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Toxic interactions of different silver forms with
freshwater green algae and cyanobacteria and
their effects on mechanistic endpoints and the
production of extracellular polymeric substances†
Cameron Taylor,a Marianne Matzke,b Alexandra Kroll,c Daniel S. Read,b
Claus Svendsenb and Alison Crossleya
This study investigated the magnitude and mechanisms of toxicity that ionic silver and two silver nanoparticles (AgNPs) of differing size and stabiliser imparted on two model organisms, green algae
(Chlamydomonas reinhardtii) and cyanobacteria (Synechococcus leopoliensis), and their effect on the extracellular polymeric substances (EPS) produced. Major silver losses were detected during the 72 h of exposure compared with introduced nominal concentrations. Cell viability and production of reactive oxygen
species (ROS) were investigated to assess mechanisms of toxicity. All silver forms had a significant effect on
viability for C. reinhardtii but only ionic silver significantly affected ROS production. For S. leopoliensis, only
ionic silver affected viability. Levels of EPS produced by both species were similar between all treatments.
Compositional differences were noted for C. reinhardtii with greater levels of lower molecular weight material produced in the presence of all silver forms. The results indicate that ionic silver has the greatest ef-
Received 21st August 2015,
Accepted 29th December 2015
fect on the toxic stress endpoints. Variations between the ionic control and the two AgNP treatments are
DOI: 10.1039/c5en00183h
possibly due to differences in the ionic release dynamics and interactions with EPS produced by microorganisms. S. leopoliensis showing greater reduction in growth rates but lower impact on viability and ROS
rsc.li/es-nano
composition.
than C. reinhardtii is likely related to differences in relevant biological properties such as size and cell wall
Nano impact
Anti-microbial properties of nanosilver have led to their increasing use in many products leading to concerns that these materials will be able to enter
environmental systems and impact aquatic microorganisms. This study investigates and builds on the knowledge base relating to the magnitude and
mechanisms of nanoparticulate silver toxicity to aquatic green algae and cyanobacteria. It was found that whilst production of ionic silver is the
predominant factor in nanosilver toxicity an additional particle-specific effect cannot be discounted. Additionally exposure to silver nanoparticles affected
the composition of extracellular polymeric species (EPS) produced by green algal species. Understanding nano-EPS interactions will help further elucidate
how nanomaterials and algal species interact in realistic exposures.
Introduction
Production of particulate silver (Ag) in the nanometer size
range has shown rapid growth in recent years due to
a
Department of Materials, Oxford University Begbroke Science Park, Begbroke
Hill, Yarnton, Oxford OX5 1PF, UK
b
NERC Centre for Ecology & Hydrology, Maclean Building, Benson Lane,
Crowmarsh Gifford, OX10 8BB, UK
c
Eawag, Swiss Federal Institute of Aquatic Science and Technology, 8600
Dübendorf, Switzerland
† Electronic supplementary information (ESI) available. See DOI: 10.1039/
c5en00183h
396 | Environ. Sci.: Nano, 2016, 3, 396–408
antimicrobial applications in numerous industries including
health and cosmetics, applied to functionalized textiles, airfiltration devices and sock fibre de-odourisers.1–5 This is due
to the unique thermal, electrical, optical and catalytic properties that arise at the nanoscale which contribute to the antibacterial properties of nanoparticulate Ag. These antibacterial
properties have been shown to have negative effects on organisms and ecosystems, particularly where they enter
through the wastewater route. Modelled levels of nanoparticulate Ag reaching the surface waters in Switzerland
have been predicted at 80 ng Ag L−1 (ref. 6) and at 127 ng Ag
L−1 for Swiss wastewater treatment plant (WWTP) effluents.7,8
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Few studies have measured Ag concentrations in situ however; using single-particle ICP-MS, nanoparticulate Ag was
recorded at an average of 100 ng Ag L−1 in the presence of
other Ag forms in effluent from a Colorado WWTP.9,10 These
values are only likely to increase with currently increasing
production of Ag containing products.11 These modelled and
measured values are average values and are much lower than
short-term EC50 values for aquatic microbes (~1–100 μg Ag
L−1 depending on species and endpoint10). However local
concentrations may vary from the averages and the fate of
AgNPs is still unknown.
Aquatic microorganisms such as algae and cyanobacteria
play an important role in nutrient cycling and the health of
freshwater ecosystems.12 They are amongst the most sensitive
species to toxicants and as such they often drive risk assessment and fate studies.12 Adverse effects of nanoparticulate
Ag on microbial growth have been noted for both Grampositive and Gram-negative bacteria,13–15 cyanobacteria16,17
and both freshwater and marine algae12,18–21 with effective
concentrations ranging between 1 μg Ag L−1 (ref. 17) and up
to 75 mg Ag L−1.13
Ionic silver (Ag+) toxicity to microorganisms acts through
inactivation of cellular proteins and thiol groups which affects ATP production, ion transport, DNA replication and respiration.22 However, the extent that toxicity mechanisms of
Ag NPs differ from ionic Ag is unclear, especially in natural
systems. Properties of nanoparticles (concentration, size, stability, morphology, and aggregation state), the surrounding
medium (composition, concentration of suspended and
dissolved species) and interactions between the two are likely
to affect their exposure and toxicity to microorganisms.
Debate has been ongoing in the scientific literature
whether the cause of AgNP toxicity to algal and cyanobacterial microorganisms is due to direct particle–cell surface
attachment, internalisation through the cell membrane, release of Ag+ through dissolution or some combination of
these.12,16,18–21,23 Some studies have linked the particle size
with the toxic impact of AgNPs through the levels of ionic silver released through the particle surface24,25 and their ability
to enter organisms through pores in the cell walls12,20
Ag NPs or ions that breach the cell wall could damage the
cell membrane and cause membrane integrity loss and cell
lysis.15,20,26–30 Oxidative stress is also thought to result from
exposure to nanoparticulate and Ag+ as high reactivity of NPs
exposed to dissolved oxygen can result in the formation of oxygen free radicals at the surface.31
Furthermore, it is known that aquatic microorganisms
produce extracellular polymeric substances (EPS) in response
to toxic stress. However the composition and the abundance
of these materials have not been researched in depth. Some
studies have assessed effects of ionic and nanoparticulate Ag
forms on bacteria in activated sludge which seemed to cause
an increase in EPS production after exposure to sub-lethal
concentrations.32 It is important to assess the effects of EPS
as an increase in these substances could lead to a decrease in
toxicity of Ag forms to aquatic microorganisms or even
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Paper
reduce Ag+ into solid nanoparticulate form.19,33–35 It is anticipated that greater levels and a different composition of EPS
will be produced at higher concentrations of toxicant and
that there will be EPS differences resulting from the different
Ag forms studied.
