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Chemiluminescent detection systems of horseradish peroxidase employing nucleophilic acylation catalysts
Analytical Biochemistry 377 (2008) 189–194
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Chemiluminescent detection systems of horseradish peroxidase employing nucleophilic acylation catalysts Ettore Marzocchi a, Stefano Grilli a, Leopoldo Della Ciana a,*, Luca Prodi b, Mara Mirasoli c, Aldo Roda c a
Cyanagen SRL, Via Stradelli Guelfi 40/c, 40138 Bologna, Italy Dipartimento di Chimica ‘‘G. Ciamician,” Via Selmi 2, Università di Bologna, 40126 Bologna, Italy c Dipartimento di Scienze Farmaceutiche, Via Belmeloro 6, Università di Bologna, 40126 Bologna, Italy b
a r t i c l e
i n f o
Article history: Received 21 December 2007 Available online 16 March 2008 Keywords: Horseradish peroxidase Luminol Chemiluminescent Enzyme assay Western blot Acylation catalyst
a b s t r a c t The light output of the peroxidase-catalyzed luminol chemiluminescent oxidation reaction can be greatly increased by incorporating different enhancers. Such an increase is attributed to the preferential oxidation of the enhancer by peroxidase intermediates and the rapid formation of enhancer radicals that, in turn, quickly oxidize luminol to its radical anion. These enhancers, which include substituted phenols, substituted boronic acids, indophenols, and N-alkyl phenothiazines, behave as electron transfer mediators. A further, very significant increase in light output was also observed by the addition of nucleophilic acylation catalyst to the enhancer/luminol/oxidant substrate. The effect of the new component is general and applicable to many of the known enhancers but is much more remarkable in association with phenothiazine enhancers (up to 10-fold light output). The addition of a nucleophilic acylation catalyst to these substrates lowered the limit of detection for horseradish peroxidase from 50 to 8 amol. Similar improvements were observed in ‘‘sandwich” enzyme-linked immunosorbent assays and Western blot assays. Ó 2008 Elsevier Inc. All rights reserved.
The horseradish peroxidase (HRP)1-catalyzed chemiluminescent oxidation of luminol is widely used in many molecular biologybased assays, such as Western blots, dot blots, and enzyme-linked immunosorbent assays (ELISAs), as well as in immunohistochemistry. Much effort has been made to improve the efficiency and analytical performance of this reaction. In particular, the addition of enhancers to the reaction substrate greatly increases light output and duration kinetics [1]. Molecules with enhancer properties are essentially redox mediators [2] capable of exchanging electrons between the peroxidase enzyme and luminol. Enhancers include substituted phenols [3], substituted boronic acids [4], indophenols, and N-alkyl phenothiazine derivatives [5]. The enhanced HRP-catalyzed oxidation of luminol is a complex multistep reaction. Although redox enhancers are useful in improving enzyme turnover and increasing the equilibrium concentration of a key intermediate, luminol radical anion, a second bottleneck may need to be removed to obtain a further increase in light output. According to a well-established mechanism [6],
* Corresponding author. Fax: +39 051534063. E-mail address: [email protected] (L. Della Ciana). 1 Abbreviations used: HRP, horseradish peroxidase; ELISA, enzyme-linked immunosorbent assay; DMAP, 4-dimethylaminopyridine; MORP, 4-morpholinopyridine; PPY, 4-pyrrolidinopyridine; THF, tetrahydrofuran; PBS, phosphate-buffered saline; SPTZ, sodium 3-(100 -phenothiazinyl)propane-1-sulfonate; HPLC, high-performance liquid chromatography; DMSO, dimethyl sulfoxide; TEA, triethanolamine; BSA, bovine serum albumin; NHE, normal hydrogen electrode. 0003-2697/$ - see front matter Ó 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.ab.2008.03.020
luminol radical anion, once formed, is subject to rapid dismutation to luminol and diazaquinone. In turn, this species is attacked by hydroperoxide anion and converted into reactive peroxides. This type of reaction is often accelerated by nucleophilic acylation catalysts [7], especially 4-aminopyridines. This study investigates the effect of these compounds on chemiluminescence light output when used in association with redox enhancers and their use in substrates for HRP chemiluminescent assays with the goal of achieving a higher detectability of HRP. This will allow the development of more sensitive bioanalytical methods and also further miniaturize chemiluminescent-based chip devices.
Materials and methods Luminol sodium salt was obtained by recrystallization of the commercially available luminol (5-amino-2,3-dihydro-1,4-phthalazinedione, Fluka) from sodium hydroxide using a procedure described previously [8]. Phenothiazine (Acros), 1,3-propanesultone (Aldrich), 4-dimethylaminopyridine (DMAP, TCI–EP), 4-morpholinopyridine (MORP, Aldrich), and 4-pyrrolidinopyridine (PPY, Fluka) were purchased from their respective suppliers. HRP (type VI-A, 797 U/mg) was purchased from Fluka. Water was freshly spilled from a Milli-Q water purification system. Dry tetrahydrofuran (THF) was obtained by distillation over Na/benzophenone. All other compounds were obtained from commercial sources and
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used without further purification. Phosphate-buffered saline (PBS) was prepared in 0.1 M concentration and pH 7.3. Instrumentation A Cary Eclipse (Varian) fluorescence spectrophotometer was used for kinetic measurements using the following setup: kinetic mode, kem = 425 nm, gate time = 5 ms, emission slits properly adjusted, Savitzky–Golay smoothing. A Luminoskan (Ascent) luminometer was used for microtiter well readings (1 s integration/well). A NightOwl LB981 (Berthold Technologies) luminograph was used to read the luminescence from Western blot membranes (120 s total integration time). The detection limits for the calibration curves were obtained graphically according to the method of Hubaux and Vos [9], that is, by plotting the regression line with the confidence interval (99%) and following the point where the upper confidence interval curve intersects the y axis horizontally until reaching the lower confidence interval and then proceeding vertically to the x axis. This point on the x axis was taken as the limit of detection. Synthesis of SPTZ Because the purity of the enhancer is of fundamental importance in the chemiluminescent process [10,11], the two-step literature procedure [12] was replaced by the following improved one-pot synthesis. Sodium hydride (60% dispersion in mineral oil, 1.76 g, 44.2 mmol) was dissolved in dry THF (40 ml) under argon atmosphere. A solution of phenothiazine (8 g, 40 mmol) in dry THF (40 ml) was added, and the mixture was stirred at room temperature for 60 min (a dark orange suspension forms) and then 90 min at 50 °C. After cooling at 0 °C, a solution of 1,3-propanesultone (4.88 g, 40 mmol) in dry THF (40 ml) was added and the resulting mixture was stirred for 30 min. The ice bath was removed, and the mixture was stirred for a further 30 min. The white precipitate was filtered off and subsequently washed with THF and Et2O and further purified by crystallization from 90% ethanol to give 3-(100 -phenothiazinyl)propane-1-sulfonate (SPTZ) as white crystals (10.0 g, 72% yield) (Fig. 1). Molecular mass of the free acid (C15H15O3S2): 321.4. MS (API-ES): 322.0 [MH]+. 1H NMR (300 MHz, D2O) d: 6.9–7.1 (m, 4H, ArH), 6.7–6.9 (m, 4H, ArH), 3.8 (t, 2H, -CH2-N-, J = 7.2 Hz), 2.9 (t, 2H, -CH2-S, J = 7.2 Hz), 1.9–2.9 (m, 2H, -CH2-CH2-CH2). The content of unreacted phenothiazine in the product was found to be less than 0.0002 parts (mol/mol) by high-performance liquid chromatography (HPLC) analysis (column: 4.6 150-mm Waters C18 XTerra; eluent: H2O/acetonitrile [3:7] at 1 ml/min; UV detector: 254 nm). Effect of different enhancers on the luminol–peroxide–peroxidase reaction
