Table of Contents Author Guidelines Submit a Manuscript
Journal of Analytical Methods in Chemistry
Volume 2016 (2016), Article ID 4306838, 10 pages
http://dx.doi.org/10.1155/2016/4306838
Research Article

Construction of a Turn Off-On-Off Fluorescent System Based on Competitive Coordination of Cu2+ between 6,7-Dihydroxycoumarin and Pyrophosphate Ion for Sensitive Assay of Pyrophosphatase Activity

1Department of Pharmacy, Xi’an Medical College, Xi’an 710021, China
2Laboratory of Analytical Chemistry for Life Science of Shaanxi Province, School of Chemistry and Chemical Engineering, Shaanxi Normal University, Xi’an 710062, China
3Beijing Research Center of Agricultural Standards and Testing, Beijing 100097, China

Received 12 June 2016; Revised 25 August 2016; Accepted 5 September 2016

Academic Editor: Jaroon Jakmunee

Copyright © 2016 Lingzhi Zhao et al. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

The detection of pyrophosphatase (PPase) activity is of great significance in diagnosing diseases and understanding the function of PPase-related biological events. This study constructed a turn off-on-off fluorescent system for PPase activity assay based on PPase-regulated competitive coordination of Cu2+ between a water-soluble fluorescent probe 6,7-dihydroxycoumarin (DHC) and pyrophosphate (PPi). The probe DHC can coordinate with Cu2+ and consequently display on-off type fluorescence response. Furthermore, the in situ formed nonfluorescent Cu2+-DHC complex can act as an effective off-on type fluorescent probe for sensing PPi due to the higher coordination reactivity between Cu2+ and PPi than that between Cu2+ and DHC. The subsequent addition of PPase to the mixture containing Cu2+, DHC, and PPi leads to the fluorescence requenching of the system again (an off state) because PPase catalyzes the hydrolysis of PPi into orthophosphate in the reaction system. Under the optimum conditions, the decrease of the fluorescence intensity of DHC-Cu2+-PPi system was linear with the increase of the PPase activity in the range from 0.1 to 0.3 U. The detection limit was down to 0.028 U PPase (). Moreover, the as-established system was also applied to evaluate PPase inhibitor. This study offers a simple yet effective method for the detection of PPase activity.

1. Introduction

As one kind of the most important biological anions, pyrophosphate (, PPi) is one of the by-products of adenosine triphosphate (ATP) hydrolysis and can be used as a potential biomarker for the clinic diagnosis and therapy of familial chondrocalcinosis or calcium pyrophosphate crystal deposition disease [14]. Inorganic pyrophosphatase (PPase) is a kind of ubiquitous enzyme that specially hydrolyzes pyrophosphate (PPi) to orthophosphate (Pi), providing a thermodynamic pull for many different biosynthetic reactions, and has been demonstrated to be directly relevant to phosphorus metabolism, carbohydrate metabolism, and evolutionary events [57]. Moreover, many diseases, such as hyperthyroidism, colorectal cancer, and lung adenocarcinomas, are associated with the expression level and the abnormal regulation of PPase [8, 9]. Therefore, there is an urgent demand for developing technically simple yet effective methods for real-time PPase activity monitoring. Up to date, various analytical methods and techniques have been developed for PPase activity assay, such as radioactive label-based chromatography, colorimetry assay, and fluorescence methods a [1016]. Most of these methods are based on monitoring the change in the amount of the PPi substrate or Pi product. For instance, Deng et al. [13] demonstrated a new colorimetric method for real-time PPase activity assay based on reversible tuning of the dispersion/aggregation states of gold nanoparticles (Au-NPs) by controlling the coordination of Cu2+ between cysteine and PPi with PPase. Zhang et al. [14] developed a convenient colorimetric strategy for PPase activity detection based on an inhibition effect of PPi on the Cu2+-catalyzed ABTS-H2O2 reaction. Therefore, the key to developing PPase detection methods is the development of effective methods for the sensing of the PPi substrate or Pi product.

For PPi detection, since PPi is usually chelated with metal ions to form coordination complexes, hence, metal complex-based chemosensors based on metal ion-ligand coordination interaction are highly interesting to researchers [1727]. This strategy is based on the metal ion displacement of the metal complex by PPi due to the stronger coordination interaction between metal ion and PPi. Lin et al. [24] presented an electrochemical method for the determination of PPi that is based on the competitive coordination of Cu2+ between a nanofilm of cysteine and dissolved PPi. It was reported that the stability constant () of the complex formed by Cu2+ and PPi is [28], suggesting that the coordination ability of PPi with Cu2+ is stronger than that PPi with other divalent metal ions. Therefore, in this study, with the purpose to simplify the assay process for manufacturing portable and affordable devices, we constructed a turn off-on-off fluorescent system in aqueous media on the basis of competitive coordination of Cu2+ between water-soluble organic dyes 6,7-dihydroxycoumarin (DHC) and PPi for quantitatively analyzing PPase activity.

