A Bacterial Biosensor for Oxidative Stress Using the Constitutively Expressed Redox-Sensitive Protein roGFP2

Abstract

A highly specific, high throughput-amenable bacterial biosensor for chemically induced cellular oxidation was developed using constitutively expressed redox-sensitive green fluorescent protein roGFP2 in E. coli (E. coli-roGFP2). Disulfide formation between two key cysteine residues of roGFP2 was assessed using a double-wavelength ratiometric approach. This study demonstrates that only a few minutes were required to detect oxidation using E. coli-roGFP2, in contrast to conventional bacterial oxidative stress sensors. Cellular oxidation induced by hydrogen peroxide, menadione, sodium selenite, zinc pyrithione, triphenyltin and naphthalene became detectable after 10 seconds and reached the maxima between 80 to 210 seconds, contrary to Cd(2+), Cu(2+), Pb(2+), Zn(2+) and sodium arsenite, which induced the oxidation maximum immediately. The lowest observable effect concentrations (in ppm) were determined as 1.0 × 10(-7) (arsenite), 1.0 × 10(-4) (naphthalene), 1.0 × 10(-4) (Cu(2+)), 3.8 × 10(-4) (H(2)O(2)), 1.0 × 10(-3) (Cd(2+)), 1.0 × 10(-3) (Zn(2+)), 1.0 × 10(-2) (menadione), 1.0 (triphenyltin), 1.56 (zinc pyrithione), 3.1 (selenite) and 6.3 (Pb(2+)), respectively. Heavy metal-induced oxidation showed unclear response patterns, whereas concentration-dependent sigmoid curves were observed for other compounds. In vivo GSH content and in vitro roGFP2 oxidation assays together with E. coli-roGFP2 results suggest that roGFP2 is sensitive to redox potential change and thiol modification induced by environmental stressors. Based on redox-sensitive technology, E. coli-roGFP2 provides a fast comprehensive detection system for toxicants that induce cellular oxidation.

Full text

Sensors 2010, 10, 6290-6306; doi:10.3390/s100706290
sensors
ISSN 1424-8220
www.mdpi.com/journal/sensors
Article
A Bacterial Biosensor for Oxidative Stress Using the
Constitutively Expressed Redox-Sensitive Protein roGFP2
Carlos R. Arias-Barreiro
1
, Keisuke Okazaki
1
, Apostolos Koutsaftis
1,2
,
Salmaan H. Inayat-Hussain
3
, Akio Tani
1
, Maki Katsuhara
1
, Kazuhide Kimbara
1,4
and Izumi C. Mori
1,
*
1
Institute of Plant Science and Resources, Okayama University, Kurashiki 710-0046, Japan;
E-Mails: cr_arias@rib.okayama-u.ac.jp (C.R.A.-B.); okazakikeis@gmail.com (K.O.);
apostolosk@gmail.com (A.K.); atani@rib.okayama-u.ac.jp (A.T.);
kmaki@rib.okayama-u.ac.jp (M.K.); tkkimba@ipc.shizuoka.ac.jp (K.K.)
2
Environmental Risk Management Authority, PO BOX 131, Wellington 6140, New Zealand
3
Faculty of Allied Health Sciences/UKM Medical Molecular Biology Institute, Universiti
Kebangsaan Malaysia, Jalan Raja Muda Abdul Aziz, Kuala Lumpur 50300, Malaysia;
E-Mail: salmaan@streamyx.com (S.H.I.-H.)
4
Department of Materials Science and Chemical Engineering, Shizuoka University, Hamamatsu
432-8561, Japan
* Author to whom correspondence should be addressed; E-Mail: imori@rib.okayama-u.ac.jp;
Tel.: +81-86-434-1215; Fax: +81-86-434-1249.
Received: 17 May 2010; in revised form: 8 June 2010 / Accepted: 21 June 2010 /
Published: 24 June 2010
Abstract: A highly specific, high throughput-amenable bacterial biosensor for chemically
induced cellular oxidation was developed using constitutively expressed redox-sensitive
green fluorescent protein roGFP2 in E. coli (E. coli-roGFP2). Disulfide formation between
two key cysteine residues of roGFP2 was assessed using a double-wavelength ratiometric
approach. This study demonstrates that only a few minutes were required to detect
oxidation using E. coli-roGFP2, in contrast to conventional bacterial oxidative stress
sensors. Cellular oxidation induced by hydrogen peroxide, menadione, sodium selenite,
zinc pyrithione, triphenyltin and naphthalene became detectable after 10 seconds and
reached the maxima between 80 to 210 seconds, contrary to Cd
2+
, Cu
2+
, Pb
2+
, Zn
2+
and
sodium arsenite, which induced the oxidation maximum immediately. The lowest
observable effect concentrations (in ppm) were determined as 1.0 × 10
−7
(arsenite),
1.0 × 10
−4
(naphthalene), 1.0 × 10
−4
(Cu
2+
), 3.8 × 10
−4
(H
2
O
2
), 1.0 × 10
−3
(Cd
2+
),
OPEN ACCESS

