Analytica Chimica Acta 442 (2001) 15–23
Enzyme-based biosensor for the direct detection of
fluorine-containing organophosphates
A.L. Simonian a,∗ , J.K. Grimsley a , A.W. Flounders b , J.S. Schoeniger b ,
Tu-Chen Cheng c , J.J. DeFrank c , J.R. Wild a
a
Biochemistry and Biophysics Department, Texas A&M University, College Station, TX 77843-2128, USA
b Chemical and Radiation Detection Laboratory, Sandia National Laboratories, Livermore, CA, USA
c US Army Edgewood Chemical Biological Center, Livermore, CA, USA
Received 8 December 2000; received in revised form 3 May 2001; accepted 18 May 2001
Abstract
The ability of the enzyme organophosphorus acid anhydrolase (OPAA) to selectively hydrolyze the P–F bond of fluorine
containing organophosphates has been used to develop a biosensor for specific detection of these compounds. Hydrolysis
rate of diisopropyl fluorophosphate (DFP), paraoxon and demeton-S, by soluble and immobilized OPAA was measured.
These compounds were selected as representative substrates of OPAA hydrolysis of P–F, P–O and P–S bonds, respectively.
Results indicate that hydrolysis of phosphofluoridates such as DFP is dominant while hydrolysis of phosphotriesters such
as paraoxon or of phosphothiolates such as demeton-S, is negligible. Two experimental approaches were used for biosensor
development. In the first, OPAA was covalently immobilized on silica gel and used in batch-mode measurements with flat
glass pH electrode to detect pH changes due to P–X bond cleavage. In the second approach, the enzyme was covalently
immobilized to the porous silica modified gate insulator of a pH-sensitive field effect transistor (pH-FET) and changes in pH
relative to a second non-enzyme coated pH-FET were measured in stop-flow mode. Concentrations of DFP down to 25 ␮M
with the glass electrode and 20 ␮M with the pH-FET were readily detected. No sensor response was observed with paraoxon
or demeton-S indicating that such OPAA-based biosensors could be useful for direct and discriminative detection of fluorine
containing organophosphorus neurotoxins (such as the G-type chemical warfare agents sarin GB and soman GD) in samples
also containing multiple organophosphate pesticides. © 2001 Elsevier Science B.V. All rights reserved.
Keywords: Organophosphorus acid anhydrolase; OPAA; pH-FET; ENFET; Biosensor; Diisopropyl fluorophosphate; Paraoxon; Demeton-S
1. Introduction
The detection of phosphofluoridate neurotoxins in
field collected water and soil samples is an extremely
difficult challenge. Soil and water samples are very
likely to contain organophosphate pesticides due to
∗
Corresponding author. Tel.: +1-979-845-6840;
fax: +1-979-845-9274.
E-mail address: [email protected] (A.L. Simonian).
heavy agricultural use of these compounds. It is essential that phosphofluoridate neurotoxins can be readily
and unequivocally distinguished from chemically similar agricultural chemicals; ubiquitous organophosphate pesticides must never be interpreted as false
positive indication of chemical warfare (CW) agents.
Sensitive biosensors based on acetylcholinesterase
(AChE) or butyryl cholinesterase (BChE) inhibition
have been developed and used for environmental monitoring [1–5]. However, cholinesterase-based sensors
0003-2670/01/$ – see front matter © 2001 Elsevier Science B.V. All rights reserved.
PII: S 0 0 0 3 - 2 6 7 0 ( 0 1 ) 0 1 1 3 1 - X
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A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
provide no discrimination between organophosphate
pesticides and organophosphate CW agents and produce uncertain and often contradictory signals in response to multiple classes of cholinesterase inhibitors
(e.g. organophosphate and carbamate) [6–8]. One
approach to improve the specificity of cholinesterase
sensors has been to monitor inhibition with both
butyryl and acetyl cholinesterase [9] or to analyze inhibition patterns obtained with several acetyl
cholinesterase enzymes from diverse sources [10].
We believe the preferred strategy to overcome
the disadvantages of inhibited cholinesterase-based
sensors is to investigate enzymes capable of selective recognition and hydrolysis of organophosphates.
