Biochem. J. (2012) 441, 813–821 (Printed in Great Britain)
813
doi:10.1042/BJ20111252
Induction of oxidative stress, lysosome activation and autophagy
by nanoparticles in human brain-derived endothelial cells
Blanka HALAMODA KENZAOUI*, Catherine CHAPUIS BERNASCONI*, Seher GUNEY-AYRA* and
Lucienne JUILLERAT-JEANNERET†1
University of Lausanne (UNIL), Rue du Bugnon 25, CH-1011 Lausanne, Switzerland, and †Centre Hospitalier Universitaire Vaudois (CHUV), Rue de Bugnon 21, CH-1011, Lausanne,
Switzerland
Different types of NPs (nanoparticles) are currently under
development for diagnostic and therapeutic applications in
the biomedical field, yet our knowledge about their possible
effects and fate in living cells is still limited. In the present
study, we examined the cellular response of human brainderived endothelial cells to NPs of different size and structure:
uncoated and oleic acid-coated iron oxide NPs (8–9 nm core),
fluorescent 25 and 50 nm silica NPs, TiO2 NPs (21 nm mean
core diameter) and PLGA [poly(lactic-co-glycolic acid)]-PEO
[poly(ethylene oxide)] polymeric NPs (150 nm). We evaluated
their uptake by the cells, and their localization, generation of
oxidative stress and DNA-damaging effects in exposed cells.
We show that NPs are internalized by human brain-derived
endothelial cells; however, the extent of their intracellular uptake
is dependent on the characteristics of the NPs. After their
uptake by human brain-derived endothelial cells NPs are
transported into the lysosomes of these cells, where they enhance
the activation of lysosomal proteases. In brain-derived endothelial
cells, NPs induce the production of an oxidative stress after
exposure to iron oxide and TiO2 NPs, which is correlated with
an increase in DNA strand breaks and defensive mechanisms that
ultimately induce an autophagy process in the cells.
INTRODUCTION
Oxidative stress may induce DNA damage, lysosomal activation
and autophagy, which may also represent a feedback mechanism
to limit ROS-mediated cell activation by removing oxidatively
damaged molecules and cell structure [7]. Autophagy is the only
cell process able to degrade large components, therefore it can be
postulated that NPs may be processed by cells using autophagy.
NPs-mediated autophagy may be an adaptive cellular response,
aiding in the degradation and clearance of nanomaterials, but
may also cause harmful cellular dysfunction. Some previous
publications have described NPs-related autophagy processes [8–
14], but none has compared the autophagy-inducing properties
with the oxidative stress-inducing properties of several NPs, and
none have evaluated these processes in brain vascular cells.
The BBB (blood–brain barrier) separating the brain from the
bloodstream is represented by very specialized endothelial cells.
The presence of tight junctions, of drug resistance mechanisms
and of detoxification systems at the level of this vascular system
protects the brain not only from the uncontrolled entry of bloodborne chemicals, but also from the entry of therapeutics, including
NPs [16,17]. Thus, depending on the context, the potential of NPs
to be taken up by brain-derived endothelial cells and the cellular
consequences of such interactions may induce dysfunction of
these cells important for brain functions and homoeostasis.
As the chemical composition, the solubility of the components
and surface reactivity, such as surface charges and redox NP
surface state are important for NP uptake by cells, cytotoxicity
and oxidative stress induction, different types of NPs need to be
compared with defined consequences of their interactions with
cells. In the present work, we compared the cell response of
New diagnostic and therapeutic technologies have been developed
in nanomedicine to improve the detection and treatment of human
diseases. Therefore it is becoming necessary to better understand
the mechanisms of interaction of nanomaterials, including
NPs (nanoparticles), with living tissues in order to assess the
biological consequences associated with nanotechnologies.
The physicochemical and biochemical properties of the NPs, like
surface properties, charge, size or the adsorption of biological
components, are important factors mediating their interactions
with cells, including cell-stress reactions and the biological
characteristics of the particular cells [1,2]. Two types of NPs,
either ‘solid core’ NPs, such as metal oxide-based NPs, or ‘soft
core’ NPs, such as polymer-based NPs, are suitable for therapeutic
use, chemically functionalized polymer-coated solid (metallic)
core NPs and polymer–copolymer NPs encapsulating therapeutic
agents. NPs may be internalized by cells and the solid core of
these NPs or their polymer components may be found in different
cellular localization.