Sensors 2010, 10 6291
1.0 × 10
−3
(Zn
2+
), 1.0 × 10
−2
(menadione), 1.0 (triphenyltin), 1.56 (zinc pyrithione),
3.1 (selenite) and 6.3 (Pb
2+
), respectively. Heavy metal-induced oxidation showed unclear
response patterns, whereas concentration-dependent sigmoid curves were observed for
other compounds. In vivo GSH content and in vitro roGFP2 oxidation assays together with
E. coli-roGFP2 results suggest that roGFP2 is sensitive to redox potential change and thiol
modification induced by environmental stressors. Based on redox-sensitive technology,
E. coli-roGFP2 provides a fast comprehensive detection system for toxicants that induce
cellular oxidation.
Keywords: oxidative biosensor; redox-sensitive GFP; ratiometric measurement; ROS;
environmental stressors
1. Introduction
Physicochemical analysis provides accurate information on the composition of complex
environmental samples. Nevertheless, it is widely accepted that this approach by itself fails to provide
thorough information on the deleterious effects on living organisms because of its inability to correctly
evaluate bioavailability [1]. Biosensor- or bioassay-based toxicity tests intend to complement
physicochemical analysis by using biological responses/endpoints to evaluate toxicity related effects.
On this note, biosensor technologies for environmental monitoring have been under constant
evolution [2].
A wide variety of environmental stressors can alter the intracellular redox status by means of
oxidative damage ultimately leading to cell death [3]. The homeostatic imbalance of the redox
potential is generated as a consequence of the cell’s inability to cope with the cumulative generation of
the reactive oxygen species (ROS): hydrogen peroxide (H
2
O
2
), hydroxyl radical (OH•) and superoxide
anion (O
2
•
–
). Among environmental toxicants, metal ions and metalloid oxyanions have been
documented to exert their toxic effect through alteration of the cellular redox status [4-7].
On this note, bacteria have been preferably used for the rapid detection of ROS due to their fast
response, high growth rate, and low cost [8-12]. Whole-cell bacterial biosensing systems rely
extensively on the use of promoter-reporter expression systems, which comprise a transcription
regulator plus promoter/operator with an open reading frame for proteins of measurable activity.
Exposure to xenobiotics activates the promoter through stress signal transduction events resulting in
protein expression. Detectable expression times for these systems range from tens of minutes to hours.
In order to develop more sophisticated oxidative stress biosensors, high levels of specificity,
sensitivity, speed, as well as high-throughput amenability have become desirable features. In this
respect, since the design by Hanson et al. [13], several studies have profited from redox-sensitive
green fluorescent proteins (roGFPs), for the live redox monitoring of intracellular oxidation state, c.f.,
yeast [14,15], plants [16,17] and mammalian cells [18,19]. These GFP mutants allow ratiometrical
quantification of the redox potential based on the formation of disulfide bonds between
surface-exposed cysteine residues. Fluorescence patterns of excitation, which represent oxidized and
reduced forms of the protein denote the extent of oxidation. In terms of analytical advantage, errors

Sensors 2010, 10 6292
resulting from variations of the indicator concentration, photobleaching and variable cell thickness are
reduced or eliminated [18]. Furthermore, when compared to the promoter-reporter approach, the
measurement benefits from a reduction of the analysis time because it is based on the conformational
changing velocity of a constitutively expressed protein, as opposed to the time required for
its expression.
Despite its beneficial characteristics, an integrating approach for live redox monitoring of
chemically induced oxidative stress in bacterial cells has not yet been undertaken. In the present study,
we developed a ratiometric fluorescence assay using roGFP2 (GFP mutant C48S/S147C/Q204C/
S65T/Q80R) [13] expressed in Escherichia coli, to monitor chemically induced oxidative stress. The
redox-sensitive properties of the roGFPs provided the basis for the engineering of a bioassay for
cellular oxidation. Furthermore, the experiments are shown to be high-throughput amenable, to acquire
data on a scale of seconds to minutes.
2. Results
2.1. Autofluorescence normalization and background oxidation
Fluorescence was detectable in roGFP2-transformed E. coli cells and non-transformants. Compared
fluorescence spectra of roGFP2-expressing cells and the vector control (autofluorescence) are shown
in Appendix Figure 1A Subtraction of vector-control fluorescence generated the distinctive roGFP2
spectrum (Appendix Figure 1B) in accordance with previous data of purified roGFP2 protein [13].
Shortly after preparation of the cell suspension, increasing spontaneous cellular oxidation was
observed in the absence of toxicants (Appendix Figure 2, 0 h). However, standing for 1 hour at room
temperature reduced the spontaneous oxidation rate (Appendix Figure 2, 1 h). Further stagnation
resulted in a slight reduction of the rate (Appendix Figure 2, 2 h). The spontaneous oxidation was
sometimes not clearly observed in multi-well plate reader experiments as seen in Figure 3. This might
be due to difference in shaking procedures: continuous stirring for spectroscopy (Appendix Figure 2
and Figure 1E), and intermittent shaking for multi-well plate analysis (Figure 3).
2.2. Response of the biosensor to H
2
O
2
Hydrogen peroxide is known to evoke oxidation in prokaryotic and eukaryotic cells. roGFP2 has
been found to successfully indicate cell oxidation by H
2
O
2
in mammalian [13] and plant cells [16].
Here, E. coli cells constitutively expressing roGFP2 (E. coli-roGFP2) were challenged with H
2
O
2
to
examine spectral and ratiometric fluorescence brought about by cellular redox changes.
Figure 1 shows the spectral response of the biosensor to H
2
O
2
. Compared to the control (Figure 1A),
0.1 and 1 mM of H
2
O
2
induced spectral pattern changes within 6 minutes (Figure 1B and C).
Fluorescence spectra displayed two distinctive excitation maxima at 400 (F
ex400
) and 490 nm (F
ex490
)
related to its oxidized and reduced form, respectively. A decrease of F
ex490
induced by H
2
O
2
became
apparent at 30 seconds and progressed further in time (>2 minutes) (Figure 1B and C). Also, an
increase of F
ex400
was observed, although the magnitude of the change was smaller than that of F
ex490
.
Subtraction of the basal spectrum at t = –2 minutes (before H
2
O
2
addition) from the other spectra