Fluorescence methods possess innate advantages over other methods, such as designability of optical probes, the simplicity of implementation, high sensitivity, excellent selectivity, and fast response time. Herein, an effective fluorescence method for highly sensitive assay of PPase activity has been developed in the physiological buffer, which utilizes a simple competitive coordination reactivity of Cu2+ between DHC and PPi. As shown in Figure 1, initially, the strong fluorescence emission peak of DHC was observed at 475 nm. Upon the presence of Cu2+, the complex between Cu2+ and DHC was formed through the coordination interaction of the O atom in hydroxy groups of DHC and Cu2+, leading to the fluorescence quenching of DHC (an off state). Further addition of PPi to the dispersion of Cu2+-DHC complex enables the fluorescence emission intensity of DHC restored substantially (an on state) owing to the higher binding affinity between PPi and Cu2+ than that between DHC and Cu2+. On the other hand, the addition of PPase to the Cu2+-PPi complex containing DHC obviously leads to requenching the fluorescence (an off state), because PPase could catalyze the hydrolysis of PPi into Pi (herein, Pi must be in pH 6.5), which cannot chelate Cu2+ to form the complex and thus Cu2+ was released. To the best of our knowledge, the as-designed turn off-on-off fluorescence sensing system is convenient without complicated protocols and exhibits excellent PPi sensing selectivity over other anions and structural analogues and excellent performances for sensitive assay of PPase activity in the physiological buffer compared to that of the existing methods, which paves a fluorescence route to the fast and simple clinic detection of PPi, the hydrolytic activity of PPases, and screening of PPase inhibitors.

Figure 1: (a) Schematic illustration of the turn off-on-off fluorescent strategy based on competitive coordination of Cu2+ between DHC and PPi for PPase activity assay. (b) Fluorescence spectra of the mixtures prepared by separate addition of 500 μM DHC; 500 μM DHC and 200 μM Cu2+; 500 μM DHC and 200 μM Cu2+ and 200 μM PPi; 500 μM DHC and 200 μM Cu2+ and 200 μM PPi and 0.25 U PPase. Buffer, HEPES buffer aqueous solution (10 mM, pH 6.5); the reaction temperature, 25°C; cuvette width: 1 mm.  nm.

2. Experimental

2.1. Materials and Instrumentation

6,7-Dihydroxycoumarin (DHC), adenosine 5′-triphosphate disodium salt hydrate (ATP), adenosine 5-diphosphate sodium salt (ADP), adenosine 5-monophosphate monohydrate (AMP), glutathione (GSH), L-cysteine (Cys), and homocysteine (Hcy) were purchased from Aladdin Company (China). Potassium pyrophosphate (PPi), Na2SO4, NaCl, KNO3, Na2SO3, Na2S2O3, NaH2PO4, Na2HPO4 (Pi), Na3PO4, Zn(NO3)2, Fe(NO3)3, Cu(NO3)2, CaCl2, MgCl2, NaF, and other inorganic salts were purchased from Sinopharm Chemical Reagent Co., Ltd. (China). Bovine serum albumin (BSA) was obtained from Shanghai Sangon Biological Engineering Technology and Services Co., Ltd. (China). Prostate specific antigen (PSA) was obtained from Fitzgerald Industries International, Inc. (USA). Human immunoglobulin G (IgG) was obtained from Beijing Middle Mountain Gold Bridge Biological Technology Co., Ltd. (China). Albumin chicken egg protein was obtained from Sino-American Biotechnology Co., Ltd. (China). Baker’s yeast inorganic pyrophosphatase (PPase, EC 3.6.1.1) was purchased from Sigma (USA). One unit of Baker’s yeast PPase liberates 1.0 μmol of inorganic orthophosphate/min at pH 7.2 and 25°C. All other chemicals were at least analytical grade reagents and used without further purification. HEPES buffer aqueous solution (10 mM, pH 6.5) was prepared by adjusting commercial 10.0 mM HEPES to its pH of 6.5 using 2 M NaOH. All aqueous solutions were prepared with Milli-Q water (18.2 M Ω·cm−1). Unless otherwise pointed out, all experiments were carried out at room temperature.

The absorbance spectra and photoluminescence (PL) spectra were recorded on a UV-Vis spectrophotometer (UV-2450, Shimadzu Corporation, Japan) and a Cary Eclipse fluorescence spectrophotometer (Varian, USA), respectively.

2.2. UV-Vis and Fluorescence Experiments

The detection measurements were conducted as follows: first, the stock solution of each reactant was prepared by dissolving the compound in HEPES buffered aqueous solution (10 mM, pH 6.5). After that, the solution of DHC, Cu2+, PPi, or other reactants was sequentially added; the final volume is 500 μL. The resulting mixture was then incubated in a 25°C water bath for 30 min. The UV-Vis and fluorescence spectra were recorded immediately. The present concentration of each reactant was the final concentration.

2.3. PPase Activity Assay

PPase activity assay was performed under the following procedures. A 5 μL volume of PPase with different activities ranging from 0.1 to 1 U was added to the aqueous dispersion consisting of 500 μM DHC, 200 μM Cu2+, 200 μM PPi, and 500 μM Mg2+. Fluorescence spectra of the dispersions were consecutively recorded in the different time at 25°C. The final volume is 500 μL.

2.4. Phosphate Assay for PPase Activity Based on Colorimetric Method of Molybdenum Blue [29]

The reaction was started by mixing 0.25 U PPase and the substrate PPi and activating agent Mg2+ in a 1.5 mL microfuge tube (the final concentration of PPi and Mg2+ is 500 μM). In order to measure effects of Cu2+ and DHC on PPase activity, 500 μM DHC or 200 μM Cu2+ was added in the above-mentioned reactant solution. After 30 min, the reaction was stopped by adding 35 μL of SDS-EDTA (13.3% SDS, 0.12 M EDTA, pH 6.5). And then, 265 μL of color development reagent (0.5% Fe2SO4, 0.5% ammonium molybdate, and 0.5 M H2SO4) was added into the tube. The final volume is 400 μL. Color was allowed to develop for 15 min and read in a microplate reader at 650 nm (EMax Plus Microplate Reader, Molecular devices, USA).

