Accepted Manuscript Title: Stability and toxicity of ZnO quantum dots: interplay between nanoparticles and bacteria Author: Xavier Bellanger Patrick Billard Rapha¨el Schneider Lavinia Balan Christophe Merlin PII: DOI: Reference:
S0304-3894(14)00751-1 http://dx.doi.org/doi:10.1016/j.jhazmat.2014.09.017 HAZMAT 16261
To appear in:
Journal of Hazardous Materials
Received date: Revised date: Accepted date:
20-4-2014 28-8-2014 1-9-2014
Please cite this article as: X. Bellanger, P. Billard, R. Schneider, L. Balan, C. Merlin, Stability and toxicity of ZnO quantum dots: interplay between nanoparticles and bacteria, Journal of Hazardous Materials (2014), http://dx.doi.org/10.1016/j.jhazmat.2014.09.017 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
1 Highlights: Dilution of aminosilane-capped ZnO QDs dramatically increases their dissolution Bacteria limit Zn2+ leakage from ZnO QDs in a physiological-dependent process.
ip t
Implementation of biosensors for assessing free metal promotes QDs instability
Ac ce p
te
d
M
an
us
cr
Dialysis combined to ICP allow studying QDs stability without prior dilution
Page 1 of 32
2 Stability and toxicity of ZnO quantum dots: interplay between nanoparticles and bacteria.
ip t
Authors:
cr
Xavier Bellangera, Patrick Billardb, Raphaël Schneiderc, Lavinia Baland, Christophe Merlina *
Affiliation:
Université de Lorraine and CNRS, Laboratoire de Chimie Physique et Microbiologie pour
us
a
l’Environnement (LCPME), UMR 7564, 15 Avenue du Charmois, 54500 Vandœuvre-lès-
Université de Lorraine and CNRS, Laboratoire Interdisciplinaire des Environnements
M
b
an
Nancy, France.
Continentaux (LIEC), UMR 7360, Boulevard des Aiguillettes, Faculté des Sciences et
Université de Lorraine and CNRS, Laboratoire Réactions et Génie des Procédés (LRGP),
te
c
d
Techniques, BP 70239, 54506 Vandœuvre-lès-Nancy, France.
UMR 7274, 1 rue Grandville, BP 20451, 54001 Nancy Cedex, France. Institut de Science des Matériaux de Mulhouse (IS2M), UMR 7361, CNRS, 15 rue Jean
Ac ce p
d
Starcky, 68093 Mulhouse, France
E-mail addresses:
[email protected] [email protected]
[email protected] [email protected] [email protected] (* Correspondence and reprints)
Page 2 of 32
3 Abstract
The toxicity of quantum dots (QDs) has been commonly attributed to the release of metal ions
ip t
from the core as well as to the production of reactive oxygen species. However, the information related to the stability of the nanoparticles are relatively scarce although this
cr
parameter may strongly influence their toxicity. The stability of aminosilane-capped ZnO QDs, here used as model nanoparticles, was investigated by inductively coupled plasma-
us
optical emission spectrometer (ICP-OES) and whole cell biosensors using a dialysis setup to separate the QDs from the leaked Zn2+ ions. The integrity of the ZnO QDs appeared strongly
an
affected by their dilution in aqueous medium, whereas the nanoparticles were slightly
M
stabilized by bacteria. Our results demonstrate some inadequacy between the implementation
Ac ce p
Keywords
te
d
and use of whole cell biosensors, and the monitoring of metal release from QDs.
ZnO QDs stability, Zn2+bioavailability, QDs toxicity, Escherichia coli, Cupriavidus metallidurans
Page 3 of 32
4 1. Introduction
The great progresses made in nanotechnologies have led to an exponential increase of the use
ip t
of engineered nanoparticles in many domains of life, going from consumer products to essential components of new technologies [1,2]. The developments of nanotechnologies have
cr
been so fast that studies regarding their possible advert effects on health and the environment are lagging behind the speed of progress, making them relatively scarce if not inexistent. This
us
lack of hindsight has attracted increasing concerns regarding the fate of the nanomaterials in the environment and the toxicity specifically linked to their small size. If their nanoscale and
an
the great surface to volume ratio associated are the essence of their outstanding properties,
M
they are also critical in terms of toxicity as they increase the bioavailability of the nanoparticles. Therefore, processes controlling the behavior of nanoparticles with respect to
d
dissolution or aggregation should necessarily have an impact on their persistence and their
te
toxicity in the environment [3,4]. With this respect, any risk assessment associated to nanoparticles release in the environment depends on the environment itself, with all its
Ac ce p
complexity, as well as on the nature and on the chemistry of the nanoparticles considered. Over the past fifteen years, semiconductor nanocrystals (quantum dots, QDs) have received considerable attention because of their unique optical properties. Their narrow and tunable fluorescence emission associated to broad absorption spectra, high quantum yields and good resistance to chemical degradation and photobleaching, have made QDs interesting nanoparticles for various applications ranging from bioimaging to optoelectronics [5-7]. CdSe- and CdTe-core QDs have been extensively studied because of the quality and the flexibility of their optical properties. However, the use of Cd-containing QDs is constrained by their dual toxicity originating from their Cd2+ content and their ability to generate deleterious reactive oxygen species (ROS) [8-10]. Both types of toxicity are controlled by the
Page 4 of 32
5 stability of the particle and have deep consequences on the QDs application outputs in terms of biological alteration whether it concerns purposely exposure, in the context of bioimaging for instance, or accidental environmental exposure, thus raising the problem of waste
ip t
management. Recently, great efforts have been made to reduce the toxicity of QDs with alternative semiconductors composed of less toxic metals, such as zinc oxide, while keeping
cr
interesting optical properties thanks to elaborated particle synthesis chemistry [11]. Still, the toxicity of the particles remains deeply associated to their state and therefore their stability
us
with respect to metal release (dissolution), reactivity (ROS production), or bioavailability of the particle (nanoscale versus aggregated state). The integrity of nanoparticles is relatively
an
easy to assess in vitro using conventional methods, such as filtration followed by inductively
M
coupled plasma (ICP) analyses that allow the quantification of the dissolved metals. Metal leakage can also be assessed using biological-based methods with whole cell biosensors for
d
instance. Such biosensors consist of microorganisms that have been genetically modified to
te
produce a quantifiable signal (e.g. fluorescence, bioluminescence) in response to specific environmental stimuli including metals [12,13]. Moreover, whole cell biosensors provide the
Ac ce p
specific advantage of measuring the bioavailable fraction of the analyte of interest as opposed to the total content as determined by chemical means. Biosensors were used on several occasions to quantify metal release from nanoparticles [14], including QDs [10,15], and therefore allow monitoring their stability with respect to dissolution. Additionally, specific whole cell biosensors allow evaluating more subtle particle-induced stresses such as oxidative stress linked to ROS production [10,16]. Biosensors are highly sensitive, selective, easy to implement, and only require the cells to reach a particular physiological state and to be exposed to serial dilutions of the sample to be analyzed. However, caution must be taken when evaluating nanoparticle dissolution using biosensor-based assays as the impact of the biosensor cells itself on the stability of the studied material has to be considered. To the best
Page 5 of 32
6 of our knowledge, the influence of either biosensor cells or the implementation of such biosensors on nanoparticles has not been documented yet. This study aimed to evaluate the influence of the Cupriavidus metallidurans and Escherichia
ip t
coli sensor strains on the stability of ZnO core QDs functionalized with 3aminopropyltrimethoxysilane (APTMS). ZnO@APTMS QDs were here chosen as model
cr
particles as the APTMS functionalization allows the ZnO core to become dispersible in
aqueous biological environment and, in the meantime, provide a good colloidal stability to the
us
nanoparticles as we reported previously [15]. We report an original approach, combining dialysis, ICP analyses and biosensing, for determining the relative contributions of the
an
bacterial cells and the experimental set up on the stability of the QDs. Zn2+ release, which is
M
the main source of ZnO QDs induced toxicity in the present study, appeared to be influenced
te
2. Materials and methods
d
by the presence of bacteria, their physiological state and the species considered.
Ac ce p
2.1 Synthesis and characterization of ZnO quantum dots
Triethyleneglycol-capped ZnO QDs were prepared by a sol-gel route using LiOH as base [17]. A LiOH/Zn(OAc)2 molar ratio of 2 was used for the synthesis. A first ligand exchange with oleic acid was undertaken to transfer the dots in weakly polar organic solvents. ZnO@oleate QDs were next transferred in aqueous solution by treatment with APTMS in DMF. In a typical experiment, under a argon atmosphere, 7.6 µL of APTMS were added to 50 mg of ZnO QDs dispersed in 7 mL of anhydrous DMF. The mixture was stirred 5 min at room temperature and next heated at 120°C for 15 min. After cooling and centrifugation, the
Page 6 of 32
7 white precipitate was washed with DMF (2 x 5 mL) and ethanol (2 x 5 mL) and dried at room temperature. The pellet was readily re-dispersed in aqueous solution for further study. The Zn content of QDs was measured by inductively coupled plasma optical emission
ip t
spectrometer (ICP-OES) (Varian) as being of 4.05 mM of Zn/g of QDs, i.e. Zn accounts for 26.5% of the QD mass. ZnO QD concentrations were expressed in molar Zn equivalent (MZn
cr
eq.)
us
2.2 Bacterial strains
an
Two wild-type bacterial strains, Escherichia coli MG1655 [18] and Cupriavidus
M
metallidurans CH34 [19] were used in this study to assess the stability and toxicity of ZnO QDs. Both strains were transformed according to Choi et al. [20] with the luxCDABE operon
d
bearing plasmid pUCD607 [21], which confers a constitutive luminescent phenotype and
te
allows evaluating the nanoparticle toxicity based on bioluminescence extinction. MG1655(pUCD607) and CH34(pUCD607) transformants were maintained on LB agar
Ac ce p
medium (LB Broth Miller, DifcoTM) supplemented with kanamycin at 50 and 500 µg/mL, respectively. Bioavailability of Zn2+ was measured using the C. metallidurans biosensor strain AE1433 deriving from CH34 [22].