For example, the well characterized metalloenzyme
organophosphorus hydrolase (OPH; EC 3.1.8.1) originally isolated from Pseudomonas diminuta, is able
to cleave P–O, P–F, P–S, and P–CN bonds via an
SN 2-type mechanism, resulting in hydrolysis products
which change solution pH [11–13]. That capability
has been used to develop new biosensors for the direct
detection of organophosphates based on pH monitoring of OPH activity with conventional pH electrodes,
pH-sensitive field effect transistors (pH-FETs), or
pH-sensitive fluorescent dyes [14–17]. This strategy
can be extended to further discriminate between different classes of OP neurotoxins by monitoring additional
organophosphate hydrolysis products. For example,
hydrolysis of phosphofluoridates yields changes in pF
as well as pH which can be detected with a fluoride
specific ion-selective electrode [18]. Detection selectivity has also been enhanced by genetically modifying OPH enzyme substrate specificity [19]. Finally,
OPH has also been combined with the inhibited AChE
strategy to produce a multi-enzyme biosensor with
the unique ability to monitor pesticide mixtures and
discriminate between different pesticide classes [6].
In the present study, another well characterized hydrolytic enzyme-organophosphorus acid anhydrolase,
OPAA (EC 3.1.8.2), has been investigated for direct
detection of organophosphate compounds. OPAA was
originally isolated from a halophilic bacterial isolate
designated JD6.5 and demonstrated high levels of
DFP hydrolysis [20]. The bacterial isolate was identified as a species of the genus Altermonas and DFP
hydrolyzing activity has been observed in several
Altermonas species [21]. Subsequently, the gene responsible for the DFP-hydrolyzing capability of the
JD6.5 strain was isolated, cloned into Escherichia coli
and sequenced [22]. The effects of buffer choice, pH,
temperature, storage conditions and reconstitution on
enzyme activity have also been reported [23]. As with
OPH, the native function of OPAA is not yet known.
However, OPAA has been unambiguously identified
as a prolidase (EC 3.4.13.9), a type of dipeptidase
hydrolyzing dipeptides with a prolyl residue in the
carboxyl-terminal position [24]. Significant effort has
focused upon development of an OPAA overproducing recombinant cell-line and use of OPAA for enzyme mediated nerve agent decontamination [24,25].
Fig. 1 compares the chemical structure of five
organophosphate compounds; all are toxic cholinesterase inhibitors and all are hydrolyzed by OPH [12].
Diisopropyl fluorophosphate (DFP) is a phosphofluoridate containing a P–F bond. The structural similarity
between DFP and the two CW nerve agents sarin and
soman makes it an appropriate analogue for the development of determination and destruction technologies
for these compounds. DFP is frequently used as a
surrogate for the fluorine containing chemical warfare
agents due to its reduced (but far from negligible)
human neurotoxicity. Paraoxon and demeton-S are
commercially available organophosphate pesticides.
Paraoxon is representative of pesticides with a P–O
bond while demeton-S is representative of pesticides
with a P–S bond.
One of the main parameters which characterizes a
biosensor is analyte specificity. Enzymes with broad
substrate specificity are suitable for detection of multiple analytes belonging to the same chemical class,
while enzymes with high specificity for a single
substrate are best for discriminative detection of a
single target analyte. Initial investigations of OPAA
substrate selectivity [20,21] indicated that the enzyme
is capable of cleaving P–F bonds, while P–O or P–S
bond hydrolysis is minimal. Since organophosphate
pesticides, such as paraoxon and demeton-S, typically
possess P–O and P–S bond structures (Fig. 1), such
unique substrate preference makes OPAA very attractive for discriminative detection of fluorine containing
organophosphates.
In this paper, use of OPAA as the recognition
component of a highly specific organophosphate
biosensor was investigated. Results demonstrated that
OPAA immobilized on silica gel enabled detection of
DFP down to 25 ␮mol in a batch mode measurement
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
17
Fig. 1. Chemical structures of some OP neurotoxins.
system with a standard pH electrode and OPAA immobilized to the exposed gate insulator of a pH-FET
enabled detection of DFP down to 20 ␮mol. Of
greatest interest was the lack of signal when either sensor system was challenged with paraoxon or
demeton-S supporting the strategy of OPAA recognition for exclusive detection of fluorine containing
organophosphates.