in 0.1 M Tris–HCl (pH 8.6). This solution was then split into portions, and to each portion different enhancers were added from previously prepared stock solutions in dimethyl sulfoxide (DMSO); for example, the final concentration of enhancer in the working solution was as follows: (a) p-coumaric acid (0.68 mM), (b) p-iodophenyl-boronic acid (0.77 mM), (c) p-iodophenol (0.62 mM), (d) SPTZ (0.75 mM), and (e) reference (no enhancer). The final DMSO concentration was 1%. Then 2 ml of each working solution was tested in a 3-ml polymethylmethacrylate disposable cuvette by adding 10 ll of HRP solution (2 lg/ml) and vortexing for a few seconds. Then, 30 s after enzyme addition, light emission was monitored for 30 min using the fluorescence spectrophotometer. Effect of DMAP association with different enhancers Working solutions were prepared as described above. Each solution was tested by adding either 10 ll of Tris buffer or 10 ll of DMAP (200 mM) in Tris buffer. The same enzyme addition protocol described above was used. Values were calculated integrating the signal during the first 10-min run of a kinetic measurement using the fluorescence spectrophotometer. Effect of nucleophilic acylation catalysts on SPTZ-enhanced substrates The working solution was freshly prepared using the following concentrations: 5 mM luminol sodium salt, 1.5 mM SPTZ, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). This solution was split into portions, and the following compounds were added to each portion to a final concentration of 1.5 mM: (a) pyridine, (b) 4-picoline, (c) PPY, (d) DMAP, (e) MORP, (f) triethanolamine (TEA), and (g) reference (no addition of acylation catalyst). The same enzyme addition protocol described above was used. Measurements were carried out with the fluorescence spectrophotometer (15 min total acquisition time). Light emission from working solutions containing MORP: Effect of pH The working solutions NoMORP and MORP were freshly prepared. The NoMORP working solution contained 5 mM luminol, 1.5 mM SPTZ, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). The MORP working solution contained 5 mM luminol, 1.5 mM SPTZ, 1.5 mM MORP, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). The pH for each working solution was adjusted using negligible amounts of HCl (5 M) or NaOH (5 M) in the pH 8 to 10 interval. The same enzyme addition protocol described above was used. Measurements were carried out with the fluorescence spectrophotometer (10 min total acquisition time). Standard curve for HRP
The working solution was freshly prepared using the following concentrations: 1.25 mM luminol sodium salt and 2.50 mM H2O2
Samples (50 ll) of HRP diluted in Tris buffer (0.1 M, pH 8.6) in the range from 0 to 100 pg/well were analyzed in a microtiter plate using alternatively NoMORP and MORP working solutions (100 ll) prepared as described above. Light emission was measured with the luminometer (15 min total acquisition time). Application of the MORP–SPTZ–luminol chemiluminescent reaction in a sandwich ELISA
Fig. 1. Sodium 3-(100 -phenothiazinyl)propane-1-sulfonate (SPTZ) structure.
A noncompetitive enzyme immunoassay for pathogen bacterium Yersinia enterocolitica previously developed in our laboratory was used to evaluate the analytical performance of the new chemiluminescent cocktail. A mouse monoclonal antibody specific for Y. enterocolitica was immobilized into the wells of a white 96-well microtiter plate (Microlite 2+, Thermo Scientific). The antibody was
Chemiluminescent detection systems of HRP / E. Marzocchi et al. / Anal. Biochem. 377 (2008) 189–194
diluted 1:1000 (v/v) in 0.05 M carbonate/bicarbonate buffer (pH 9.6), and then 100 ll was dispensed in the wells and incubated overnight at 4 °C. After washing, a second coating was carried out with 2% bovine serum albumin (BSA) in PBS, and then the wells were dried under vacuum and stored at 4 °C until use. To perform the assay, 100 ll of serial dilutions of Y. enterocolitica (ATCC 23715) lysate in PBS (ranging from 1.5 104 to 3.0 107 cells/ml) were incubated in duplicate in the wells for 1 h at 37 °C. After washing with 4% Tween in PBS, wells were incubated for 1 h at 37 °C with 100 ll of rabbit polyclonal anti-Y. enterocolitica antibody diluted 1:2000 (v/v) in PBS/Tween and washed again. The plate was then incubated for 30 min at room temperature in the dark with HRPconjugated anti-rabbit antibody (Sigma) diluted 1:8000 (v/v) in PBS and washed again. Calibration curves prepared in parallel were produced by adding either NoMORP or MORP working solution (see protocol above) and by measuring light emission with the luminometer. Light intensity was monitored for a period of 15 min. Application of MORP-added luminol–SPTZ solution in Western blot assays Serial dilutions of albumin from chicken egg white (Sigma) were electrophoresed on 6% polyacrylamide gel and then transferred to a nitrocellulose membrane. The blot was blocked in 2% dry milk for 1 h at room temperature and then washed with PBS twice for 15 min. The blot was then incubated with rabbit antichicken egg albumin (whole anti-serum, 1:4000 [v/v] dilution) for 1 h at room temperature and washed as described previously. The blot was then incubated with HRP-conjugated anti-rabbit antibody (1:2000 dilution) for 1 h at room temperature and washed again. Two blots were prepared and treated alternatively with NoMORP and MORP working solutions (see protocol above). Light emission from the membrane was measured with the luminograph. Results and discussion The spontaneous chemiluminescent oxidation of luminol (LH2) by peroxides occurs according to a well-established mechanism [6]. In neutral or alkaline solution, the monoanion of luminol (LH–) is oxidized to the corresponding radical anion (L–). Dismutation of L- regenerates LH– while producing a two-electron oxidized luminol species, a diazaquinone (L). The diazaquinone is then attacked by hydrogen peroxide anion HO2 . An intermediate peroxide species is formed and then collapses, with loss of nitrogen, to 3-aminophthalate in its excited state, AP*. The decay of AP* to aminophthalate, AP, is responsible for the chemiluminescent light emission at 425 nm. Although the spontaneous oxidation of luminol in aqueous solution is very slow, it can be catalyzed by peroxidases such as HRP. The intensity of the light emission can be further increased by adding redox mediators such as phenols, indophenols, phenothiazines, and substituted boronic acids. A quantitative mechanism of the enhanced luminescence has been proposed [2]. Although this model was developed specifically for phenolic compounds, its general features are applicable to other classes of enhancers. According to the model, the native Fe(III) enzyme (HRP) is oxidized by peroxide to HRP-I in a two-electron oxidation (see Eq. (1)). HRP-I then returns to its native state, HRP, by reaction with the enhancer (E) in two one-electron transfer steps, with HRP-II as intermediate (see Eqs. (2) and (3)). In each of these steps, the enhancer is oxidized to its radical form (E): HRP þ H2 O2 ! HRP I þ H2 O
ð1Þ
HRP I þ E ! HRP II þ E
ð2Þ
HRP II þ E ! HRP þ E :
ð3Þ
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The reaction of luminol with HRP-II is approximately 100-fold less rapid than the reaction of luminol with HRP-I. Luminescence enhancers react more rapidly with HRP-II than does luminol, thereby accelerating enzyme turnover. In the next step, the enhancer radical E oxidizes luminol anion LH– to the key intermediate L-. This reaction is reversible: E þ LH ¡E þ L :
ð4Þ
