Detection of organophosphorus compounds by covalently immobilized organophosphorus hydrolase

Article abstract of DOI:10.1021/ac061118m

As a consequence of organophosphorus (OP) toxins posing a threat to human life globally, organophosphorus hydrolase (OPH) has become the enzyme of choice to detoxify such compounds. Organophosphorus hydrolase was covalently immobilized onto a quartz subst

Full text of DOI:10.1021/ac061118m

Anal. Chem. 2006, 78, 7016-7021
Detection of Organophosphorus Compounds by
Covalently Immobilized Organophosphorus
Hydrolase
†
†
‡
‡
Jhony Orbulescu, Celeste A. Constantine, Vipin K. Rastogi, Saumil S. Shah,
‡
,†
Joseph J. DeFrank, and Roger M. Leblanc*
Department of Chemistry, University of Miami, Coral Gables, Florida, 33146, and U.S. Army Edgewood Chemical Biological
Center, Biotechnology Team, Research and Technology Directorate, Aberdeen Proving Ground, Maryland 21010-5423
As a consequence of organophosphorus (OP) toxins
posing a threat to human life globally, organophosphorus
hydrolase (OPH) has become the enzyme of choice to
detoxify such compounds. Organophosphorus hydrolase
was covalently immobilized onto a quartz substrate for
utilization in paraoxon detection. The substrate was
cleaned and modified prior to chemical attachment. Each
modification step was monitored by imaging ellipsometry
as the thickness increased with each modification step.
The chemically attached OPH was labeled with a fluores-
cent dye (7-isothiocyanato-4-methylcoumarin) for the
detection of paraoxon in aqueous solution, ranging from
Organophosphorus hydrolase (OPH) enzyme can hydrolyze
a large variety of organophosphorus compounds by producing less
toxic products such as p-nitrophenol and diethyl phosphate. OPH
5
is one of the most studied enzymes related to its activity toward
pesticides and nerve agents. It has been shown that OPH cata-
lyzes hydrolysis reactions that induce cleavage in P-O, P-S, P-F,
6
and P-CN bonds within organophosphorus neurotoxins. Numer-
ous analytical methods have been devised to detect the pres-
7
,8
ence of OP compounds, e.g., gas chromatography, high-per-
9,10
formance liquid chromatography, and other methods of detec-
tion, such as electrochemical.1
1-14
Most of these techniques are
time-consuming, expensive, and complex. To overcome these dis-
advantages, enzymatic bioassays have been designed for en-
hanced speed of detection, high efficiency, sensitivity, and cost-
effectiveness.15
-
9
-5
1
0
to 10 M. UV-visible spectra were also acquired
for the determination of the hydrolysis product of para-
oxon, namely p-nitrophenol.
Several viable methodologies of enzyme immobilization are
now explored.16 Immobilization is important because it stabilizes
the enzyme, and in many cases, the enzyme can be recovered
Organophosphorus (OP) derivatives are harmful compounds
found in insecticides and pesticides and are widely utilized around
17
and its activity regenerated. These include physical immobili-
1
the world. These OP compounds are structurally similar to nerve
18-20
21-23
24,25
zation,
encapsulation,
cross-linking,
and chemical bind-
gases and are well-known inhibitors of acetylcholinesterase, which
is responsible for transmitting nerve impulses across synaptic
(5) Munnecke, D. M. Biotechnol. Bioeng. 1979, 21, 2247-2261.
(6) Russell, R.; Pishko, M.; Simonian, A. S.; Wild, J. R. Anal. Chem. 1999, 71,
2
junctions. As a consequence of the acute toxicity of these
4909-4912.
neurotoxins, stringent environmental monitoring programs have
been implemented to ensure that both groundwater and soil
concentrations of OP derivatives remain below harmful toxic
(
(
(
7) Mendoza, C. E. Thin-layer chromatography. In Pesticide Analysis; Dumas,
K. G., Ed.; Marcel Dekker: New York, 1981; pp 1-44.
8) Das, K. G.; Kulkarni, P. S. Gas-liquid chromatography. In Pesticide Analysis;
Dumas, K. G., Ed.; Marcel Dekker: New York, 1981.