Sensors 2010, 10 6293
showed that the F
ex400
increase and F
ex490
decrease (Figure 1D), providing evidence that fluorescence
change unequivocally indicated oxidation of the biosensor.
Figure 1E shows increasing F
ex400
/F
ex490
ratio at high H
2
O
2
concentrations. An oxidation maximum
at 100 µM H
2
O
2
was reached in 4 minutes (Figure 1E), while approximately a period of 15 minutes
was necessary for mammalian cells to reach the same state [18]. These results demonstrate the
capability of roGFP2 to monitor oxidation in E. coli cells, as previously reported for mammalian [18]
and plant cells [16].
Figure 1. Spectroscopic analysis of E. coli-roGFP2 after H
2
O
2
-induced cellular oxidation.
Fluorescent excitation spectra of a typical kinetic experiment starting at −2 minutes (before
the addition of H
2
O
2
), 0 minute (addition), up to 6 minutes after addition of the chemical
with water (A), 0.1 mM H
2
O
2
(B) and 1 mM H
2
O
2
(C). Clarified spectral behavior after
treatment with 1 mM H
2
O
2
and subsequent subtraction of basal fluorescence at −2 minutes
(D). Kinetic analysis of the 400/490 nm ratiometric response (E) (open squares: 1 mM,
closed triangles: 0.1 mM, open triangles 10 mM, closed circles: 1 mM, and open circles:
water control). Error bars show ±S.E.M.
2.3. Spectral changes of the biosensor upon exposure to toxic metal compounds
Monitoring of cell oxidation was carried out after exposure of E. coli-roGFP2 to the
environmentally relevant heavy metal (HM) cations Cd
2+
, Cu
2+
, Pb
2+
and
+
Zn
2+
; and the oxyanions
AsO
2
–
and SeO
3
2–
.
Figure 2 shows the effect of HM cations on the fluorescence of the biosensor. Fluorescence changes,
indicating oxidation of the biosensor, were observed at 4 minutes by addition 0.5 ppm Cu
2+
, 0.5 ppm
Zn
2+
, 1 ppm Cd
2+
and 2.5 ppm Pb
2+
(Figure 2B-E). Ten ppm SeO
3
2–
and 0.25 ppm AsO
2
–
showed a
comparable response to HM cations upon exposure (Figure 2G and H). Increase of F
ex400
and decrease
of F
ex490
were clearly observed, when subtracted from the t = 0 minute data, demonstrating oxidation
of roGFP2 (Figures 2F and I). The data provide evidence on the capabilities of the biosensor to detect
the HM and metalloid-evoked cell oxidation.

Sensors 2010, 10 6294
Figure 2. Spectroscopic analysis of E. coli-roGFP2 after induction of cellular oxidation by
heavy metal compounds and oxyanions. Fluorescent excitation spectra of kinetic
experiments after exposure of 4 minutes to water (A), 1 ppm CdCl
2
(B), 0.5 ppm CuCl
2
(C), 2.5 ppm PbCl
2
(D) and 0.5 ppm ZnCl
2
(E). Overall comparison among control and
cations at 4 minutes after subtraction from 0 minute (E). Kinetic data obtained
after 4 minutes exposure to 0.25 ppm NaAsO
2
(F) and 10 ppm Na
2
SeO
3
(G). Overall
comparison among oxyanions at 4 minutes after subtraction from 0 minute (I).
2.4. Ratiometric characterization of the biosensor
Kinetic analyses were performed with a multiwell plate reader to further characterize oxidation
patterns of the E. coli-roGFP2 biosensor in a high throughput fashion. F
ex400
and F
ex490
were measured
every 30 seconds during 7 minutes (Figure 3). The first data point was obtained at 10 seconds after the
addition of chemicals due to the measurement lag time.
Figure 3. Multi-well ratiometric kinetic analysis of cellular oxidation using E. coli-roGFP2.
Exposure to several concentrations of H
2
O
2
(A), menadione (B), CdCl
2
(C), CuCl
2
(D),
Pb(CH
3
COO)
2
(E), ZnCl
2
(F), Na
2
SeO
3
(G), NaAsO
2
(H), ZnPT (I), TPT (J) and
naphthalene (K). Each data point represents the average of six replicates. Measurements
were made every 30 seconds during 7 minutes. Results shown correspond to a typical
experiment, where each data point represents the quotient of the excitation fluorescence
relative value at 400 and 490 nm.

Sensors 2010, 10 6295
Figure 3. Cont.
2.4.1. Detection of oxidation upon exposure to H
2
O
2
and menadione
A gradual increase of the F
ex400
/F
ex490
ratio was observed with 1 and 10 mM H
2
O
2
reaching the
plateau within 100–200 seconds (Figure 3A). In order to assess concentration dependency of biosensor
oxidation, the average of F
ex400
/F
ex490
ratio from 10–430 seconds was plotted against concentrations of
added chemicals (Figure 4). The concentration-dependency curve showed a gradual increase
proportionally to H
2
O
2
concentration, and the lowest observed effective concentration (LOEC) was
determined as low as 10 nM (p < 0.01) (Figure 4A).
Figure 3B shows the maximum oxidation was reached at approximately 200 seconds after exposure
to menadione (0.1–100 ppm). The concentration-response correlation demonstrated the LOEC to
be 0.01 ppm at p < 0.01 (Figure 4B). Conclusively, sigmoid response curves were observed for both
H
2
O
2
and menadione treatments (Figures 4A and B) further establishing oxidation causality under
increasing concentrations of the chemicals.

Sensors 2010, 10 6296
Figure 4. Concentration-response curves as shown by fluorescent ratiometric changes.
Exposure to several concentrations of H
2
O
2
(A), menadione (B), CdCl
2
(C), CuCl
2
(D), Pb(CH
3
COO)
2
(E), ZnCl
2
(F), Na
2
SeO
3
(G), NaAsO
2
(H), ZnPT (I), TPT (J) and
naphthalene (K). Each data point represents the ratiometric average of the dataset
during 7 minutes (10–430 seconds) for a given concentration. Significant differences
between control (concentration=0) and treated groups are indicated by * (p < 0.05) and
** (p < 0.01) using Dunnett’s test followed by ANOVA. Error bars show ±S.E.M. Error
bars in panels A, B, I and J are too small to be seen.
2.4.2. Effects of HM cations on E. coli-roGFP2
Treatments with Cd
2+
, Cu
2+
, Pb
2+
and Zn
2+
showed an irregular kinetic behaviour together with
positive ratiometric responses (Figure 3C-F), unlike H
2
O
2
and menadione, which increased gradually
over time. Moreover, ratiometric changes were smaller compared to H
2
O
2
and menadione. An
exception was observed, however, at 100 ppm of Pb(CH
3
COO)
2
where an almost linear increase was
triggered. Although significant differences with respect to the controls were found,
concentration-dependent behaviour did not show typical sigmoid curves in any of the treatments but
rather fluctuating ones, following immediate oxidation after exposure to HMs (Figure 4C-F). This
behaviour may be explained by the coordination to side chains of cysteine residues and cross-link of
thiol groups with HM cations due to their thiophilic nature [20]. The affinity to Cd
2+
, Cu
2+
and Pb
2+
,
and in a lesser degree Zn
2+
, alters the conformation of roGFP2. Although initial exposure to metal