2.5. Inhibitor Efficiency Evaluation

For investigating the inhibition efficiency of NaF on PPase activity, 5 μL of PPase (0.25 U) was initially added into 60 μL of aqueous solutions of NaF with different concentrations (100 nM, 1 μM, 5 μM, 10 μM, 50 μM, 100 μM, and 500 μM) in the presence of 500 μM Mg2+ in a 25°C water bath for 15 min. After 15 min, the NaF-treated PPase was added to the aqueous dispersion consisting of DHC (500 μM), Cu2+ (200 μM), and PPi (200 μM). After 30 min, Fluorescence measurements of the dispersions were obtained. The final volume is 500 μL.

3. Results and Discussion

3.1. Principle of Fluorometric Assay of Inorganic Pyrophosphatase Activity

To begin with, as displayed in Figure 1(a), the fluorescence spectrum of 6,7-dihydroxycoumarin (DHC) dispersion in HEPES buffer (10 mM, pH 6.5) was obtained by excitation of the fluorophore at 375 nm, and a strong emission peak was observed at 475 nm (an on state) (Figure 1(b), blue curve). Intriguingly, the addition of Cu2+ to this aqueous dispersion dramatically quenched the fluorescence of DHC through the coordination chemistry between Cu2+ and DHC (an off state) (Figure 1(b), black curve) and the formation of Cu2+-DHC complex. Further addition of PPi to the dispersion of Cu2+-DHC complex enabled the peak intensity of the emission spectrum of DHC restored substantially (an on state) (Figure 1(b), red curve) because of a stronger coordination between PPi and Cu2+ than that between DHC and Cu2+. On the other hand, the addition of PPase to the Cu2+-PPi complex containing DHC obviously leads to the fluorescence quenching again (an off state) (Figure 1(b), green curve) because Pi, which is produced from PPase-catalyzed hydrolysis of PPi, cannot chelate Cu2+ to form the complex. The large difference in the capability between PPi and other phosphate anions to compete with DHC in the coordination with Cu2+ will be investigated in subsequent fluorescence spectrum study, which actually forms the basis for construction of the turn off-on-off fluorescent system for sensitive assay of PPase activity as demonstrated in Figure 1.

3.2. The Feasibility of Fluorometric Assay

At first, the interaction of DHC with Cu2+ was investigated by UV-Vis spectrophotometric titration in HEPES buffer (10 mM, pH 6.5). Initially, the UV-Vis absorption spectrum of DHC displayed a strong absorbance band centered at about 340 nm, as shown in Figure 2(a), which is the characteristic absorption profile of DHC. The addition of Cu2+ to this aqueous dispersion produced a new absorption peak at about 375 nm. This phenomenon was elucidated by the coordination chemistry between Cu2+ and DHC; the O atom in two hydroxy groups of DHC can coordinate with Cu2+ to form the Cu-DHC complex. Such a coordination interaction eventually leads to the red shift of the absorption peak of DHC. In order to judge the coordination number of Cu-DHC complex, Cu2+ titration experiment was performed. As shown in Figure 2(a), upon the addition of increasing amounts of Cu2+, a new absorption peak at 375 nm increased and the absorption peak of DHC at 340 nm gradually decreased in the concentration of introduced Cu2+ (0-1 equiv. of Cu2+). Appearance of an isosbestic point at 355 nm also demonstrated the formation of a well-defined complex between DHC and Cu2+. When the concentration of Cu2+ increased from 1 to 10 equiv. (Figure 2(a), inset), the absorbance band was no longer changed noticeably, suggesting 1 : 1 stoichiometric complexation of DHC with Cu2+.

Figure 2: (a) UV-Vis absorption responses of DHC (500 μM) upon the addition of different concentrations of Cu2+ (0, 0.1, 0.2, 0.3, 0.4, 0.5, 0.6, 0.7, 0.8, 0.9, 1.0, 2.0, 3.0, 4.0, 5.0, 7.0, and 10.0 equiv.) in HEPES buffer aqueous solution (10 mM, pH 6.5). Inset: the corresponding changes of absorbance of DHC at 375 nm in the presence of different concentrations of Cu2+. (b) Fluorescence titration of DHC (500 μM) with Cu2+ (0, 10, 30, 50, 70, 100, and 200 μM) in HEPES aqueous buffer (10 mM, pH 6.5). Inset shows fluorescence intensity changes of DHC at 475 nm as a function of the Cu2+ concentration. Other reaction conditions are the same as Figure 1.

DHC exhibited the maximum emission at a wavelength of 475 nm in HEPES buffer (10 mM, pH 6.5) under the optimal fluorescence excitation at a wavelength of 365 nm (Figure 2(b)). It is clearly seen that the distinct fluorescence peak of DHC decreased markedly in the presence of Cu2+. This fluorescence quenching might be attributed to the coordination interaction between Cu2+ and DHC and the aggregation of DHC after Cu2+ reacted with DHC. To quantitatively evaluate the fluorescence response of DHC toward Cu2+, fluorescence titration with Cu2+ in varying concentrations was conducted. As depicted in Figure 2(b), with the increasing concentration of Cu2+, the fluorescence emission intensity of DHC decreased gradually and was directly proportional to the concentrations of Cu2+ in the range from 10 to 200 μM (, ). The fluorescence of DHC (500 μM) was quenched by approximately 95% with the presence of 200 μM Cu2+. Therefore, 200 μM Cu2+ was chosen in subsequent experiments.