9) Hanks, A. R.; Colvin, B. M. High-performance liquid chromatography. In
Pesticide Analysis; Dumas, K. G., Ed.; Marcel Dekker: New York, 1981; pp
3
levels. Paraoxon is the oxidative desulfuration product of the
4
pesticide parathion when in contact with oxygen. OP compounds
99-174.
are poisonous when ingested, inhaled, or absorbed through the
skin. Therefore, development of sensors capable of determining
harmful concentrations of theses compounds would be beneficial
(
10) Barcelo, D.; Lawrence, J. F. Residue analysis of organophosphorus pesticides.
In Emerging Strategies for Pesticide Analysis; Charins, T., Sherma, J., Eds.;
CRC Press: Boca Raton, FL, 1992; pp 127-150.
3
(11) Palchetti, I.; Cagnini, A.; Del Carlo, M.; Coppi, C.; Mascini, M.; Turner, A.
to both human and livestock populations.
P. F. Anal. Chim. Acta 1997, 337, 315-321.
(12) Liu, G.; Lin, Y. Anal. Chem. 2005, 77, 5894-5901.
*
Corresponding author. Tel: +1-305-284-2194. Fax: +1-305-284-6367.
(13) Liu, G.; Lin, Y. Anal. Chem. 2006, 78, 835-843.
(14) Lei, Y.; Mulchandani, P.; Wang, J.; Chen, W.; Mulchandani, A. Environ. Sci.
Technol. 2005, 39, 8853-8857.
(15) Dumschat, C.; Muller, H.; Stein, K.; Schwede, G. Anal. Chim. Acta 1991,
252, 7-9.
(16) Liu, Y.; Wang, C.; Hsiung, K. Anal. Biochem. 2001, 299, 130-135.
(17) Mesthrige, K. W.; Amro, N.; Liu, G. Scanning 2000, 22, 380-388.
(18) Ihalainen, P.; Peltonen, J. Langmuir 2002, 18, 4953-4962.
(19) Yan, A. X.; Li, X. W.; Ye, Y. H. Appl. Biochem. Biotechnol. 2002, 101, 113-
129.
(20) Lee, J. G.; Lee, W. C. Biotechnol. Appl. Biochem. 1998, 27, 225-230.
(21) Decher, G. Science 1997, 277, 1232-1237.
E-mail: rml@miami.edu.
†
University of Miami.
U.S. Army Edgewood Chemical Biological Center.
‡
(
(
1) Munnecke, D. J. Agric. Food Chem. 1980, 28, 105-111.
2) Quinn, D. M.; Selwood, T.; Pryor, A. N.; Lee, B. H.; Leu, L. S.; Acheson, S.
A.; Silman, I.; Doctor, B. P.; Rosenberry, T. L. In Multidisciplinary Approaches
to Cholinesterase Functions; Shafferman, A., Velan, B., Eds.; Plenum Press:
New York, 1992.
3) Mulchandani, A.; Pan, S.; Chen, W. Biotechnol. Prog. 1999, 15, 130-134.
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IL, 1972.
(
(
7016 Analytical Chemistry, Vol. 78, No. 19, October 1, 2006
10.1021/ac061118m CCC: $33.50 © 2006 American Chemical Society
Published on Web 08/10/2006

ing to a solid substrate.26-30 Covalent immobilization results in
an increased stability since the attachment is not reversed by
factors such as pH, ionic strength, substrate, solvent, or tem-
during paraoxon hydrolysis. Coumarin is a widely used fluoro-
phore; this UV-excitable dye possesses excellent spectroscopic
properties, e.g., large Stokes shift (larger than 80 nm) and medium
quantum efficiency of fluorescence (Φ ) 0.5). These key char-
acteristics make coumarin derivatives very attractive for use as
labels.4 In addition, the emission of the coumarin dye (∼430
nm) can overlap with the absorbance of the hydrolysis product,
namely, p-nitrophenol (∼400 nm), leading to fluorescence reso-
nance energy transfer (FRET) that can quench the coumarin
emission.