2.3 QD suspensions and ZnCl2 solutions
ZnCl2 was used to get standard solutions of known Zn2+ ion concentrations and was chosen over other zinc mineral salts for its very good solubility in water. Solutions of ZnCl2 and suspensions of QDs were prepared in RM mineral medium, a MOPS-based mineral culture medium previously designed to minimize metal chelation and precipitation [22]. The stock
Page 7 of 32
8 suspensions of QDs were prepared at 3.10-3 M and sonicated at 130 W for 1 min to be dispersed in the medium. QDs suspensions were vortexed prior each utilization. Freshly made
ip t
ZnCl2 solutions and QDs suspensions were prepared before each replicate experiment.
cr
2.4 ZnCl2 and QD toxicity assays
A single colony of either E. coli MG1655(pUCD607) or C. metallidurans CH34(pUCD607)
us
was picked from a LB selective plate and used to inoculate 10 mL of LB broth containing kanamycin in a 100 mL conical flask. The culture was grown at 30°C with continuous
an
shaking at 160 rpm for 16 to 18h. Bacterial cells were harvested by centrifugation (3,000 rpm,
M
3 min), washed with 10 mL of RM supplemented with SL7 trace elements solution [23] and 0.2 % gluconate as a carbon source [22], and then centrifuged again in the same conditions.
d
Cell pellets were re-suspended in 10 mL of RM - SL7 gluconate 0.2 % and allowed to adapt
te
in the medium at 30°C with continuous shaking (160 rpm) for one hour. The OD600 of the cell suspensions were then adjusted to 0.3 for E. coli MG1655(pUCD607) and to 0.54 for
Ac ce p
C. metallidurans CH34(pUCD607) in the same medium, and dispensed (180 µL for E. coli and 100 µL for C. metallidurans) in clear bottom black microtiter plates (Isoplate-96F, PerkinElmer), previously filled with various dilutions of either ZnCl2 or QDs to reach a 200 µL final volume. Microplates were maintained at 25°C in a VICTOR3 Multilabel Counter plate reader (Perkin-Elmer), where light emission and OD600 were recorded every 3 min, with a 5 s and 1 s measurement time, respectively, preceded by a 10 s double orbital shaking. For both parameters, background provided by RM-SL7 gluconate medium was subtracted, and the luminescence was normalized to the corresponding OD600. Normalized luminescence values were averaged among quadruplicate and induction factors were calculated by dividing the normalized luminescence values by those obtained for the untreated control. The maximal No
Page 8 of 32
9 Observed Effect Concentration (NOEC) and the half maximal Inhibitory Concentration (IC50) were determined 120 minutes after exposure.
ip t
2.5 Quantification of bioavailable Zn2+
cr
Bioavailability of Zn2+ was quantified using the whole cell biosensor Cupriavidus
metallidurans AE1433 that has been genetically engineered to emit a bioluminescence signal
us
when sensing bioavailable Zn2+ [22]. The protocol used is slightly modified from Corbisier et al [22]. Briefly, strain AE1433 was cultured in LB medium until stationary phase (OD600 =
an
1.5) and cells were washed twice by alternating centrifugation (10,000 g for 2 min at 20°C)
M
and cell pellet dispersion in RM medium supplemented with gluconate 0.01 %. The cell density was finally adjusted to 0.54 and 100 µl of the suspension was distributed in black
d
microtiter plates with clear bottom, previously filled with 100 µl of various dilutions of either
te
ZnCl2, for calibration curves, or QDs, for testing. Bioluminescence and OD600 were recorded as described above at 25°C in a VICTOR3 Multilabel Counter plate reader (Perkin-Elmer)
Ac ce p
and each test was carried out in 3 independent replicates. For each series of experiments, luminescence induction factors were plotted against ZnCl2 concentrations to set a standard curve from which linear regression could be obtained for the non-toxic part of the curve, and were used to deduce the apparent concentrations of bioavailable Zn2+ leaking from QDs.
2.6 QD dialysis in the presence or absence of bacterial cells
One milliliter of QD suspension was introduced in a 1 mL ready-to-use dialysis device with a molecular weight cut-off of 8-10 kD (Float-A-Lyzer G2; Spectra/Por) previously conditioned as indicated by the manufacturer. QD suspensions were dialyzed for 24h, in the dark, under
Page 9 of 32
10 agitation (160 rpm), and at 30°C against 24 mL of RM gluconate 0.01% medium in sterile capped tubes. Dialysates and retentates were further analyzed for their Zn content by ICPOES or using the AE1433 biosensor (Fig. 1).
ip t
When testing the effect of cells on QD stability, bacterial suspensions in different physiological states were added together with the nanoparticle in the ready-to-use dialysis
cr
device. Briefly, strains E. coli MG1655 or C. metallidurans CH34 were grown at 30°C under agitation (160 rpm) in conical flasks filled with 1/5th of LB medium. Cultures were either
us
stopped at OD600 = 0.3 to obtain exponentially growing cells, or were incubated for 16 to 18 h to obtain cells in late stationary phase of growth. Cells were then washed twice with RM
an
gluconate 0.01 % and the concentration of the cell suspension was finally adjusted to OD600 =
M
30. Ten microliters of concentrated cell suspension were added to the QD suspension within the Float-A-Lyzer G2 to obtain 1 mL of QD suspension with cells at final OD600 = 0.3. For
d
one tests series, the influence of dead material was evaluated with stationary phase cells boiled for 10 minutes before being added to QD suspensions in the dialysis device. For each
te
condition, the amount of Zn2+ released from the QDs was deduced from ICP-OES
Ac ce p
measurement in the dialysate.
2.7 Statistics
The IC50 was calculated by solving the fitted equation for relative luminescence equal to 50% by using the online curve fitting web site ZunZun.com (http://zunzun.com/Equation/2/Sigmoidal/Boltzmann Sigmoid A/). Statistical analysis was performed using the web tool BiostaTGV (http://marne.u707.jussieu.fr/biostatgv/) and as described by Cumming et al [24]. All websites were last accessed on 27 February 2014.
Page 10 of 32
11 3. Results
ip t
3.1 ZnO QD toxicity towards E. coli and C. metallidurans
With the rapid development of QD-based technologies, an increasing attention is given to a
cr
better control of the toxicity associated to these metal-containing particles. If the toxicity of QDs is now recognized to involve both the liberation of metal ions from the core and the
us
production of ROS, the relative contribution of the two phenomena can vary from one particle to another, and both are impacted by the particle stability. This work presents an integrated
an
study regarding the toxicity and the stability of ZnO@APTMS QDs, used as model
M
nanoparticles. These ZnO QDs were synthesized using a sol-gel route and were easily dispersed in aqueous solution after surface functionalization with APTMS [15]. Transmission
d
electron microscopy and X-ray diffraction experiments showed that ZnO QDs have an
te
average diameter of ca. 4 nm and a cubic zinc blende structure, respectively (Fig. 2a and 2b). The photoluminescence spectrum of colloidal ZnO@APTMS QDs shows a broad emission
Ac ce p
centered at 530 nm, which is typical for ZnO QDs prepared via a sol-gel process (Fig. 2c). The electrical potential of these dots, referred to as the zeta potential, was measured to be +27.4 ± 4 mV at pH = 7.0.
The toxicity of APTMS-coated ZnO QDs towards E. coli and C. metallidurans was determined using two bacterial strains that have been genetically modified to emit light constitutively, namely E. coli MG1655(pUCD607) and C. metallidurans CH34(pUCD607). Any toxicity interfering with the bacterial metabolism will result in the reduction of the emitted bioluminescence. For both strains, bioluminescence extinction curves are presented in Fig. 3. It can be seen that ZnO QDs and ZnCl2 were more toxic towards E. coli MG1655 (IC50 = 1.610-5 M and 2.710-5 M for ZnCl2 and ZnO QDs, respectively) than towards
Page 11 of 32
12 C. metallidurans CH34 (IC50 = 2.810-4 M and 1.210-3 M for ZnCl2 and ZnO QDs, respectively). Strain CH34 is well-known for its resistance to several metals cations including Zn2+ [19], which may account for such strain-specific differences in metal/QDs sensitivity.
ip t
For both strains, the ZnO QDs appeared less toxic than their Zn2+ content. Considering that all the toxicity experiments were carried out in the dark, photo-induced ROS should not be
cr
considered, therefore ruling out the involvement of the QDs photoreactivity in the observed extinction of bioluminescence. Thus, the toxicity observed should mainly result from the
us
liberation of Zn2+ cations from the ZnO QD cores. Interestingly, for E. coli the toxicity curves
an
for ZnCl2 and ZnO QDs showed similar shapes and were quite close to each other as seen by the two IC50 values of the same range (1.610-5 M and 2.710-5 M, respectively), thus tending
M
to show that ZnO QDs were almost, if not completely, dissolved. In contrast, for C. metallidurans, the two toxicity curves appeared well separated, with almost one order of
d
magnitude between the two IC50 values (2.810-4 M and 1.210-3 M for ZnCl2 and ZnO QDs,
te
respectively), suggesting that the QDs only partially dissolved. Although at this stage it is not yet possible to explain the differences observed between the ZnCl2 and the ZnO QDs toxicity
Ac ce p
curves, it should be noted that (i) the different concentrations at which the QDs toxicity was observed, from one bacteria to the other, may affect the dissolution equilibrium of the particle, and (ii) that the two bacterial strains may possibly exert different effects on the QDs stability.
3.2 Determining bioavailable Zn2+ leaked from ZnO QDs
The concentration-dependent dissolution of ZnO QDs was first investigated using a wholecell Zn2+ biosensor. Strain AE1433 of C. metallidurans, emitting light when sensing Zn2+, was employed to determine the amount of bioavailable Zn2+ ions released from freshly
Page 12 of 32
13 prepared dispersions of ZnO QDs. Dilutions of QDs ranging from 710-6 to 210-3 MZn eq. were exposed to C. metallidurans AE1433 whose luminescence was compared to that obtained with a ZnCl2 standard. For QDs concentrations ranging from 710-6 to 310-5 MZn the apparent concentrations of bioavailable Zn2+ fitted the Zn2+ content of the QDs, thus
ip t
eq.,
indicating a complete dissolution of the particle (Fig. 4), which is in agreement with what was
cr
suspected while studying the QDs toxicity on E. coli (Fig. 3). At higher QD concentrations,
the apparent concentrations of bioavailable Zn2+ reached a maximum at around 40 µM, which
an
proportional anymore due to metal-induced toxicity.
us
coincides with the Zn2+ concentration limit over which the response of the biosensor is not
In order to study Zn2+ leaking at higher QDs concentrations and, in the mean time, to take into
M
consideration a possible contact-effect between the biosensor cells and the nanoparticles, a dialysis-based approach was setup. In a first attempt, 1 mL of a 310-3 M Zn eq QDs
d
suspension was dialyzed for 24 hours against 24 mL of RM gluconate 0.01 % medium. Strain
te
AE1433 was used to estimate and compare the apparent concentration of bioavailable Zn2+ ions in both dialysis compartments (the retentate and the dialysate) at two different times (t =
Ac ce p
0 and t = 24 h). Experimentally, the content of both compartments was sampled, serially diluted and assayed with the biosensor as described above. A set of dilutions producing a biosensor response within the range of usable concentrations (i.e. linear relationship between light emission and Zn2+ concentration) was selected and used to determine the bioavailable Zn2+ in each compartment. Fig. 5 shows that the amounts of bioavailable Zn2+ measured did not significantly differ at t = 0 and t = 24 h. In both cases, the apparent concentrations of bioavailable Zn2+ in the retentates appeared about a thousand times more concentrated than those of the dialysates. If such a difference could be expected before equilibrium (at t = 0), the concentrations of bioavailable Zn2+ should equilibrate upon dialysis, unless the bioavailable Zn2+ ions are released from the QDs after the dialysis process, during the exposure to the
Page 13 of 32
14 biosensor cells. Therefore, it can be assumed that ZnO QDs are relatively stable over time at elevated concentrations but further dilution and/or incubation with the biosensor are likely to be responsible for the liberation of bioavailable Zn2+ ions detected in biosensing assays.