2. Experimental
2.1. Reagents and buffers
Paraoxon (diethyl-p-nitrophenyl phosphate), DFP,
3-amino-propyl triethoxysilane glutaraldehyde (grade
1), and all buffer reagents (NH4 Cl, NaCl, MnCl2 ) were
obtained from Sigma Chemical Company (St. Louis,
MO, USA); demeton-S (O,O-dimethyl-S-2-ethylthiolethyl phosphorothioate) was from Chem Service (West
Chester, PA, USA). All solutions were prepared using
18 MW cm ultrapure water (Milli-Q Plus, Millipore,
Bedford, MA, USA).
2.2. Enzyme and immobilization
2.2.1. Enzyme
OPAA (EC 3.1.8.2) from Altermonas sp. strain
JD6.5 was isolated and purified to homogeneity [23]
at the US Army Edgewood Chemical Biological Center and obtained as a lyophilized enzyme powder
(0.31 mg protein/mg powder). Since OPAA activity
is enhanced with ammonia-based buffers with low
concentrations of MnCl2 [23], OPAA solutions were
prepared in 50 mM NH4 Cl, pH 8.5, with 0.1 mM
MnCl2 , 100 mM NaCl (standard buffer).
2.2.2. Immobilization on silica gel
OPAA was immobilized on silica gel particles [26]
by modification of the method reported previously
[18]. All chemical treatments were followed by thorough rinsing with deionized water. Gel particles were
cleaned by washing with 1 N HCl, then 20% H2 O2 at
room temperature. Particles were silanized in a 10%
aqueous solution of 3-aminopropyl triethoxysilane for
6 h at 85◦ C followed by treatment with a 10% aqueous solution of glutaraldehyde for 2 h at room temperature. Finally, 1 g of activated particles was reacted
with 1 ml of OPAA (3.12 mg/ml) in standard buffer at
4◦ C for 12 h with gentle stirring. The resulting biocatalyst was stored in standard buffer at 4◦ C.
2.2.3. Immobilization on chip
OPAA was immobilized to sensor chips using the
same procedure reported previously [17] and almost
identical to the silica immobilization described above.
Enzyme (3.12 mg/ml in standard buffer) was applied to
only one device gate of a dual pH-FET chip (described
below) by pipetting into only one epoxy defined well.
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A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
The immobilization was carried out at room temperature for 2 h. After extensive rinsing, chips were stored
in standard buffer at 4◦ C.
2.3. Apparatus and procedures
A batch mode measurement chamber (5 ml)
developed at the Yerevan Physics Institute (Yerevan,
Armenia) [18] was fitted with a pH (FTP-2, Lazar
Research Laboratories, Los Angeles, CA, USA)
and pF (Model 96-09, Orion, Beverly, MA, USA)
ion-selective electrode and used to obtain kinetic characteristics of soluble OPAA. Enzyme and substrate
were added to the chamber then pH or pF signals
were collected by a digital pH-meter (Ion Analyzer
350, Corning, NY, USA), registered by computer
(ProComm, Datastorm Technologies, Colombia, MO,
USA) and analyzed (KaleidaGraph, Synergy Software, Reading, PA, USA).
Sensor chips were prototype pH-FETs (SenDx
Medical, Inc., Carlsbad, CA, USA). Each chip contained two discrete depletion mode, n-channel transistors with a non-metallized gate insulator stack of
thermal silicon dioxide and chemical vapor deposited
(CVD) silicon nitride. A porous ceramic matrix was
formed at the gate surface by dip coating the chip
prior to packaging into a solution of 200 nm silica
microspheres formed via a sol–gel process. This procedure has been described in detail previously [17].
Chips were diced, packaged and installed in a differential control circuit [17]; a simplified schematic of
the chip control circuit is shown in Fig. 2. The control circuit was configured as a classical differential
pair amplifier and used to monitor the gate to source
voltage of the enzyme-modified device relative to the
non-enzyme-modified device. The main advantage of
the differential pair is that common mode variations
such as temperature and bulk pH changes are eliminated while local pH changes at the FET gate with
immobilized enzyme are amplified.
2.3.1. Batch-mode assay with conventional
pH electrode
The temperature-controlled (25◦ C) measurement
cell was filled with modified standard buffer (1 mM
NH4 Cl), and the pH electrode-stopper assembly was
plugged carefully to avoid air bubble formation. For
analysis of soluble enzyme kinetics, substrate was
injected into the measurement cell and enzymatic
reactions were initiated by 10 ␮l of OPAA (0.098 mg/
ml solution). In the immobilized enzyme experiments,
500 mg of carrier with immobilized OPAA was injected into the measurement cell and the reaction was
initiated by injection of different concentrations of
substrate. The velocity of pH or pF changes (in mV/s)
was calculated for each concentration of substrate
and the results were used for determination of kinetic
parameters using the Michaelis–Menten equation
V =
Vmax S
KM + S
where V is the reaction rate, S the substrate concentration, Vmax the maximum reaction rate, and KM the
apparent Michaelis constant. To remove adsorbed fluoride, the electrode, stir bar, and chamber were rinsed
between calibration readings and between reactions in
standard buffer with 20% methanol (v/v) followed by
a thorough rinsing with methanol-free standard buffer.