The aims of this study were to firstly gauge the effects of
two stabilised silver nanoparticles (AgNPs) with different
properties (size and stabiliser) on freshwater microbes. Secondly we wished to investigate differences in response between prokaryotic and eukaryotic species using two model
species representing different classes of photosynthetic
microorganisms; the green algae Chlamydomonas reinhardtii
and the cyanobacterium Synechococcus leopoliensis. Finally
we wished to analyse if EPS (extracellular polymeric substances) production by algal and cyanobacterial species is affected by Ag and AgNP exposure.
Experimental
Chemicals
Two AgNP products were investigated in this study. The first,
commercially available and referred to as AG1 (Amepox, Poland), had an advertised primary particle size of 3–8 nm and
was suspended in an alkane stabiliser. The second, referred
to as AG2 (Institut Català de Nanotecnologia, Spain), was a
powder, had an advertised primary particle size of 50 nm (silver 81.08%) and was coated with polyvinylpyrrolidone (PVP
9.73%) and further stabilised with tannic acid (9.19%). All
other chemicals were purchased from Sigma Aldrich if not
stated otherwise. Stock suspensions of 500 mg L−1 were created via a modified protocol36 by weighing out powder and
suspending in Milli-Q water. Silver nitrate (AgNO3) powder
dissolved in Milli-Q water (stock concentration 50 mg L−1)
was used as an Ag+ control to distinguish between particle
specific effects and effects caused by Ag+. Fluorescence stains
(fluorescein diacetate: FDA, dihydroethidium: HE and 2′,7′dichlorodihydrofluorescein diacetate: H2DCFDA) were purchased from Sigma-Aldrich and Molecular Probes.
Nanoparticle characterization
Characterization of NP dispersions was performed at 1 mg
L−1 for optimal measurement quality based upon the concentration limits of the different techniques. These stock suspensions were diluted to 1 mg L−1 in MBL Woods Hole medium
with and without suspended extracellular polymeric substances (unstressed C. reinhardtii and S. leopoliensis: CR− and
SL− and stressed C. reinhardtii: CR+). Extracellular polymeric
substances (EPS) were extracted using the same method as
described below for EPS characterisation. To analyse the influence of EPS on the AgNPs behaviour under realistic conditions EPS containing MBL Woods Hole medium was produced by growing algae and cyanobacteria for the testing
period of 72 hours. After this period the organisms were
harvested by centrifugation to gain a pure suspension
containing only medium and EPS for the exposure of the
AgNPs. AgNPs exposed in MBL Woods Hole medium with
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and without EPS (for a control), were characterised for hydrodynamic diameter using dynamic light scattering (DLS,
Zetasizer Nano ZS, Malvern Instruments), nanoparticletracking analysis (NTA, Nanosight LM10, NTA 2.0) and differential centrifugal sedimentation (DCS, CPS disc centrifuge).
Both DLS and NTA measure hydrodynamic diameters (dH)
and particle size distributions (PSDs) of individual dHs. However, the former technique gives an intensity derived
Z-average signal while the latter gives a number based distribution which means larger particles are weighted stronger in
DLS than in NTA if samples are not monodisperse.37 DCS
measures PSD and stokes diameter (dS). Zeta potential (ZP)
was calculated from measurements of electrophoretic mobility (Malvern Zetasizer Nano ZS, Malvern Instruments) and pH
was assessed using calibrated Sartorius Professional pH meter PP-25. Each suspension was measured at 0 h (within 10
minutes of sample preparation), 24 h, 48 h, 72 h, 168 h and
336 h in triplicates. Measurements at 168 h and 336 h,
greater than the 72 h toxicity test, were assessed to understand long term stability.
For the assessment of size and morphology, samples of AG1
and AG2 were deposited on a holey carbon coated Cu TEM grid
(Agar Scientific: 0 h and 336 h) and dried at room temperature
for several hours before transmission electron microscopy
(TEM) measurements were taken. An analytical TEM (JEOL
2010) with a LaB6 electron gun, operation range 80–200 kV, resolution of 0.19 nm, electron probe size down to 0.5 nm and
maximum specimen tilt of ±10° along both axes was used for
measurements. Images were analysed using GatanDigital
Micrograph (Gatan Inc.) for particle size and aspect ratio data.
Details on how many particles were counted (between 17 and
40 depending on the sample) are given in the ESI† (Table S1).
Growth inhibition tests with freshwater green algae and
cyanobacteria
C. reinhardtii (strain number CCAP 11/45) was obtained from
the Culture Collection of Algae and Protozoa (CCAP, Scottish
Marine Institute, Oban, United Kingdom); S. leopoliensis
(strain number 1402–1) was obtained from the Culture Collection of Algae, University of Gottingen, Germany. Organisms were cultured in MBL Woods Hole medium from CSIRO
adapted from Nichols (Nichols 1973) for freshwater organisms. Cells were cultivated in batch culture of 50 ml of MBL
Woodshole medium (composition in Table S2†) in a controlled temperature room in cell culture flasks (TC easy flasks
for suspension culture, 75 cm3, VWR, United Kingdom). Cultures were exposed to 16 : 8 hours light : dark regime, 9000 lux
light intensity at 22 °C and agitated at 80–95 rpm. To keep
the cell growth in the exponential phase the test cultures
were regularly diluted 3 times per week to a cell density of 5
× 104 cells per mL. The cell numbers were determined using
flow cytometry (Beckman Coulter Gallios equipped with blue
(488 nm) and red (638 nm) solid state diode lasers).