ip t
Indeed, bacteria have a variety of properties that can affect metal solubility [25] and initially insoluble forms of heavy metals salts may become bioavailable due to the direct contact with
cr
bacterial biosensor [26,27]. The present experiments show that the procedure associated to the
an
experiment in terms of Zn2+ leakage quantification.
us
use of a whole cell biosensor affect the state of the particle, and therefore the outcome of the
M
3.3 Stability of ZnO QD upon dilution
In order to dissociate the concentration effect (dissolution equilibrium) from the cell exposure effect (biodegradation for instance) regarding the fate of ZnO QDs, the QDs stability was
te
d
further studied using the same dialysis setting but with direct quantification of the Zn2+ content of each compartment without prior dilution. Series of 1 mL of QDs suspensions at
Ac ce p
concentrations ranging from 310-5 to 310-3 M Zn eq. were dialyzed as previously for 24 hours against 24 mL of RM gluconate 0.01 % medium. Both compartments were analyzed by ICPOES and the Zn2+ content in the dialysates was used to estimate the amount of free Zn2+ ions having leaked from the QDs. At the highest QDs concentration tested (310-3 M Zn eq), the concentration of free Zn2+ was estimated to be 1.610-5 M accounting for 12% of the initial QDs content (Fig. 6). In contrast, at low QD concentration, close to 10-4 M Zn eq., the amount of Zn2+ leaking from the QDs increased to 100%. Therefore, the QDs stability appears to vary according to its initial concentration within the dialysis device, which likely reflects the QDs dissociation equilibrium in aqueous solution. Furthermore, it should be noted that for an initial QDs concentration of 2.710-5 M Zn eq., corresponding to the IC50 observed for E. coli
Page 14 of 32
15 (Fig. 3), 100% of the particles are dissolved, thus explaining the closeness of the QDs and ZnCl2 toxicity curves. Finally, at 1.210-3 M Zn eq., the IC50 obtained for C. metallidurans, ca. 40% of the Zn2+ ions have leaked from the QDs, which agrees well with the apparent lower
ip t
toxicity of particles compared to their Zn content for this bacterium.
cr
3.3 Impact of the bacteria on the stability of ZnO QDs
us
Having demonstrated the influence of QDs dilution on their dissolution, QDs stability was
an
further investigated to evaluate the influence of the bacterial cells. For this purpose, the nanoparticles were co-incubated with bacterial cells in the dialysis device at three significant
M
ZnO QDs concentrations: 310-3, 310-4 and 310-5 M Zn eq., presenting 12%, 70%, and 100% QDs dissolution in terms of Zn2+ release, respectively (Fig. 6). The QDs were independently
d
incubated with either E. coli MG1655 or C. metallidurans CH34, from cultures in different
te
metabolic states (exponential or stationary growth phase), or even to dead cells resulting from boiled cultures. The release of Zn2+ from QDs was quantified by ICP-OES measurements of
Ac ce p
the dialysates and the retentates after 24 hours. The addition of bacteria to QDs at the two extreme concentrations (310-3 and 310-5 MZn eq.) did not significantly modify the behaviors of the nanoparticles upon dialysis regardless the state of the cells (Fig. 7a and 7c). Nevertheless, for a QDs concentration of 310-4 M Zn eq., at which the nanoparticles displayed an intermediate dissolution state, the presence of bacteria significantly reduced the release of free Zn2+ in some instances (Fig. 7b). For both strains, dead cells have statistically no effect on QDs stability. In contrast, metabolically active bacteria added in exponential growth phase significantly reduced the release of free Zn2+ from 70% to 20% and 30% of the QDs Zn content for C. metallidurans and E. coli, respectively. An intermediate effect could be observed for C. metallidurans added in stationary phase of growth. Because the release of free
Page 15 of 32
16 Zn2+ is quantified on the basis of the amount of Zn2+ able to cross the dialysis membrane, a reduction of free Zn2+ in the dialysate could also originate from the Zn2+ sequestration by the bacterial cells in the retentate compartment. To examine this possibility, the dialysis
ip t
experiment was run with various concentrations of ZnCl2 co-incubated or not with bacterial cells. In each case, Zn2+ concentrations determined by ICP-OES in the dialysates after 24
cr
hours of dialysis did not differ whether or not bacterial cells were present, and whatever their physiological state (data not shown), therefore ruling out the possible retention of Zn2+ by
us
bacterial cells. Thus, against all odd, bacteria seem to be able to limit the Zn2+ leakage from
an
ZnO QDs in a way that appeared to be both strain- and physiological state-dependent.
M
4. Discussion and conclusion
d
This work demonstrates that both the dilution of ZnO@APTMS QDs and the presence of bacterial cells interfere in opposite directions with the particles stability regarding the
te
liberation of Zn2+ ions. Toxicity studies with QDs often oppose the liberation of metal to the
Ac ce p
ROS production and scarcely include the problem posed by the nanoparticle stability. However, this stability is of prime importance to dictate QDs toxicity either towards the ROS production or towards the liberation of metal. Studying QDs toxicity often implies exposing the biological material to dilution series of the nanoparticles in order to discover critical concentrations affecting biological functions. The dilution, by promoting the particles dissolution, is likely to orientate the QDs toxicity solely towards their metal content, thus biasing our perception of the QDs toxicity linked to the reactivity of the particles. This observation also stands for the use whole-cell metal biosensors. Because these biosensors can only operate within a define range of concentrations, from the detection limit to toxic concentrations, they often require working on dilution series of the samples. If the sample
Page 16 of 32
17 stability is sensitive to dilution in terms of metal release, biosensing is likely to overestimate the bioavailability of metals from unstable samples. This points out that great care should be given to preliminary study regarding the stability of nanoparticles such as QDs while studying
ip t
their toxicity in relation to metal release. In this context, we present an original dialysis-based approach to study the stability of QDs, which allows maintaining the nanoparticles in various
cr
conditions, including in the presence of bacterial cells, while analyzing the metal released in
another compartment. With this approach, we can also expect to limit biases associated with
us
the centrifugation procedure commonly used to measure metal ion release in nanotoxicology [28].
an
We demonstrated that, within the limit of the experiments presented, bacteria reduced the
M
metal release from ZnO@APTMS QDs. Cultures of Pseudomonas aeruginosa were also reported to stabilize and limit the dissolution CdSe QDs. However, the mechanism by which
d
such protection occurred was not investigated further [29]. We also showed that the
te
physiological state of these bacteria could modulate the stability of the dots. The simple fact that dead bacteria do not significantly alter QDs stability, and that cells in stationary phase
Ac ce p
stabilize less than cells in log phase, suggests that the stabilization is an active process rather than being solely linked to a structural component of the bacteria. With this respect, the membrane potential has been shown to vary according to the physiological state [30]. On the other hand, QDs have been shown to interfere with the bacterial membrane potential [31], which raises the question of QDs stability in a biologically generated electron flux.
Acknowledgments
This work is supported by the Agence Nationale pour la Recherche (ANR CESA 2011, project NanoZnOTox) and the Zone Atelier Moselle. We thank Hervé Marmier (LIEC,
Page 17 of 32
18 Université de Lorraine) for ICP-OES analyses and are grateful to Hélène Guilloteau (LCPME, Université de Lorraine) for technical support and to Boussad Arroua for his contribution to the toxicity experiments. We also thank Ghouti Medjahdi (IJL, UMR CNRS 7198, Université
ip t
de Lorraine) for XRD analyses.
cr
References
us
[1] J.R. Peralta-Videa, L. Zhao, M.L. Lopez-Moreno, G. de la Rosa, J. Hong, J.L.GardeaTorresdey, Nanomaterials and the environment: a review for the biennium 2008-2010, J.
an
Hazard. Mater. 186 (2011) 1-15.
M
[2] M.C. Roco, M.C. Mirkin, C.A. Hersam, Nanotechnology research directions for societal needs in 2020, Springer, Netherlands, 2011.
d
[3] S.J. Klaine, P.J. Alvarez, G.E. Batley, T.F. Fernandes, R.D. Handy, D.Y. Lyon, S.
te
Mahendra, M.J. McLaughlin, J.R. Lead, Nanomaterials in the environment: behavior, fate,
Ac ce p
bioavailability, and effects, Environ. Toxicol. Chem. 27 (2008) 1825-1851. [4] G.E. Batley, J.K. Kirby, M.J. McLaughlin, Fate and risks of nanomaterials in aquatic and terrestrial environments, Acc. Chem. Res. 46 (2013) 854-862. [5] T. Jamieson, R. Bakhshi, D. Petrova, R. Pocock, M. Imani, A.M. Seifalian, Biological applications of quantum dots, Biomaterials 28 (2007) 4717-4732. [6] D. Bera, L. Qian, T.K. Tseng, P.H. Holloway, Quantum dots and their multimodal applications: a review, Materials 3 (2010) 2260-2345. [7] P. V. Kamat, Quantum dot solar cells, The next big thing in photovoltaics, J. Phys. Chem. Lett. 4 (2013) 908-918.
Page 18 of 32
19 [8] J.M. Tsay, X. Michalet, New light on quantum dot cytotoxicity, Chem. Biol.12 (2005) 1159-1161. [9] R. Hardman, A toxicologic review of quantum dots: toxicity depends on physicochemical
F.A. Kauffer, C. Merlin, L. Balan, R. Schneider, Incidence of the core composition on
cr
[10]
ip t
and environmental factors. Environ. Health Perspect. 114 (2006) 165-172.
the stability, the ROS production and the toxicity of CdSe quantum dots, J. Hazard. Mater.
T. Pons, E. Pic, N. Lequeux, E. Cassette, L. Bezdetnaya, F. Guillemin, F. Marchal, B.
an
[11]
us
268 (2014) 246-255.
Dubertret, Cadmium-free CuInS2/ZnS quantum dots for sentinel lymph node imaging with
[12]
M
reduced toxicity, ACS Nano 4 (2010) 2531-2538.
S. Daunert, G. Barrett, J.S. Feliciano, R. Shetty, S. Shrestha, W. Smith-Spencer,
d
Genetically engineered whole-cell sensing systems: coupling biological recognition with
S. Köhler, S. Belkin, R.D. Schmid, Reporter gene bioassays in environmental analysis,
Ac ce p
[13]
te
reporter genes, Chem. Rev. 100 (2000) 2705-2738.
Fresenius J. Anal. Chem. 366 (2000) 769 – 779. [14]
A. Kahru, H.C. Dubourguier, I. Blinova, A. Ivask, K. Kasemets, Biotests and
biosensors for ecotoxicology of metal oxide nanoparticles: a minireview, Sensors 8 (2008) 5153-5170. [15]
A. Aboulaich, C.M. Tilmaciu, C. Merlin, C. Mercier, H. Guilloteau, G. Medjahdi, R.
Schneider, Physicochemical properties and cellular toxicity of (poly) aminoalkoxysilanesfunctionalized ZnO quantum dots, Nanotechnology 23 (2012) 335101.