Sensors 2010, 10 6297
cations elicit the production of ROS, it is possible that the formation of roGFP-sulfide complexes can
inhibit the structural changes that account for the F
ex400
/F
ex490
ratiometric response.
To evaluate the involvement of HMs and sulfhydryl-rich groups, we next determined the content of
reduced glutathione (GSH) in E. coli-roGFP2 cells under representative concentrations of HMs
(Figure 5). The exposure induced reduction of GSH contents by 4.0 and 16.8% at 2 minutes, and 10.6
and 14.2% at 7 minutes, for Cd
2+
and Pb
2+
respectively. Interestingly, H
2
O
2
did not induce a significant
reduction of GSH (0.9 and 1.1%). Control (H
2
O) values were kept below 1% for both timepoints.
Menadione showed essentially the same result as the metals, with 16.8 and 24.0% reduction
at 2 and 7 minutes, respectively (Figure 5). Hydrogen peroxide and menadione effects were in
accordance with previous data obtained in E. coli by Smirnova et al. [21].
Figure 5. Effect of Cd
2+
, Pb
2+
, H
2
O
2
and menadione on intracellular levels of GSH.
Changes in concentration of GSH as quantified as described in Materials and methods are
shown at 2 and 7 minutes after exposure to 0.1 ppm CdCl
2
, 1 ppm Pb(CH
3
COO)
2
, 1 ppm
menadione and 1 mM H
2
O
2
. Control corresponds to cells treated with H
2
O. Six replicates
were used for each dataset. Error bars indicate standard deviation.
Additionally, ratiometric changes of isolated roGFP2 in vitro were determined to observe the direct
interaction of roGFP2 protein and chemicals (Figure 6).
Figure 6. Effect of Cd
2+
, Pb
2+
, H
2
O
2
and menadione and on roGFP2 in vitro.
A 1.39 µg/mL solution of roGFP2 was exposed to 0.1 ppm CdCl
2
, 1 ppm Pb(CH
3
COO)
2
,
1 ppm menadione and 1 mM H
2
O
2
. Changes in 400/490 nm fluorescence ratio were
monitored for 5 minutes with a multi-well plate reader and expressed in percentage
compared to the value at t = 0 minute. Three replicates were used for each dataset and error
bars indicate standard deviation.

Sensors 2010, 10 6298
A steady increase was observed for 0.1 ppm Cd
2+
and 1 ppm Pb
2+
ranging from 1.8 to 8.3% and 6.9
to 12.6%, respectively. These increments were smaller than those observed with 1-ppm menadione and
1-mM H
2
O
2
treatments at 9.1 to 29.4% and 10.5 to 42.4%, respectively. Moreover, a net reduction of
the fluorescence signal was not observed while ratiometric response increased (data not shown).
The reduced effects on the direct oxidation of roGFP2 by HMs, when compared to H
2
O
2
and
menadione, may be accounted by the coordination bond formation between HMs and key cysteine
residues (C147 and C204) near the chromophore.
Formation of the disulfide bond occurs between both residues deriving in the characteristic
fluorescence shift when redox potential is altered. Reduction of GSH, coupled with production of ROS
would favour an oxidizing environment while initial sensitivity and affinity of roGFP2 would be
hindered resulting in the irregular kinetic and concentration-dependent patterns. These counteracting
effects might also explain the large variations observed in Figure 4C–F. Despite the lack of regular
response patterns, the HM data can be differentiated from responses of other oxidative chemicals,
based on their particular ratiometric patterns. Notwithstanding the anomalous responses, LOECs could
be determined at 1 × 10
−3
, 1 × 10
−4
, 6.3 and 1 × 10
−3
ppm at p < 0.05 for Cd
2+
, Cu
2+
, Pb
2+
and Zn
2+
,
respectively (Figure 4).
2.4.3. Oxidation detected upon exposure to selenite and arsenite
Selenite and arsenite are environmentally relevant pollutants and known to induce oxidative stress
as a mechanism of their toxicity. Thus, we next examined arsenite and selenite-induced cell oxidation
using the E. coli-roGFP2 biosensor.
Exposure of the biosensor to SeO
3
2–
induced a gradual increase of the F
ex400
/F
ex490
ratio
within 150–200 seconds at 12.5–100 ppm (Figure 3G), contrary to HM that displayed irregular kinetics.
Response did not increase at concentrations higher than 50 ppm SeO
3
2–
(Figure 4G).
In arsenite-induced oxidation, an immediate rise of the F
ex400
/F
ex490
ratio occurred within 10 seconds,
followed by a progressive reduction when 1 ppm AsO
2
–
was added (Figure 3H). Notably, intensity of
F
ex490
was reduced gradually by the same concentration suggesting that the ratiometric reduction is
most likely due to its strong toxic effects. For lower concentrations (1 × 10
−6
–0.1 ppm) maximum
responses were reached within the first 100 seconds.
Both SeO
3
2−
and AsO
2
−
treatments showed a concentration dependent response (Figures 4G and H)
with a great difference in their sensitivity, i.e., the LOECs for SeO
3
2−
and AsO
2
−
were determined as
low as 3.1 ppm (p < 0.05) and 1x10
−7
ppm (p < 0.01), respectively (Figures 4G and H).
2.4.4. Detection of oxidation caused by organometallic compounds
Figures 3I and 3J show the results of oxidation of the biosensor by two organometallic compounds
which are used as biocides, zinc pyrithione (ZnPT) and triphenyltin (TPT), respectively. Exposure of
cells to ZnPT generated gradual increases in the fluorescence ratios (12.5–100 ppm). Similar gradual
time-dependent increases were observed for TPT at 1, 10 and 100 ppm (Figure 3J). Both compounds
reached a maximum level of oxidation between 250 and 310 seconds after the exposure. Furthermore,