72 h growth inhibition tests were performed with C.
reinhardtii and S. leopoliensis covering a concentration–response
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Table 1 Nominal concentration ranges (μg L−1) of AgNO3, AG1 and AG2
used in dose–response growth rate-inhibition experiments on
C. reinhardtii and S. leopoliensis
Nominal concentrations (μg L−1)
Species
AgNO3
AG1
AG2
C. reinhardtii
S. leopoliensis
0.047–500
0.1–500
0.5–800
0.5–8000
0.5–8000
0.5–20 000
curve for each toxicant (AG1, AG2, AgNO3, see Table 1 for
concentration ranges) according to a modified version of the
OECD 201 Guideline.38
The OECD test protocol 201 was modified using a light :
dark cycle to increase ecological realism, a different test medium than the one recommended in the guideline (MBL
Woodshole medium) and the use of the green algae
Chlamydomonas reinhardtii instead of Pseudokirchneriella
subcapitata to be able to compare these results to previous
studies from our lab (publication in preparation). Samples
were prepared in sterile cell culture flasks (TC easy flasks for
suspension culture, 25 cm2, VWR, United Kingdom) by addition of 1 ml of culture (final cell density in the test vessels:
2–7 104 cells per ml) to 9 ml of MBL Woodshole medium
dosed with toxicant and incubated for 72 hours. Each substance was tested at least in three independent experiments
with 8 different concentrations in 2 replicates with 6
untreated controls to ensure reproducibility of the data.
Growth inhibition was calculated as percentage difference
between the control growth rate and sample growth rate as
calculated from changes in population density between cultures exposed to a toxicant and that of an untreated control.
Cell numbers were determined using a Beckman Coulter
Gallios flow cytometer equipped with blue (488 nm) and red
(638 nm) solid state diode lasers. Dot plots of chlorophyll
fluorescence (FL4 – 695/30 nm, 488 nm excitation) and phycocyanin fluorescence (FL6 – 660/20 nm, 635 nm excitation),
both against side scatter (SS) were employed to quantify the
algal populations. Counts were obtained by gating the area
around an expected algal population. Because in a flow
cytometer the flow rate of the sheath fluid is variable the addition of number calibrated latex beads (1.568 ± 0.026 μm
carboxylate latex microspheres (Fluoresbrite™, PolySciences
Inc, UK)) is necessary to be able to accurately determine the
cell numbers (given as counts per mL) in the sample. The exact number calibration of these beads was done using
Beckmann Flow Count Fluorespheres (High Wycombe, UK),
100 μL of the calibrated beads were added to each sample.
Concentration-response curves were created by plotting
growth rate inhibition against toxicant concentration using R
(details see below).
Analysis of cell viability and ROS formation by flow cytometry
Further analysis of toxic action was assessed using fluorescence staining followed by flow cytometry. For analysis of cell
viability, fluorescein diacetate (FDA) was used as a viability
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probe that measures both enzymatic activity and cellmembrane integrity. FDA was supplied by Sigma Aldrich. For
the analysis of reactive oxygen species (ROS), dihydroethidium (HE) and 2′,7′-dichlorodihydrofluorescein diacetate
(H2DCFDA) were used (Molecular Probes).
FDA is non-fluorescent, non-polar and lipophilic, and enters cells through cell walls and membranes. It is deacetylated to yield hydrophilic and polar fluorescein which is
unable to leave the cell unless there is damage to cell
membranes. HE is oxidized by free oxygen radicals such as
H2O2 and NO3− reducing blue HE to read hydroethidine39
which associates with acids in the cell nucleus and fluoresces.40,41
H2DCFDA (2′,7′-dichlorodihydrofluorescein diacetate) is deacetylated intracellularly by esterases yielding non-fluorescent
H2DCF (2′,7′-dichlorodihydrofluorescein). Upon interaction
with ROS, it is oxidized to fluorescent DCF (2′,7′dichlorofluorescein).42–44
C. reinhardtii and S. leopoliensis (6 ml) were exposed at cell
density of 2 × 106 cells per ml to the EC70 obtained from the
growth inhibition assays (Table S3:† concentration at which
70% inhibition of growth occurs assessed from dose–response data) of each toxicant for 60 minutes before addition
of fluorescent stains for the required time to reach optimum
staining of the samples which was 10 minutes for FDA and
HE and 80 minutes for H2DCFDA. All stains were prepared in
DMSO (dimethyl sulfoxide) and diluted with ultrapure water
before being added to the samples according to instructions.
Cells were stained with the following final concentrations:
FDA 240 μM, HE 5 μM and H2DCFDA 10 μM. The efficacy of
each stain was tested using a H2O2 positive control at a concentration of 1% for same exposure time as the Ag
treatments.45
Counts were obtained of populations within a pre-defined
gated area using Gallios Cytometry List Mode Data Acquisition and Analysis Software. FDA and DCF stained samples
were recorded on a plot of green fluorescence (525/40 nm,
488 nm excitation) versus side scatter while HE was recorded
on a plot of yellow fluorescence (575/30 nm, 488 nm excitation) versus side scatter. For each measurement the nonstressed control population exposed to the stain was gated
and the level of response was gauged by movement of the
population out of the pre-defined gate. The population in the
‘non-stressed’ control gate or the ‘stressed’ gate were plotted
as a percentage against the total cells measured to assess the
effect of each toxicant on the endpoints defined by the stains
(cell viability and ROS generation) (example shown in Fig.
S1†). Each experiment (individual algal population) had three
replicates and experiments were repeated a minimum of
three times to create the data set.
Significant differences between each treatment and the
control in a data set were analysed using the raw data. Ratios
of stressed to unstressed alga for each treatment were natural
log-transformed (response variable) and compared to the control using a standard linear model with treatment and replicate as predictor variables. The fit of the model was assessed
using R. The p-values indicated the probability that no
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relationship exists between the treatment and the control:
<0.05 indicated significance and therefore likely a difference
between variables.
Extraction and characterization of extracellular polymeric
substances (EPS)
Extracellular polymeric substances EPS were extracted from
C. reinhardtii and S. leopoliensis after 72 h of exposure in
growth medium with or without AgNPs and AgNO3. Exposure
conditions were as described for the growth inhibition experiments. Nominal Ag exposure concentrations for C. reinhardtii
were 50 and 969 μg L−1 (AG1), 1624 and 3500 μg L−1 (AG2), 5
and 23 μg L−1 (AgNO3) while for S. leopoliensis exposure concentrations were 300 μg L−1 (AG1), 600 μg L−1 (AG2) and 9.2
μg L−1 (AgNO3). The extraction procedure was done according
to Kroll et al.35 Three replicates of 10 ml from each control
and exposed culture were centrifuged at 3647g for 10 min
(room temperature). Biomass was re-suspended a second
time in 5 mL fresh growth medium and treated as described
above. The resulting biomass pellets were frozen at −20 °C
and subsequently lyophilised to determine dry weight. All
supernatants were sequentially filtered (1 μm glass fibre
[VWR], 0.45 μm polypropylene [PALL], and 0.22 μm PES
[Millipore] filters). Filters were washed with nanopure water
(18.1 MΩ cm, Milli-Q) prior to use. EPS extracts were stored at
4 °C (0.03% (w/v) NaN3). All glassware used for EPS extraction
was heat treated in a muffle furnace before use (450 °C, 4 h).