Page 19 of 32
20 [16]
A. Ivask, O. Bondarenko, N. Jepihhina, A. Kahru, Profiling of the reactive oxygen
species-related ecotoxicity of CuO, ZnO, TiO2, silver and fullerene nanoparticles using a set of recombinant luminescent Escherichia coli strains: differentiating the impact of particles
[17]
ip t
and solubilised metals, Anal. Bioanal. Chem. 398 (2010) 701-716. H.M. Xiong, R.Z. Ma, S.F. Wang, Y.Y. Xia, Photoluminescent ZnO nanoparticles
cr
synthesized at the interface between air and triethylene glycol, J. Mater. Chem. 21 (2011)
[18]
us
3178-3182.
F.R. Blattner, G. Plunkett, C.A. Bloch, N.T. Perna, V. Burland, M. Riley, J. Collado-
an
Vides, J.D. Glasner, C.K. Rode, G.F. Mayhew, J. Gregor, N.W. Davis, H.A. Kirkpatrick, M.A. Goeden, D.J. Rose, B. Mau,Y. Shao, The complete genome sequence of Escherichia
M. Mergeay, D. Nies, H.G. Schlegel, J.Gerits, P. Charles, F. Van Gijsegem,
d
[19]
M
coli K-12, Science 277 (1997) 1453-1462.
te
Alcaligenes eutrophus CH34 is a facultative chemolithotroph with plasmid-bound resistance
Ac ce p
to heavy metals, J. Bacteriol. 162 (1985) 328-334. [20]
K.H. Choi, A. Kumar, H.P. Schweizer, A 10-min method for preparation of highly
electrocompetent Pseudomonas aeruginosa cells: application for DNA fragment transfer between chromosomes and plasmid transformation, J. Microbiol. Meth. 64 (2006) 391-397. [21]
J.J. Shaw, C.I. Kado, Development of a Vibrio bioluminescence gene-set to monitor
phytopathogenic bacteria during the ongoing disease process in a nondisruptive manner, Nat. Biotechnol. 4 (1986) 560-564. [22]
P. Corbisier, D. van der Lelie, B. Borremans, A. Provoost, V. de Lorenzo, N.L.
Brown, J.R. Loyd, J.L. Hobamn, E. Csöregi, G. Johansson, B. Mattiasson,Whole cell- and
Page 20 of 32
21 protein-based biosensors for the detection of bioavailable heavy metals in environmental samples, Anal. Chim. Acta 387 (1999) 235–244. [23]
H. Biebl, N. Pfennig, Isolation of members of the family Rhodospirillaceae, in M. P.
ip t
Starr, J. Stolp, H. G. Truper, A. Balows, and H. G. Schlegel (Eds.), The prokaryotes, a handbook on habitats, isolation and identification of bacteria, vol. 1. Springer-Verlag, Berlin,
G. Cumming, F. Fidler, D.L. Vaux, Error bars in experimental biology, J. Cell. Biol.
us
[24]
cr
1981, pp. 267-273.
[25]
an
177 (2007) 7-11.
G.M. Gadd, Metals, minerals and microbes: geomicrobiology and bioremediation,
[26]
M
Microbiology 156 (2010) 609-643.
A. Kahru, A. Ivask, K. Kasemets, L. Põllumaa, I. Kurvet, M. Francois, H.C.
d
Dubourguier, Biotests and biosensors in ecotoxicological risk assessment of field soils
O. Bondarenko, T. Rõlova, A. Kahru, A. Ivask, Bioavailability of Cd, Zn and Hg in
Ac ce p
[27]
te
polluted with zinc, lead and cadmium, Environ. Toxicol. Chem. 24 (2005) 2973-2982.
soil to nine recombinant luminescent metal sensor bacteria, Sensors 8 (2008) 6899-6923. [28]
M. Xu, J. Li, N. Hanagata, H. Su, H. Chen, D. Fujita, Challenge to assess the toxic
contribution of metal cation released from nanomaterials for nanotoxicology-the case of ZnO nanoparticles, Nanoscale 5 (2013) 4763-4769. [29]
J.H. Priester, P.K. Stoimenov, R.E. Mielke, S.M. Webb, C. Ehrhardt, J.P. Zhang, G.D.
Stucky, P.A. Holden, Effects of soluble cadmium salts versus CdSe quantum dots on the growth of planktonic Pseudomonas aeruginosa, Environ. Sci. Technol. 43 (2009) 2589-2594.
Page 21 of 32
22 [30]
P. Monfort, B. Baleux, Cell cycle characteristics and changes in membrane potential
during growth of Escherichia coli as determined by a cyanine fluorescent dye and flow cytometry, J. Microbiol. Meth. 25 (1996) 79-86. Y. Yang, H. Zhu, V.L. Colvin, P.J. Alvarez, Cellular and transcriptional response of
ip t
[31]
Pseudomonas stutzeri to quantum dots under aerobic and denitrifying conditions, Environ.
us
cr
Sci. Technol. 45 (2011) 4988-4994.
an
Figure captions
M
Fig. 1: Experimental setup for the quantification of Zn2+ released from ZnO QDs. QDs were incubated in the dialysis device in presence or absence of bacterial cells. The relative
d
amount of Zn2+ released was calculated from ICP-OES or biosensing measurement made in
te
the dialysate. Total Zn engaged in the experiment was obtained by combining ICP
Ac ce p
measurement in the retentate plus the dialysate.
Fig. 2: (a) TEM micrograph, (b) XRD pattern and (c) absorption (black) and photoluminescence (red) spectra of ZnO@APTMS QDs. Absorption and photoluminescence spectra were recorded in water (excitation wavelength of 330 nm).
Fig. 3: Effect ZnCl2 or ZnO QD concentrations on the relative luminescence emitted by E. coli MG1655(pUCD607) and C. metallidurans CH34(pUCD607). Squares and diamonds represent experiments carried out with E. coli and C. metallidurans, respectively. Continuous and dotted lines represent the experiments performed with ZnCl2 and ZnO QDs, respectively.
Page 22 of 32
23 Each data point is the mean of three independent experiments (with four repeats for each) and errors bars correspond to standard error of the mean.
ip t
Fig. 4: AE1433 biosensing of apparent bioavailable Zn2+ released by the ZnO QDs. The inset presents the results obtained at low QD concentrations (from 6.7×10-7 to 6.7×10-5 MZn
for which a linear fit could be obtained (equation: y = 0.99x; coefficient of determination
cr
eq.)
R2 = 0.996). Each data point is the mean of three independent experiments (with four repeats
us
for each) and errors bars correspond to standard error of the mean.
an
Fig. 5: AE1433 biosensing of apparent bioavailable Zn2+ released by the ZnO QDs
M
before and after a 24 hours dialysis. One mL of QD suspension at a concentration of 3×10-3 MZn eq. was dialyzed against 24 mL of RM gluconate 0.01% medium. The dotted line indicates
d
the theoretical initial QD concentration within the dialysis device at t=0. Each data point is
te
the mean of at four replicates and errors bars correspond to standard error of the mean.
Ac ce p
Fig. 6: Quantification of the Zn2+ released as a function of the initial ZnO QD concentration after 24h of dialysis. One mL of QD suspension was dialyzed against 24 mL of RM gluconate 0.01% medium. Total amounts of Zn2+ released were extrapolated from ICP-OES quantification of the dialysate. The dotted line indicates a log fit between the initial QD concentration (from 2×10-4 to 3×10-3 MZn eq.) and the relative amount of Zn2+ released (the equations and coefficients of determination corresponding to the trend lines are indicated). Each data point is the mean of at least three independent experiments and errors bars correspond to standard error of the mean.
Page 23 of 32
24 Fig. 7: Quantification of the Zn2+ released by ZnO QDs co-incubated 24h in the dialysis device with bacterial cells. Experiments were performed at three initial QDs concentrations: 3×10-3 (a), 3×10-4 (b) and 3×10-5 MZn eq. (c). Suspensions of cells in log phase (Log) or late
ip t
stationary phase (Stat.) were added at final OD600=0.3. Boiled cells were prepared from bacteria in late stationary phase. Each data point is the mean of at least three independent
cr
experiments and errors bars correspond to standard error of the mean. Black stars indicate values that are statistically different from control (QDs without cells) at p≤0.01. ND: not
Ac ce p
te
d
M
an
us
done.
Page 24 of 32
ip t
Graphical Abstract (for review)
cr
dilution effect Functional nanoparticle
Kd
us
Dissolved nanoparticle
an
ZnO QDs
stabilization
bacterium
Ac
ce
pt
ed
metaldependent toxicity
M
Zn2+
Page 25 of 32
Figure (revised)
us
cr
ip t
Figure 1
Zn2⁺ ZnO QDs
Zn2⁺
ed
Zn2⁺
Dialysate
M
Zn2⁺
an
Float-A-Lyzer G2 dialysis device (cut-off of 8-10 kD)
± bacteria
Biosensing ICP-OES
Ac
ce
pt
Retentate
Page 26 of 32
ip t
Figure 2
an
us
cr
(a)
0
40
2 (degree)
(200)
(103)
60
80
ce
20
pt
(102)
2000
(110)
ed
Intensity (a.u.)
(100) (002)
4000
M
(101)
(b)
(c)
1,0
Absorbance (a.u.)
PL Intensity (a.u.)
Ac
0,8
0,6
0,4
0,2
0,0
300
400
500
600
700
Wavelength (nm)
Page 27 of 32
cr
ip t
Figure 3
us
140 120
an
100 80 60 40 20 0 10-7
10-5
10-4
10-3
10-2
ed
10-6
M
Luminescence (% of control)
160
Ac
ce
pt
Concentration of ZnCl2 or ZnO QDs (MZn eq.)
Page 28 of 32
cr
50
us
40
40 20
30
0 210-5
410-5
an
0
20 10 0 10-8
10-7
10-6
M
Apparent bioavailable [Zn2+] (µM)
ip t
Figure 4
10-5
10-4
10-3
10-2
Ac
ce
pt
ed
QD concentration (MZn eq.)
Page 29 of 32
cr us
10-2
an
10-3 10-4
10-6 Retentate
M
10-5
Dialysate
ed
Apparent bioavailable [Zn2+] (M)
ip t
Figure 5
Dialysate
t = 24 h
Ac
ce
pt
t=0h
Retentate
Page 30 of 32
us
y = -62.2 Log(x) - 144.1 R2 = 0.996
100 80
an
60 40 20
M
Total Zn2+ released (%)
120
cr
ip t
Figure 6
0 10-5
10-4
10-3
10-2
Ac
ce
pt
ed
Initial QD concentration in dialysis device (MZneq)
Page 31 of 32
ip t
Figure 7
cr
25
15 10 5 ND
0 No cells
Boiled Stat.
CH34
60
20 0
No cells
ce Total Zn2+ released (%)
Ac
Log
Boiled Stat.
Boiled Stat.
*
*
Log
Log
MG1655
*
ed
40
ND
M
80
pt
Total Zn2+ released (%)
(b)
(c)
us
20
an
Total Zn2+ released (%)
(a)
Boiled Stat.
CH34
MG1655
ND
ND
Log
100 80 60 40 20
0 No cells
Boiled Stat.