The position of the equilibrium is determined by the difference between the reduction potential of the redox couples, L-/LH– and E/E. With phenolic enhancers, it was found that the rate of reaction with HRP-II is also determined by the reduction potential of the E/E couple. For these enhancers, a bell-shaped dependence of the luminescence efficiency on the reduction potential of E was observed, with its maximum at approximately 0.8 V versus normal hydrogen electrode (NHE), roughly the same potential of luminol, 0.82 V versus NHE. Fine tuning of the reduction potential for E is required to avoid loss of reaction efficiency. N-Alkyl-substituted phenothiazine enhancers were first described by Sugiyama in the patent literature [5]. A water-soluble derivative, SPTZ (Fig. 1), was employed most often in these studies. However, no major improvement over the performance of phenolic enhancers was demonstrated. A few years later, much better results in terms of luminescent intensity and duration were reported by using a carefully purified sample of SPTZ [11]. In fact, we found that the two-step, low-yield method used for the synthesis of SPTZ [12] is largely responsible for the presence of impurities, especially unreacted phenothiazine, that quench L– and drastically reduce light output. Unfortunately, these impurities are not easily removed by conventional purification methods [11]. For these reasons, we developed a new one-step synthesis of SPTZ. Under our conditions, the SPTZ enhancer spontaneously crystallizes out of the reaction solution in high yield and extreme purity. Considering now the system in more detail, it is worth observing that the reduction potential of SPTZ (SPTZ+/SPTZ) is 0.83 V versus NHE [12], nearly identical to the value reported for luminol and close to the optimal value observed for phenolic enhancers. On the other hand, phenothiazine itself, with a reduction potential (PTZ+/ PTZ) of 0.62 V versus NHE [13], is much easier to oxidize than is SPTZ and effectively quenches the production of luminescence even when present in trace amount. Next, the performance of SPTZ and other commonly used enhancers—p-coumaric acid, p-iodophenylboronic acid, and piodophenol—was evaluated. For each enhancer, the literature concentration was used [3,4,11]. All other conditions were kept constant, including luminol and oxidant concentrations, pH, and temperature. The best performance, in terms of both luminosity and duration, was obtained with SPTZ (Fig. 2). In the next experiment, DMAP, a powerful acylation catalyst, was added to each substrate solution. Whereas a very slight signal increase ( 10%) was observed for p-iodophenol and 4-iodophenylboronic acid, the effect was much more significant for SPTZ (> 800%) (Table 1). Therefore, all further studies were focused on the SPTZ system. Because this enhancer is a salt, it is much more water soluble than other enhancers and its concentration can be raised to higher levels (e.g., 1.5 mM). An improved SPTZ substrate solution is obtained by increasing luminol and peroxide concentrations to 5 and 4 mM, respectively. In addition, raising the pH from 8.5 to 9.0 further improves the duration of luminescence and total light output. In practice, the fully optimized SPTZ substrate produces a signal twice as intense as does the unoptimized system. This optimized substrate was used to evaluate a series of pyridine acylation catalysts. Pyridines, and especially 4-dialkylaminopyrines, are wellknown nucleophilic acylation catalysts and are widely used in organic synthesis [7]. For some of these compounds, the following
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100
60
40
a b c d e f g
400
Intensity (a.u.)
80 Intensity (a.u.)
500
a b c d e
300 200 100
20 0 0
2
0 0
5
10
15
20
25
4
6 8 10 Time (min)
12
14
30
Time (min) Fig. 2. Kinetic measurement of light output intensity from luminol (1.25 mM)/H2O2 (2.5 mM)/HRP (227 pM) systems with different enhancers: (a) d, coumaric acid (0.68 mM); (b) s, p-iodophenylboronic acid (0.77 mM); (c) ., p-iodophenol (0.62 mM); (d) D, SPTZ (0.75 mM); (e) j, reference (no addition of enhancer). Data are the means of triplicate assays.
Fig. 3. Kinetic measurement of light output intensity from luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM)/HRP (227 pM) systems with 1.5 mM of different nucleophilic acylation catalysts: (a) d, pyridine; (b) s, 4-picoline; (c) ., PPY; (d) D, DMAP; (e) j, MORP; (f) h, TEA; (g) , reference (no addition of catalyst). Data are the means of triplicate assays.
1600
1200 Intensity (a.u.)
data were reported [7] (where RE is the relative effectiveness as acylation catalyst): PPY (RE 100, pKa 9.9) > DMAP (RE 40, pKa 9.7) > MORP (RE 8, pKa 8.7) >> 4-picoline (RE 0.02, pKa 6.1) > pyridine (RE 0.002, pKa 5.3). When they are added to the SPTZ substrate, the same overall reactivity is observed with some exceptions: MORP > DMAP > PPY >> 4-picoline > pyridine (Fig. 3). The apparent reversal observed with PPY, DMAP, and MORP is likely to be due at least partially to their differences in pKa. At pH 9.0, the active free base form is 66% of the total for MORP, 17% for DMAP, and only 11% for PPY. A basic nonnucleophilic amine such as TEA had practically no effect on light output. Participation in the enhanced reaction of the free base form of the acylation catalysts is confirmed by a pH dependence study whereby the optimal value for light output shifts to higher pH values. Fig. 4 shows a comparison between a working solution based on luminol/SPTZ/oxidant (NoMORP) and one with added MORP. The highest signal was obtained at pH 8.4 for NoMORP. Similar optimal pH values were observed for well-known enhancers such as p-iodophenol [3] and p-iodophenylboronic acid [4]. The addition of MORP not only increased the signal but also shifted the optimal pH to a 9.1 to 9.3 interval. A similar shift was also observed for PPY and DMAP, with both having the optimal pH in the same interval (data not shown). To understand the effect of these catalysts, it is important to observe that they do not act themselves as enhancers (no effect on light output is observed when used alone). Although it is not possible, at this stage, to exclude a synergistic effect on the enhancer, an alternative explanation is proposed by considering in more detail the reaction steps from diazaquinone (L) to a luminol peroxide intermediate [6] (Fig. 5). This reaction involves nucleophilic attack on one of the diazaquinone carbonyls by hydrogen peroxide monoanion. A hydroperoxide species that can rearrange to the endoperoxide is formed.
a b
1400
1000 800 600 400 200 0 8.0
8.5 9.0 9.5 Working solution pH
10.0
Fig. 4. Comparison of working solution pH optimal value for a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM)/HRP (227 pM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). Each point is calculated integrating the signal during the first 10-min run of a kinetic measurement.
Either compound is very unstable and collapses to 3-aminophthalate in its excited state. Perhaps pyridine acylation catalysts could facilitate hydrogen peroxide attack by converting L into a more reactive intermediate. For analytical applications, the nontoxic acylation catalyst MORP is preferred over the highly toxic DMAP and PPY. The analytical performance of MORP and NoMORP working solutions was explored by comparing results obtained in two bioassays: a sandwich ELISA for Y. enterocolitica and a Western blot assay for chicken egg white albumin. Both methods belong to the class of noncompetitive ‘‘sandwich-type” formats given that all of the analyte is captured and then revealed by means of an excess of detection
Table 1 Effect of DMAP association with different enhancers
Without DMAP With DMAP
No enhancer
p-Coumaric acid
p-Iodophenyl-boronic acid
p-Iodophenol
SPTZ
6.77 ± 0.13 6.66 ± 0.06
22.25 ± 2.73 21.99 ± 0.79
100.73 ± 6.88 109.65 ± 0.13
153.12 ± 2.41 172.09 ± 3.16
227.5 ± 13.45 2201.23 ± 317.16
Note. Data are means of triplicate measures of integrated signal intensities in arbitrary units.