To gain insight into the sensing properties of DHC, the selectivity of DHC for sensing Cu2+ was evaluated by comparing its fluorescence response toward Cu2+ with those of 13 other kinds of conventional metal ions including Fe3+, Fe2+, Zn2+, Al3+, Cr3+, Ag+, Pb2+, Cd3+, Co2+, Mg2+, Na+, K+, and Ca2+. The emission characteristics were examined in the presence of various metal species. As shown in Figure 3(b), most cations caused minor fluorescence intensity changes (the red columns in Figure 3(b)), and the quenching efficiency for 200 μM Cu2+ came to 95% while that for 6 other kinds of metal ions did not exceed 15%. 200 μM Fe3+ and Fe2+ caused the largest fluorescence intensity quenching and the quenching efficiency reached about 50%. But in the physiological conditions, such as in human serum, the concentration of Fe3+ and Fe2+ is quite low (5–30 μM), which did not interfere with our detection system. In contrast, Na+, K+, Mg2+, and Ca2+ (500 μM) induced fluorescence enhancement to a certain degree. Since Cu2+ demonstrated the largest fluorescence intensity quenching among these conventional metal ions, Cu2+ was employed to construct the on-off fluorescent system for PPase activity assay.

Figure 3: (a) Fluorescence changes of DHC (500 μM) in the absence (curve A) and the presence of various cations (curves B to O) in HEPES buffer aqueous solution (10 mM, pH 6.5). (b) Fluorescence responses of DHC (500 μM) at 475 nm in the absence (black bars) and presence of different cations (1 to 15: Cu2+, Fe3+, Fe2+, Zn2+, and Al3+ 200 μM; Cr3+, Pb2+, Ni2+, Cd3+, Co2+, and Ag+, 100 μM; Na+, K+, Mg2+, and Ca2+, 500 μM) (red bars). Other reaction conditions are the same as Figure 1.

To investigate the different concentration of PPi sensing in Cu2+-DHC complex system, fluorescence responses of Cu2+-DHC complex with different concentrations of PPi in HEPES buffer solutions (10 mM, pH 6.5) were recorded, as showed in Figure 4; it can be seen that, with the increasing concentration of PPi, the fluorescence emission intensity of DHC was restored gradually. This observation can be attributed to the fact that the coordination effect of PPi with Cu2+ is stronger than that of DHC with Cu2+ and DHC was released from Cu2+-DHC complex as demonstrated in Figure 1(a). The fluorescence emission intensity had the dependence on the concentrations of PPi in the range from 10 to 200 μM with the detection limit of 580 nM, which fit well for the detection PPi in some biological fluids, such as plasma and serum in which PPi was found to be around the micromolar range. Therefore, these results demonstrated that the in situ formed Cu2+-DHC complex could be a promising “off-on” type fluorescence sensor for the PPi based on the displacement approach. Additionally, 200 μM PPi can induce the fluorescence 90% recovery and thus was chosen in further assays.

Figure 4: Fluorescence responses of DHC (500 μM) (curve A) and Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex with PPi (curves B to G: 0, 10, 30, 50, 100, and 200 μM) in HEPES aqueous buffer (10 mM, pH 6.5). Inset shows fluorescence changes of Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex at 475 nm as a function of PPi concentration. Other reaction conditions are the same as Figure 1.

To further examine the feasibility of the as-designed sensing method, the interference from the products produced from the PPase-based biocatalytic reaction was studied. The direct product of PPase-catalyzed hydrolysis of PPi is , and the protonation states of including , , and are present in the different pH. According to acid dissociation constants of H3PO4 (, , and ), in HEPES buffer (10 mM, pH 6.5), (namely, Pi) exists dominantly. In our designed strategy, the addition of PPase to the Cu2+-PPi complex containing DHC obviously leads to the fluorescence quenching again because Pi, which is produced from PPase-catalyzed hydrolysis of PPi, cannot chelate Cu2+ to form the complex. Therefore, in order to define chelation interaction between Pi and Cu2+, fluorescence titration with Pi in varying concentrations was conducted in Cu2+-DHC complex system. As shown in Figure 5(a), with the increasing concentration of Pi, the fluorescence emission intensity of DHC was not restored gradually as well as PPi, suggesting no chelation interaction between Pi and Cu2+. Furthermore, all the phosphate-related anions, including , , , AMP, ADP, and ATP, were also investigated in this study. As shown in Figure 5(b), all the phosphate-related anions except PPi restored the fluorescence intensity of DHC weakly, essentially demonstrating that these substances do not interfere with the PPase activity assay because of their weaker coordination capability with Cu2+. This feature actually forms the basis for the development of the effective fluorescence method for sensitive PPase activity assay, as demonstrated in Figures 25.