CH34
Log
Boiled Stat.
Log
MG1655
Page 32 of 32
S0304-3894(14)00751-1 http://dx.doi.org/doi:10.1016/j.jhazmat.2014.09.017 HAZMAT 16261
To appear in:
Journal of Hazardous Materials
Received date: Revised date: Accepted date:
20-4-2014 28-8-2014 1-9-2014
Please cite this article as: X. Bellanger, P. Billard, R. Schneider, L. Balan, C. Merlin, Stability and toxicity of ZnO quantum dots: interplay between nanoparticles and bacteria, Journal of Hazardous Materials (2014), http://dx.doi.org/10.1016/j.jhazmat.2014.09.017 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
1 Highlights: Dilution of aminosilane-capped ZnO QDs dramatically increases their dissolution Bacteria limit Zn2+ leakage from ZnO QDs in a physiological-dependent process.
ip t
Implementation of biosensors for assessing free metal promotes QDs instability
Ac ce p
te
d
M
an
us
cr
Dialysis combined to ICP allow studying QDs stability without prior dilution
Page 1 of 32
2 Stability and toxicity of ZnO quantum dots: interplay between nanoparticles and bacteria.
ip t
Authors:
cr
Xavier Bellangera, Patrick Billardb, Raphaël Schneiderc, Lavinia Baland, Christophe Merlina *
Affiliation:
Université de Lorraine and CNRS, Laboratoire de Chimie Physique et Microbiologie pour
us
a
l’Environnement (LCPME), UMR 7564, 15 Avenue du Charmois, 54500 Vandœuvre-lès-
Université de Lorraine and CNRS, Laboratoire Interdisciplinaire des Environnements
M
b
an
Nancy, France.
Continentaux (LIEC), UMR 7360, Boulevard des Aiguillettes, Faculté des Sciences et
Université de Lorraine and CNRS, Laboratoire Réactions et Génie des Procédés (LRGP),
te
c
d
Techniques, BP 70239, 54506 Vandœuvre-lès-Nancy, France.
UMR 7274, 1 rue Grandville, BP 20451, 54001 Nancy Cedex, France. Institut de Science des Matériaux de Mulhouse (IS2M), UMR 7361, CNRS, 15 rue Jean
Ac ce p
d
Starcky, 68093 Mulhouse, France
E-mail addresses:
[email protected] [email protected]
[email protected] [email protected] [email protected] (* Correspondence and reprints)
Page 2 of 32
3 Abstract
The toxicity of quantum dots (QDs) has been commonly attributed to the release of metal ions
ip t
from the core as well as to the production of reactive oxygen species. However, the information related to the stability of the nanoparticles are relatively scarce although this
cr
parameter may strongly influence their toxicity. The stability of aminosilane-capped ZnO QDs, here used as model nanoparticles, was investigated by inductively coupled plasma-
us
optical emission spectrometer (ICP-OES) and whole cell biosensors using a dialysis setup to separate the QDs from the leaked Zn2+ ions. The integrity of the ZnO QDs appeared strongly
an
affected by their dilution in aqueous medium, whereas the nanoparticles were slightly
M
stabilized by bacteria. Our results demonstrate some inadequacy between the implementation
Ac ce p
Keywords
te
d
and use of whole cell biosensors, and the monitoring of metal release from QDs.
ZnO QDs stability, Zn2+bioavailability, QDs toxicity, Escherichia coli, Cupriavidus metallidurans
Page 3 of 32
4 1. Introduction
The great progresses made in nanotechnologies have led to an exponential increase of the use
ip t
of engineered nanoparticles in many domains of life, going from consumer products to essential components of new technologies [1,2]. The developments of nanotechnologies have
cr
been so fast that studies regarding their possible advert effects on health and the environment are lagging behind the speed of progress, making them relatively scarce if not inexistent. This
us
lack of hindsight has attracted increasing concerns regarding the fate of the nanomaterials in the environment and the toxicity specifically linked to their small size. If their nanoscale and
an
the great surface to volume ratio associated are the essence of their outstanding properties,
M
they are also critical in terms of toxicity as they increase the bioavailability of the nanoparticles. Therefore, processes controlling the behavior of nanoparticles with respect to
d
dissolution or aggregation should necessarily have an impact on their persistence and their
te
toxicity in the environment [3,4]. With this respect, any risk assessment associated to nanoparticles release in the environment depends on the environment itself, with all its
Ac ce p
complexity, as well as on the nature and on the chemistry of the nanoparticles considered. Over the past fifteen years, semiconductor nanocrystals (quantum dots, QDs) have received considerable attention because of their unique optical properties. Their narrow and tunable fluorescence emission associated to broad absorption spectra, high quantum yields and good resistance to chemical degradation and photobleaching, have made QDs interesting nanoparticles for various applications ranging from bioimaging to optoelectronics [5-7]. CdSe- and CdTe-core QDs have been extensively studied because of the quality and the flexibility of their optical properties. However, the use of Cd-containing QDs is constrained by their dual toxicity originating from their Cd2+ content and their ability to generate deleterious reactive oxygen species (ROS) [8-10]. Both types of toxicity are controlled by the
Page 4 of 32
5 stability of the particle and have deep consequences on the QDs application outputs in terms of biological alteration whether it concerns purposely exposure, in the context of bioimaging for instance, or accidental environmental exposure, thus raising the problem of waste
ip t
management. Recently, great efforts have been made to reduce the toxicity of QDs with alternative semiconductors composed of less toxic metals, such as zinc oxide, while keeping
cr
interesting optical properties thanks to elaborated particle synthesis chemistry [11]. Still, the toxicity of the particles remains deeply associated to their state and therefore their stability
us
with respect to metal release (dissolution), reactivity (ROS production), or bioavailability of the particle (nanoscale versus aggregated state). The integrity of nanoparticles is relatively
an
easy to assess in vitro using conventional methods, such as filtration followed by inductively
M
coupled plasma (ICP) analyses that allow the quantification of the dissolved metals. Metal leakage can also be assessed using biological-based methods with whole cell biosensors for
d
instance. Such biosensors consist of microorganisms that have been genetically modified to
te
produce a quantifiable signal (e.g. fluorescence, bioluminescence) in response to specific environmental stimuli including metals [12,13]. Moreover, whole cell biosensors provide the
Ac ce p
specific advantage of measuring the bioavailable fraction of the analyte of interest as opposed to the total content as determined by chemical means. Biosensors were used on several occasions to quantify metal release from nanoparticles [14], including QDs [10,15], and therefore allow monitoring their stability with respect to dissolution. Additionally, specific whole cell biosensors allow evaluating more subtle particle-induced stresses such as oxidative stress linked to ROS production [10,16]. Biosensors are highly sensitive, selective, easy to implement, and only require the cells to reach a particular physiological state and to be exposed to serial dilutions of the sample to be analyzed. However, caution must be taken when evaluating nanoparticle dissolution using biosensor-based assays as the impact of the biosensor cells itself on the stability of the studied material has to be considered. To the best
Page 5 of 32
6 of our knowledge, the influence of either biosensor cells or the implementation of such biosensors on nanoparticles has not been documented yet. This study aimed to evaluate the influence of the Cupriavidus metallidurans and Escherichia
ip t
coli sensor strains on the stability of ZnO core QDs functionalized with 3aminopropyltrimethoxysilane (APTMS). ZnO@APTMS QDs were here chosen as model
cr
particles as the APTMS functionalization allows the ZnO core to become dispersible in
aqueous biological environment and, in the meantime, provide a good colloidal stability to the
us
nanoparticles as we reported previously [15]. We report an original approach, combining dialysis, ICP analyses and biosensing, for determining the relative contributions of the
an
bacterial cells and the experimental set up on the stability of the QDs. Zn2+ release, which is
M
the main source of ZnO QDs induced toxicity in the present study, appeared to be influenced
te
2. Materials and methods
d
by the presence of bacteria, their physiological state and the species considered.
Ac ce p
2.1 Synthesis and characterization of ZnO quantum dots
Triethyleneglycol-capped ZnO QDs were prepared by a sol-gel route using LiOH as base [17]. A LiOH/Zn(OAc)2 molar ratio of 2 was used for the synthesis. A first ligand exchange with oleic acid was undertaken to transfer the dots in weakly polar organic solvents. ZnO@oleate QDs were next transferred in aqueous solution by treatment with APTMS in DMF. In a typical experiment, under a argon atmosphere, 7.6 µL of APTMS were added to 50 mg of ZnO QDs dispersed in 7 mL of anhydrous DMF. The mixture was stirred 5 min at room temperature and next heated at 120°C for 15 min. After cooling and centrifugation, the
Page 6 of 32
7 white precipitate was washed with DMF (2 x 5 mL) and ethanol (2 x 5 mL) and dried at room temperature. The pellet was readily re-dispersed in aqueous solution for further study. The Zn content of QDs was measured by inductively coupled plasma optical emission
ip t
spectrometer (ICP-OES) (Varian) as being of 4.05 mM of Zn/g of QDs, i.e. Zn accounts for 26.5% of the QD mass. ZnO QD concentrations were expressed in molar Zn equivalent (MZn
cr
eq.)
us
2.2 Bacterial strains
an
Two wild-type bacterial strains, Escherichia coli MG1655 [18] and Cupriavidus
M
metallidurans CH34 [19] were used in this study to assess the stability and toxicity of ZnO QDs. Both strains were transformed according to Choi et al. [20] with the luxCDABE operon
d
bearing plasmid pUCD607 [21], which confers a constitutive luminescent phenotype and
te
allows evaluating the nanoparticle toxicity based on bioluminescence extinction. MG1655(pUCD607) and CH34(pUCD607) transformants were maintained on LB agar
Ac ce p
medium (LB Broth Miller, DifcoTM) supplemented with kanamycin at 50 and 500 µg/mL, respectively. Bioavailability of Zn2+ was measured using the C. metallidurans biosensor strain AE1433 deriving from CH34 [22].