2.3.2. Stop-flow assay with pH-FET
The flow system consisted of a peristaltic pump
and a flow cell mounted over packaged chips which
provided a head space of approximately 75 ␮l
(Fig. 2). The pH response of the device was tested by
Fig. 2. Schematic of the constant current differential sensor circuit.
See details in Section 2.3.
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
monitoring changes in the common source voltage
(Vcs , Fig. 2) with pH 4, 7, and 10 standard buffers
before and after enzyme immobilization. Enzyme
specific responses were measured as changes in differential signal (Vdiff , Fig. 2). A silver/silver chloride
flow-through reference electrode (Microelectrodes
16-702, New London, CT, USA) was incorporated in
the inlet stream manifold. Measurements were taken
with flow stopped to avoid streaming potential interference. Each experimental point was the average of
3–5 measurements.
19
3.2. Batch-mode assay with conventional
pH electrode
Fig. 3 compares the change in pH generated by
OPAA immobilized to silica gel in response to varying
concentrations of each organophosphate compound
tested. Response to DFP was very good; DFP concentrations as low as 20 mM were detectable and a linear
response was observed from 20 to 500 mM. The response to paraoxon was extremely low, and no activity
was detected for demeton-S (Fig. 3, curves 3 and 4).
3.3. Stop-flow assay with pH-FET chip
3. Results and discussion
3.1. Soluble OPAA kinetic characteristics
Table 1 presents measured kinetic characteristics of
OPAA, as well as previously reported characteristics
of OPH [19]. The comparison of OPAA and OPH
kinetic parameters emphasizes the unique recognition
capability of OPAA relative to OPH. OPAA displays
a strong preference for P–F bond hydrolysis making
it suitable for discriminative detection of fluorine
containing organophosphates, while OPH displays
significant activity for both P–O and P–F bond hydrolysis making it suitable for detection of multiple
organophosphates.
Apparent kinetic parameters for soluble OPAA
with DFP as a substrate, obtained by pH elecapp
app
trode, are Vmax = 2.70 ± 0.28 mV/s and Km =
0.54 ± 0.08 mM. The same parameters in the presapp
ence of 1 mM paraoxon are Vmax = 2.54 ± 0.30 mV/s
app
and Km = 0.55 ± 0.23 mM and in the presence of
app
1 mM demeton-S are 2.63 ± 0.30 mV/s and Km =
0.57 ± 0.23 mM.
A typical FET-chip sensor response to different concentration of DFP is presented in Fig. 4. Detection
time required for 90% of the total signal change was
between 100 and 250 s, depends on concentration of
DFP. Fig. 5 presents response of the OPAA-modified
pH-sensitive chips as a function of substrate concentration for different substrates. As expected, based on
OPAA kinetic characteristics, differences between responses to DFP and other substrates are quite large.
Paraoxon generated barely measurable responses, even
at high concentrations; there was no detectable signal for demeton-S. Good linearity was observed for
DFP concentrations from 12.5 to 500 mM (Fig. 5b).
In addition to excellent discrimination, the presence
of 0.5 mM paraoxon plus 0.5 mM demeton-S had little
effect on sensor response to DFP (Fig. 5).
3.4. pH dependence
The previously reported pH optimum for OPAA acapp
tivity (kcat ) with DFP as a substrate was 8.5, while
Table 1
Comparative kinetic constants for OPH and OPAA enzymes
Substrate/hydrolyzed bond
Enzyme
DFP [P–F]a
OPAA
OPH
Paraoxon [P–O]b
OPAA
OPH
Demeton-S [P–S]b
OPAA
OPH
a
b
Detected by fluoride-specific electrode.