3
1-33
perature.
The orientation can be predicted if small molecules
3
4
or receptors are attached. Metal oxide surfaces contain surface
hydroxyl groups that are useful for the coupling of organic
1,42
3
5
materials. Carbon electrodes, after chemical treatment, possess
different surface-associated functional groups such as carboxylic,
carbonyl, lactone, and hydroxyl groups, which make the covalent
attachment of enzymes possible.36,37
The number of ways an enzyme can be immobilized onto the
solid surface is restricted by the groups that can be modified and
present on the exterior of the enzyme “shell”. Based on this
restriction, we choose the modification of the free amine groups
from any peripheral amino acids of the enzyme as is well known
that amines can be modified easily without requiring harsh
conditions that might affect the enzyme activity. In this investiga-
tion, quartz slides are functionalized with (3-aminopropyl)tri-
methoxysilane. The terminal amine group is then modified with
diethylthiocarbamoyl chloride to form the isothiocyanate group.
Isothiocyanates are known to react with amine groups in very mild
conditions. OPH through the amine groups of the lysine, glutamic,
In the present investigation, covalent binding of OPH to a
silanized quartz slide was performed. Diethylthiocarbamoyl chlo-
ride was employed as a reagent for modifying the amino-modified
quartz slide surface, which was then used for covalently attaching
enzymes. OPH was then labeled with a coumarin derivative
fluorophore. This sensing device was utilized in monitoring the
paraoxon hydrolysis reaction using UV-visible and fluorescence
spectroscopies.
EXPERIMENTAL SECTION
The quartz slides were obtained from Hellma Inc. (Plainview,
NY). (3-Aminopropyl)trimethoxysilane used for the silanization
of the quartz slide and diethylthiocarbamoyl chloride were
obtained from Aldrich (Milwauke, WI). Carbonate-bicarbonate
buffer capsules, pH 9.6, were purchased from Sigma (St. Louis,
MO). Spectroscopic grade solvents were obtained from Fischer
Scientific (Fair Lawn, NJ). The 7-isothiocyanato-4-methylcoumarin
was synthesized from the commercially available 7-amino-4-
19,38,39
or aspartic residues on the periphery of the enzyme
can react
easily with the isothiocyanate groups from the substrate surface.
Because the reaction occurs on the “outside” of the enzyme
molecules, the binding sites of the OPH molecules remain
4
0
available to interact with an external substrate.
An enzyme can be utilized as a bioassay if it can provide
qualitative and quantitative data regarding the reaction of interest,
hydrolysis of OP compounds in this case. The hydrolysis product
of paraoxon is p-nitrophenol, which has absorbance in UV region,
but even with its relatively high extinction coefficient (ꢀ400 ) 18 000
4
3
methylcoumarin obtained from Aldrich (Milwaukee, WI). OPH
85-90%) (EC 3.1.8.1) was isolated, extracted, and purified at the
(
U.S. Army Laboratory (Edgewood Chemical and Biological Center,
MD). A stock solution of OPH (1.8 mg/mL) was prepared in 100
mM bis-tris-propane, pH 7.3, containing 10 µM Co2 . The stock
solution was frozen at -4 °C. The stock solution was diluted to a
concentration of 0.18 mg/mL prior to use. Pure water was
provided from the Modulab 2020 water purification system
+
-1
-1
M
‚cm ), only fairly concentrated solutions of p-nitrophenol can
be measured by UV-visible spectroscopy. It is well known that
fluorescence spectroscopy can detect much lower concentrations
than UV-visible spectroscopy. Based on this fact, we decided to
label the OPH enzyme with a reliable fluorophore whose emission
can be affected by the formation in solution of the p-nitrophenol
(Continental Water Systems Corp., San Antonio, TX). The resis-
tance and surface tension of pure water were 18 MΩ‚cm and 72.6
mN/m at 20.0 ( 0.5 °C, respectively. Attenuated total reflectance
(
ATR)-FT-IR spectra were recorded on an Equinox 55 FT-IR
(
(
(
22) Wei, Y.; Xu, J. G.; Feng, Q. W. J. Nanosci. Nanotechnol. 2001, 1, 83-93.