Sensors 2010, 10 6299
both compounds significantly induced cell oxidation with similar LOECs of 1.56 ppm at p < 0.01
and 1 ppm at p < 0.01 for ZnPT and TPT exposures, respectively (Figures 4I and 4J).
2.4.5. Detection of oxidation after exposure to naphthalene
To further characterize the biosensor, cells were exposed to naphthalene, a two-ring hydrocarbon.
Figure 3K shows that a maximum oxidation level was reached within 100 seconds of exposure
with 10 and 100 ppm. The LOEC was found to be 1 × 10
−4
ppm at p < 0.05 (Figure 4K). The fact that
the biosensor responded to naphthalene indicates versatility for detecting highly lipophilic chemicals,
such as poly aromatic hydrocarbons (PAHs).
3. Discussion
In this study, chemically induced oxidation of E. coli was successfully assessed using roGFP2.
Fluorescent ratiometric changes, which denote interchange between oxidized and reduced forms of
roGFP2, allow a nearly live measurement of the intracellular redox conditions. Moreover, data showed
that roGFP2 expressed in E. coli was subject to changes brought about by chemically induced
alteration of the intracellular redox status.
Measuring fluorescence of the constitutively expressed protein in cells allowed a very rapid
evaluation, reaching maxima within 2 to 6 minutes for most chemicals tested. This high-speed
characteristic of the biosensor may allow the analysis of a large number of samples in a short time
frame and facilitate time-consuming processes, such as the identification of toxicants.
Genes belonging to regulons SoxRS (induced by O
2
•
−
) [11], OxyRS (induced by H
2
O
2
and
OH•) [8,10] or both [9,12] fused to the luxCDABE operon have provided the fusions of choice for the
development of oxidative stress biosensors. Note that specificity of regulons to ROS is different.
Comprehensive oxidative biosensors should be able to detect the activity of chemicals inducing both
oxidative stress regulons. In this study, detection of cellular oxidation was not based on the induction
of gene expression, but through the increase of the intracellular redox potential. The E. coli-roGFP2
biosensor detected both H
2
O
2
- and menadione-induced oxidation, indicating broad specificity of the
biosensor for ROS. Differences between H
2
O
2
and menadione may be attributed to the ROS produced
by the latter, which includes, O
2
•
–
and H
2
O
2
[21].
Thiophilic activity may also be corroborated by the speed of the oxidative reaction induced by HMs.
Transition divalent cations react immediately with GSH to form bisglutathionate complexes [22]. The
presence of cysteine residues should onset a similar reaction velocity in roGFP2, prompting the
alteration of its fluorescent properties. A disturbance in protein activity due to HMs is also observed in
enzymes containing sulfhydryl groups in their active sites [23]. Although Pb is not a transition metal, it
induces oxidative stress through interaction with GSH [5], which might account for the
observed effects.
Different kinetic patterns of biosensor oxidation were observed (Figure 3), indicating that oxidation
is influenced by the nature of the utilized xenobiotics. The heavy metals Cd
2+
, Cu
2+
, Pb
2+
and Zn
2+
only showed 1.6, 3.4, 3.4 and 1.9% oxidation increases, respectively; which were apparently lower
than others i.e., (in %) H
2
O
2
, 16.5; menadione, 37.4; SeO
3
2–
, 7.6; AsO
2
–
, 5.9; ZnPT, 19.2; TPT, 21.1

Sensors 2010, 10 6300
and naphthalene, 12 (Figure 3). Soft Lewis acids such as Cd
2+
and Pb
2+
, and in a lesser degree
intermediates Cu
2+
and Zn
2+
, display a marked preference for coordination to thiol compounds, such as
cysteine and GSH [20]. After reaction with GSH, the formed bisglutathionate complex can then react
with molecular oxygen rendering the metal cation, H
2
O
2
and oxidized bisglutahione (GSSG). The
latter is in turn reduced again in a NADPH-dependent reaction, and metal cations immediately bind
another two glutathione molecules, producing oxidative stress [24].
Approximately 90% of the reduced thiol concentration in E. coli corresponds to GSH [25]. It is thus
reasonable to consider GSH as the main target of HMs tested. After exposure, an increase in the
GSSG/GSH ratio would alter the redox potential and ultimately cause stress. However, disulfide
formation between C147 and C20 of roGFP2 would also be hindered. The GSSG/GSH ratio,
would
thus fail to show the true extent of change of the redox equilibrium as suggested by the anomalous
kinetics and non-sigmoid concentration dependency patterns observed for Cd
2+
, Pb
2+
, Cu
2+
and Zn
2+
.
Recently, roGFP1-R12 was expressed in Saccharomyces cerevisiae, and oxidative stress was
monitored after exposure to H
2
O
2
, NaAsO
2
and Pb(NO
3
)
2
[14]. The authors were, however, unable to
perform ratiometric analysis for unclear reasons. The lack of a ratiometric feature would be of critical
importance to evaluate the extent of oxidation in roGFP-expressing cells.
E. coli-roGFP2 represented bioavailable fractions of a wide variety of environmental oxidants.
Interestingly, detection limits for inorganic compounds like AsO
2
–
fall within environmentally relevant
concentrations [26]. Among organic chemicals, menadione, a naphthoquinone used as an alternative
biocide substituting for other nonselective oxidants [27], was detected at the ppb level. Naphthalene, a
toxicologically and environmentally relevant PAH, capable of inducing ROS production and oxidative
stress [28] also evoked cellular oxidation. Naphthalene is metabolized in mammalian cells to
naphthoquinone indicating that a similar process might be involved in bacteria, as the toxicity of
naphthoquinones is related to disturbances in intracellular GSH concentrations [29].
Zinc pyrithione is known for its antifouling properties, in addition to its bactericidal and fungicidal
activities in cosmetic products [30]. Although reports on the toxicity of ZnPT on various organisms are
relatively abundant [31, and references therein], the mechanism of action remains unclear. Here, we
showed the redox changes caused by ZnPT in E. coli that may provide a probable model for
Gram-negative bacteria. This information is of particular importance for ecotoxicological implications.
Triphenyl tin, a biocide and pesticide organotin [32], also induced oxidation. This finding is the first
report for this mode of action in bacteria according to our best knowledge, and is supported by
modification of critical thiol groups in mitochondria leading to cell death [33,34]. It could be
speculated that organometallic biocides commonly induce cellular oxidation in Gram-negative bacteria.
However, a more thorough examination is required.
A series of results presented in the study suggest a possible application of E. coli-roGFP2 biosensor
to ecotoxicological evaluation of a wide variety of hazardous chemicals. Further testing should
determine the correlation between toxicity and chemical-induced oxidation. Finally, complementing
E. coli-roGFP2 data with Toxicity Identification Evaluation could provide a valuable tool in the
determination of the chemical sources of oxidative stress in environmental samples.