The cellular reaction to NP uptake may result in the
modification of cellular functions, such as repression/activation
of genes and activation of specific cell organelles [3]. Cellular
stress, due to redox imbalance, is also an important expression
of cytotoxicity, which will occur at early stages of interaction.
Cellular oxidative stress and the production of ROS (reactive
oxygen species) upon cell exposure to NPs have previously
been shown to be a common property of NPs, relevant to
assess their potential negative effects on cell functions [2,4–7].
Key words: autophagy, DNA damage, lysosome, nanoparticle,
oxidative stress.
Abbreviations used: aminoPVA, poly(vinyl alcohol/vinylamine); BBB, blood–brain barrier; carboxy-H2 DCFDA, 5/6-carboxy-2,7-dichlorodihydrofluorescein diacetate; CNS, central nervous system; DAPI, 4 ,6 -diamidino-2-phenylindole; DLS, dynamic light scattering; DMEM, Dulbecco’s modified
Eagle’s medium; FBS, fetal bovine serum; HBSS, Hanks balanced salt solution; HCEC, human cerebral endothelial cell; MTT, 3-(4,5-dimethylthiazol-2yl)-2,5-diphenyl-2H -tetrazolium bromide; NEM, N -ethylmaleimide; NP, nanoparticle; PEO, poly(ethylene oxide); PLGA, poly(lactic-co-glycolic acid); PVA,
poly(vinyl alcohol); ROS, reactive oxygen species; TEM, transmission electron microscopy; USPIO, ultrasmall superparamagnetic iron oxide.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2012 Biochemical Society
814
Table 1
B. Halamoda Kenzaoui and others
Primary characteristics of the NPs
NPs
Core size*† (nm)
Hydrodynamic size† (nm)
ζ -potential‡ (mV)
Initial concentration (mg/ml)
Uncoated USPIO
Oleic acid-coated USPIO
TiO2
Fluorescent silica
8
8
21
25
50
143
–
14–15
–
–
–
–
+ 15 (10 mM NaCl); − 23 (DMEM)
− 30
− 30
− 40
− 40
− 39
18§
206§
2
17
12
10
PLGA-PEO
*Size determined by TEM.
†Size determined by DLS.
‡ζ -potential is provided at pH 7 unless otherwise specified.
§Determined by quantitative Prussian Blue assay [3,8].
Determined by thermogravimetric analysis.
human brain-derived endothelial cells as models of the BBB with
exposure to iron-, silica- and titanium-oxide-based solid core NPs
and one polymeric NP as models for NPs with different properties.
We show that defined NPs induced an oxidative stress, DNA
damage, the activation of lysosomal proteases and an autophagy
reaction in these cells, which we hypothesize to be a defensive
mechanism of the cerebral endothelial cells to protect the brain.
EXPERIMENTAL
NPs
All NPs used in the present study, with the exception
of aminoPVA [poly(vinyl alcohol/vinylamine)]-coated USPIO
(ultrasmall superparamagnetic iron oxide) NPs [18], are
commercially available and were provided with physicochemical
characterization by the provider (Table 1). Some further
characterization of the commercial NPs was performed (D.
Bilanicova and G. Pojana, University of Venice, Italy; and M.
Dusinska and the NanoTEST Consortium), when requested for
the present study. AminoPVA-coated USPIO NPs were prepared
and characterized according to the protocol previously described
[18]. Briefly, ferrofluid was prepared by alkaline co-precipitation
of ferric and ferrous chlorides, refluxed in nitric oxide-ferrous
nitrate and dialysed, providing iron oxide NPs (ferrofluid) of 9 nm.
To obtain aminoPVA-USPIO NPs, the ferrofluid was mixed with
PVA [poly(vinyl alcohol)] and aminoPVA at a ratio of polymer
to iron of 10 and a ratio of PVA to aminoPVA copolymer of 45
(mass ratios) [18], resulting in USPIO NPs with a hydrodynamic
diameter of 25–30 nm and a positive ζ -potential of + 25 mV.