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a b
10
40
8
Intensity (a.u.)
6 4
30
2 0 0
20
2
4
6
8 10 12 14
10 Fig. 5. Reaction of hydrogen peroxide with diazaquinone to form the hydroperoxide intermediate.
0 antibody. The detectability of the enzyme label is one of the most important factors determining the detection limit of noncompetitive assays provided that a low degree of nonspecific binding of the detection reagents occurs [14]. A comparison of the dose–response curve for HRP using the SPTZ-enhanced luminol/peroxide substrate with and without MORP is shown in Fig. 6. The detection limit for HRP, calculated as described previously, was 0.35 pg (8 amol)/well for the reaction containing MORP, as compared with 2.20 pg (50 amol)/well for the NoMORP substrate. Next, a sandwich chemiluminescent enzyme immunoassay for the determination of Y. enterocolitica was carried out to compare the performance of MORP and NoMORP chemiluminescent reagents. Calibration curves obtained by plotting signals subtracted of the blank (Fig. 7) show that the detection cocktail containing MORP produced for each bacteria concentration a signal approximately 1 order of magnitude higher than that obtained with NoMORP, thereby providing a steeper calibration curve. Indeed, MORP reagent allowed reaching a limit of detection of 0.6 106 cells/ml, as compared with that of NoMORP cocktail, which allowed reaching a limit of detection of 5.0 106 cells/ml. The linear range extended up to 7.5 106 cells/ml for MORP and up to 1.5 107 cells/ml for NoMORP. Finally, Fig. 8 shows Western blots for chicken egg albumin detected with an HRP conjugate using the MORP and NoMORP reagents. The much stronger signal produced by the MORP-catalyzed
a b
200
1400
150
1200 Intensity (a.u.)
100
1000
50
800
0
0 2 4 6 8 10 12 14
0
5 10 15 20 25 30 10 -6 x Bacteria Concentration (cells/ml)
Fig. 7. Sandwich ELISA tests for the detection of Y. enterocolitica using a couple of specific mouse monoclonal and rabbit polyclonal antibodies and an anti-rabbit HRP-conjugated antibody. The comparison is between a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). The inset shows the linear range with the 99% confidence interval from which the detection limits were calculated. Each point is calculated integrating the signal during the first 15-min run of a kinetic measurement. Data are the means of quadruplicate assays subtracted of the blank. Error bars: ± 1 SD.
Fig. 8. Western blots for albumin from chicken egg white detected using HRPconjugated antibody. The comparison is between a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/ SPTZ (1.5 mM)/perborate (4 mM): (a) NoMORP (no catalyst added, left to right: 19.0, 12.7, 8.4, 5.6, 3.8, 2.5, 1.7, and 1.1 pg albumin); (b) MORP (1.5 mM, left to right: 2.9, 1.9, 1.3, 0.8, 0.6, 0.4, 0.3, and 0.2 pg albumin). Light emission was detected with a charge-coupled device (CCD)-based instrument with 2 min acquisition.
substrate allowed the detection of more diluted protein bands, with a clear advantage in the Western blot technique. In conclusion, the incorporation of an acylation catalyst in enhancer/luminol/oxidant HRP substrates is highly advantageous, especially in the case of N-alkylated phenothiazine enhancers. Although the exact mechanism of action of acylation catalysts has not yet been fully investigated, it is clear from the results of this study that the very significant increase in light output observed in their presence can be translated into a corresponding improvement in sensitivity of chemiluminescent assays. Acknowledgments
600 400 200 0 0
20
40
60
80
100
HRP Amount (pg/well) Fig. 6. Comparison of HRP dose–response curves for a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/ SPTZ (1.5 mM)/perborate (4 mM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). The inset shows the linear range with the 99% confidence interval from which the detection limits were calculated. Each point is calculated integrating the signal during the first 15-min run of a kinetic measurement. Data are the means of quadruplicate assays subtracted of the blank. Error bars: ± 1 SD.
Thanks are due to Luisa Stella Dolci and Manuela Rizzoli for molecular biology-based assays and to Matteo Beltrame for synthesis and purification of luminol. We are also grateful to Monica Musiani and Elisabetta Manaresi for the Western blot tests at the Department of Clinical and Experimental Medicine, Section of Microbiology, University of Bologna. References [1] L.J. Kricka, J.C. Voyta, I. Bronstein, Chemiluminescent methods for detecting and quantitating enzyme activity, Methods Enzymol. 305 (2000) 370–390. [2] P.M. Easton, A.C. Simmonds, A. Rakishev, A.M. Egorov, L.P. Candeias, Quantitative model of the enhancement of peroxidase-induced luminol luminescence, J. Am. Chem. Soc. 118 (1996) 6619–6624. [3] G.H.G. Thorpe, L.J. Kricka, Enhanced chemiluminescent reactions catalyzed by horseradish peroxidase, Methods Enzymol. 133 (1986) 331–353.
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[4] L.J. Kricka, M. Cooper, X. Ji, Synthesis and characterization of 4-iodophenylboronic acid: A new enhancer for the horseradish peroxidase-catalyzed chemiluminescent oxidation of luminol, Anal. Biochem. 240 (1996) 119–125. [5] M. Sugiyama, Method of the chemiluminescence assays of the activity of peroxidase, U.S. patent no. 5,171,668 (1992). [6] F. McCapra, Chemical generation of excited states: The basis of chemiluminescence and bioluminescence, Methods Enzymol. 305 (2000) 3– 47. [7] G. Höfle, W. Steglich, H. Vorbrüggen, 4-Dialkylaminopyridines as highly active acylation catalysts, Angew. Chem. Intl. Ed. Engl. 17 (1978) 569–583. [8] G. Ham, R. Belcher, L.J. Kricka, T.J.N. Carter, Stability of trace iodine solutions, Anal. Lett. 12 (1979) 535–541.
[9] A. Hubaux, G. Vos, Decision and detection limits for linear calibration curves, Anal. Chem. 42 (1970) 849–855. [10] M.A. Motsenbocker, K. Kondo, Improvements to enhanced horseradish peroxidase detection sensitivity, J. Biolumin. Chemilumin. 9 (1994) 15–20. [11] P. D. Davis, Assay of peroxidase activity, U.S. patent no. 6,432,662 (2002). [12] Y. Kawanishi, N. Kitamura, S. Tazuke, Coulombic effect of photoinduced electron-transfer reactions between phenothiazines and viologens, J. Phys. Chem. 90 (1986) 2469–2475. [13] C. Bodea, I. Silberg, Recent advances in the chemistry of phenothiazines, Adv. Heterocycl. Chem. 9 (1968) 321–460. [14] A. Roda, M. Guardigli, E. Michelini, M. Mirasoli, P. Pasini, Analytical bioluminescence and chemiluminescence, Anal. Chem. 75 (2003) 463–470.