Sensors 2010, 10 6301
4. Materials and Methods
4.1. Chemicals
Tested compounds comprised sodium arsenite, cadmium chloride and copper (II) chloride (Wako
Pure Chemicals Industries, Osaka, Japan), zinc chloride, lead (II) chloride and lead (II) acetate
(Nacalai Tesque, Inc, Kyoto, Japan), menadione (2-methyl-1,4-naphtoquinone) (Mitsuwa Chemical
Co., Ltd, Japan), naphthalene (Sigma Aldrich, Tokyo, Japan), disodium selenite and the 30% hydrogen
peroxide (Santoku Chemical Industries Co., Ltd., Tokyo, Japan), zinc pyrithione (ZnPT) (Sigma
Aldrich, St. Louis, USA) and triphenyltin (TPT) (Fluka, Hong Kong). All chemicals were of the
highest purity available. Stocks were prepared using Milli-Q water (Nihon Millipore KK, Tokyo,
Japan) or analytical grade dimethyl sulfoxide (DMSO) from Sigma-Aldrich. After preliminary tests,
we tested a range of concentrations from non-observable effect concentrations up to 100 ppm, unless
otherwise stated.
4.2. Bacterial strain
Escherichia coli strain DH5α™ (Invitrogen, Tokyo, Japan) cells were transformed with plasmid
pRSET-roGFP2 [13], which allows constitutive expression of roGFP2 protein. We chose roGFP2
because of its higher sensitivity over other roGFP variants [13,18]. Ampicillin (50 µg L
−1
) was used to
assure plasmid maintenance on Luria-Bertani agar and liquid media.
4.3. Spectrophotometric measurement of roGFP2 fluorescence
Five hundred µL of overnight-precultured E. coli cells harboring pRSET-roGFP2 (E. coli-roGFP2)
were inoculated in 50-mL LB liquid medium and further cultivated at 37 °C until optical density at
600 nm reached 1.3. Cells were washed twice in 5 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic
acid (HEPES) buffer containing 171 mM NaCl (pH 7.0). Resuspended cells in 10 mL of the
suspension buffer were left to settle at 20 °C for 1 hour prior testing.
Intensity of fluorescence was measured spectrophotometrically (RF-5300PC, Shimadzu, Tokyo,
Japan). Emission wavelength was fixed at 525 nm and excitation wavelength was scanned
from 350 to 500 nm. Bandwidths of excitation and emission were set at 3 and 10 nm, respectively. Cell
suspension in the cuvette was continuously stirred with a small magnetic bar and an add-on magnetic
stirrer. The fluorescence excitation ratios (400/490 nm) were used as index for cellular oxidation [18].
4.4. Microplate-based measurement of roGFP2 fluorescence
Cell suspension was prepared identically as described above for the spectrophotometric
measurements. Aliquots (99 µL) of cell suspension were placed in the wells of a black flat
bottom 96-well assay plates (BD Falcon, New Jersey, USA), followed by settling the cell suspension
for 1 hour. One microliter of the selected chemical solution was added in the wells before the
beginning of the measurement.

Sensors 2010, 10 6302
Fluorescence intensity was recorded every 30 seconds using excitation filters 400 and 490 nm
(10 and 20 nm bandwidth, respectively). A 528 nm emission filter (20 nm bandwidth) was accordingly
used. Measurements were performed in a multi-detection microplate reader (Powerscan HT,
Dainippon Sumitomo Pharma, Osaka, Japan).
4.5. Measurement of cellular GSH
The content of reduced glutathione (GSH) in E. coli-roGFP2 cells was determined according to
Ellman [35] with slight modifications as described by Inayat-Hussain et al. [36]. Cells were prepared
as described above in the ice-cold suspension buffer. Aliquots of 990 µL were distributed in 1.5 mL
tubes and kept on ice until used. Ten microliters of the desired chemical solution were added to the
tubes followed by incubation at 37 °C. At the desired time (0, 2 and 7 minutes), cells were centrifuged
for 5 minutes (13,000 rpm) at 4 °C. Subsequently, pelleted cells were permealized by incubation in the
lysis buffer containing 50 mM K
2
HPO
4
, 1 mM EDTA, and 0.1% v/v Triton X-100 (pH 6.5) on ice
for 15 minutes. The crude lysates were cleared by centrifugation at 13,000 rpm for 15 minutes at 4 °C.
Then, 50 µL of supernatant were mixed with 50 µL solution of 80 mM Na
2
HPO
4
, 0.8 mM EDTA,
pH 8.0, and 0.8 mg/mL 5,5’-dithiobis-2-nitrobenzoic acid followed by a 15-minutes incubation
at 37 °C. Absorbance at 405 nm was measured using a microplate reader. The concentration of free
thiols in the samples was calculated based on reduced GSH as the standard.
4.6. In vitro effects of chemicals on roGFP2
After culturing E. coli-roGFP2 as mentioned in Section 2.2, cells were resuspended in a buffer
containing 50 mM HEPES (pH 7.9), 300 mM NaCl and 10% glycerol, followed by sonication on ice
for 7 minutes (VC505, Sonics & Materials, Inc., Newton, CT, USA). After discarding the supernatant,
roGFP2 protein, which has a His tag, was purified and concentrated using a Ni
2+
-nitriloacetic
acid-agarose resin (Qiagen, Hilden, Germany) essentially as previously described by
Hanson et al. (2004). The isolated roGFP2 was fully oxidized. In order to reduce roGFP2,
dithiothreitol (DTT) was added to a final concentration of 1 mM, mixed and incubated for 5 minutes.
The reduced roGFP2 was subsequently subjected to gel filtration (Sephadex G-25 column, Pharmacia)
to replace the buffer to 75 mM HEPES (pH 7.0) and 140 mM NaCl, and remove the remaining DTT.
Protein concentration was 1.39 µg mL
-1
, and fluorescence was measured as described in Section 4.3.
4.7. Data analysis
In order to determine the lowest observed effect concentrations (LOECs), datasets were compared
by one-way ANOVA followed by Dunnett multiple comparison post hoc test. For spectrophotometric
measurements, presented data points are the mean of 5 measurements. On the other hand, time points
of the 96-well assay data represent the mean of 6 measurements, unless otherwise stated.