The size distribution of organic carbon and nitrogen
compounds was measured by size-exclusion chromatography–organic carbon detection–organic nitrogen detection
(LC-OCD-OND) as described previously.35,46 Samples were
diluted 1 : 10 with nanopure water (18.1 MΩ cm, Milli-Q)
directly before measurements. A size exclusion column (250 ×
20 mm, Toyopearl TSK HW-50S) was used to separate EPS
compounds. The mobile phase was phosphate buffer (24
mM, pH 6.6) and the acidification solution was phosphoric acid
(60 mM, pH 1.2). The software FIFFIKUS was used to quantify
total organic carbon (TOC), dissolved organic carbon (DOC),
and chromatographable DOC compounds (cDOC). The chromatograms obtained from LC-OCD-OND were normalized to
the dry weight of the samples and integrated between 28 and
75 min retention time to determine the amount of organic
carbon in different size fractions of the EPS.
Quantification of total and dissolved Ag in the exposure
medium
C. reinhardtii and S. leopoliensis dose–response curves (Fig.
S2 and S3†) were used to select a range of concentrations that
were measured for total and dissolved Ag (Tables S4 and
S5†). Total and dissolved Ag were analysed at 0 hours and 72
hours in each treatment using ICP-MS (inductively coupled
plasma mass spectrometry).
For total Ag measurements, 1 ml from each treatment was
acidified by adding 0.15 ml of 69% HNO3 and stored at 4 °C
until analysis. For quantification of dissolved Ag, 1.5 ml of
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sample was centrifuged at 20 000g for 3 hour to separate particles (>3.5 nm) and dissolved ions (25 °C, Ag, ρAg 10.5 g
cm−3). After centrifugation, 1 mL of supernatant was acidified
with 0.15 ml 69% HNO3 and stored at 4 °C until analysis. All
samples were treated by microwave digestion prior to ICP-MS
analysis. 0.5 mL of each sample were digested with 4 mL of
65% HNO3 and 0.5 mL of 30% H2O2 in a microwave digestion
unit (MLS ultraClave 4; 10 min at 180 °C/100 bar, 14 min at
210 °C/100 bar) and diluted 1 : 100 with nanopure water (18.1
MΩ cm, Milli-Q). One sample per run contained only HNO3
and H2O2 to determine the background concentration of Ag.
Ag concentrations were measured by HR-ICP-MS (Element 2
High Resolution Sector Field ICP-MS; Thermo Finnigan). The
instrument was calibrated with a multi-element mass standard (Merck, 1113550100). The calibration curve for data
analysis was made with the calibration standard SCP-33-MS
(140-130-321, PlasmaCAL) in the concentration range 0–20 μg
L−1. A reference with a concentration within the calibration
range was measured every 10 samples, the calibration samples were measured every 40 samples. The LOQ of Ag was
0.01 μg L−1.
Statistics and data analysis
Characterisation data from DLS, NTA, DCS, and TEM techniques was analysed using Microsoft Excel 2010. Error bars
in charts were set to one standard deviation of the mean
obtained from at least 3 replicates. Flow cytometry data was
converted into an excel spreadsheet and was analysed in R.47
A log-logarithmic four parameter curve was fitted to each plot
to obtain ECx values using the drc R package.48,49 EC20, EC50
and EC70 of each toxicant for each species are given in Table
S3.† The number of experiments (N) and the total number of
data points (n) used to create each concentration–response
curve varied for each curve and toxicant (Table S3†). Tests
were repeated at least 3 times, two replicates were used per
individual concentration point in each experiment. ECx
values are given as nominal concentration ± 1 standard error
and concentration–response curves can be found in ESI†
(Fig. S6 and S7). R was also used to fit a linear model to compare the mechanistic toxicity effects of each treatment to the
control in the fluorescent staining experiments. Data on EPS
composition were analysed with one-way ANOVA and
Bonferroni's post-test (α = 0.05) and were plotted with
GraphPad Prism 5 for Windows.
Results and discussion
Characterisation data indicate little change in size and particle size distribution of both AG1 and AG2 particles over the
duration of the experiments as measured by TEM, NTA, DLS
and DCS (ESI:† Tables S6, S1, S7. Fig. S4, S5). Zeta Potential
showed high to moderate electrostatic stability for the duration of the experiment (−15 to −50 mV)50 (Table S6†). AG1
and AG2 primary particles in all suspensions (1 mg L−1 in
MBL Woods Hole with and without suspended EPS) were
400 | Environ. Sci.: Nano, 2016, 3, 396–408
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shown to be ∼7.7–10.4 nm and ∼43–75 nm, respectively,
through TEM imaging.
Total and dissolved Ag in the exposure medium
Background Ag levels in the MBL Woods Hole medium were
0.065–0.102 μg L−1. Measured total Ag levels for AG1 and AG2
in both C. reinhardtii and S. leopoliensis exposures at 0 and
72 hours indicate large losses compared to the nominal concentrations (Fig. S6, S7; Tables S4, S5†). In general, the loss
rates AgNO3 were between 10 and 50% (depending on the
tested species and concentration) smaller than for the AgNPs.
Here for both particles loss rates were ranging approximately
between 92 and 98.5%, again depending on tested species
and concentration. Consistent silver losses for all nanoparticle treatments likely indicate that absorption to container occurred during the experiments, possibly even during the
preparation steps. As a consequence, the true exposure concentrations in growth inhibition rate, toxic mechanism and
EPS analysis are much lower than the stated nominal values.
Differences between measured and nominal total Ag concentrations were used to calculate factors to correct the ECx
values derived from the concentration–response curves to assess the realistic Ag concentrations in the test exposures (correction factors shown in Table S5†). The factors were based
on the measured total silver at the test start divided by the
nominal concentration. Because measurements were only
done at test start and after 72 hours (test end) the dynamics
in the system, i.e. further losses of silver could not be taken
into account and we therefore focused on the real concentrations in the system when the exposures started. Nominal
concentrations used in EPS experiments were also corrected
(Table S9†).
Measured dissolved Ag for AgNO3 samples decreased compared to total measured Ag after 72 h in all treatments (∼87–
96% decrease at lower concentrations, 66–69% decrease at
higher concentrations: Table S4†).