23) Wei, Y.; Xu, J. G.; Feng, Q. W. Mater. Lett. 2000, 44, 6-11.
24) Carbone, K.; Casarci, M.; Varrone, M. J. Appl. Polym. Sci. 1999, 74, 1881-
spectrometer (Bruker Optics Inc., Billerica, MA) with a 25
reflection variable angle ATR P/N1 1000 (Specac Inc., Smyrna,
GA) accessory. The modulation frequency was set at 1666 cm
and 300 scans were collected for each spectrum at a resolution of
4 cm . KRS-5 crystal was used, and the incidence angle was set
at 60°. The changes in thickness of the films upon modification
were measured using an I-Elli2000 imaging ellipsometer (Nano-
film, G o¨ ttingen, Germany). A Lambda 900 UV-visible-NIR
spectrophotometer (Perkin-Elmer, Boston, MA) using a quartz
cuvette of 1-cm optical path length was used to measure the
absorption spectra. The fluorescence spectra were obtained using
a Spex Fluorolog 1680 spectrophotometer (Jobin Yvon, Inc.,
Edison, NJ).
1
889.
-1
,
(25) Ulbricht, M.; Papra, A. Enzyme Microb. Technol. 1997, 20, 61-68.
(26) Norde, W.; Zoungrana, T. Biotechnol. Appl. Biochem. 1998, 28, 133-143.
(27) Alkota, I.; Garbisu, C.; Llama, M. J. Enzyme Microb. Technol. 1996, 18,
-
1
1
41-146.
(28) Wang, P.; Dai, S.; Waezsada, S. D. Biotechnol. Bioeng. 2001, 74, 249-255.
(29) Chae, H.; Kim, E. Appl. Biochem. Biotechnol. 1998, 73, 195-204.
(30) Mateo, C.; Fernandez-Lorente, G.; Abian, O. Biomacromolecules 2000, 1,
7
39-745.
(
(
31) Guilbault, G. Analytical Uses of Immobilized Enzymes; Marcel Dekker: New
York, 1984.
32) Bayramoglu, G.; Kacar, Y.; Denizli, A.; Arica, M. J. Food Eng. 2002, 52,
3
67-374.
(33) Willner, I.; Katz, E. Angew. Chem., Int. Ed. 2000, 39, 1180-1218.
(34) Novak, I.; Kovac, B. J. Electron Spectrosc. Relat. Phenom. 2000, 113, 9-13.
(35) Finklea, H.; Vithanage, R. J. Phys. Chem. 1982, 86, 3621-3626.
(36) Razumas, J.; Jasaitis, J. Bioelectrochem. Bioenerg. 1984, 12, 297-322.
(37) Puri, B. R. In Chemistry and Physics of Carbon; Walker, P. L., Jr., Ed.; Marcel
Dekker: New York, 1970.
Covalent Immobilization of OPH-Coumarin. Sample sub-
strates were cleaned by immersion in a chromic mixture and
(41) Kele, P.; Orbulescu, J.; Calhoun, T. L.; Leblanc, R. M. Langmuir 2002, 18,
8523-8526.
(42) Kele, P.; Orbulescu, J.; Calhoun, T. L.; Leblanc, R. M. Tetrahedron Lett.
(
(
38) Katz, E.; Riklin, A.; Willner, I. J. Electroanal. Chem. 1993, 354, 129-144.
39) Willner, I.; Riklin, A.; Shoham, B.; Rivenzon, D.; Katz, E. Adv. Mater. 1993,
5, 912-915.
2002, 43, 4413-4416.
(
40) Gregorius, K.; Theisen, M. Anal. Biochem. 2001, 299, 84-91.
(43) Kele, P., Ph.D. Thesis, University of Miami, 2002.
