/Rho-Arg-2/papain system with “direct” fluorescence-based detection, assays were carried out in which the change in fluorescence intensity was determined after a 3-min incubation period (ΔI_flr) for the PPECO2/Rho-Arg-2/papain and Rho-Arg-2/papain systems. In both experiments, the substrate concentration was the same (333 nM), but the papain concentration was varied (3.5 nM and 7.0 nM). The parameters used on the fluorescence spectrometer were the same, except the fluorescence excitation/detection wavelengths were at 400/460 nm for PPECO2 and at 500/520 nm for Rho-Arg-2. Fig. 4 compares the results of the assays, and in both cases it can be seen that the change in intensity for the PPECO2-based papain activity sensor is six to ten times larger than when the fluorescence from Rho-Arg is detected directly. Because of the difference in spectrometer wavelengths it is not possible to provide a quantitative measure of the increase in sensitivity afforded by addition of the CPE to the assay (mainly because of the wavelength dependence of the excitation source intensity), it is evident from this comparison that signal amplification is achieved with the CPE system, and, consequently, the overall sensitivity of the polymer-based system is better than that afforded by direct fluorescence detection of the Rho-Arg product. Note that rhodamine dyes are widely used in high-sensitivity, fluorescence-based assays, primarily because of their high fluorescence quantum efficiency (and good photostability). Thus, the fact that signal amplification relative to the Rho-Arg-2 benchmark is observed suggests that the CPE-based turn-off sensor provides a signal that is higher than the best fluorescence-based enzyme activity sensors that are presently available.

Conclusions and Outlook

This work demonstrates the application of fluorescent CPEs to real-time detection of proteolytic enzyme activity. The turn-on sensor allows direct measurement of enzyme-catalyzed reaction kinetics, even at very low substrate and enzyme concentrations. The turn-off sensor affords sensitivity similar to the turn-on sensor, and benchmarking of this system relative to the “conventional” assay based on the Rho-Arg-2 substrate indicates that the CPE affords at least a 10-fold signal enhancement.

Although this work demonstrates the method by using specific proteolytic enzymes and substrates, the method is quite general and has many benefits. Specifically, the turn-on method works with p-NA peptide derivatives; these derivatives are easily prepared and many are commercially available because they provide the basis for colorimetric assays. In addition, the method can be generalized to p-NA peptides that are negatively charged by using a positively charged CPE (32–34). The turn-on approach is not limited to p-NA peptides, because a variety of species strongly quench CPE fluorescence, e.g., p-nitrophenyl esters, azobenzene dyes, and diimides. This leads to the possibility of using different types of substrates for the CPE-based enzyme activity sensor. Finally, the method is clearly not limited to detection of protease activity. Thus, it can be extended to develop assays for a variety of enzyme-catalyzed processes, e.g., esterases, kinases, phosphatases, lipases, oxidases, etc. Because of the broad applicability of the sensor, it can also likely be used to amplify the optical signal generated in ELISA immunoassays.