The dissolved fraction in AG1 remained in the same range
for the 72 h duration of the experiment (∼6–20%) in the presence of both species whereas the dissolved fraction in AG2
increased from ∼2–3% to ∼4–44% (Table S4†).
In sum, exposure conditions regarding the fraction of
dissolved silver were distinctly different between AgNO3, AG1,
and AG2 at the start of the experiment for both species
(AgNO3: ∼80–100%, Ag NPs: ∼0.6–16%), but approximated
each other over time. An exception was AG1 in C. reinhardtii
exposures where fraction of dissolved silver remained fairly
constant with only slight changes from an average of 1.5 μg
L−1 for t0 to 1.6 μg L−1 for t72 (for nominal 100 μg L−1, Table
S4†) and 1.9 μg L−1 (t0) to 2.1 μg L−1 (t72) (nominal concentration 500 μg L−1, Table S4†).
Comparison between measured total Ag and dissolved Ag
indicated that whilst dissolution of NPs was low, it was concentration dependent with higher concentrations of both
AG1 and AG2 exhibiting less dissolution per total Ag level
than lower concentrations (Fig. S2†). Carbonate used as
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stabilizing agent for AgNPs has been shown to increasingly
disassociate from the NPs with higher dilution resulting in
higher dissolution of Ag.51 This process could apply to the
AgNP used in this study. Further, experiments on the abovementioned carbonate-stabilized AgNP and AgNO3 showed
that AgNP may increase in size and are formed from AgNO3
by EPS-enhanced photoreduction.35 The measured concentrations of total and dissolved Ag may thus also be the result of
a concentration-dependent equilibrium of oxidation and reduction of Ag.
Effect of nanoparticles on growth-inhibition of C. reinhardtii
and S. leopoliensis
The EC20, EC50 and EC70 values of each toxicant based on
nominal concentrations and on corrected concentrations
show the magnitude of toxicity of each toxicant to each species (Table S3†). All forms of Ag in this experiment exhibited
toxicity to both microorganisms so that dose-response curves
could be recorded for all tested compounds and species.
Results based upon nominal concentrations indicated that
whilst AgNO3 had the strongest effect on growth rate inhibition no difference between species was noted [EC50 (±SE) C.
reinhardtii: 43 ± 7.7 and S. leopoliensis: 66.9 ± 65 μg L−1] (Table S3†). However, nominal concentrations of AG1 and AG2
caused both species and NP specific effects. EC50 (±SE) of
AG1 and AG2 in S. leopoliensis (164 ± 27 and 184 ± 59.8) were
∼60% higher than AgNO3 EC50. In C. reinhardtii EC50 AG1
was about half the AG2 EC50 and both were 20–50 times
higher than EC50 of AgNO3.
In contrast, based on concentrations corrected for measured amounts of Ag, there is little difference in toxicity between the three toxicants to either species (Table S3†). No difference between species was noted for AgNO3 or EC50 and
EC70 for AG1 and EC70 for AG2 based on the actual measured
silver concentrations. However, some corrected ECx values for
S. leopoliensis are lower than those in C. reinhardtii (AG1:
EC20, AG2: EC20 and EC50) indicating that cyanobacteria may
be more sensitive at lower ECx values to the NPs than the
green algae. Corrected concentrations of AgNO3, AG1 and
AG1 were in the same range, between 25–61 μg L−1 for
C. reinhardtii and with greater variation (1–124 μg L−1) in
S. leopoliensis exposures (Table S3†).
ECx values were corrected using dissolved silver levels
taken at 0 h in the analytical experiments (Table S8†) however the fraction of dissolved silver in relation to total silver
changes from 0 h to 72 h indicating changing dissolution/
reprecipitation kinetics during the experiment. It is possible
that toxicity of the toxicants to both species is influenced by
these dynamics and further study is required.
The differences between nominal and measured concentrations indicate the necessity of analytical verification of exposure concentrations. It is likely that the organisms were
not exposed to the nominal concentrations of Ag. Corrected
concentrations indicate little difference between the
This journal is © The Royal Society of Chemistry 2016
Paper
magnitude of concentration that each toxicant imparts on
each organism i.e. no particle specific effect was noted.
The corrected EC50 values of AgNO3 (44.6 μg L−1 ± 8 SD)
and AG1 (38.5 μg L−1 ± 0.3 SD) were in the same range as
green algae species P. subcapitata (AgNO3 33.79 ± 2.96 μg L−1
SE, AG1 32.40 ± 2.09 μg L−1 SE) in another study33 although
nominal values for AG1 were much higher.
Another study using photosynthetic yield as end point and
shorter exposure periods (1–5 hours) found similar EC50
values for AgNO3 to the current study (21.25–31.94 μg L−1)
but higher EC50 for carbonate coated 25 ± 13 nm AgNPs
(89.42 μg L−1 at 5 hours, 356 μg L−1 at 1 hour) of which ∼1%
of total Ag was in dissolved form.20
Studies investigating NP toxicity to cyanobacteria have
noted values much higher than the corrected EC70 concentrations for AG1 and AG2 in this study (Table S3.† 13.1 ± 3 and
22.6 ± 11.4 μg L−1) to M. aeruginosa17 (87% growth inhibition
from 1 mg L−1 tannic acid synthesised AgNPs) and
Synechococcus sp., the same genus as S. leopoliensis.16
The possible higher sensitivity of S. leopoliensis to the
AgNPs than C. reinhardtii in the present study could be due
to both physical and biological reasons. S. leopoliensis being
smaller organisms, have a greater surface to volume ratio for
interaction or uptake of toxicants and also thinner cell walls
(∼10 nm for Synechococcus species).17 Cell walls of C.
reinhardtii have been suggested to be important when gauging AgNP toxicity.27 The biological makeup of each organism's cell wall could result in differences in affinity and nonspecific interactions with Ag forms. Green algae cell walls are
mainly cellulose while for cyanobacteria they consist mainly
of peptides, peptidoglycan in particular.17
Smaller AG1 primary particles (∼7.7–10 nm for the experiment duration) could pass through an algae cell wall which
tend to range from 5–20 nm.52 However, no difference in toxicity was noted between the two particles for corrected concentration values. There are only few studies investigating algae and cyanobacterial cell wall pore sizes, but the few
available data indicate typical sizes between 5 and 20
nm.53,54
Pores could also become damaged by NP interaction and
become large enough for bigger particles to become internalised20 due to the high affinity of silver and sulphur
containing algal cell walls.55 This cannot be discounted for
AG2 NPs.