Figure 5: (a) Fluorescence responses of DHC (500 μM) (curve A) and Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex with (curves B to F: 0, 10, 30, 50, 100, and 200 μM) in HEPES aqueous buffer (10 mM, pH 6.5). (b) Fluorescence changes of DHC (500 μM, curve A) and Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex in the absence (curve B) and presence of various anions (curves C to I: , , , AMP, ADP, ATP, and PPi, 200 μM) in HEPES buffer aqueous solution (10 mM, pH 6.5). Other reaction conditions are the same as Figure 1.
3.3. Interference Study for PPi and PPase

The selectivity of the as-designed sensing method for PPi was also evaluated by comparing the fluorescence response toward PPi (200 μM) with 9 other kinds of anions, including Cl, F, Br, I, , , , , and (200 μM) in HEPES buffer aqueous solution (10 mM, pH 6.5) containing 500 μM DHC and 200 μM Cu2+ under the same detection condition as described in Section 3.2. As shown in Figure 6(a), the results demonstrated that these anions caused minor fluorescence intensity changes of Cu2+-DHC complex at 475 nm (the blue columns in Figure 6(b)) while PPi restored about 90% fluorescence intensity of DHC. Thus, the in situ generated Cu2+-DHC complex can behave as a high selective fluorescence “off-on” sensor for PPi. Additionally, the selectivity of DHC for sensing Cu2+ was evaluated by comparing its fluorescence response toward Cu2+ with those of 17 other kinds of conventional anions including Cl, F, Br, I, , , , , , ATP, AMP, ADP, PPi, , , , and PPi (200 μM), as shown in Figure 7. The results demonstrated that these tested anions induced negligible fluorescence intensity changes of DHC. Therefore, the proposed strategy has a high selectivity for PPi, allowing potential applications in complex samples.

Figure 6: (a) Fluorescence changes of DHC (500 μM, curve A) and Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex in the absence (curve B) and presence of various anions (curves C to I: Cl, F, Br, I, , , , , , and PPi; 200 μM) in HEPES buffer aqueous solution (10 mM, pH 6.5). (b) Fluorescence responses of DHC (500 μM) (black bars) and Cu2+-DHC (Cu2+: 200 μM, DHC: 500 μM) complex at 475 nm in the absence (red bars) and presence of different anions (1 to 10: PPi, Cl, F, Br, I, , , , , and ; 200 μM) (blue bars). Other reaction conditions are the same as Figure 1.
Figure 7: Fluorescence changes of DHC (500 μM, curve A) in the absence (curve A) and presence of various anions (curves B to P: Cl, F, Br, I, , , , , , ATP, AMP, ADP, PPi, , , , and PPi, 200 μM) in HEPES buffer aqueous solution (10 mM, pH 6.5). Other reaction conditions are the same as Figure 1.

Since the proposed fluorescent system was to be applied for the measurements of PPi and PPase in biological samples, it was necessary to validate whether there were interferences on this sensing system from ammonium ion, EDTA, and sulfhydryl compounds including GSH, Cys, and Hcy, all of which can coordinate with Cu2+. As shown in Figures  S-2 and S-3 (see Supplementary Material available online at http://dx.doi.org/10.1155/2016/4306838), the results showed that five substances as interferences had less observable fluorescent responses and can restore the fluorescence emission intensity of DHC compared with PPi. On the other hand, EDTA does not exist in body fluids, suggesting that EDTA had no interference on this fluorescent sensing system for the application in biological samples. Furthermore, the above-mentioned physiological substances, including , GSH, Cys, and Hcy, keep at the low concentrations, which do not exceed 50 μM in biological samples. In such the concentration range, these four physiological substances had the slight fluorescent responses, indicating that these substances within a certain range of their concentration related to physiological changes have no interference on this fluorescent sensing system for the application in biological samples.

3.4. Quantitative Monitoring of PPase Activity

It was reported that PPase is highly active in catalyzing the cleavage reaction of PPi to Pi in the presence of Mg2+ ( [30]). Herein, we attempted to apply the as-designed turn off-on-off fluorescent system for PPase activity assay in the presence of 500 μM Mg2+. Typically, PPase with various concentrations was added into HEPES buffer aqueous solution containing 500 μM DHC, 200 μM Cu2+, 200 μM PPi, and 500 μM Mg2+. Figure 8(a) depicts the typical time-dependent fluorescence spectra of the detection system. The spectra were consecutively recorded every 1 min at 25°C immediately after the addition of PPase to this system in the first 10 minutes and every 5 min in the later 30 minutes. For comparison, the time-dependent fluorescence spectrum of the detection system without the addition of PPase was constant (Figure 8(a), black curve). The interaction time required for optimum fluorescence assay in the presence of PPase was investigated in Figure 8(a). It is clearly seen that the emission peak at 475 nm sharply decreases from 0 to 10 min and then slightly decreases and finally keeps a plateau after 30 min. This suggests that PPase-catalyzed hydrolysis of PPi has finished within 30 min. Therefore, fluorescence responses were recorded at a time point of 30 min after the addition of PPase.

Figure 8: (a) Kinetic plots of time-dependent fluorescence responses versus different activities of PPase present. (b) Fluorescence responses of DHC (500 μM) (curve A) and Cu2+-PPi (Cu2+: 200 μM, PPi: 200 μM) complex with DHC (500 μM) were recorded at a time point of 30 min after the addition of PPase (curves B to M: 0, 0.1, 0.12, 0.15, 0.17, 0.2, 0.22, 0.25, 0.27, 0.3, 0.35, and 0.4 U) in HEPES aqueous buffer (10 mM, pH 6.5) in the presence of 500 μM Mg2+. The final volume of each resulting mixture was adjusted to 500 μL. (c) Fluorescence responses versus . Error bars were estimated from three replicate measurements. Other reaction conditions are the same as Figure 1.