Uncoated USPIO NPs (Fe3 O4 , uncoated USPIO NPs) were
obtained from PlasmaChem as a ∼3 % nanosuspension in water,
average NP size 8 +
− 3 nm [as determined by DLS (dynamic light
scattering), intensity (Z) average], ζ -potential + 15 mV in 10 mM
NaCl and − 23 mV in DMEM (Dulbecco’s modified Eagle’s
medium). The iron content was determined to be 18 mg iron/ml
by quantitative Prussian Blue reaction according to a previously
described protocol [18]. Oleic acid-coated USPIO NPs (Fe3 O4 ,
3 % oleic acid coating) were obtained from PlasmaChem as
an ∼7 % nanosuspension in water, average particle iron oxide
core size 8 +
− 3 nm, hydrodynamic size 14–15 nm (determined
by DLS) and ζ -potential − 30 mV at pH 7. The iron content
was determined to be 206 mg iron/ml by quantitative Prussian
Blue reaction. TiO2 NPs (Aeroxide® TiO2 P25) was obtained
from Evonik Degussa and were provided by M. Whelan, JRC,
Ispra, average particle size 21 nm, large dispersion size and
ζ -potential − 30 mV. The size distribution of the solid core
NPs was controlled by TEM (transmission electron microscopy).
c The Authors Journal compilation c 2012 Biochemical Society
Polymeric NPs consisting of a mixture of PLGA [poly(lacticco-glycolic acid)] and PEO [poly(ethylene oxide)] (PLGA-PEO
NPs) were obtained from Advancell Nanosystems as a 10 mg/ml
suspension in water, average particle size 143 nm and ζ -potential
− 39 mV. Fluorescent rhodamine-labelled silica nanospheres
(25 nm silica NPs and 50 nm silica NPs) were obtained from
Corpuscular as a 25 mg/ml suspension in water [17 and 12 mg/ml
respectively as measured by thermogravimetric analysis by H.
Hofmann and P. Bowen (EPFL, Lausanne, Switzerland] with a
ζ -potential of − 40 mV. The size of the two coated aminoPVAUSPIO NPs [18] and of oleic acid-coated USPIO NPs are of the
order of 35 nm in cell culture medium and the size of uncoated
USPIO NPs and TiO2 NPs was determined in cell culture medium
to be 910 and 650 nm respectively by DLS, with large size
dispersion. The size of other NPs did not notably change in the
cell culture medium.
Preparation of NP suspensions
Uncoated and oleic acid-coated USPIO NPs in suspension in
water were diluted in PBS to obtain a final concentration of 2 mg
of iron/ml. Immediately before use aliquots were vortex-mixed
for 1 min and added to cell culture medium to obtain a 75 μg
of iron/cm2 working solution, then serially diluted in the cell
culture medium. Stock suspensions of TiO2 NPs were made as a
suspension of 2 mg/ml in culture media without FBS (fetal bovine
serum) containing 15 mM Hepes, then sonicated for a total of
3 min (9×20 s) at 4 ◦ C and 60 W, vortex-mixed for 10 s, then
stored at − 20 ◦ C, according to a previously described protocol
[19]. Immediately before use the NP suspension was thawed,
vortex-mixed for 10 s, sonicated for a total of 1 min (3×20 s) at
4 ◦ C and 60 W [18], added to the cell culture medium to achieve
a 75 μg/cm2 working solution and then serially diluted. PLGAPEO NPs in suspension in water were diluted in the appropriate
cell culture medium to obtain a stock solution of 75 μg/cm2 , then
serially diluted in the cell culture medium. Fluorescent 25 nm and
50 nm silica-NPs in suspension in water were vortex-mixed for
a few minutes, diluted in the appropriate cell culture medium to
obtain a stock solution of 75 μg/cm2 , then serially diluted in the
cell culture medium.
Cell lines and culture conditions
HCECs (human cerebral endothelial cells) were a gift from
Professor D. Stanimirovic, Faculty of Medicine, University of
Ottawa, Ottawa, Ontario, Canada, and were grown in DMEM,
containing 4.5 g/l glucose, 10 % FBS and penicillin/streptomycin.
All cell culture reagents were purchased from Gibco and
NP-induced stress in cerebral endothelial cells
Invitrogen. Cells were exposed to NPs in 250 μl of complete
culture medium in 48-well plates (Costar) at decreasing
concentrations, from 235 μg/ml equivalent to 75 μg/cm2 NPs
to 0.4 μg/ml equivalent to 0.12 μg/cm2 NPs. Experiments
were performed in three wells at least twice in independent
experiments.