Effects of AgNO3 and AgNPs on cell viability
The effect of AgNPs and AgNO3 on cell membrane integrity
in C. reinhardtii and S. leopoliensis was investigated by FDA
fluorescence staining. A linear model of the percentage of
highly FDA-fluorescent cells as a function of treatment was
constructed and applied to each toxicant: log transformed ratio of unstressed to stressed cells ∼ Replicate + Treatment
where replicate states the number of repeated treatments and
treatment refers to the specific exposure, i.e. AgNO3, AG1,
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Environmental Science: Nano
Table 2 Model fit statistics (R 2, F-statistic, degrees of freedom, p-value) for application of linear model to log-transformed ratios of stressed to unstressed algal populations for each treatment (EC70 level exposures of AG1, AG2 and AgNO3) and species. Linear model applied to log-transformed ratios
of stressed to unstressed cells using R: log-transformed ratios (stressed/unstressed cells) ∼ Replicate + Treatment
Linear model: log-transformed ratio (stressed/unstressed cells) ∼ Replicate + Treatment
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Species
C. reinhardtii
S. leopoliensis
Stain
Stain
Model fit–statistics
FDA (cell viability)
HE (ROS)
DCF (ROS)
FDA (cell viability)
HE (ROS)
DCF (ROS)
R2
F-statistic
Degrees of freedom
p-value
0.7735
24.9
5 and 30
7.665 × 10−10
0.1323
2.8
5 and 54
0.0255
0.5904
14.55
5 and 42
2.824 × 10−8
0.9059
68.35
5 and 30
1.754 × 10−15
0.2803
3.727
5 and 30
0.009636
0.4564
6.877
5 and 30
0.0002208
Table 3 P-values for comparison of log-transformed ratios of stressed to unstressed algal populations against the control for each treatment (EC70
level exposures of AG1, AG2 and AgNO3) and species using a linear model. Values indicate the likelihood that no relationship exists between the treatments. Asterisks indicate the significance of the relationship: no asterisk is no significance (≥0.05), ‘*’ is low significance (≤0.05), ‘**’ is moderately significant (≤0.01), ‘***’ is highly significant (≤0.001)
Species
C. reinhardtii
S. leopoliensis
p-values
Stain
Stain
Treatment (endpoint)
FDA (cell viability)
HE (ROS)
DCF (ROS)
FDA (cell viability)
HE (ROS)
DCF (ROS)
AG1
AG2
AgNO3
1.40 × 10−10 ***
4.28 × 10−9 ***
2.38 × 10−10 ***
0.0940
0.75376
0.00151 **
0.0655
0.7780
3.08 × 10−9 ***
0.00597 **
0.06039
<2 × 10−16 ***
0.001964 **
0.1114885
0.000583 ***
0.0139 **
0.16605
0.04060 *
AG2 or control (i.e. medium only). In C. reinhardtii, the
model provided a good fit (Tables 2 and 3: F(x, y) = 0.7735).
The mean decrease in the fraction of intact cells in the
presence of all Ag forms was significant. In S. leopoliensis,
the model provided a very good fit (Tables 2 and 3, F(x, y) =
0.9059). AgNO3 produced a significant decrease in intact cells
compared to the control (Fig. 1). AG1 may have had a significant effect compared to the control while AG2 did not significantly affect membrane integrity (Fig. 1).
In C. reinhardtii, loss of cell membrane integrity is likely
an important toxicity mechanism as applying a linear model
indicated significant differences in cell viability from the control for all silver forms. However, only AgNO3 showed a significant difference compared to the control in S. leopoliensis.
This indicates that a species specific mechanistic toxic effect
is occurring.
A silver ion specific acute effect could have occurred as
Ag+ originating from AgNO3 likely damaged cell membranes
more quickly than Ag+ released from NPs as the former are
present in greater concentrations at the beginning of the experiment. Removal of Ag+ from the AgNO3 treatments over 72
h may have been caused by binding to suspended EPS or cell
surfaces. Differences in membrane integrity noted between
the two species for the NPs, AG1 showing potentially greater
damage than AG2, are interesting considering that both particles showed similar levels of dissolved Ag. It is possible that
the fraction of smaller-sized AG1 particles is sufficient to
cause damage to the two species' membranes, while in
C. reinhardtii, we speculate exposure to AG2 possibly results
in membrane damage through attraction and accumulation
of silver at the membrane surface (due to high affinity between silver and sulphur-containing cell components55
Fig. 1 Cell viability: FDA stain. Percentage of intact cells when exposed to EC70 AgNO3, AG1 and AG2 for A) C. reinhardtii cultures and B) S.
leopoliensis cultures. ‘a’ indicates significant effect compared to the control (p value ≤ 0.05) derived from fitting a standard linear model to ratios
of stressed/unstressed cells. Control values deviate from 100% as even in healthy cultures some cells fell outside the non-stressed gate.
402 | Environ. Sci.: Nano, 2016, 3, 396–408
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increasing permeability and release of cell components as
EPS. It is possible that the fraction of smaller-sized AG1 particles is sufficient to cause damage to the two species' membranes, while in C. reinhardtii, exposure to AG2 possibly results in membrane damage by abrasion.
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Effects of silver and AgNPs on ROS development
The effects of AG1, AG2 and AgNO3 on ROS production in the
cells of C. reinhardtii and S. leopoliensis were assessed by
changes in fluorescence as derived by HE (specific to H2O2)
and H2DCFDA (oxidized most efficiently by hydroxyl radical
and peroxynitrite anion). A linear model of the percentage of
highly HE or DCF-fluorescent cells as a function of treatment
was constructed and applied for each toxicant: logtransformed ratio of unstressed to stressed cells ∼ Replicate
+ Treatment.
NPs may interfere with enzyme/substrate and opticalreadout based toxicity tests,56 either by depletion of the substrate, or the reaction product, by NP-catalysed substrate
turn-over, by interfering with the required reaction, or the
readout itself. Affected measurements due to optical interference, NP-catalysis, or interference with the reaction are unlikely in this case, as we chose single-cell measurements
rather than measuring fluorescence in suspension. Adsorption of H2DCF-DA is thus the most probable type of interference. As the substrate was provided in large access (roughly
3.2 and 3.8 cm2 L−1 AG2, 21.7 and 10.2 cm2 L−1 AG1 in C.
reinhardtii and S. leopoliensis media, respectively, at EC70
concentrations based on mean particle size (TEM), and about
46 400 cm2 L−1 H2DCFDA), we argue that depletion of the
substrate was not substantial.