As typically displayed in Figure 8(b), fluorescence responses decrease with increasing PPase activity from 0.1 to 0.4 U (the final volume 500 μL). Figure 8(c) shows the fluorescence intensity at 475 nm versus the PPase concentrations. The linear equation was (U/500 μL) () in the range from 0.1 to 0.3 U. The detection limit of PPase based on a signal-to-noise ratio of 3 was calculated to be 0.028 U in 500 μL. These results demonstrated that our system was comparable and even more efficient for detection of the activity of PPase compared to those provided by other approaches [13, 14, 23]. Moreover, the selectivity of our method for PPase was verified by, respectively, testing some nonspecific proteins including BSA, egg protein, IgG, and PSA with different isoelectric points and structures (figure has not been shown). The result demonstrated that no remarkable fluorescence changes were induced by these tested nonspecific proteins, suggesting that the proposed strategy has a high selectivity in a quantitative PPase activity and allows potential applications in complex samples.

In order to evaluate the validity of the developed method, especially on interference arising from DHC and Cu2+ for PPase activity assay, a colorimetric method using molybdenum blue for PPase activity assay [29] was performed as described in Section 2.4. The results in a microplate reader showed that the absorbance at 650 nm was 0.732, 0.716, and 0.789 in the absence and the presence of 200 μM Cu2+ or 500 μM DHC, respectively. This suggests that the presence of 200 μM Cu2+ or 500 μM DHC did not affect the PPase activity. The above results of the microplate reader are also according to UV-Vis absorption responses and photos perfectly (see Figure  S-1).

3.5. Inhibitor Efficiency Evaluation

Sodium fluoride (NaF) has been reported to be an inhibitor of PPase activity through the formation of the [F−PPase] intermediate [31]. Here, NaF as a model inhibitor was investigated using the developed method in this study. We first studied whether NaF could induce the fluorescence intensity changes of DHC or Cu-DHC complex. Comparing Figure 9 with Figures 6 and 7, the sole addition of NaF to DHC or Cu2+-DHC complex caused negligible fluorescence intensity changes. This observation indicates that NaF itself could not interfere with fluorescence emission in both these sensing systems. The inhibitor efficiency evaluation was performed as described in Section 2.5. The fluorescence responses at 475 nm were restored gradually with the increasing NaF concentration from 0.1 to 1000 μM, as showed in Figure 9. This indicates that NaF could remarkably inhibit the PPase activity.

Figure 9: (a) Fluorescence responses of Cu2+-PPi (Cu2+: 200 μM, PPi: 200 μM) complex with DHC (500 μM) after adding different concentration of NaF-treated PPase (0.25 U) in HEPES aqueous buffer (10 mM, pH 6.5). The concentration of NaF (curves A to I: 0, 0.1, 1, 5, 10, 50, 100, 500, and 1000 μM). The final volume of each resulting mixture was adjusted to 500 μL. (b) Fluorescence intensity versus the NaF concentration. Error bars were estimated from three replicate measurements. Other reaction conditions are the same as Figure 1.

To further confirm that the fluorescence recovery resulted from the NaF-inhibited PPase, we also checked the inhibition effect of NaF on the fluorescence system containing Cu2+ (200 μM) and PPi (200 μM) and DHC (500 μM). The results showed that 2 mM NaF could not affect the fluorescence emission of the Cu2+-PPi-DHC system (figure has not been shown). This observation suggests that the as-developed fluorescence method in this study can be potentially used to evaluate the inhibitor efficiency of NaF for PPase activity.

It has been reported that PPase activity could be inhibited by micromolar F ion [31]. Herein, we found that the restored fluorescence emission of the tested system reached constant when 0.25 U of PPase (500 μL) was preincubated with 500 μM NaF, and the higher concentration of NaF (1000 μM) cannot restore the fluorescence intensity again. These observations suggest that the concentration of NaF less than 500 μM could remarkably inhibit the activity of 0.25 U of PPase. Moreover, in order to evaluate the inhibition effect of NaF, IC50, which represents the concentration of an inhibitor that is required for 50% inhibition of an enzyme, could be used as a parameter to evaluate the inhibitors. As can be seen from Figure 9(b), a profile was obtained by plotting the fluorescence intensity versus the NaF concentration. The IC50 value of PPase was calculated to be about μM. These results essentially demonstrated that the turn off-on-off fluorescent system could also be used to evaluate the inhibitor efficiency and may further be applied to screen other PPase inhibitors simply.

4. Conclusion

We have designed and constructed a simple and efficient turn off-on-off fluorescent system based on PPase-regulated competitive coordination of Cu2+ between a water-soluble fluorescent probe DHC and inorganic ion PPi, which shows a highly selective and sensitive fluorescence response toward PPi and PPase over other tested anions and structural analogues in a 100% aqueous medium. The developed method with good linearity and low detection limit bears advantages in the low technical and instrumental demands and could be also very promising for PPase inhibitor efficiency evaluation. Thus, the present turn off-on-off fluorescent strategy holds great potential for applications in the reliable clinic sensing of PPi in biological and environmental samples and the PPase-related mechanism research and disease diagnostics.

Competing Interests

The authors declare that they have no competing interests.

Acknowledgments

This work is financially supported by NSF of China (Grant nos. 21305109 for Lingzhi Zhao, 81202492 for Chunye Liu, and 91332101 for Chenxiao Zhang), by the Ph.D. Startup Foundation of Xi’an Medical University of China (no. 2012DOC09), and by the Scientific Research Plan Projects Foundation of Shaanxi Science and Technology Department of China (no. 2014JQ2073, 2014K02-11-01).