Analytical Chemistry, Vol. 78, No. 19, October 1, 2006 7017

Scheme 1. Reaction Steps for OPH Immobilization: (A) Surface Silanization and Activation; (B)
Enzyme Immobilization, (C) Immobilized Enzyme Labeling
sonicated for 30 min. This was followed by alternate rinsing and
sonication. The steps showing the covalent binding of OPH are
presented in Scheme 1. In the first step prior to immobilization
of OPH, the substrate cleaned and activated (OH groups on the
surface), was silanized by reaction with (3-aminopropyl)trimeth-
groups were reacted with diethylthiocarbamoyl chloride producing
the isothiocyanatopropyl-modified slide (Scheme 1A, step 2). The
isothiocyanate groups subsequently reacted with the amino groups
on the surface of OPH. The secondary structure of OPH was
investigated to confirm the presence of the enzyme (Figure 2).
The IR spectrum of the enzyme shows the amide I and II bands,
4
4
oxysilane. The amino-modified slides were reacted with dieth-
ylthiocarbamoyl chloride at 70 °C for 10 h in order to convert the
amino to isothiocyanate groups. The isothiocyanate functional
group was then used for covalent binding of OPH.4 This process
required another 10-h incubation time while the functionalized
quartz slides were immersed in an aqueous buffered solution of
OPH (0.2 mg/mL OPH in 0.05 M carbonate-bicarbonate buffer,
pH 9.6). After immobilization, the slides were washed thoroughly
to remove all unreacted and physically adsorbed OPH. The
covalently attached OPH was labeled with 7-isothiocyanato-4-
methylcoumarin for fluorescence spectroscopic studies. The
covalently immobilized OPH-coumarin quartz slides were stored
in a phosphate buffer solution (pH 8.6) prior to use.
-
1
which are in the region of 1700-1600 and 1600-1500 cm
,
respectively. The two amide regions were clearly observed. The
amide I region contains the greatest information for the analysis
of enzyme secondary structure. The peaks were assigned to the
corresponding secondary structure as shown in Table 1. It can
be seen that the covalently bound OPH film exhibited absorbance
bands for both the R-helix and â-sheet conformations. The
5
-
1
frequency peaks at 1657, 1647, 1550, and 1534 cm correspond
-
1
to the R-helix and those at 1693, 1679, 1634, and 1619 cm
correspond to the â-sheet conformations.
RESULTS AND DISCUSSION
Characterization of the covalent reaction. ATR-IR spectros-
copy was utilized to confirm the functionality of the new bonds
-
1
formed. Two bands were noted at 2853 and 2924 cm after the
first step of the reaction (Scheme 1A), and these peaks represent
the C-H symmetric and asymmetric stretching modes, respec-
tively, which is due to the presence of the alkyl chain (Figure 1).
Although the amino group could not be observed from the ATR
spectrum, the C-H vibration provides strong enough evidence
to confirm the initial silanization step. Hereafter, the quartz slide
was essentially prepared for the following steps where the amino
(
44) Zehl, A.; Cech, D. Liebigs Ann. Chem. 1997, 3, 595-600.
(
45) Moon, J.; Shin, J.; Kim, S.; Park, J. Langmuir 1996, 12, 4621-4624.
Figure 1. ATR-IR spectrum of silanized quartz slide.
7018 Analytical Chemistry, Vol. 78, No. 19, October 1, 2006

It can be seen that the surface is not covered homogeneously
with the OPH enzyme. This is very likely the effect of the mild
experimental conditions used in the OPH attachment in order to
maintain the enzyme activity unaltered. As a last step, the
chemically attached OPH was labeled with 7-isothiocyanato-4-
methylcoumarin by reacting free amine groups from the enzyme
with the isothiocyanate from the coumarin (Scheme 1C). Follow-
ing surface modification and enzyme labeling, the modified slides
called thereafter a sensing device was used for the investigation
of the paraoxon hydrolysis by the surface-attached OPH using
spectroscopic methods.