Sensors 2010, 10 6303
Acknowledgements
We are indebted to S.J. Remington, from Oregon University, OR, USA for kindly providing us the
roGFP2-expression plasmid. This study was supported by funds from Kurita Water and Environment
Foundation, Japan Science and Technology Agency—Innovative Seeds, the Okayama Foundation for
Science and Technology, Japan-Malaysia JSPS-VCC Core University Program
(NN-015-2007) and Okayama University—COE Program.
References
1. Magrisso, S.; Erel, Y.; Belkin, S. Microbial reporters of metal bioavailability. Microb. Biotechnol.
2008, 1, 320-330.
2. Farré, M.; Rodríguez-Mozaz, S.; López de Alda, M.J.; Barceló, D.; Hansen, P.D. In The
Handbook of Environmental Chemistry; Barceló, D., Hansen, P.D., Eds.; Springer-Verlag, Berlin,
Germany, 2009; Volume. 5, Chapter 1, pp. 1-32.
3. Franco, R.; Sánchez-Olea, R.; Reyes-Reyes, E.M.; Panayiotidis, M.I. Environmental toxicity,
oxidative stress and apoptosis; Ménage à trois. Mutat. Res. Genet. Toxicol. Environ. Mutagen.
2009, 674, 3-22.
4. Shen, H.; Yang, C.; Liu, J.; Ong, C. Dual role of glutathione in selenite-induced oxidative stress
and apoptosis in human hepatoma cells. Free Radic. Biol. Med. 2000, 28, 1115-1124.
5. Ercal, N.; Gurer-Orhan, H.; Aykin-Burns, N. Toxic metals and oxidative stress part I:
Mechanisms involved in metal-induced oxidative damage. Curr. Top. Med. Chem. 2001, 1,
529-539.
6. Valko, M.; Morris, H.; Cronin, M.T. Metals, toxicity and oxidative stress. Curr. Med. Chem. 2005,
12, 1161-1208.
7. Kumagai, Y.; Sumi, D. Arsenic: Signal transduction, transcription factor, and biotransformation
involved in cellular response and toxicity. Annu. Rev. Pharmacol. Toxicol. 2007, 47, 243-262.
8. Belkin, S.; Smulski, D.R.; Vollmer, A.C.; Van Dyk, T.K.; LaRossa, R.A. Oxidative stress
detection with Escherichia coli harboring a katG'::lux fusion. Appl. Environ. Microbiol. 1996, 62,
2252-2256.
9. Lee, H.J.; Gu, M.B. Construction of a sodA::luxCDABE fusion Escherichia coli: Comparison
with a katG fusion strain through their responses to oxidative stresses. Appl. Microbiol.
Biotechnol. 2003, 60, 577-580.
10. Mitchell, R.J.; Gu, M.B. Construction and characterization of novel dual stress-responsive
bacterial biosensors. Biosens. Bioelectron. 2004, 19, 977-985.
11. Kim, B.C.; Youn,
C.H.; Ahn, J.M.; Gu, M.B. Screening of target-specific stress-responsive genes
for the development of cell-based biosensors using a DNA microarray. Anal. Chem. 2005, 77,
8020-8026.
12. Niazi, J.H.; Kim, B.C.; Ahn, J.M.; Gu, M.B. A novel bioluminescent bacterial biosensor using the
highly specific oxidative stress-inducible pgi gene. Biosens. Bioelectron. 2008, 24, 670-675.