References

  1. M. S. Huang, A. P. Sage, J. Lu, L. L. Demer, and Y. Tintut, “Phosphate and pyrophosphate mediate PKA-induced vascular cell calcification,” Biochemical and Biophysical Research Communications, vol. 374, no. 3, pp. 553–558, 2008. View at Publisher · View at Google Scholar · View at Scopus
  2. F. W. L. Tsui, “Genetics and mechanisms of crystal deposition in calcium pyrophosphate deposition disease,” Current Rheumatology Reports, vol. 14, no. 2, pp. 155–160, 2012. View at Publisher · View at Google Scholar · View at Scopus
  3. A. Caswell, D. F. Guilland-Cumming, P. R. Hearn, M. K. McGuire, and R. G. Russell, “Pathogenesis of chondrocalcinosis and pseudogout. Metabolism of inorganic pyrophosphate and production of calcium pyrophosphate dihydrate crystals,” Annals of the Rheumatic Diseases, vol. 42, no. 1, pp. 27–37, 1983. View at Publisher · View at Google Scholar · View at Scopus
  4. R. A. Terkeltaub, “Inorganic pyrophosphate generation and disposition in pathophysiology,” American Journal of Physiology—Cell Physiology, vol. 281, no. 1, pp. C1–C11, 2001. View at Google Scholar · View at Scopus
  5. P. Heikinheimo, J. Lehtonen, A. Baykov, R. Lahti, B. S. Cooperman, and A. Goldman, “The structural basis for pyrophosphatase catalysis,” Structure, vol. 4, no. 12, pp. 1491–1508, 1996. View at Publisher · View at Google Scholar · View at Scopus
  6. E. Oksanen, A.-K. Ahonen, H. Tuominen et al., “A complete structural description of the catalytic cycle of yeast pyrophosphatase,” Biochemistry, vol. 46, no. 5, pp. 1228–1239, 2007. View at Publisher · View at Google Scholar · View at Scopus
  7. M. Ilias and T. W. Young, “Streptococcus gordonii soluble inorganic pyrophosphatase: an important role for the interdomain region in enzyme activity,” Biochimica et Biophysica Acta—Proteins and Proteomics, vol. 1764, no. 7, pp. 1299–1306, 2006. View at Publisher · View at Google Scholar · View at Scopus
  8. E. Koike, S. Toda, F. Yokoi et al., “Expression of new human inorganic pyrophosphatase in thyroid diseases: its intimate association with hyperthyroidism,” Biochemical & Biophysical Research Communications, vol. 341, no. 3, pp. 691–696, 2006. View at Publisher · View at Google Scholar · View at Scopus
  9. D. B. Friedman, S. Hill, J. W. Keller et al., “Proteome analysis of human colon cancer by two-dimensional difference gel electrophoresis and mass spectrometry,” Proteomics, vol. 4, no. 3, pp. 793–811, 2004. View at Publisher · View at Google Scholar · View at Scopus
  10. H. P. J. Bloemers, M. L. Stephenson, and P. C. Zamecnik, “A sensitive micro assay for inorganic pyrophosphatase,” Analytical Biochemistry, vol. 34, no. 1, pp. 66–70, 1970. View at Publisher · View at Google Scholar · View at Scopus
  11. M. Hørder, “Colorimetric determination of orthophosphate in the assay of inorganic pyrophosphatase activity,” Analytical Biochemistry, vol. 49, no. 1, pp. 37–47, 1972. View at Publisher · View at Google Scholar · View at Scopus
  12. D. H. Vance and A. W. Czarnik, “Real-time assay of inorganic pyrophosphatase using a high-affinity chelation-enhanced fluorescence chemosensor,” Journal of the American Chemical Society, vol. 116, no. 20, pp. 9397–9398, 1994. View at Publisher · View at Google Scholar · View at Scopus
  13. J. J. Deng, Q. Jiang, Y. X. Wang, L. Yang, P. Yu, and L. Mao, “Real-time colorimetric assay of inorganic pyrophosphatase activity based on reversibly competitive coordination of Cu2+ between cysteine and pyrophosphate ion,” Analytical Chemistry, vol. 85, no. 19, pp. 9409–9415, 2013. View at Publisher · View at Google Scholar · View at Scopus
  14. L. Zhang, M. Li, Y. Qin, Z. Chu, and S. Zhao, “A convenient label free colorimetric assay for pyrophosphatase activity based on a pyrophosphate-inhibited Cu2+-ABTS-H2O2 reaction,” Analyst, vol. 139, no. 23, pp. 6298–6303, 2014. View at Publisher · View at Google Scholar · View at Scopus
  15. J. Sun, F. Yang, D. Zhao, and X. Yang, “Highly sensitive real-time assay of inorganic pyrophosphatase activity based on the fluorescent gold nanoclusters,” Analytical Chemistry, vol. 86, no. 15, pp. 7883–7889, 2014. View at Publisher · View at Google Scholar · View at Scopus
  16. H. H. Deng, F. F. Wang, X. Q. Shi et al., “Water-soluble gold nanoclusters prepared by protein-ligand interaction as fluorescent probe for real-time assay of pyrophosphatase activity,” Biosensors & Bioelectronics, vol. 83, pp. 1–8, 2016. View at Publisher · View at Google Scholar