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Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio
Chemiluminescent detection systems of horseradish peroxidase employing nucleophilic acylation catalysts Ettore Marzocchi a, Stefano Grilli a, Leopoldo Della Ciana a,*, Luca Prodi b, Mara Mirasoli c, Aldo Roda c a
Cyanagen SRL, Via Stradelli Guelfi 40/c, 40138 Bologna, Italy Dipartimento di Chimica ‘‘G. Ciamician,” Via Selmi 2, Università di Bologna, 40126 Bologna, Italy c Dipartimento di Scienze Farmaceutiche, Via Belmeloro 6, Università di Bologna, 40126 Bologna, Italy b
a r t i c l e
i n f o
Article history: Received 21 December 2007 Available online 16 March 2008 Keywords: Horseradish peroxidase Luminol Chemiluminescent Enzyme assay Western blot Acylation catalyst
a b s t r a c t The light output of the peroxidase-catalyzed luminol chemiluminescent oxidation reaction can be greatly increased by incorporating different enhancers. Such an increase is attributed to the preferential oxidation of the enhancer by peroxidase intermediates and the rapid formation of enhancer radicals that, in turn, quickly oxidize luminol to its radical anion. These enhancers, which include substituted phenols, substituted boronic acids, indophenols, and N-alkyl phenothiazines, behave as electron transfer mediators. A further, very significant increase in light output was also observed by the addition of nucleophilic acylation catalyst to the enhancer/luminol/oxidant substrate. The effect of the new component is general and applicable to many of the known enhancers but is much more remarkable in association with phenothiazine enhancers (up to 10-fold light output). The addition of a nucleophilic acylation catalyst to these substrates lowered the limit of detection for horseradish peroxidase from 50 to 8 amol. Similar improvements were observed in ‘‘sandwich” enzyme-linked immunosorbent assays and Western blot assays. Ó 2008 Elsevier Inc. All rights reserved.
The horseradish peroxidase (HRP)1-catalyzed chemiluminescent oxidation of luminol is widely used in many molecular biologybased assays, such as Western blots, dot blots, and enzyme-linked immunosorbent assays (ELISAs), as well as in immunohistochemistry. Much effort has been made to improve the efficiency and analytical performance of this reaction. In particular, the addition of enhancers to the reaction substrate greatly increases light output and duration kinetics [1]. Molecules with enhancer properties are essentially redox mediators [2] capable of exchanging electrons between the peroxidase enzyme and luminol. Enhancers include substituted phenols [3], substituted boronic acids [4], indophenols, and N-alkyl phenothiazine derivatives [5]. The enhanced HRP-catalyzed oxidation of luminol is a complex multistep reaction. Although redox enhancers are useful in improving enzyme turnover and increasing the equilibrium concentration of a key intermediate, luminol radical anion, a second bottleneck may need to be removed to obtain a further increase in light output. According to a well-established mechanism [6],
* Corresponding author. Fax: +39 051534063. E-mail address: [email protected] (L. Della Ciana). 1 Abbreviations used: HRP, horseradish peroxidase; ELISA, enzyme-linked immunosorbent assay; DMAP, 4-dimethylaminopyridine; MORP, 4-morpholinopyridine; PPY, 4-pyrrolidinopyridine; THF, tetrahydrofuran; PBS, phosphate-buffered saline; SPTZ, sodium 3-(100 -phenothiazinyl)propane-1-sulfonate; HPLC, high-performance liquid chromatography; DMSO, dimethyl sulfoxide; TEA, triethanolamine; BSA, bovine serum albumin; NHE, normal hydrogen electrode. 0003-2697/$ - see front matter Ó 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.ab.2008.03.020
luminol radical anion, once formed, is subject to rapid dismutation to luminol and diazaquinone. In turn, this species is attacked by hydroperoxide anion and converted into reactive peroxides. This type of reaction is often accelerated by nucleophilic acylation catalysts [7], especially 4-aminopyridines. This study investigates the effect of these compounds on chemiluminescence light output when used in association with redox enhancers and their use in substrates for HRP chemiluminescent assays with the goal of achieving a higher detectability of HRP. This will allow the development of more sensitive bioanalytical methods and also further miniaturize chemiluminescent-based chip devices.
Materials and methods Luminol sodium salt was obtained by recrystallization of the commercially available luminol (5-amino-2,3-dihydro-1,4-phthalazinedione, Fluka) from sodium hydroxide using a procedure described previously [8]. Phenothiazine (Acros), 1,3-propanesultone (Aldrich), 4-dimethylaminopyridine (DMAP, TCI–EP), 4-morpholinopyridine (MORP, Aldrich), and 4-pyrrolidinopyridine (PPY, Fluka) were purchased from their respective suppliers. HRP (type VI-A, 797 U/mg) was purchased from Fluka. Water was freshly spilled from a Milli-Q water purification system. Dry tetrahydrofuran (THF) was obtained by distillation over Na/benzophenone. All other compounds were obtained from commercial sources and
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used without further purification. Phosphate-buffered saline (PBS) was prepared in 0.1 M concentration and pH 7.3. Instrumentation A Cary Eclipse (Varian) fluorescence spectrophotometer was used for kinetic measurements using the following setup: kinetic mode, kem = 425 nm, gate time = 5 ms, emission slits properly adjusted, Savitzky–Golay smoothing. A Luminoskan (Ascent) luminometer was used for microtiter well readings (1 s integration/well). A NightOwl LB981 (Berthold Technologies) luminograph was used to read the luminescence from Western blot membranes (120 s total integration time). The detection limits for the calibration curves were obtained graphically according to the method of Hubaux and Vos [9], that is, by plotting the regression line with the confidence interval (99%) and following the point where the upper confidence interval curve intersects the y axis horizontally until reaching the lower confidence interval and then proceeding vertically to the x axis. This point on the x axis was taken as the limit of detection. Synthesis of SPTZ Because the purity of the enhancer is of fundamental importance in the chemiluminescent process [10,11], the two-step literature procedure [12] was replaced by the following improved one-pot synthesis. Sodium hydride (60% dispersion in mineral oil, 1.76 g, 44.2 mmol) was dissolved in dry THF (40 ml) under argon atmosphere. A solution of phenothiazine (8 g, 40 mmol) in dry THF (40 ml) was added, and the mixture was stirred at room temperature for 60 min (a dark orange suspension forms) and then 90 min at 50 °C. After cooling at 0 °C, a solution of 1,3-propanesultone (4.88 g, 40 mmol) in dry THF (40 ml) was added and the resulting mixture was stirred for 30 min. The ice bath was removed, and the mixture was stirred for a further 30 min. The white precipitate was filtered off and subsequently washed with THF and Et2O and further purified by crystallization from 90% ethanol to give 3-(100 -phenothiazinyl)propane-1-sulfonate (SPTZ) as white crystals (10.0 g, 72% yield) (Fig. 1). Molecular mass of the free acid (C15H15O3S2): 321.4. MS (API-ES): 322.0 [MH]+. 1H NMR (300 MHz, D2O) d: 6.9–7.1 (m, 4H, ArH), 6.7–6.9 (m, 4H, ArH), 3.8 (t, 2H, -CH2-N-, J = 7.2 Hz), 2.9 (t, 2H, -CH2-S, J = 7.2 Hz), 1.9–2.9 (m, 2H, -CH2-CH2-CH2). The content of unreacted phenothiazine in the product was found to be less than 0.0002 parts (mol/mol) by high-performance liquid chromatography (HPLC) analysis (column: 4.6 150-mm Waters C18 XTerra; eluent: H2O/acetonitrile [3:7] at 1 ml/min; UV detector: 254 nm). Effect of different enhancers on the luminol–peroxide–peroxidase reaction