Sensors 2010, 10 6304
13. Hanson, G.T.; Aggeler, R.; Oglesbee, D.; Cannon, M.; Capaldi, R.A.; Tsien, R.Y.; Remington, S.J.
Investigating mitochondrial redox potential with redox-sensitive green fluorescent protein
indicators. J. Biol. Chem. 2004, 279, 13044-13053.
14. Yu, S.; Qin, W.; Zhuang, G.; Zhang, X.; Chen, G.; Liu, W. Monitoring oxidative stress and DNA
damage induced by heavy metals in yeast expressing a redox-sensitive green fluorescent protein.
Curr. Microbiol. 2009, 58, 504-510.
15. Delic, M.; Mattanovich, D.; Gasser B. Monitoring intracellular redox conditions in the
endoplasmic reticulum of living yeasts. FEMS Microbiol. Lett. 2010, 306, 61-66.
16. Meyer, A.J.; Brach, T.; Marty, L.; Kreye, S.; Rouhier, N.; Jacquot, J.P.; Hell, R. Redox-sensitive
GFP in Arabidopsis thaliana is a quantitative biosensor for the redox potential of the cellular
glutathione redox buffer. Plant J. 2007, 52, 973-986.
17. Jubany-Mari, T.; Alegre-Battle, L.; Jiang, K.; Feldman, L.J. Use of a redox-sensing GFP
(c-roGFP1) for real-time monitoring of cytosol redox status in Arabidopsis thaliana water-stressed
plants. FEBS Lett. 2010, 584, 889-897.
18. Dooley, C.T.; Dore, T.M.; Hanson, G.T.; Jackson, W.C.; Remington, S.J.; Tsien, R.Y. Imaging
dynamic redox changes in mammalian cells with green fluorescent protein indicators. J. Biol.
Chem. 2004, 279, 22284-22293.
19. Gutscher, M.; Pauleau, A.L.; Marty, L.; Brach, T.; Wabnitz, G.H.; Samstag, Y.; Meyer, A.J.; Dick,
T.P. Real-time imaging of the intracellular glutathione redox potential. Nat. Methods 2008, 5,
553-559.
20. Hobman, J.L.; Yamamoto, K.; Oshima, T. In Molecular Microbiology of Heavy Metals; Nies,
D.H.; Silver, S., Eds.; Springer-Verlag, Berlin, Germany, 2007; Volume. 1, Chapter 3, pp. 73-115.
21. Smirnova, G.V.; Muzyka, N.G.; Glukhovchenko, M.N.; Oktyabrsky, O.N. Effects of menadione
and hydrogen peroxide on glutathione status in growing Escherichia coli. Free Radic. Biol. Med.
2000, 28, 1009-1016.
22. Helbig, K.; Bleuel, C.; Krauss, G.J.; Nies, D.H. Glutathione and transition-metal homeostasis in
Escherichia coli. J. Bacteriol. 2008, 190, 5431-5438.
23. Quig, D. Cysteine metabolism and metal toxicity. Alter. Med. Rev. 1998, 3, 262-270.
24. Nies, D.H. Microbial heavy-metal resistance. Appl. Microbiol. Biotechnol. 1999, 51, 730-750.
25. Turner, R.J.; Weiner, J.H.; Taylor, D.E. Selenium metabolism in Escherichia coli. Biometals 1998,
11, 223-227.
26. Stocker, J.; Balluch, D.; Gsell, M.; Harms, H.; Feliciano, J.; Daunert, S.; Malik, K.A.;
van der Meer, J.R. Development of a set of simple bacterial biosensors for quantitative and rapid
measurements of arsenite and arsenate in potable water. Environ. Sci. Technol. 2003, 37,
4743-4750.
27. Wright, D.A.; Dawson, R.; Cutler, S.J.; Cutler, H.G.; Orano-Dawson, C.E.; Graneli, E.
Naphthoquinones as broad spectrum biocides for treatment of ship's ballast water: Toxicity to
phytoplankton and bacteria. Water Res. 2007, 41, 1294-1302.
28. Stohs, S.J.; Ohia, S.; Bagchi, D. Naphthalene toxicity and antioxidant nutrients. Toxicology 2002,
180, 97-105.
29. Wilson, A.S.; Davis, C.D.; Williams, D.P.; Buckpitt, A.R.; Pirmohamed, M.; Park, B.K.
Characterisation of the toxic metabolite(s) of naphthalene. Toxicology 1996, 114, 233-242.

Sensors 2010, 10 6305
30. Koutsaftis, A.; Aoyama, I. The interactive effects of binary mixtures of three antifouling biocides
and three heavy metals against the marine algae Chaetoceros gracilis. Environ. Toxicol. 2006, 21,
432-439.
31. Mochida, K.; Fujii, K. Toxicity in plankton and fish Kazuhiko Mochida and Kazunori Fujii, In
Ecotoxicology of Antifouling Biocides; Arai, T., Harino, H., Ohji, M., Langston, W.J., Eds.;
Springer: Tokyo, Japan, 2009; Volume 1, Chapter 21, pp. 365-382.
32. Appel, K.E. Organotin compounds: Toxicokinetic aspects. Drug. Metab. Rev. 2004, 36, 763-786.
33. Zazueta, C.; Reyes-Rivas, H.; Bravo, C.; Pichardo, J.; Corona, N.; Chávez, E. Triphenyltin as
inductor of mitochondrial membrane permeability transition. J. Bioenerg. Biomembr. 1994, 26,
457-462.
34. Robertson, J.D.; Orrenius, S. Molecular mechanisms of apoptosis induced by cytotoxic chemicals.
Crit. Rev. Toxicol. 2000, 30, 609-627.
35. Ellman, G.L. Tissue sulfhydryl groups. Arch. Biochem. Biophys. 1959, 82, 70-77.
36. Inayat-Hussain, S.H.; Chan, K.M.; Rajab, N.F.; Din, L.B.; Chow, S.C.; Kizilors, A.; Farzaneh, F.;
Williams, G.T. Goniothalamin-induced oxidative stress, DNA damage and apoptosis via
caspase-2 independent and Bcl-2 independent pathways in Jurkat T-cells. Toxicol. Lett. 2010, 193,
108-114.
Appendix Figure 1. Fluorescence of roGFP2 expressed in E.coli cells. (A) fluorescence
spectra of roGFP2-expressing cells (filled circle) and vector control (open circle),
(B) subtraction of vector control from the roGFP2-expressing spectrum. Each data point
was obtained within a 1 nm interval. The data presented shows the results from a single
typical experiment of 3 replicates.

Sensors 2010, 10 6306
Appendix Figure 2. Auto-oxidation measurement of roGFP2 in E. coli cells and time
dependent relief from turbulence. Fluorescence was measured spectrophotometrically,
immediately after preparing the cell suspension (0 h) and after settling the cells for 1 h
(1 h) and 2 h (2 h).
© 2010 by the authors; licensee MDPI, Basel, Switzerland. This article is an Open Access article
distributed under the terms and conditions of the Creative Commons Attribution license
(http://creativecommons.org/licenses/by/3.0/).