Evaluation of cell viability
Cells were grown in 48-well cell culture plates (Costar) until
75 % confluent, then exposed to NPs for the concentration
and time indicated, then washed in saline. Cell viability
was evaluated using the MTT [3-(4,5-dimethylthiazol-2-yl)-2,5diphenyl-2H tetrazolium bromide] assay (Sigma–Aldrich) added
to the cells in fresh complete culture medium at a 250 μg/ml
final concentration. After 2 h the supernatant was removed and
the precipitated formazan was dissolved in 0.1 M HCl in propan2-ol and quantified at 540 nm in a multiwell plate reader (iEMS
Labsystems, BioConcepts).
Evaluation of DNA synthesis
Briefly, following exposure of cells to NPs for 48 h, [3 H]thymidine
(GE Healthcare, 400 nCi/ml final concentration) was added to the
cells for 2 h. Then the cell layers were precipitated with 10 %
trichloroacetic acid and dissolved in 0.1 % SDS in 0.1 M NaOH
and scintillation cocktail (Optiphase HI-Safe, PerkinElmer) was
added. Radioactivity was counted with a β-counter (WinSpectra).
The radioactivity counts of treated cells were compared with
the radioactivity counts of untreated cells. Experiments were
conducted in triplicate wells and repeated twice. Means +
− S.D.
were calculated.
Evaluation of ROS production
Free radical formation was determined with 5/6-carboxy2,7-dichlorodihydrofluorescein. Cells were grown until confluent in 48-well plates (Costar), then preincubated for
40 min with carboxy-H2 DCFDA (5/6-carboxy-2,7-dichlorodihydrofluorescein diacetate, Molecular Probes, Invitrogen, stock
solution 25 mg/ml in DMSO, final concentration 20 μM), washed
and incubated at 37 ◦ C with the NPs diluted in enriched HBSS
(Hanks balanced salt solution; containing 1.3 mM Ca2 + , 1.1 mM
Mg2 + and 5 mM glucose, from Gibco). Then 5 mM t-butylH2 O2 (Sigma) was added as a positive control for free radical
formation. Then the plate was read at λex /λem = 485/530 nm in a
thermostatically controlled fluorescence plate reader (CytoFluor
Series 4000, PerSeptive Biosystems) at t = 0 h, then every hour
for 4 h.
Fluorescence microscopy
Cells were grown for 24 h on glass slides (Menzel-Gläser). Then
the medium was changed, and the NPs diluted in complete culture
medium were added at the desired concentrations for a further 24 h
(uptake of silica NPs) or only 4 h (oxidative stress evaluation). At
the end of the treatment, the cell layers were washed in PBS, fixed
in 4 % PFA (paraformaldehyde) for 20 min at 4 ◦ C, washed in PBS
and incubated with DAPI (4 ,6 -diamidino-2-phenylindole; Roche
Diagnostics; 1 μg/ml in PBS) for 30 min at 37 ◦ C. For oxidative
stress evaluation, the incubation with DAPI was preceded by
carboxy-H2 DCFDA staining (20 μM) for 30 min at 37 ◦ C. The
slides were washed with PBS and mounted in 20 % glycerol in
PBS, then examined by fluorescence microscopy.
815
Determination of cellular thiols
The monobromobimane assay was used to measure cellular thiol
levels. Cells were grown until half-confluent in 48-well plates and
exposed for 4, 24 or 48 h to the NPs, washed with HBSS, then
250 μl/well of 100 μM monobromobimane (Sigma–Aldrich) in
HBSS was added at room temperature (22 ◦ C) for 5 min in the
dark. The cell layers were washed with HBSS and lysed with
0.1 % Triton X-100 (Sigma–Aldrich) in HBSS. For a positive
control, cells were exposed to 100 μM NEM (N-ethylmaleimide,
Sigma–Aldrich) for 1 min before the assay. The fluorescence was
immediately read in a thermostatically controlled fluorescence
plate reader (CytoFluor) at λex /λem = 485/580 nm.