2.3 QD suspensions and ZnCl2 solutions
ZnCl2 was used to get standard solutions of known Zn2+ ion concentrations and was chosen over other zinc mineral salts for its very good solubility in water. Solutions of ZnCl2 and suspensions of QDs were prepared in RM mineral medium, a MOPS-based mineral culture medium previously designed to minimize metal chelation and precipitation [22]. The stock
Page 7 of 32
8 suspensions of QDs were prepared at 3.10-3 M and sonicated at 130 W for 1 min to be dispersed in the medium. QDs suspensions were vortexed prior each utilization. Freshly made
ip t
ZnCl2 solutions and QDs suspensions were prepared before each replicate experiment.
cr
2.4 ZnCl2 and QD toxicity assays
A single colony of either E. coli MG1655(pUCD607) or C. metallidurans CH34(pUCD607)
us
was picked from a LB selective plate and used to inoculate 10 mL of LB broth containing kanamycin in a 100 mL conical flask. The culture was grown at 30°C with continuous
an
shaking at 160 rpm for 16 to 18h. Bacterial cells were harvested by centrifugation (3,000 rpm,
M
3 min), washed with 10 mL of RM supplemented with SL7 trace elements solution [23] and 0.2 % gluconate as a carbon source [22], and then centrifuged again in the same conditions.
d
Cell pellets were re-suspended in 10 mL of RM - SL7 gluconate 0.2 % and allowed to adapt
te
in the medium at 30°C with continuous shaking (160 rpm) for one hour. The OD600 of the cell suspensions were then adjusted to 0.3 for E. coli MG1655(pUCD607) and to 0.54 for
Ac ce p
C. metallidurans CH34(pUCD607) in the same medium, and dispensed (180 µL for E. coli and 100 µL for C. metallidurans) in clear bottom black microtiter plates (Isoplate-96F, PerkinElmer), previously filled with various dilutions of either ZnCl2 or QDs to reach a 200 µL final volume. Microplates were maintained at 25°C in a VICTOR3 Multilabel Counter plate reader (Perkin-Elmer), where light emission and OD600 were recorded every 3 min, with a 5 s and 1 s measurement time, respectively, preceded by a 10 s double orbital shaking. For both parameters, background provided by RM-SL7 gluconate medium was subtracted, and the luminescence was normalized to the corresponding OD600. Normalized luminescence values were averaged among quadruplicate and induction factors were calculated by dividing the normalized luminescence values by those obtained for the untreated control. The maximal No
Page 8 of 32
9 Observed Effect Concentration (NOEC) and the half maximal Inhibitory Concentration (IC50) were determined 120 minutes after exposure.
ip t
2.5 Quantification of bioavailable Zn2+
cr
Bioavailability of Zn2+ was quantified using the whole cell biosensor Cupriavidus
metallidurans AE1433 that has been genetically engineered to emit a bioluminescence signal
us
when sensing bioavailable Zn2+ [22]. The protocol used is slightly modified from Corbisier et al [22]. Briefly, strain AE1433 was cultured in LB medium until stationary phase (OD600 =
an
1.5) and cells were washed twice by alternating centrifugation (10,000 g for 2 min at 20°C)
M
and cell pellet dispersion in RM medium supplemented with gluconate 0.01 %. The cell density was finally adjusted to 0.54 and 100 µl of the suspension was distributed in black
d
microtiter plates with clear bottom, previously filled with 100 µl of various dilutions of either
te
ZnCl2, for calibration curves, or QDs, for testing. Bioluminescence and OD600 were recorded as described above at 25°C in a VICTOR3 Multilabel Counter plate reader (Perkin-Elmer)
Ac ce p
and each test was carried out in 3 independent replicates. For each series of experiments, luminescence induction factors were plotted against ZnCl2 concentrations to set a standard curve from which linear regression could be obtained for the non-toxic part of the curve, and were used to deduce the apparent concentrations of bioavailable Zn2+ leaking from QDs.
2.6 QD dialysis in the presence or absence of bacterial cells
One milliliter of QD suspension was introduced in a 1 mL ready-to-use dialysis device with a molecular weight cut-off of 8-10 kD (Float-A-Lyzer G2; Spectra/Por) previously conditioned as indicated by the manufacturer. QD suspensions were dialyzed for 24h, in the dark, under
Page 9 of 32
10 agitation (160 rpm), and at 30°C against 24 mL of RM gluconate 0.01% medium in sterile capped tubes. Dialysates and retentates were further analyzed for their Zn content by ICPOES or using the AE1433 biosensor (Fig. 1).
ip t
When testing the effect of cells on QD stability, bacterial suspensions in different physiological states were added together with the nanoparticle in the ready-to-use dialysis
cr
device. Briefly, strains E. coli MG1655 or C. metallidurans CH34 were grown at 30°C under agitation (160 rpm) in conical flasks filled with 1/5th of LB medium. Cultures were either
us
stopped at OD600 = 0.3 to obtain exponentially growing cells, or were incubated for 16 to 18 h to obtain cells in late stationary phase of growth. Cells were then washed twice with RM
an
gluconate 0.01 % and the concentration of the cell suspension was finally adjusted to OD600 =
M
30. Ten microliters of concentrated cell suspension were added to the QD suspension within the Float-A-Lyzer G2 to obtain 1 mL of QD suspension with cells at final OD600 = 0.3. For
d
one tests series, the influence of dead material was evaluated with stationary phase cells boiled for 10 minutes before being added to QD suspensions in the dialysis device. For each
te
condition, the amount of Zn2+ released from the QDs was deduced from ICP-OES
Ac ce p
measurement in the dialysate.
2.7 Statistics
The IC50 was calculated by solving the fitted equation for relative luminescence equal to 50% by using the online curve fitting web site ZunZun.com (http://zunzun.com/Equation/2/Sigmoidal/Boltzmann Sigmoid A/). Statistical analysis was performed using the web tool BiostaTGV (http://marne.u707.jussieu.fr/biostatgv/) and as described by Cumming et al [24]. All websites were last accessed on 27 February 2014.
Page 10 of 32
11 3. Results
ip t
3.1 ZnO QD toxicity towards E. coli and C. metallidurans
With the rapid development of QD-based technologies, an increasing attention is given to a
cr
better control of the toxicity associated to these metal-containing particles. If the toxicity of QDs is now recognized to involve both the liberation of metal ions from the core and the
us
production of ROS, the relative contribution of the two phenomena can vary from one particle to another, and both are impacted by the particle stability. This work presents an integrated
an
study regarding the toxicity and the stability of ZnO@APTMS QDs, used as model
M
nanoparticles. These ZnO QDs were synthesized using a sol-gel route and were easily dispersed in aqueous solution after surface functionalization with APTMS [15]. Transmission
d
electron microscopy and X-ray diffraction experiments showed that ZnO QDs have an
te
average diameter of ca. 4 nm and a cubic zinc blende structure, respectively (Fig. 2a and 2b). The photoluminescence spectrum of colloidal ZnO@APTMS QDs shows a broad emission
Ac ce p
centered at 530 nm, which is typical for ZnO QDs prepared via a sol-gel process (Fig. 2c). The electrical potential of these dots, referred to as the zeta potential, was measured to be +27.4 ± 4 mV at pH = 7.0.
The toxicity of APTMS-coated ZnO QDs towards E. coli and C. metallidurans was determined using two bacterial strains that have been genetically modified to emit light constitutively, namely E. coli MG1655(pUCD607) and C. metallidurans CH34(pUCD607). Any toxicity interfering with the bacterial metabolism will result in the reduction of the emitted bioluminescence. For both strains, bioluminescence extinction curves are presented in Fig. 3. It can be seen that ZnO QDs and ZnCl2 were more toxic towards E. coli MG1655 (IC50 = 1.610-5 M and 2.710-5 M for ZnCl2 and ZnO QDs, respectively) than towards
Page 11 of 32
12 C. metallidurans CH34 (IC50 = 2.810-4 M and 1.210-3 M for ZnCl2 and ZnO QDs, respectively). Strain CH34 is well-known for its resistance to several metals cations including Zn2+ [19], which may account for such strain-specific differences in metal/QDs sensitivity.
ip t
For both strains, the ZnO QDs appeared less toxic than their Zn2+ content. Considering that all the toxicity experiments were carried out in the dark, photo-induced ROS should not be
cr
considered, therefore ruling out the involvement of the QDs photoreactivity in the observed extinction of bioluminescence. Thus, the toxicity observed should mainly result from the
us
liberation of Zn2+ cations from the ZnO QD cores. Interestingly, for E. coli the toxicity curves
an
for ZnCl2 and ZnO QDs showed similar shapes and were quite close to each other as seen by the two IC50 values of the same range (1.610-5 M and 2.710-5 M, respectively), thus tending
M
to show that ZnO QDs were almost, if not completely, dissolved. In contrast, for C. metallidurans, the two toxicity curves appeared well separated, with almost one order of
d
magnitude between the two IC50 values (2.810-4 M and 1.210-3 M for ZnCl2 and ZnO QDs,
te
respectively), suggesting that the QDs only partially dissolved. Although at this stage it is not yet possible to explain the differences observed between the ZnCl2 and the ZnO QDs toxicity
Ac ce p
curves, it should be noted that (i) the different concentrations at which the QDs toxicity was observed, from one bacteria to the other, may affect the dissolution equilibrium of the particle, and (ii) that the two bacterial strains may possibly exert different effects on the QDs stability.
3.2 Determining bioavailable Zn2+ leaked from ZnO QDs
The concentration-dependent dissolution of ZnO QDs was first investigated using a wholecell Zn2+ biosensor. Strain AE1433 of C. metallidurans, emitting light when sensing Zn2+, was employed to determine the amount of bioavailable Zn2+ ions released from freshly
Page 12 of 32
13 prepared dispersions of ZnO QDs. Dilutions of QDs ranging from 710-6 to 210-3 MZn eq. were exposed to C. metallidurans AE1433 whose luminescence was compared to that obtained with a ZnCl2 standard. For QDs concentrations ranging from 710-6 to 310-5 MZn the apparent concentrations of bioavailable Zn2+ fitted the Zn2+ content of the QDs, thus
ip t
eq.,
indicating a complete dissolution of the particle (Fig. 4), which is in agreement with what was
cr
suspected while studying the QDs toxicity on E. coli (Fig. 3). At higher QD concentrations,
the apparent concentrations of bioavailable Zn2+ reached a maximum at around 40 µM, which
an
proportional anymore due to metal-induced toxicity.
us
coincides with the Zn2+ concentration limit over which the response of the biosensor is not
In order to study Zn2+ leaking at higher QDs concentrations and, in the mean time, to take into
M
consideration a possible contact-effect between the biosensor cells and the nanoparticles, a dialysis-based approach was setup. In a first attempt, 1 mL of a 310-3 M Zn eq QDs
d
suspension was dialyzed for 24 hours against 24 mL of RM gluconate 0.01 % medium. Strain
te
AE1433 was used to estimate and compare the apparent concentration of bioavailable Zn2+ ions in both dialysis compartments (the retentate and the dialysate) at two different times (t =
Ac ce p
0 and t = 24 h). Experimentally, the content of both compartments was sampled, serially diluted and assayed with the biosensor as described above. A set of dilutions producing a biosensor response within the range of usable concentrations (i.e. linear relationship between light emission and Zn2+ concentration) was selected and used to determine the bioavailable Zn2+ in each compartment. Fig. 5 shows that the amounts of bioavailable Zn2+ measured did not significantly differ at t = 0 and t = 24 h. In both cases, the apparent concentrations of bioavailable Zn2+ in the retentates appeared about a thousand times more concentrated than those of the dialysates. If such a difference could be expected before equilibrium (at t = 0), the concentrations of bioavailable Zn2+ should equilibrate upon dialysis, unless the bioavailable Zn2+ ions are released from the QDs after the dialysis process, during the exposure to the
Page 13 of 32
14 biosensor cells. Therefore, it can be assumed that ZnO QDs are relatively stable over time at elevated concentrations but further dilution and/or incubation with the biosensor are likely to be responsible for the liberation of bioavailable Zn2+ ions detected in biosensing assays.