in 0.1 M Tris–HCl (pH 8.6). This solution was then split into portions, and to each portion different enhancers were added from previously prepared stock solutions in dimethyl sulfoxide (DMSO); for example, the final concentration of enhancer in the working solution was as follows: (a) p-coumaric acid (0.68 mM), (b) p-iodophenyl-boronic acid (0.77 mM), (c) p-iodophenol (0.62 mM), (d) SPTZ (0.75 mM), and (e) reference (no enhancer). The final DMSO concentration was 1%. Then 2 ml of each working solution was tested in a 3-ml polymethylmethacrylate disposable cuvette by adding 10 ll of HRP solution (2 lg/ml) and vortexing for a few seconds. Then, 30 s after enzyme addition, light emission was monitored for 30 min using the fluorescence spectrophotometer. Effect of DMAP association with different enhancers Working solutions were prepared as described above. Each solution was tested by adding either 10 ll of Tris buffer or 10 ll of DMAP (200 mM) in Tris buffer. The same enzyme addition protocol described above was used. Values were calculated integrating the signal during the first 10-min run of a kinetic measurement using the fluorescence spectrophotometer. Effect of nucleophilic acylation catalysts on SPTZ-enhanced substrates The working solution was freshly prepared using the following concentrations: 5 mM luminol sodium salt, 1.5 mM SPTZ, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). This solution was split into portions, and the following compounds were added to each portion to a final concentration of 1.5 mM: (a) pyridine, (b) 4-picoline, (c) PPY, (d) DMAP, (e) MORP, (f) triethanolamine (TEA), and (g) reference (no addition of acylation catalyst). The same enzyme addition protocol described above was used. Measurements were carried out with the fluorescence spectrophotometer (15 min total acquisition time). Light emission from working solutions containing MORP: Effect of pH The working solutions NoMORP and MORP were freshly prepared. The NoMORP working solution contained 5 mM luminol, 1.5 mM SPTZ, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). The MORP working solution contained 5 mM luminol, 1.5 mM SPTZ, 1.5 mM MORP, and 4 mM sodium perborate in 0.125 M Tris–HCl (pH 9.0). The pH for each working solution was adjusted using negligible amounts of HCl (5 M) or NaOH (5 M) in the pH 8 to 10 interval. The same enzyme addition protocol described above was used. Measurements were carried out with the fluorescence spectrophotometer (10 min total acquisition time). Standard curve for HRP
The working solution was freshly prepared using the following concentrations: 1.25 mM luminol sodium salt and 2.50 mM H2O2
Samples (50 ll) of HRP diluted in Tris buffer (0.1 M, pH 8.6) in the range from 0 to 100 pg/well were analyzed in a microtiter plate using alternatively NoMORP and MORP working solutions (100 ll) prepared as described above. Light emission was measured with the luminometer (15 min total acquisition time). Application of the MORP–SPTZ–luminol chemiluminescent reaction in a sandwich ELISA
Fig. 1. Sodium 3-(100 -phenothiazinyl)propane-1-sulfonate (SPTZ) structure.
A noncompetitive enzyme immunoassay for pathogen bacterium Yersinia enterocolitica previously developed in our laboratory was used to evaluate the analytical performance of the new chemiluminescent cocktail. A mouse monoclonal antibody specific for Y. enterocolitica was immobilized into the wells of a white 96-well microtiter plate (Microlite 2+, Thermo Scientific). The antibody was
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diluted 1:1000 (v/v) in 0.05 M carbonate/bicarbonate buffer (pH 9.6), and then 100 ll was dispensed in the wells and incubated overnight at 4 °C. After washing, a second coating was carried out with 2% bovine serum albumin (BSA) in PBS, and then the wells were dried under vacuum and stored at 4 °C until use. To perform the assay, 100 ll of serial dilutions of Y. enterocolitica (ATCC 23715) lysate in PBS (ranging from 1.5 104 to 3.0 107 cells/ml) were incubated in duplicate in the wells for 1 h at 37 °C. After washing with 4% Tween in PBS, wells were incubated for 1 h at 37 °C with 100 ll of rabbit polyclonal anti-Y. enterocolitica antibody diluted 1:2000 (v/v) in PBS/Tween and washed again. The plate was then incubated for 30 min at room temperature in the dark with HRPconjugated anti-rabbit antibody (Sigma) diluted 1:8000 (v/v) in PBS and washed again. Calibration curves prepared in parallel were produced by adding either NoMORP or MORP working solution (see protocol above) and by measuring light emission with the luminometer. Light intensity was monitored for a period of 15 min. Application of MORP-added luminol–SPTZ solution in Western blot assays Serial dilutions of albumin from chicken egg white (Sigma) were electrophoresed on 6% polyacrylamide gel and then transferred to a nitrocellulose membrane. The blot was blocked in 2% dry milk for 1 h at room temperature and then washed with PBS twice for 15 min. The blot was then incubated with rabbit antichicken egg albumin (whole anti-serum, 1:4000 [v/v] dilution) for 1 h at room temperature and washed as described previously. The blot was then incubated with HRP-conjugated anti-rabbit antibody (1:2000 dilution) for 1 h at room temperature and washed again. Two blots were prepared and treated alternatively with NoMORP and MORP working solutions (see protocol above). Light emission from the membrane was measured with the luminograph. Results and discussion The spontaneous chemiluminescent oxidation of luminol (LH2) by peroxides occurs according to a well-established mechanism [6]. In neutral or alkaline solution, the monoanion of luminol (LH–) is oxidized to the corresponding radical anion (L–). Dismutation of L- regenerates LH– while producing a two-electron oxidized luminol species, a diazaquinone (L). The diazaquinone is then attacked by hydrogen peroxide anion HO2 . An intermediate peroxide species is formed and then collapses, with loss of nitrogen, to 3-aminophthalate in its excited state, AP*. The decay of AP* to aminophthalate, AP, is responsible for the chemiluminescent light emission at 425 nm. Although the spontaneous oxidation of luminol in aqueous solution is very slow, it can be catalyzed by peroxidases such as HRP. The intensity of the light emission can be further increased by adding redox mediators such as phenols, indophenols, phenothiazines, and substituted boronic acids. A quantitative mechanism of the enhanced luminescence has been proposed [2]. Although this model was developed specifically for phenolic compounds, its general features are applicable to other classes of enhancers. According to the model, the native Fe(III) enzyme (HRP) is oxidized by peroxide to HRP-I in a two-electron oxidation (see Eq. (1)). HRP-I then returns to its native state, HRP, by reaction with the enhancer (E) in two one-electron transfer steps, with HRP-II as intermediate (see Eqs. (2) and (3)). In each of these steps, the enhancer is oxidized to its radical form (E): HRP þ H2 O2 ! HRP I þ H2 O
ð1Þ
HRP I þ E ! HRP II þ E
ð2Þ
HRP II þ E ! HRP þ E :
ð3Þ
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The reaction of luminol with HRP-II is approximately 100-fold less rapid than the reaction of luminol with HRP-I. Luminescence enhancers react more rapidly with HRP-II than does luminol, thereby accelerating enzyme turnover. In the next step, the enhancer radical E oxidizes luminol anion LH– to the key intermediate L-. This reaction is reversible: E þ LH ¡E þ L :
ð4Þ