  17. Q. L. Yue, Y. N. Hou, S. Z. Yue et al., “Construction of an off-on fluorescence system based on carbon dots for trace pyrophosphate sensing,” Journal of Fluorescence, vol. 25, no. 3, pp. 585–594, 2015. View at Publisher · View at Google Scholar · View at Scopus
  18. X. Huang, Z. Guo, W. Zhu, Y. Xie, and H. Tian, “A colorimetric and fluorescent turn-on sensor for pyrophosphate anion based on a dicyanomethylene-4H-chromene framework,” Chemical Communications, no. 41, pp. 5143–5145, 2008. View at Publisher · View at Google Scholar · View at Scopus
  19. X. L. Feng, Y. X. An, Z. Y. Yao, C. Li, and G. Shi, “A turn-on fluorescent sensor for pyrophosphate based on the disassembly of Cu2+-mediated perylene diimide aggregates,” ACS Applied Materials & Interfaces, vol. 4, no. 2, pp. 614–618, 2012. View at Publisher · View at Google Scholar · View at Scopus
  20. Z. Liu, J. Xiao, X. Wu et al., “Switch-on fluorescent strategy based on N and S co-doped graphene quantum dots (N-S/GQDs) for monitoring pyrophosphate ions in synovial fluid of arthritis patients,” Sensors and Actuators, B: Chemical, vol. 229, pp. 217–224, 2016. View at Publisher · View at Google Scholar · View at Scopus
  21. J.-R. Lin, C.-J. Chu, P. Venkatesan, and S.-P. Wu, “Zinc(II) and pyrophosphate selective fluorescence probe and its application to living cell imaging,” Sensors & Actuators B: Chemical, vol. 207, no. 2, pp. 563–570, 2015. View at Publisher · View at Google Scholar · View at Scopus
  22. L. P. Lin, X. H. Song, Y. Y. Chen et al., “One-pot synthesis of highly greenish-yellow fluorescent nitrogen-doped graphene quantum dots for pyrophosphate sensing via competitive coordination with Eu3+ ions,” Nanoscale, vol. 7, no. 37, pp. 15427–15433, 2015. View at Publisher · View at Google Scholar · View at Scopus
  23. J. Deng, P. Yu, L. Yang, and L. Mao, “Competitive coordination of Cu2+ between cysteine and pyrophosphate ion: toward sensitive and selective sensing of pyrophosphate ion in synovial fluid of arthritis patients,” Analytical Chemistry, vol. 85, no. 4, pp. 2516–2522, 2013. View at Publisher · View at Google Scholar · View at Scopus
  24. Y. Q. Lin, L. L. Hu, L. B. Li, K. Wang, Y. Ji, and H. Zou, “Electrochemical determination of pyrophosphate at nanomolar levels using a gold electrode covered with a cysteine nanofilm and based on competitive coordination of Cu(II) ion to cysteine and pyrophosphate,” Microchimica Acta, vol. 182, no. 11-12, pp. 2069–2075, 2015. View at Publisher · View at Google Scholar · View at Scopus
  25. Y. X. Lin, Q. Zhou, J. Li et al., “Magnetic graphene nanosheet-based microfluidic device for homogeneous real-time electronic monitoring of pyrophosphatase activity using enzymatic hydrolysate-induced release of copper ion,” Analytical Chemistry, vol. 88, no. 1, pp. 1030–1038, 2016. View at Publisher · View at Google Scholar · View at Scopus
  26. H. Deng, F. Wang, X. Shi et al., “Water-soluble gold nanoclusters prepared by protein-ligand interaction as fluorescent probe for real-time assay of pyrophosphatase activity,” Biosensors and Bioelectronics, vol. 83, pp. 1–8, 2016. View at Publisher · View at Google Scholar
  27. J. Sun, B. Wang, X. Zhao, Z.-J. Li, and X. Yang, “Fluorescent and colorimetric dual-readout assay for inorganic pyrophosphatase with Cu2+-triggered oxidation of o-phenylenediamine,” Analytical Chemistry, vol. 88, no. 2, pp. 1355–1361, 2016. View at Publisher · View at Google Scholar
  28. J. B. English, A. E. Martell, R. J. Motekaitis, and I. Murase, “Molecular interaction of pyrophosphate with 1,13-dioxa-4,7,10,16,20,24-hexaazacyclohexacosane (OBISDIPEN) and its mononuclear and dinuclear copper(II) complexes,” Inorganica Chimica Acta, vol. 258, no. 2, pp. 183–192, 1997. View at Publisher · View at Google Scholar · View at Scopus
  29. M. Yamazaki, Q. W. Shen, and D. R. Swartz, “Tripolyphosphate hydrolysis by bovine fast and slow myosin subfragment 1 isoforms,” Meat Science, vol. 85, no. 3, pp. 446–452, 2010. View at Publisher · View at Google Scholar · View at Scopus
  30. T. Miyajima and S. Ohashi, “Application of gel chromatography to the determination of the stability constants of a magnesium complex with diphosphate,” Bulletin of the Chemical Society of Japan, vol. 51, no. 9, pp. 2543–2547, 1978. View at Publisher · View at Google Scholar
  31. A. A. Baykov, I. P. Fabrichniy, P. Pohjanjoki, A. B. Zyryanov, and R. Lahti, “Fluoride effects along the reaction pathway of pyrophosphatase: evidence for a second enzyme·pyrophosphate intermediate,” Biochemistry, vol. 39, no. 39, pp. 11939–11947, 2000. View at Publisher · View at Google Scholar · View at Scopus