UV-Visible Absorption Studies of Covalently Immobilized
OPH. To ensure that the covalently attached OPH was labeled
with the coumarin derivative, a UV-visible absorption spectrum
of the sensing device was measured and compared to the one of
7
-isothiocyanato-4-methylcoumarin in aqueous solution. Figure 4
Figure 2. ATR-IR spectrum of covalently bound OPH.
shows spectrum 1 of the coumarin derivative in aqueous solution
with an absorption band situated at 341 nm characteristic of the
coumarin derivatives. The presence of the isothiocyanate group
at position 7 shifts the absorption maximum to shorter wavelength
Table 1. IR Band Assignment Indicating the Secondary
Structure of OPH Covalently Bound to the Quartz Slide
frequency band
-
1
47
position (cm
)
assignment
as shown by spectrum 2. This spectrum confirms the presence
of the coumarin attached to the OPH enzyme.
1
1
1
1
1
1
1
1
693
679
657
647
634
619
550
534
antiparallel â-sheet or pleated turn
â-sheet
R-helix
amide I
â-sheet
â-sheet
R-helix
amide II
The covalently attached OPH-coumarin was exposed to
paraoxon solution, which was hydrolyzed by OPH into its products
diethyl phosphate and p-nitrophenol. The UV-visible absorption
-
6
spectrum of paraoxon solution (1 × 10 M) is shown in Figure
5, curve 1 with an absorption maximum at 274 nm. After recording
the absorption spectrum of the paraoxon solution in a 1-cm optical
path length cuvette, the sensing device was placed into the
solution for 30 s. The sensing device was then removed and the
absorption spectrum of the solution measured again. A new band
situated at 400 nm was observed and corresponds to the absorp-
tion maximum of the reaction product, namely, p-nitrophenol
Imaging ellipsometry was also used to characterize each step
of the reaction scheme (see Scheme 1). The thickness for each
step was calculated directly from the ellipsometric angles where
the incident angle was set up at 53°. To calculate the thickness
(Figure 5, curve 2). The process of placing and removing the
(
d) for each step, an optical model was used that consisted of
sensing device from the paraoxon solution at different time periods
permitted us to follow the kinetics of p-nitrophenol formation. As
shown in the inset in Figure 5, the decrease in paraoxon
absorption at 274 nm was paralleled by an increase in p-nitrophenol
absorption at 400 nm as exposure time of the sensing device
increased. The symmetry (inset in Figure 5) between the decrease
in absorbance for the paraoxon and the increase in absorbance
for the generated p-nitrophenol is a direct proof that an amount
of paraoxon is hydrolyzed by the OPH enzyme and a similar
amount of p-nitrophenol is formed, and no other effects are
involved in the changes in the absorbance.
An isosbestic point at 318 nm in the absorption spectrum as
shown in Figure 5 was observed, and this suggests the presence
of two species, namely, paraoxon as reactant and p-nitrophenol
as product; no other intermediates were involved in the hydrolysis
reaction. The meaning of the isosbestic point is that the total
concentration of the two species is a constant and when the
concentration of paraoxon is decreasing a similar increase in the
p-nitrophenol concentration is observed. In addition, the molar
absorptivity of the two species is constant at 318 nm. The sensing
device’s fast detection time was shown by the ability to detect
several parameters. The refractive index (n) and extinction
coefficient (k) were 1.46 and 0, respectively, for the substrate used.
The refractive index for air and for the film was 1.0 and 1.50,
respectively. The amplitude (∆) and the phase (ψ) of the wave
was 171.13 and 4.97, respectively. The wavelength of the laser
was at 532 nm. The average thickness generated in the 2-D
ellipsometric images in Figure 3A and B are indicated by the red
color. Figure 3A presents the thickness map generated in both
-D and 3-D for the silanization step (Scheme 1A, step 1). The
average thickness of the silanization step was determined to be
10 Å. This was in close agreement with the CPK value of 8 Å.