Detected by pH-sensitive electrode.
kcat (s−1 )
KM (mM)
kcat /KM (M−1 s−1 )
770∗
3500
2.81
1.42
2.7 × 105
2.4 × 106
4.64
11500
0.6
0.12
7.75 × 103
9.6 × 107
3.5
4.0
8.0
1.2 × 103
0.028
4.8
20
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
Fig. 3. Calibration curves for batch-mode biosensor with a flat pH electrode: (1) fresh immobilized OPAA with DFP as a substrate; (2)
the same, after 5 months; (3) fresh immobilized OPAA with paraoxon as a substrate; (4) with demeton-S.
Fig. 4. Biosensor response for 1, 0.5 and 0.25 mM of DFP. Stop-flow mode with pH-sensitive field effect transistor. Measurement condition:
1 mM NH4 Cl buffer, containing 0.1 mM MnCl2 and 100 mM NaCl.
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
21
Fig. 5. Signal versus concentration for DFP (black squares) paraoxon (inverted triangles), and mixture of 0.5 mM of paraoxon, 0.5 mM
of demeton-S and DFP (triangles). (a) Stop-flow assay with pH-sensitive field effect transistor (full curves). (b) The linear ranges of the
calibration curves; f (x)DFP = 43.5 + 1469x, R = 0.99976; f (x)mix = 42.5 + 1255x, R = 0.98891; f (x)PX = 13.9 + 77.1x, R = 0.85367.
22
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
Fig. 6. pH profile for DFP biosensor response. Stop-flow mode with pH-sensitive field effect transistor. Measurement condition: 1 mM
NH4 Cl buffer, containing 0.1 mM MnCl2 and 100 mM NaCl. DFP concentration 1 mM.
app
app
the highest catalytic efficiency (kcat /Km ) was observed at pH 6.8 [20]. However, enzyme immobilization can influence enzyme operational capability and
change the pH optimum, depending on which sites on
the enzyme were involved in binding to a surface. To
determine the pH optimum for the OPAA-modified
pH-FET sensor, response to a fixed concentration of
DFP (1 mM) at pH 6.2–9.0 was measured (Fig. 6).
Maximum sensor response was obtained at pH 8.5,
however, the sensor response was not as sensitive to
pH as the soluble enzyme [20] and satisfactory signal
was obtained from pH 7.5 to 9.0.
3.5. Long-term stability
The enzyme stability was significantly different for
the two sensor formats. OPAA immobilized on silica
gel retained about 60% of the starting activity after
5 months (Fig. 3, curve 2). However, OPAA on the
pH-sensitive chip lost 60% of its activity in 22 days,
and there was no response after 30 days. This was
especially surprising since previous results demonstrated that the silica microspheres modification of
the pH-sensitive chips enhanced enzyme stability and
enabled stable sensor response for more than 2 months
[17]. Such dramatic differences in long-term stability
may be due to the significant differences in enzyme
capacity of the two sensor formats; the silica gel has a
much greater surface area than the pH-sensitive chip
surface.
4. Conclusions
The results obtained in this study demonstrate the
ability of the enzyme OPAA to serve as a highly
discriminative biorecognition element for organophosphate biosensors. OPAA can uniquely hydrolyze
fluorine containing organophosphates, such as DFP
(k cat = 1650 s−1 ) or the chemical warfare agents
sarin (k cat = 611 s−1 ) and soman (k cat = 3145 s−1 )
[24], without responding to non-fluorine containing
organophosphate pesticides (this study). OPAA was
tested in a batch mode system by immobilizing to
silica gel and monitoring changes in solution pH
as a function of DFP concentration with a standard
pH electrode. In addition, OPAA was covalently immobilized to the silica microspheres modified gate
A.L. Simonian et al. / Analytica Chimica Acta 442 (2001) 15–23
insulator of a pH-FET and changes in pH relative to
a second non-enzyme coated pH-FET were measured
in stop-flow mode. Concentrations of DFP down to
25 ␮M with the standard pH electrode and 20 ␮M
with the pH-FET were readily detected. Little or no
sensor signal was generated in response to paraoxon
or demeton-S. Such excellent discrimination between
compounds with such dramatic differences in application yet within the same chemical class is highly
desirable. OPAA should also prove valuable in combination with other organophosphate hydrolyzing
enzymes, such as OPH, so as to enable construction
of an array of biosensors able to discriminate between
several classes of neurotoxins.