Based on HE fluorescence, no significant increase in
intracellular H2O2 could be observed for either species owing
to the models' poor fit (Fig. S8, Tables 2 and 3: C. reinhardtii:
F(x, y) = 0.1323, S. leopoliensis: F(x, y) = 0.2803). For DCF fluorescence, the linear model provided a good fit for
C. reinhardtii exposed to AgNO3 indicating a significant increase in ROS production (Tables 2 and 3: F(x, y) = 0.5904).
AG1 and AG2 did not have a significant effect on this endpoint (Fig. 2). For S. leopoliensis, the model did not produce
a good fit, and of the three toxicants, only AG1 may have had
Paper
an effect compared to the control (Fig. 2, Table 3: F(x, y) =
0.4564).
Out of all the treatments for both species the only significant increase in ROS was noted for AgNO3 exposure to C.
reinhardtii after H2DCFDA staining (Fig. 2), indicating that
Ag+ produces a hydroxyl radical response. As no significant
effect was noted for either NP this is likely due to the greater
fraction of AgNO3 total silver in dissolved form compared
with the NPs as seen in the analytical experiments. As such,
no species specific or a particle specific response was able to
be discerned from this data.
ROS generation has previously been shown to be concentration dependent and species specific in freshwater and marine algae (C. vulgaris and D. tertiolecta)27 exposed to 1–10
mg L−1 AgNPs over 24 hours. Lack of ROS production for AG1
and AG2 may be due to short exposure times (60 minutes)
and different AgNPs used in this study.
Agglomeration of NPs has been suggested to lower surface
area available for ROS production (uncoated CuO NPs57).
Whilst no additional agglomeration occurred over the duration of the experiment of either NP, clusters of AG1 particles
were present in suspension and lack of ROS due to agglomeration cannot be discounted.
Microalgae/cyanobacteria have both enzymatic and nonenzymatic oxygen scavenging defence mechanisms to prevent
ROS from damaging cellular structures58 which have been
shown to account for differences in H2O2-stimulated ROS
detected by DCF in M. aeruginosa and P. subcapitata59 and
which may account for low ROS in S. leopoliensis in this
study. Lower S. leopoliensis ROS levels compared to C.
reinhardtii may also relate to lower penetration of the DCF
stain into thinner and biologically different cell walls of the
cyanobacteria resulting in lower levels of H2DCFDA entering
the cell.59,60 It is also possible that ROS effect was mitigated
by exuded EPS as these have been reported to protect against
DNA damage and lipid peroxidation in other cyanobacterial
species by complexing with AgNPs.61
Toxicity driving factor: nanoparticles or ions?
Cellular mechanisms of toxicity of AgNPs on aquatic microorganisms are still not fully understood. Ag+ are among the
Fig. 2 Reactive oxygen species production: DCF stain. Percentage of high DCF-fluorescent cells when exposed to EC70 of AgNO3, AG1 and AG2
in A) C. reinhardtii and B) S. leopoliensis. Means of 3 replicates of 4 (C. reinhardtii) and 3 experiments (S. leopoliensis) respectively. ‘a’ indicates significant difference compared to the control (p value ≤ 0.05) derived from fitting a standard linear model to ratios of stressed/unstressed cells.
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Paper
most toxic metal ions to aquatic microorganisms due to their
positive charge and strong interactions with ligands present
in aquatic environments.62 Levels of dissolved Ag for most
AG1 and AG2 particle treatments over the duration of the experiment changed little when exposed to both C. reinhardtii
and S. leopoliensis (Table S5†). These values were much lower
than both the nominal and the corrected ECx values for the
AgNO3 control indicating that ions are not playing a large
part in the toxicity of these particles.
However, it is not known whether the EPS present in the
suspensions bound to Ag+ released from AgNPs resulting in
lower bioavailable Ag+ concentrations. The dissolved Ag concentrations for the AgNO3 control for both species indicate
that Ag+ is being removed from suspension although this is
shown to be concentration dependent with not all Ag+ removed at higher concentration exposures. The cause for the
Ag+ removal is unclear. It was hypothesised that EPS would
bind with the ionic forms and removes them from suspension. However, as discussed below total EPS levels do not
change indicating another mechanism is likely the cause.
Furthermore, the concentration-dependent dissolution results for AgNO3 could indicate that at higher concentrations
AgNPs are reforming through ion–EPS interactions. Further
study is needed to elucidate the dynamics of these
interactions.
Some studies have shown that Ag+ release from AgNPs was
the determinant factor in toxicity to freshwater algae, including C. reinhardtii,20 P. subcapitata,18 marine green algae such
as P. tricornutum18 and the marine diatom Thalassiosira
weissflogii.19 However, other studies on photosynthetic
microorganisms such as early phase C. reinhardtii,21 P.
subcapitata,63 a marine diatom (T. pseudonana), and cyanobacteria (Synechococcus sp.16) as well as higher trophic organisms such as daphnids and zebrafish63 have proposed a dual
free Ag+ and Ag particle effect. The interplay between
suspended EPS and the low levels of released ions from the
NPs still needs to be elucidated and more work is needed to
establish a particle or ionic specific effect.
Effects of Ag+ and AgNPs on EPS
To understand whether the different silver species affected
the EPS produced by C. reinhardtii and S. leopoliensis, EPS
was isolated from exposed and control cultures and analysed
by LC-OCD-OND. To exclude that changes in amount or composition of EPS in the cultures are due to physical depletion
of EPS components by Ag+ or AgNP, we mixed EPS isolated
from control C. reinhardtii and S. leopoliensis cultures with
the three different silver forms analogue to exposure conditions. We did not detect changes in EPS concentration or
composition in the supernatant of the AgNP or AgNO3
(Fig. S9†).
Chromatograms of all C. reinhardtii EPS extracts obtained
from exposed and control cultures showed the four expected
major features which relate to different retention times and
therefore different molecular weight (Mr) fractions:46 high Mr
404 | Environ. Sci.: Nano, 2016, 3, 396–408
Environmental Science: Nano
biopolymers, building blocks of humics, low Mr acids and
amphiphilic/neutral substances as shown in Fig. S10.† These
were converted into dry masses for each fraction and
converted into percentages of total EPS (Tables S10–S14, Fig.
S11–S14†).
In C. reinhardtii, 3500 μg L−1 AG2 reduced the total
amount of EPS per dry weight to about 40% (Table S10†).