ip t
Indeed, bacteria have a variety of properties that can affect metal solubility [25] and initially insoluble forms of heavy metals salts may become bioavailable due to the direct contact with
cr
bacterial biosensor [26,27]. The present experiments show that the procedure associated to the
an
experiment in terms of Zn2+ leakage quantification.
us
use of a whole cell biosensor affect the state of the particle, and therefore the outcome of the
M
3.3 Stability of ZnO QD upon dilution
In order to dissociate the concentration effect (dissolution equilibrium) from the cell exposure effect (biodegradation for instance) regarding the fate of ZnO QDs, the QDs stability was
te
d
further studied using the same dialysis setting but with direct quantification of the Zn2+ content of each compartment without prior dilution. Series of 1 mL of QDs suspensions at
Ac ce p
concentrations ranging from 310-5 to 310-3 M Zn eq. were dialyzed as previously for 24 hours against 24 mL of RM gluconate 0.01 % medium. Both compartments were analyzed by ICPOES and the Zn2+ content in the dialysates was used to estimate the amount of free Zn2+ ions having leaked from the QDs. At the highest QDs concentration tested (310-3 M Zn eq), the concentration of free Zn2+ was estimated to be 1.610-5 M accounting for 12% of the initial QDs content (Fig. 6). In contrast, at low QD concentration, close to 10-4 M Zn eq., the amount of Zn2+ leaking from the QDs increased to 100%. Therefore, the QDs stability appears to vary according to its initial concentration within the dialysis device, which likely reflects the QDs dissociation equilibrium in aqueous solution. Furthermore, it should be noted that for an initial QDs concentration of 2.710-5 M Zn eq., corresponding to the IC50 observed for E. coli
Page 14 of 32
15 (Fig. 3), 100% of the particles are dissolved, thus explaining the closeness of the QDs and ZnCl2 toxicity curves. Finally, at 1.210-3 M Zn eq., the IC50 obtained for C. metallidurans, ca. 40% of the Zn2+ ions have leaked from the QDs, which agrees well with the apparent lower
ip t
toxicity of particles compared to their Zn content for this bacterium.
cr
3.3 Impact of the bacteria on the stability of ZnO QDs
us
Having demonstrated the influence of QDs dilution on their dissolution, QDs stability was
an
further investigated to evaluate the influence of the bacterial cells. For this purpose, the nanoparticles were co-incubated with bacterial cells in the dialysis device at three significant
M
ZnO QDs concentrations: 310-3, 310-4 and 310-5 M Zn eq., presenting 12%, 70%, and 100% QDs dissolution in terms of Zn2+ release, respectively (Fig. 6). The QDs were independently
d
incubated with either E. coli MG1655 or C. metallidurans CH34, from cultures in different
te
metabolic states (exponential or stationary growth phase), or even to dead cells resulting from boiled cultures. The release of Zn2+ from QDs was quantified by ICP-OES measurements of
Ac ce p
the dialysates and the retentates after 24 hours. The addition of bacteria to QDs at the two extreme concentrations (310-3 and 310-5 MZn eq.) did not significantly modify the behaviors of the nanoparticles upon dialysis regardless the state of the cells (Fig. 7a and 7c). Nevertheless, for a QDs concentration of 310-4 M Zn eq., at which the nanoparticles displayed an intermediate dissolution state, the presence of bacteria significantly reduced the release of free Zn2+ in some instances (Fig. 7b). For both strains, dead cells have statistically no effect on QDs stability. In contrast, metabolically active bacteria added in exponential growth phase significantly reduced the release of free Zn2+ from 70% to 20% and 30% of the QDs Zn content for C. metallidurans and E. coli, respectively. An intermediate effect could be observed for C. metallidurans added in stationary phase of growth. Because the release of free
Page 15 of 32
16 Zn2+ is quantified on the basis of the amount of Zn2+ able to cross the dialysis membrane, a reduction of free Zn2+ in the dialysate could also originate from the Zn2+ sequestration by the bacterial cells in the retentate compartment. To examine this possibility, the dialysis
ip t
experiment was run with various concentrations of ZnCl2 co-incubated or not with bacterial cells. In each case, Zn2+ concentrations determined by ICP-OES in the dialysates after 24
cr
hours of dialysis did not differ whether or not bacterial cells were present, and whatever their physiological state (data not shown), therefore ruling out the possible retention of Zn2+ by
us
bacterial cells. Thus, against all odd, bacteria seem to be able to limit the Zn2+ leakage from
an
ZnO QDs in a way that appeared to be both strain- and physiological state-dependent.
M
4. Discussion and conclusion
d
This work demonstrates that both the dilution of ZnO@APTMS QDs and the presence of bacterial cells interfere in opposite directions with the particles stability regarding the
te
liberation of Zn2+ ions. Toxicity studies with QDs often oppose the liberation of metal to the
Ac ce p
ROS production and scarcely include the problem posed by the nanoparticle stability. However, this stability is of prime importance to dictate QDs toxicity either towards the ROS production or towards the liberation of metal. Studying QDs toxicity often implies exposing the biological material to dilution series of the nanoparticles in order to discover critical concentrations affecting biological functions. The dilution, by promoting the particles dissolution, is likely to orientate the QDs toxicity solely towards their metal content, thus biasing our perception of the QDs toxicity linked to the reactivity of the particles. This observation also stands for the use whole-cell metal biosensors. Because these biosensors can only operate within a define range of concentrations, from the detection limit to toxic concentrations, they often require working on dilution series of the samples. If the sample
Page 16 of 32
17 stability is sensitive to dilution in terms of metal release, biosensing is likely to overestimate the bioavailability of metals from unstable samples. This points out that great care should be given to preliminary study regarding the stability of nanoparticles such as QDs while studying
ip t
their toxicity in relation to metal release. In this context, we present an original dialysis-based approach to study the stability of QDs, which allows maintaining the nanoparticles in various
cr
conditions, including in the presence of bacterial cells, while analyzing the metal released in
another compartment. With this approach, we can also expect to limit biases associated with
us
the centrifugation procedure commonly used to measure metal ion release in nanotoxicology [28].
an
We demonstrated that, within the limit of the experiments presented, bacteria reduced the
M
metal release from ZnO@APTMS QDs. Cultures of Pseudomonas aeruginosa were also reported to stabilize and limit the dissolution CdSe QDs. However, the mechanism by which
d
such protection occurred was not investigated further [29]. We also showed that the
te
physiological state of these bacteria could modulate the stability of the dots. The simple fact that dead bacteria do not significantly alter QDs stability, and that cells in stationary phase
Ac ce p
stabilize less than cells in log phase, suggests that the stabilization is an active process rather than being solely linked to a structural component of the bacteria. With this respect, the membrane potential has been shown to vary according to the physiological state [30]. On the other hand, QDs have been shown to interfere with the bacterial membrane potential [31], which raises the question of QDs stability in a biologically generated electron flux.
Acknowledgments
This work is supported by the Agence Nationale pour la Recherche (ANR CESA 2011, project NanoZnOTox) and the Zone Atelier Moselle. We thank Hervé Marmier (LIEC,
Page 17 of 32
18 Université de Lorraine) for ICP-OES analyses and are grateful to Hélène Guilloteau (LCPME, Université de Lorraine) for technical support and to Boussad Arroua for his contribution to the toxicity experiments. We also thank Ghouti Medjahdi (IJL, UMR CNRS 7198, Université
ip t
de Lorraine) for XRD analyses.
cr
References
us
[1] J.R. Peralta-Videa, L. Zhao, M.L. Lopez-Moreno, G. de la Rosa, J. Hong, J.L.GardeaTorresdey, Nanomaterials and the environment: a review for the biennium 2008-2010, J.
an
Hazard. Mater. 186 (2011) 1-15.
M
[2] M.C. Roco, M.C. Mirkin, C.A. Hersam, Nanotechnology research directions for societal needs in 2020, Springer, Netherlands, 2011.
d
[3] S.J. Klaine, P.J. Alvarez, G.E. Batley, T.F. Fernandes, R.D. Handy, D.Y. Lyon, S.
te
Mahendra, M.J. McLaughlin, J.R. Lead, Nanomaterials in the environment: behavior, fate,
Ac ce p
bioavailability, and effects, Environ. Toxicol. Chem. 27 (2008) 1825-1851. [4] G.E. Batley, J.K. Kirby, M.J. McLaughlin, Fate and risks of nanomaterials in aquatic and terrestrial environments, Acc. Chem. Res. 46 (2013) 854-862. [5] T. Jamieson, R. Bakhshi, D. Petrova, R. Pocock, M. Imani, A.M. Seifalian, Biological applications of quantum dots, Biomaterials 28 (2007) 4717-4732. [6] D. Bera, L. Qian, T.K. Tseng, P.H. Holloway, Quantum dots and their multimodal applications: a review, Materials 3 (2010) 2260-2345. [7] P. V. Kamat, Quantum dot solar cells, The next big thing in photovoltaics, J. Phys. Chem. Lett. 4 (2013) 908-918.
Page 18 of 32
19 [8] J.M. Tsay, X. Michalet, New light on quantum dot cytotoxicity, Chem. Biol.12 (2005) 1159-1161. [9] R. Hardman, A toxicologic review of quantum dots: toxicity depends on physicochemical
F.A. Kauffer, C. Merlin, L. Balan, R. Schneider, Incidence of the core composition on
cr
[10]
ip t
and environmental factors. Environ. Health Perspect. 114 (2006) 165-172.
the stability, the ROS production and the toxicity of CdSe quantum dots, J. Hazard. Mater.
T. Pons, E. Pic, N. Lequeux, E. Cassette, L. Bezdetnaya, F. Guillemin, F. Marchal, B.
an
[11]
us
268 (2014) 246-255.
Dubertret, Cadmium-free CuInS2/ZnS quantum dots for sentinel lymph node imaging with
[12]
M
reduced toxicity, ACS Nano 4 (2010) 2531-2538.
S. Daunert, G. Barrett, J.S. Feliciano, R. Shetty, S. Shrestha, W. Smith-Spencer,
d
Genetically engineered whole-cell sensing systems: coupling biological recognition with
S. Köhler, S. Belkin, R.D. Schmid, Reporter gene bioassays in environmental analysis,
Ac ce p
[13]
te
reporter genes, Chem. Rev. 100 (2000) 2705-2738.
Fresenius J. Anal. Chem. 366 (2000) 769 – 779. [14]
A. Kahru, H.C. Dubourguier, I. Blinova, A. Ivask, K. Kasemets, Biotests and
biosensors for ecotoxicology of metal oxide nanoparticles: a minireview, Sensors 8 (2008) 5153-5170. [15]
A. Aboulaich, C.M. Tilmaciu, C. Merlin, C. Mercier, H. Guilloteau, G. Medjahdi, R.
Schneider, Physicochemical properties and cellular toxicity of (poly) aminoalkoxysilanesfunctionalized ZnO quantum dots, Nanotechnology 23 (2012) 335101.