The position of the equilibrium is determined by the difference between the reduction potential of the redox couples, L-/LH– and E/E. With phenolic enhancers, it was found that the rate of reaction with HRP-II is also determined by the reduction potential of the E/E couple. For these enhancers, a bell-shaped dependence of the luminescence efficiency on the reduction potential of E was observed, with its maximum at approximately 0.8 V versus normal hydrogen electrode (NHE), roughly the same potential of luminol, 0.82 V versus NHE. Fine tuning of the reduction potential for E is required to avoid loss of reaction efficiency. N-Alkyl-substituted phenothiazine enhancers were first described by Sugiyama in the patent literature [5]. A water-soluble derivative, SPTZ (Fig. 1), was employed most often in these studies. However, no major improvement over the performance of phenolic enhancers was demonstrated. A few years later, much better results in terms of luminescent intensity and duration were reported by using a carefully purified sample of SPTZ [11]. In fact, we found that the two-step, low-yield method used for the synthesis of SPTZ [12] is largely responsible for the presence of impurities, especially unreacted phenothiazine, that quench L– and drastically reduce light output. Unfortunately, these impurities are not easily removed by conventional purification methods [11]. For these reasons, we developed a new one-step synthesis of SPTZ. Under our conditions, the SPTZ enhancer spontaneously crystallizes out of the reaction solution in high yield and extreme purity. Considering now the system in more detail, it is worth observing that the reduction potential of SPTZ (SPTZ+/SPTZ) is 0.83 V versus NHE [12], nearly identical to the value reported for luminol and close to the optimal value observed for phenolic enhancers. On the other hand, phenothiazine itself, with a reduction potential (PTZ+/ PTZ) of 0.62 V versus NHE [13], is much easier to oxidize than is SPTZ and effectively quenches the production of luminescence even when present in trace amount. Next, the performance of SPTZ and other commonly used enhancers—p-coumaric acid, p-iodophenylboronic acid, and piodophenol—was evaluated. For each enhancer, the literature concentration was used [3,4,11]. All other conditions were kept constant, including luminol and oxidant concentrations, pH, and temperature. The best performance, in terms of both luminosity and duration, was obtained with SPTZ (Fig. 2). In the next experiment, DMAP, a powerful acylation catalyst, was added to each substrate solution. Whereas a very slight signal increase ( 10%) was observed for p-iodophenol and 4-iodophenylboronic acid, the effect was much more significant for SPTZ (> 800%) (Table 1). Therefore, all further studies were focused on the SPTZ system. Because this enhancer is a salt, it is much more water soluble than other enhancers and its concentration can be raised to higher levels (e.g., 1.5 mM). An improved SPTZ substrate solution is obtained by increasing luminol and peroxide concentrations to 5 and 4 mM, respectively. In addition, raising the pH from 8.5 to 9.0 further improves the duration of luminescence and total light output. In practice, the fully optimized SPTZ substrate produces a signal twice as intense as does the unoptimized system. This optimized substrate was used to evaluate a series of pyridine acylation catalysts. Pyridines, and especially 4-dialkylaminopyrines, are wellknown nucleophilic acylation catalysts and are widely used in organic synthesis [7]. For some of these compounds, the following
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100
60
40
a b c d e f g
400
Intensity (a.u.)
80 Intensity (a.u.)
500
a b c d e
300 200 100
20 0 0
2
0 0
5
10
15
20
25
4
6 8 10 Time (min)
12
14
30
Time (min) Fig. 2. Kinetic measurement of light output intensity from luminol (1.25 mM)/H2O2 (2.5 mM)/HRP (227 pM) systems with different enhancers: (a) d, coumaric acid (0.68 mM); (b) s, p-iodophenylboronic acid (0.77 mM); (c) ., p-iodophenol (0.62 mM); (d) D, SPTZ (0.75 mM); (e) j, reference (no addition of enhancer). Data are the means of triplicate assays.
Fig. 3. Kinetic measurement of light output intensity from luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM)/HRP (227 pM) systems with 1.5 mM of different nucleophilic acylation catalysts: (a) d, pyridine; (b) s, 4-picoline; (c) ., PPY; (d) D, DMAP; (e) j, MORP; (f) h, TEA; (g) , reference (no addition of catalyst). Data are the means of triplicate assays.
1600
1200 Intensity (a.u.)
data were reported [7] (where RE is the relative effectiveness as acylation catalyst): PPY (RE 100, pKa 9.9) > DMAP (RE 40, pKa 9.7) > MORP (RE 8, pKa 8.7) >> 4-picoline (RE 0.02, pKa 6.1) > pyridine (RE 0.002, pKa 5.3). When they are added to the SPTZ substrate, the same overall reactivity is observed with some exceptions: MORP > DMAP > PPY >> 4-picoline > pyridine (Fig. 3). The apparent reversal observed with PPY, DMAP, and MORP is likely to be due at least partially to their differences in pKa. At pH 9.0, the active free base form is 66% of the total for MORP, 17% for DMAP, and only 11% for PPY. A basic nonnucleophilic amine such as TEA had practically no effect on light output. Participation in the enhanced reaction of the free base form of the acylation catalysts is confirmed by a pH dependence study whereby the optimal value for light output shifts to higher pH values. Fig. 4 shows a comparison between a working solution based on luminol/SPTZ/oxidant (NoMORP) and one with added MORP. The highest signal was obtained at pH 8.4 for NoMORP. Similar optimal pH values were observed for well-known enhancers such as p-iodophenol [3] and p-iodophenylboronic acid [4]. The addition of MORP not only increased the signal but also shifted the optimal pH to a 9.1 to 9.3 interval. A similar shift was also observed for PPY and DMAP, with both having the optimal pH in the same interval (data not shown). To understand the effect of these catalysts, it is important to observe that they do not act themselves as enhancers (no effect on light output is observed when used alone). Although it is not possible, at this stage, to exclude a synergistic effect on the enhancer, an alternative explanation is proposed by considering in more detail the reaction steps from diazaquinone (L) to a luminol peroxide intermediate [6] (Fig. 5). This reaction involves nucleophilic attack on one of the diazaquinone carbonyls by hydrogen peroxide monoanion. A hydroperoxide species that can rearrange to the endoperoxide is formed.
a b
1400
1000 800 600 400 200 0 8.0
8.5 9.0 9.5 Working solution pH
10.0
Fig. 4. Comparison of working solution pH optimal value for a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM)/HRP (227 pM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). Each point is calculated integrating the signal during the first 10-min run of a kinetic measurement.
Either compound is very unstable and collapses to 3-aminophthalate in its excited state. Perhaps pyridine acylation catalysts could facilitate hydrogen peroxide attack by converting L into a more reactive intermediate. For analytical applications, the nontoxic acylation catalyst MORP is preferred over the highly toxic DMAP and PPY. The analytical performance of MORP and NoMORP working solutions was explored by comparing results obtained in two bioassays: a sandwich ELISA for Y. enterocolitica and a Western blot assay for chicken egg white albumin. Both methods belong to the class of noncompetitive ‘‘sandwich-type” formats given that all of the analyte is captured and then revealed by means of an excess of detection
Table 1 Effect of DMAP association with different enhancers
Without DMAP With DMAP
No enhancer
p-Coumaric acid
p-Iodophenyl-boronic acid
p-Iodophenol
SPTZ
6.77 ± 0.13 6.66 ± 0.06
22.25 ± 2.73 21.99 ± 0.79
100.73 ± 6.88 109.65 ± 0.13
153.12 ± 2.41 172.09 ± 3.16
227.5 ± 13.45 2201.23 ± 317.16
Note. Data are means of triplicate measures of integrated signal intensities in arbitrary units.