The scale of -5 to +5 nm, on the right side of the image, has no
physical meaning and it was used only for a better illustration in
terms of colors for the measured thickness. The 3-D map (Figure
2
∼
3
1
B) shows a homogeneous surface. The second step (Scheme
A, step 2) was not imaged because the resolution of the
instrument is (1 Å and a theoretical value of 9 Å is expected,
which is ∼1 Å thicker than the first step. The third step (Scheme
1
B), which indicates the covalent attachment of OPH, is shown
in Figure 3B. The average thickness generated is between 50 and
0 Å. An expected value of 58 Å was calculated based on the X-ray
crystalline structure of OPH with an overall dimension of ap-
6
(
(
46) Benning, M. M.; Kuo, J. M.; Raushel, F. M.; Holden, H. M. Biochemistry
994, 33, 15001-15007.
47) Haugland, R. P. Handbook of fluorescent probes and research chemicals, 6th
1
4
6
proximately 51 Å × 55 Å × 51 Å. There is a good correlation
between the experimental average value and the theoretical one.
ed.; Molecular Probes, Inc.: Eugene, OR, 1996.
Analytical Chemistry, Vol. 78, No. 19, October 1, 2006 7019

Figure 3. Ellipsometric characterization: (A) and (B) 2-D and 3-D images, respectively, of silanized quartz slide; (C) and (D) 2-D and 3-D
images, respectively, of covalently bound OPH.
Figure 4. UV-visible spectra of 7-isothiocyanato-4-methylcoumarin
-
6
in solution (1 × 10 M, 1-cm optical path length) (9) and as a label
on immobilized OPH (0).
Figure 5. UV-visible absorption spectra of the hydrolysis product
p-nitrophenol at different time intervals for the detection of paraoxon
-
6
solution (1) 1 × 10 M. Incubation time of the sensing device (see
last modification step in Scheme 1) in paraoxon solution: (2) 30 s
and (3-7) 1, 2, 5, 10, and 15 min. Inset: decrease in paraoxon
absorption (1) and the corresponding increase in p-nitrophenol
absorption (4); 1-cm optical path length.
p-nitrophenol after only 30 s of exposure. We interpret the waiting
time of 30 s as time needed for the accumulation of sufficient
p-nitrophenol in solution for detection. Our reasoning is based
on the previous work of Benning et al.48 and Omburo et al.,49 who
(
48) Benning, M. M.; Kuo, J. M.; Raushel, F. M.; Holden, H. M. Biochemistry
995, 34, 7973-7978.
49) Omburo, G. A.; Kuo, J. M.; Millins, L. S.; Raushel, F. M. J. Biol. Chem. 1992,
9, 13278-13283.
showed that the paraoxon hydrolysis occurs very fast, as sug-
1
gested by their kinetic measurements, i.e., K
m
) 45 ( 5, kcat )
(
1
1520 ( 40, and kcat/K ) 3.4 ( 0.3. The mechanism and kinetics
m
7020 Analytical Chemistry, Vol. 78, No. 19, October 1, 2006

bound enzyme hydrolyses paraoxon into p-nitrophenol. The
mechanism responsible for the quenching of the fluorescence
intensity is attributed to the F o¨ rster FRET. The inset of Figure 6
presents the spectral overlap between the absorption of p-nitro-
phenol (acceptor) and fluorescence of the coumarin fluorophore
(donor). As paraoxon is hydrolyzed, excited-state energy from the
initially excited 7-isothiocyanato-4-methylcoumarin is transferred
to p-nitrophenol, thereby leading to the observed quenching
phenomenon. After enzyme-bound quartz slides were used, they
2 4
were washed in water and KH PO -NaOH buffer solution (pH
8.6). Curve 7 shows the fluorescence intensity after recovery. This
process removed some of the product from the sensing device.