Page 19 of 32
20 [16]
A. Ivask, O. Bondarenko, N. Jepihhina, A. Kahru, Profiling of the reactive oxygen
species-related ecotoxicity of CuO, ZnO, TiO2, silver and fullerene nanoparticles using a set of recombinant luminescent Escherichia coli strains: differentiating the impact of particles
[17]
ip t
and solubilised metals, Anal. Bioanal. Chem. 398 (2010) 701-716. H.M. Xiong, R.Z. Ma, S.F. Wang, Y.Y. Xia, Photoluminescent ZnO nanoparticles
cr
synthesized at the interface between air and triethylene glycol, J. Mater. Chem. 21 (2011)
[18]
us
3178-3182.
F.R. Blattner, G. Plunkett, C.A. Bloch, N.T. Perna, V. Burland, M. Riley, J. Collado-
an
Vides, J.D. Glasner, C.K. Rode, G.F. Mayhew, J. Gregor, N.W. Davis, H.A. Kirkpatrick, M.A. Goeden, D.J. Rose, B. Mau,Y. Shao, The complete genome sequence of Escherichia
M. Mergeay, D. Nies, H.G. Schlegel, J.Gerits, P. Charles, F. Van Gijsegem,
d
[19]
M
coli K-12, Science 277 (1997) 1453-1462.
te
Alcaligenes eutrophus CH34 is a facultative chemolithotroph with plasmid-bound resistance
Ac ce p
to heavy metals, J. Bacteriol. 162 (1985) 328-334. [20]
K.H. Choi, A. Kumar, H.P. Schweizer, A 10-min method for preparation of highly
electrocompetent Pseudomonas aeruginosa cells: application for DNA fragment transfer between chromosomes and plasmid transformation, J. Microbiol. Meth. 64 (2006) 391-397. [21]
J.J. Shaw, C.I. Kado, Development of a Vibrio bioluminescence gene-set to monitor
phytopathogenic bacteria during the ongoing disease process in a nondisruptive manner, Nat. Biotechnol. 4 (1986) 560-564. [22]
P. Corbisier, D. van der Lelie, B. Borremans, A. Provoost, V. de Lorenzo, N.L.
Brown, J.R. Loyd, J.L. Hobamn, E. Csöregi, G. Johansson, B. Mattiasson,Whole cell- and
Page 20 of 32
21 protein-based biosensors for the detection of bioavailable heavy metals in environmental samples, Anal. Chim. Acta 387 (1999) 235–244. [23]
H. Biebl, N. Pfennig, Isolation of members of the family Rhodospirillaceae, in M. P.
ip t
Starr, J. Stolp, H. G. Truper, A. Balows, and H. G. Schlegel (Eds.), The prokaryotes, a handbook on habitats, isolation and identification of bacteria, vol. 1. Springer-Verlag, Berlin,
G. Cumming, F. Fidler, D.L. Vaux, Error bars in experimental biology, J. Cell. Biol.
us
[24]
cr
1981, pp. 267-273.
[25]
an
177 (2007) 7-11.
G.M. Gadd, Metals, minerals and microbes: geomicrobiology and bioremediation,
[26]
M
Microbiology 156 (2010) 609-643.
A. Kahru, A. Ivask, K. Kasemets, L. Põllumaa, I. Kurvet, M. Francois, H.C.
d
Dubourguier, Biotests and biosensors in ecotoxicological risk assessment of field soils
O. Bondarenko, T. Rõlova, A. Kahru, A. Ivask, Bioavailability of Cd, Zn and Hg in
Ac ce p
[27]
te
polluted with zinc, lead and cadmium, Environ. Toxicol. Chem. 24 (2005) 2973-2982.
soil to nine recombinant luminescent metal sensor bacteria, Sensors 8 (2008) 6899-6923. [28]
M. Xu, J. Li, N. Hanagata, H. Su, H. Chen, D. Fujita, Challenge to assess the toxic
contribution of metal cation released from nanomaterials for nanotoxicology-the case of ZnO nanoparticles, Nanoscale 5 (2013) 4763-4769. [29]
J.H. Priester, P.K. Stoimenov, R.E. Mielke, S.M. Webb, C. Ehrhardt, J.P. Zhang, G.D.
Stucky, P.A. Holden, Effects of soluble cadmium salts versus CdSe quantum dots on the growth of planktonic Pseudomonas aeruginosa, Environ. Sci. Technol. 43 (2009) 2589-2594.
Page 21 of 32
22 [30]
P. Monfort, B. Baleux, Cell cycle characteristics and changes in membrane potential
during growth of Escherichia coli as determined by a cyanine fluorescent dye and flow cytometry, J. Microbiol. Meth. 25 (1996) 79-86. Y. Yang, H. Zhu, V.L. Colvin, P.J. Alvarez, Cellular and transcriptional response of
ip t
[31]
Pseudomonas stutzeri to quantum dots under aerobic and denitrifying conditions, Environ.
us
cr
Sci. Technol. 45 (2011) 4988-4994.
an
Figure captions
M
Fig. 1: Experimental setup for the quantification of Zn2+ released from ZnO QDs. QDs were incubated in the dialysis device in presence or absence of bacterial cells. The relative
d
amount of Zn2+ released was calculated from ICP-OES or biosensing measurement made in
te
the dialysate. Total Zn engaged in the experiment was obtained by combining ICP
Ac ce p
measurement in the retentate plus the dialysate.
Fig. 2: (a) TEM micrograph, (b) XRD pattern and (c) absorption (black) and photoluminescence (red) spectra of ZnO@APTMS QDs. Absorption and photoluminescence spectra were recorded in water (excitation wavelength of 330 nm).
Fig. 3: Effect ZnCl2 or ZnO QD concentrations on the relative luminescence emitted by E. coli MG1655(pUCD607) and C. metallidurans CH34(pUCD607). Squares and diamonds represent experiments carried out with E. coli and C. metallidurans, respectively. Continuous and dotted lines represent the experiments performed with ZnCl2 and ZnO QDs, respectively.
Page 22 of 32
23 Each data point is the mean of three independent experiments (with four repeats for each) and errors bars correspond to standard error of the mean.
ip t
Fig. 4: AE1433 biosensing of apparent bioavailable Zn2+ released by the ZnO QDs. The inset presents the results obtained at low QD concentrations (from 6.7×10-7 to 6.7×10-5 MZn
for which a linear fit could be obtained (equation: y = 0.99x; coefficient of determination
cr
eq.)
R2 = 0.996). Each data point is the mean of three independent experiments (with four repeats
us
for each) and errors bars correspond to standard error of the mean.
an
Fig. 5: AE1433 biosensing of apparent bioavailable Zn2+ released by the ZnO QDs
M
before and after a 24 hours dialysis. One mL of QD suspension at a concentration of 3×10-3 MZn eq. was dialyzed against 24 mL of RM gluconate 0.01% medium. The dotted line indicates
d
the theoretical initial QD concentration within the dialysis device at t=0. Each data point is
te
the mean of at four replicates and errors bars correspond to standard error of the mean.
Ac ce p
Fig. 6: Quantification of the Zn2+ released as a function of the initial ZnO QD concentration after 24h of dialysis. One mL of QD suspension was dialyzed against 24 mL of RM gluconate 0.01% medium. Total amounts of Zn2+ released were extrapolated from ICP-OES quantification of the dialysate. The dotted line indicates a log fit between the initial QD concentration (from 2×10-4 to 3×10-3 MZn eq.) and the relative amount of Zn2+ released (the equations and coefficients of determination corresponding to the trend lines are indicated). Each data point is the mean of at least three independent experiments and errors bars correspond to standard error of the mean.
Page 23 of 32
24 Fig. 7: Quantification of the Zn2+ released by ZnO QDs co-incubated 24h in the dialysis device with bacterial cells. Experiments were performed at three initial QDs concentrations: 3×10-3 (a), 3×10-4 (b) and 3×10-5 MZn eq. (c). Suspensions of cells in log phase (Log) or late
ip t
stationary phase (Stat.) were added at final OD600=0.3. Boiled cells were prepared from bacteria in late stationary phase. Each data point is the mean of at least three independent
cr
experiments and errors bars correspond to standard error of the mean. Black stars indicate values that are statistically different from control (QDs without cells) at p≤0.01. ND: not
Ac ce p
te
d
M
an
us
done.
Page 24 of 32
ip t
Graphical Abstract (for review)
cr
dilution effect Functional nanoparticle
Kd
us
Dissolved nanoparticle
an
ZnO QDs
stabilization
bacterium
Ac
ce
pt
ed
metaldependent toxicity
M
Zn2+
Page 25 of 32
Figure (revised)
us
cr
ip t
Figure 1
Zn2⁺ ZnO QDs
Zn2⁺
ed
Zn2⁺
Dialysate
M
Zn2⁺
an
Float-A-Lyzer G2 dialysis device (cut-off of 8-10 kD)
± bacteria
Biosensing ICP-OES
Ac
ce
pt
Retentate
Page 26 of 32
ip t
Figure 2
an
us
cr
(a)
0
40
2 (degree)
(200)
(103)
60
80
ce
20
pt
(102)
2000
(110)
ed
Intensity (a.u.)
(100) (002)
4000
M
(101)
(b)
(c)
1,0
Absorbance (a.u.)
PL Intensity (a.u.)
Ac
0,8
0,6
0,4
0,2
0,0
300
400
500
600
700
Wavelength (nm)
Page 27 of 32
cr
ip t
Figure 3
us
140 120
an
100 80 60 40 20 0 10-7
10-5
10-4
10-3
10-2
ed
10-6
M
Luminescence (% of control)
160
Ac
ce
pt
Concentration of ZnCl2 or ZnO QDs (MZn eq.)
Page 28 of 32
cr
50
us
40
40 20
30
0 210-5
410-5
an
0
20 10 0 10-8
10-7
10-6
M
Apparent bioavailable [Zn2+] (µM)
ip t
Figure 4
10-5
10-4
10-3
10-2
Ac
ce
pt
ed
QD concentration (MZn eq.)
Page 29 of 32
cr us
10-2
an
10-3 10-4
10-6 Retentate
M
10-5
Dialysate
ed
Apparent bioavailable [Zn2+] (M)
ip t
Figure 5
Dialysate
t = 24 h
Ac
ce
pt
t=0h
Retentate
Page 30 of 32
us
y = -62.2 Log(x) - 144.1 R2 = 0.996
100 80
an
60 40 20
M
Total Zn2+ released (%)
120
cr
ip t
Figure 6
0 10-5
10-4
10-3
10-2
Ac
ce
pt
ed
Initial QD concentration in dialysis device (MZneq)
Page 31 of 32
ip t
Figure 7
cr
25
15 10 5 ND
0 No cells
Boiled Stat.
CH34
60
20 0
No cells
ce Total Zn2+ released (%)
Ac
Log
Boiled Stat.
Boiled Stat.
*
*
Log
Log
MG1655
*
ed
40
ND
M
80
pt
Total Zn2+ released (%)
(b)
(c)
us
20
an
Total Zn2+ released (%)
(a)
Boiled Stat.
CH34
MG1655
ND
ND
Log
100 80 60 40 20
0 No cells
Boiled Stat.
CH34
Log
Boiled Stat.
Log
MG1655
Page 32 of 32