Acknowledgements
The authors gratefully acknowledge the support of
the US Department of Energy, Office of Nonproliferation Research and Engineering, NN-20, and the
US Army Medical Research and Materiel Command
(JRW). We thank Joanne Volponi of Sandia National
Laboratories for chip preparation and pH testing. Sandia is a multiprogram laboratory operated by Sandia
Corporation, a Lockheed Martin Company, for the
United States Department of Energy under contract
DE-AC04-94AL85000.
References
[1] G.A. Evtugyn, H.C. Budnikov, E.B. Nikolskaya, Analyst 121
(1996) 1911.
[2] N. Mionetto, J.-L. Marty, I. Karube, Biosens. Bioelectron. 9
(1994) 463.
[3] G. Palleschi, M. Bernabei, C. Cremisini, M. Mascini, Sens.
Actuators B 7 (1992) 513.
[4] P.SkladalM. Mascini, Biosens. Bioelectron. 7 (1992) 335.
[5] J.J. Kulys, E.J. D’Costa, Biosens. Bioelectron. 6 (1991) 109.
23
[6] A.L. Simonian, E.I. Rainina, J. R Wild, Anal. Lett. 30 (14)
(1997) 2453.
[7] A. Herbert, L. Guilhermino, H.C. Da Silva De Assis, P.-D.
Hasen, Z. Angew. Zool. 3 (1995–1996) 1–15.
[8] M. Brufani, M. Marta, M. Pomponi, Eur. J. Biochem. 157
(1986) 115–120.
[9] A.M. Nyamsi Hendji, N. Jaffrezic-Renault, C. Martelet,
P. Clechet, A.A. Shul’ga, V.I. Srtkha, L.I. Netchiporuk,
A.P. Soldatkin, W.B. Wolodarski, Anal. Chim. Acta 281
(1993) 3.
[10] T.T. Bachmann, R.D. Schmid, Anal. Chim. Acta 401 (1999)
95.
[11] D.P. Dumas, J.R. Wild, F.M. Raushel, Biotech. Appl.
Biochem. 11 (1989) 235.
[12] D.P. Dumas, H.D. Durst, W.G. Landis, F.M. Raushel, J.R.
Wild, Arch. Biochem. Biophys. 277 (1990) 155.
[13] K. Lai, N.J. Stolowich, J.R. Wild, Arch. Biochem. Biophys.
318 (1995) 59.
[14] E. Rainina, A. Simonian, E. Efremenko, S. Varfolomeyev, J.
Wild, Biosens. Bioelectron. 11 (1996) 991–1000.
[15] A. Mulchandani, I. Kaneva, W. Chen, Anal. Chim. Acta 70
(1998) 4140–4145.
[16] R. Russell, M. Pishko, A. Simonian, J. Wild, Anal. Chem.
71 (1999) 4909–4912.
[17] A.W. Flounders, A.K. Singh, J.V. Volponi, S.C. Carichner, K.
Wally, A.L. Simonian, J.R. Wild, J.S. Schoeniger, Biosens.
Bioelectron. 14 (1999) 715–722.
[18] A.L. Simonian, B.D. diSioudi, J.R. Wild, Anal. Chim. Acta
389 (1999) 189–196.
[19] B.D. diSioudi, C.E. Miller, K. Lai, J.K. Grimsley, J.R. Wild,
Chem.-Biol. Interact. 119/120 (1999) 211–223.
[20] J.J. DeFrank, T.-C. Cheng, J. Bacteriol. 173 (1991) 1938–
1943.
[21] J.J. DeFrank, W.T. Beaudry, T.-C. Cheng, S.P. Harvey, A.N.
Stroup, L.L. Szafraniec, Chem.-Biol. Interact. 87 (1993) 141–
148.
[22] T.-C. Cheng, S.P. Harvey, G.L. Chen, Appl. Environ.
Microbiol. 62 (1996) 1636–1641.
[23] T.-C. Cheng, J.J. Calomiris, Enzyme Microbial Technol. 18
(1996) 597–601.
[24] T.-C. Cheng, J.J. DeFrank, V.K. Rastogi, Chem.-Biol. Interact.
119/120 (1999) 455–462.
[25] T.-C. Cheng, V.K. Rastogi, J.J. DeFrank, G.P. Sawiris, Ann.
New York Acad. Sci. 864 (1998) 253–258.
[26] A.L. Simonian, G.E. Khachatrian, S.Sh. Tatikian, Ts.M.
Avakian, I.E. Badalian, Biosens. Bioelectron. 6 (1991) 93–99.