Removal of the product allowed the coumarin label to regain 60%
of its fluorescence intensity. Possible reasons for only 60% recovery
of the fluorescence are related to the fact that the enzyme-bound
quartz slide used for coumarin labeling was not sonicated but
rinsed with solvent, and secondary structure alteration can occur
if sonication is used. It is possible that some physically adsorbed
enzyme was present on the surface. Following the experiments,
storage of the modified slide in buffer solution could remove the
adsorbed enzyme; thus, the recovery would give a lower percent
that the initial emission of the labeled attached enzyme measured
before the experiments. Second and most probable is the presence
of some coumarin derivative that interacted with the enzyme
surface but not chemically bonded. If so, storing the slide in buffer
solution could remove the coumarin molecules adsorbed onto the
enzyme surface by hydrogen bonding or π-π stacking between
the aromatic ring of the coumarin and aromatic side chains of
tyrosine, tryptophan, or phenylalanine present on the enzyme
surface.
Figure 6. Fluorescence spectra of coumarin-labeled immobilized
OPH in the presence of different concentrations of paraoxon solution
(
λexc ) 340 nm, 1-cm optical path length): (1) solid substrate labeled
-
9
-8
-7
with OPH-coumarin, (2) 5 × 10 , (3) 5 × 10 , (4) 5 × 10 , (5) 1
-6
-5
×
10 , and (6) 1 × 10 M, and (7) recovery of slide. Inset: overlap
between the absorption of the acceptor (p-nitrophenol) and the
fluorescence of the donor (coumarin-labeled enzyme).
of paraoxon hydrolysis were recently studied in detail by Aubert
5
0
et al., who showed that among four substrates investigated the
kinetics appeared faster for paraoxon. It was also reported by
51
Brise n˜ o-Roa et al. that, among the organophosphorus compounds
studied, the hydrolysis reaction was faster for paraoxon. Since
the catalysis step is fast, the 30 s was needed to accumulate
p-nitrophenol at a concentration measurable by UV-visible
absorption spectroscopy. This interpretation is in agreement with
CONCLUSION
5
2
Silanization was successfully performed to functionalize a
the data reported by Caldwell et al. about the effect of solvent
viscosity and structure-reactivity relationships.
quartz substrate in order to provide an environment suitable for
the covalent binding of OPH. Rapid detection of paraoxon can be
carried out using both UV-visible and fluorescence spec-
troscopies. Fluorescence spectroscopy is more sensitive and allows
Fluorescence Emission Spectroscopic Studies of Co-
valently Immobilized OPH. Fluorescence measurements were
recorded at different concentrations of paraoxon solution when
exposed to the sensing device. From the UV-visible spectroscopic
data, it was determined that the analyte required 30 s of exposure
to the sensing device in order to detect the hydrolysis product,
p-nitrophenol. Increasing concentration of paraoxon solutions were
exposed to the sensing device, and the results are presented in
Figure 6. The initial fluorescence intensity of the fluorophore is
shown by curve 1 (λEx ) 350 nm, λEm ) 445 nm). On exposure to
paraoxon solution, there is a slight decrease of the fluorescence
intensity (curves 1-4) followed by a more drastic quenching of
the fluorescence intensity (curves 5 and 6) as the chemically
-
9
a lower detection limit (5 × 10 M) than UV-visible spectros-
copy. The results were reproducible and show that the fluores-
cence intensity of the immobilized OPH-coumarin was partially
recovered (60%). Based on the sensitivity and high efficiency of
the sensing device, it could lead to a prototype for an industrial
biosensor that can be implemented for environmental monitoring
of organophosphorus compounds.
ACKNOWLEDGMENT
This work was supported by a grant from the U.S. Army
Research Office under contract DAAD19-03-1-0131.
(
(
50) Aubert, S. D.; Li, Y. C.; Raushel, F. M. Biochemistry 2004, 44, 5707-5715.
51) Brise n˜ o-Roa, L.; Hill, J.; Notman, S.; Sellers, D.; Smith, A. P.; Timperley,.M.;
Wetherell, J.; Williams, N. H.; Williams, G. R.; Fersht, A. R.; Griffiths, A. D.
J. Med. Chem. 2006, 49, 246-255.
Received for review June 20, 2006. Accepted July 14,
2006.
(52) Caldwell, S. R.; Newcomb, J. R.; Schlecht, K. A.; Rauschel, F. M. Biochemistry
1991, 30, 7438-7444.
AC061118M
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