The EPS composition was changed to higher concentrations
in low molecular weight compounds and lower concentrations in high molecular weight compounds (Fig. S11–S14†).
This effect was significant in 969 μg L−1 AG1, 23 μg L−1
AgNO3 and 1624 μg L−1 AG2.
Total EPS isolated from S. leopoliensis was in the same
range as in C. reinhardtii (0.047–0.055 mg EPS per mg dry
weight in controls: data not shown), however, the variability
in the Ag exposed cultures was very high and no significant
differences were detected. EPS composition was also very variable in treated S. leopoliensis cultures and no significant differences could be calculated (data not shown). Follow-up
work would be required to understand the interaction of
these organisms with Ag forms.
The interaction of EPS with Ag has been suggested to decrease Ag toxicity to microorganisms.19,34,64 Accordingly,
Escherichia coli and bacteria in activated sludge showed an
increase of EPS production when exposed to sub lethal concentrations of AgNP.34,65 Exposure to AgNP increased the
amount of EPS produced in a marine diatom.19 In benthic
microbial biofilms, EPS per dry weight did not significantly
change within three weeks of exposure to AgNP/AgNO3, however, the C : N ratio decreased indicating a higher protein
concentration.35 Depending on the microbial species and duration of exposure to silver forms, EPS amount and/or composition are thus sensitive endpoints. The present study
showed that the correspondence between the variables is
probably not a sigmoidal concentration–response relationship
and still needs to be investigated.
Summary of observed responses
S .leopoliensis had lower ECx values (nominal and corrected)
than C. reinhardtii and was thus more sensitive to all toxicants (AgNO3, AG1 and AG2). All toxicants had a significant
effect on C. reinhardtii cell viability while only AgNO3 had a
significant impact on S. leopoliensis. Reactive oxygen species
(ROS) generation was measured using two different fluorescent stains, however, the model fit on the data for the HE
stain was poor for both species exposures and no significant
effect could be detected. The DCF stain indicated that of the
three toxicants only AgNO3 produced ROS in C. reinhardtii exposures. EPS production when exposed to the three toxicants
was similar between the two species, however, variability between replicates meant that in the S. leopoliensis exposures
no conclusion could be made. Similarly, the composition results were also variable for S. leopoliensis. In C. reinhardtii,
less high molecular weight and greater levels of low molecular weight material is produced.
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These changes are complex and the mechanisms behind
this behaviour require greater study. Despite showing little
difference in cell viability and ROS production from the control the high suppression of S. leopoliensis growth in nanoparticle exposures likely indicates another mechanism may
be occurring. Further study could indicate that energy normally used to promote growth and replication could be
diverted to another task such as membrane repair or ROS
amelioration. It is known that some algae and bacterial species have inherent anti-oxidant mechanisms.66 The difference
in the type of EPS produced by C. reinhardtii when stressed
could also indicate that this energy is possibly rerouted to altered volume and composition of exuded EPS and study into
the composition of cyanobacterial EPS could yield more
answers.
Conclusions
The aim of this study was to investigate the effects of two different AgNPs on the growth rate inhibition, mechanisms of
action and production of extracellular polymeric substances
(EPS) to the green algae Chlamydomonas reinhardtii and the
cyanobacterium Synechococcus leopoliensis, a cyanobacteria
species. Two AgNPs with differing size distributions and capping agents showed little difference in inhibition of growth
rate in either species after correction for Ag loss indicating
the necessity for analytical verification of nominal concentrations during exposures. Dissolution of AG1 and AG2 NPs
remained around 0.16–6.25 μg L−1, much lower than the
nominal concentrations. It is not possible to say that low
levels of dissolved Ag were due to binding by EPS as EPS
levels did not change. Dissolved Ag from AgNO3 was noted to
decrease over the experimental duration at a concentration
dependent level and more work is needed to understand
these interactions.
It should be stated that whilst the 72 h exposure period
for the toxicity tests in this study is recommended in the
standard OECD protocol the silver dissolution dynamics may
not have stabilized over this time. Stevenson et al. investigated the interaction of citrate-coated AgNPs with C.
reinhardtii for up to two weeks and showed that for these particles silver dissolution was slower and silver ions interacted
with EPS produced by the algae.21 Whilst the size and stability of the particles was mostly unchanged for two weeks,
more information is needed for particle dissolution kinetics
after 72 h to further understand these dynamics in longer
timeframes.
Both AgNPs had a significant effect on cell membrane
integrity in C. reinhardtii but not in S. leopoliensis while neither particle showed a significant effect on ROS generation.
All silver species affect the composition of EPS with a concentration dependent response to AgNO3 and AG1 and an inverse effect to AG2.
AG1 NPs used in this study are used in industry and act as
a useful example of particles likely to be released into the environment. Whilst little difference in AG1 toxicity compared
This journal is © The Royal Society of Chemistry 2016
Paper
with AG2 after correction occurred, behaviour and effects of
industry standard particles must be measured to accurately
model NP toxicity in natural environments. With increasing
production levels of engineered nanomaterials, concentrations in the environment are expected to rise.
Currently, estimations of silver concentrations in European surface waters are in the range of 0.01–127 ng Ag
L−1,7,8,11 over two orders of magnitude lower than the EC20
values calculated in this study. Whilst this indicates that it is
unlikely that high acute toxicity will occur to green algae and
cyanobacteria at modelled concentrations these are averages.
It may be that actual concentrations are much higher or
lower locally and as such similar concentrations should be
further assessed in future studies in both laboratory and realistic environments. In realistic environmental scenarios, the
EPS produced by aquatic microorganisms likely play a large
role in mitigating the toxic effect of both ionic and particulate silver and the effect of toxicants on production as well as
the countering effect of EPS on the form of the toxicant is a
relationship which warrants further study.
Finally, the silver losses noted in this study indicate the
necessity of analytical verification of samples and is important to consider during the nanomaterial production process
in particular as silver levels will affect the values that are input into risk assessment and will affect the calculation of
safety factors.
Acknowledgements
The authors are grateful for funding through: the EU-project
NanoFate (Nanoparticle Fate Assessment and Toxicity in the
Environment), NMP4-SL-2010-24773, the SNSF Ambizione
grant PZ00P2_142533 to A. Kroll, FP7-PEOPLE-2011-IEF Micronanotox (PIEF-GA-2011-303140) to M. Matzke. The authors declare that they have no conflict of interest.
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