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a b
10
40
8
Intensity (a.u.)
6 4
30
2 0 0
20
2
4
6
8 10 12 14
10 Fig. 5. Reaction of hydrogen peroxide with diazaquinone to form the hydroperoxide intermediate.
0 antibody. The detectability of the enzyme label is one of the most important factors determining the detection limit of noncompetitive assays provided that a low degree of nonspecific binding of the detection reagents occurs [14]. A comparison of the dose–response curve for HRP using the SPTZ-enhanced luminol/peroxide substrate with and without MORP is shown in Fig. 6. The detection limit for HRP, calculated as described previously, was 0.35 pg (8 amol)/well for the reaction containing MORP, as compared with 2.20 pg (50 amol)/well for the NoMORP substrate. Next, a sandwich chemiluminescent enzyme immunoassay for the determination of Y. enterocolitica was carried out to compare the performance of MORP and NoMORP chemiluminescent reagents. Calibration curves obtained by plotting signals subtracted of the blank (Fig. 7) show that the detection cocktail containing MORP produced for each bacteria concentration a signal approximately 1 order of magnitude higher than that obtained with NoMORP, thereby providing a steeper calibration curve. Indeed, MORP reagent allowed reaching a limit of detection of 0.6 106 cells/ml, as compared with that of NoMORP cocktail, which allowed reaching a limit of detection of 5.0 106 cells/ml. The linear range extended up to 7.5 106 cells/ml for MORP and up to 1.5 107 cells/ml for NoMORP. Finally, Fig. 8 shows Western blots for chicken egg albumin detected with an HRP conjugate using the MORP and NoMORP reagents. The much stronger signal produced by the MORP-catalyzed
a b
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150
1200 Intensity (a.u.)
100
1000
50
800
0
0 2 4 6 8 10 12 14
0
5 10 15 20 25 30 10 -6 x Bacteria Concentration (cells/ml)
Fig. 7. Sandwich ELISA tests for the detection of Y. enterocolitica using a couple of specific mouse monoclonal and rabbit polyclonal antibodies and an anti-rabbit HRP-conjugated antibody. The comparison is between a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/SPTZ (1.5 mM)/perborate (4 mM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). The inset shows the linear range with the 99% confidence interval from which the detection limits were calculated. Each point is calculated integrating the signal during the first 15-min run of a kinetic measurement. Data are the means of quadruplicate assays subtracted of the blank. Error bars: ± 1 SD.
Fig. 8. Western blots for albumin from chicken egg white detected using HRPconjugated antibody. The comparison is between a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/ SPTZ (1.5 mM)/perborate (4 mM): (a) NoMORP (no catalyst added, left to right: 19.0, 12.7, 8.4, 5.6, 3.8, 2.5, 1.7, and 1.1 pg albumin); (b) MORP (1.5 mM, left to right: 2.9, 1.9, 1.3, 0.8, 0.6, 0.4, 0.3, and 0.2 pg albumin). Light emission was detected with a charge-coupled device (CCD)-based instrument with 2 min acquisition.
substrate allowed the detection of more diluted protein bands, with a clear advantage in the Western blot technique. In conclusion, the incorporation of an acylation catalyst in enhancer/luminol/oxidant HRP substrates is highly advantageous, especially in the case of N-alkylated phenothiazine enhancers. Although the exact mechanism of action of acylation catalysts has not yet been fully investigated, it is clear from the results of this study that the very significant increase in light output observed in their presence can be translated into a corresponding improvement in sensitivity of chemiluminescent assays. Acknowledgments
600 400 200 0 0
20
40
60
80
100
HRP Amount (pg/well) Fig. 6. Comparison of HRP dose–response curves for a MORP-catalyzed system and a noncatalyzed system, both having the working solution based on luminol (5 mM)/ SPTZ (1.5 mM)/perborate (4 mM): (a) d, NoMORP (no catalyst added); (b) s, MORP (1.5 mM). The inset shows the linear range with the 99% confidence interval from which the detection limits were calculated. Each point is calculated integrating the signal during the first 15-min run of a kinetic measurement. Data are the means of quadruplicate assays subtracted of the blank. Error bars: ± 1 SD.
Thanks are due to Luisa Stella Dolci and Manuela Rizzoli for molecular biology-based assays and to Matteo Beltrame for synthesis and purification of luminol. We are also grateful to Monica Musiani and Elisabetta Manaresi for the Western blot tests at the Department of Clinical and Experimental Medicine, Section of Microbiology, University of Bologna. References [1] L.J. Kricka, J.C. Voyta, I. Bronstein, Chemiluminescent methods for detecting and quantitating enzyme activity, Methods Enzymol. 305 (2000) 370–390. [2] P.M. Easton, A.C. Simmonds, A. Rakishev, A.M. Egorov, L.P. Candeias, Quantitative model of the enhancement of peroxidase-induced luminol luminescence, J. Am. Chem. Soc. 118 (1996) 6619–6624. [3] G.H.G. Thorpe, L.J. Kricka, Enhanced chemiluminescent reactions catalyzed by horseradish peroxidase, Methods Enzymol. 133 (1986) 331–353.
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Chemiluminescent detection systems of HRP / E. Marzocchi et al. / Anal. Biochem. 377 (2008) 189–194
[4] L.J. Kricka, M. Cooper, X. Ji, Synthesis and characterization of 4-iodophenylboronic acid: A new enhancer for the horseradish peroxidase-catalyzed chemiluminescent oxidation of luminol, Anal. Biochem. 240 (1996) 119–125. [5] M. Sugiyama, Method of the chemiluminescence assays of the activity of peroxidase, U.S. patent no. 5,171,668 (1992). [6] F. McCapra, Chemical generation of excited states: The basis of chemiluminescence and bioluminescence, Methods Enzymol. 305 (2000) 3– 47. [7] G. Höfle, W. Steglich, H. Vorbrüggen, 4-Dialkylaminopyridines as highly active acylation catalysts, Angew. Chem. Intl. Ed. Engl. 17 (1978) 569–583. [8] G. Ham, R. Belcher, L.J. Kricka, T.J.N. Carter, Stability of trace iodine solutions, Anal. Lett. 12 (1979) 535–541.
[9] A. Hubaux, G. Vos, Decision and detection limits for linear calibration curves, Anal. Chem. 42 (1970) 849–855. [10] M.A. Motsenbocker, K. Kondo, Improvements to enhanced horseradish peroxidase detection sensitivity, J. Biolumin. Chemilumin. 9 (1994) 15–20. [11] P. D. Davis, Assay of peroxidase activity, U.S. patent no. 6,432,662 (2002). [12] Y. Kawanishi, N. Kitamura, S. Tazuke, Coulombic effect of photoinduced electron-transfer reactions between phenothiazines and viologens, J. Phys. Chem. 90 (1986) 2469–2475. [13] C. Bodea, I. Silberg, Recent advances in the chemistry of phenothiazines, Adv. Heterocycl. Chem. 9 (1968) 321–460. [14] A. Roda, M. Guardigli, E. Michelini, M. Mirasoli, P. Pasini, Analytical bioluminescence and chemiluminescence, Anal. Chem. 75 (2003) 463–470.