Evaluation of DNA damage
Induction of DNA strand breaks by the NPs in HCECs was
determined using the alkaline single-cell gel electrophoresis
(Comet assay) [20]. Briefly, cells were grown for 24 h in a 24well plate (Costar), then exposed to the NPs for the concentration
and time indicated. Cells were detached using trypsin/EDTA
(TrypLE Express, Invitrogen) and the cell suspension (0.8×106
cells/ml) in 1 % low-melting-point agarose (Sigma–Aldrich) was
deposited on glass slides coated with 1 % normal-melting-point
agarose (Eurobio). After 5 min at 4 ◦ C, to allow solidification
of the cell layer, slides were immersed in lysis buffer (2.5 M
NaCl, 100 mM EDTA, 10 mM Tris-base and 1 % Triton X-100,
pH 10.0) for 1 h at 4 ◦ C, then a 40 min unwinding process at
4 ◦ C and electrophoresis at 25 V for 30 min were performed, both
under alkaline conditions (300 mM NaOH and 1 mM EDTA,
pH 13.0). Following washing with ice-cold PBS and ice-cold
nanopure water, slides were stained with 1 μg/ml DAPI (Roche)
in PBS. Cells treated with 100 μM H2 O2 for 5 min at 4 ◦ C
were used as a positive control. Analysis of Comet appearance
was performed with a Zeiss Axioplan 2 Imaging microscope
at a ×200 magnification and λex /λem = 365/420 nm. For each
experiment, 100 cells were analysed twice randomly per treatment
using the Comet assay image analysis software (Comet Visual,
NILU, Health Effects Laboratory). Cells were scored visually and
assigned to one of five classes according to the Comet tail size
(from undamaged 0 to maximally damaged 4) giving expression
in arbitrary units from 0 (completely undamaged) to 400 (with
maximal damage) [20].
Western-blot experiments
Cells were grown in 10-cm-diameter Petri dishes and exposed
to the NPs for the indicated time and concentration. After
treatment, the cell layers were washed with ice-cold PBS
and lysed in 200 μl of lysis buffer (150 mM NaCl, 2 mM
EDTA, 0.5 % Triton X-100, 50 mM Tris/HCl, 2 mM sodium
orthovanadate and 50 mM NaF, pH 7.2) and 10 μl of protease
inhibitor cocktail (Sigma–Aldrich), scraped with a cell scraper,
extracted by three cycles of freeze/thawing and centrifuged at
10 000 rev./min at 4 ◦ C for 10 min on an Eppendorf 5415R
centrifuge. Supernatants were applied to SDS/PAGE and
transferred on to a nitrocellulose membrane (Whatman). The
membranes were blocked with 5 % non-fat dried skimmed
milk powder in PBS, washed in 0.05 % Tween 20 (Sigma–
Aldrich) in PBS and incubated overnight at 4 ◦ C with anti-human
LC3 (Novus Biologicals; diluted 1:2000 in 1 % non-fat dried
skimmed milk powder and 0.05 % Tween 20 in PBS), anticathepsin B (BioVision; diluted 1:5000), anti-cathepsin D (H-75)
(Santa Cruz Biotechnology; diluted 1:2000) or anti-SQSTM1/p62
(Cell Signaling Technologies; diluted 1:3000) rabbit polyclonal
c The Authors Journal compilation c 2012 Biochemical Society
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B. Halamoda Kenzaoui and others
antibodies, and then exposed for 60 min to horseradish
peroxidase-conjugated anti-rabbit antibody (Promega; diluted
1:5000) and visualized using chemoluminescence (ECL® ,
GE Healthcare). Transferrin receptor/CD71 was determined
with the goat polyclonal anti-CD71 antibody (Santa Cruz
Biotechnology; diluted 1:3000), and then exposed for 60 min
to horseradish peroxidase-conjugated anti-goat antibody (Sigma;
diluted 1:10 000) and visualized using chemoluminescence. To
control for loading, the membranes were stripped by successive
incubation in 0.1 M glycine, pH 2.3, 1 M NaCl in PBS and 0.05 %
Tween 20 in PBS, blocked for 1 h with 5 % non-fat dried skimmed
milk powder in PBS and exposed to a monoclonal anti-chicken
α-tubulin mouse antibody (Abcam; diluted 1:5000) for 1 h at
room temperature followed by 1 h incubation with horseradish
peroxidase-conjugated anti-mouse antibody (Sigma) and treated
as described above. For densitometric analysis, the band areas
of cathepsin B, cathepsin D and α-tubulin were evaluated using
ImageJ software (NIH), then the ratio of the area of the cathepsin
band to the α-tubulin band was calculated.
TEM
Cells were grown in 10-cm-diameter Petri dishes (Nunclon) until
70–80 % confluent, then exposed to NPs for the concentration
and time indicated. At the end of the incubation, the cells
were washed twice with PBS, detached from the Petri dishes
in trypsin/EDTA and centrifuged, then the cell pellets were
fixed in 2.3 % cacodylate-buffered glutaraldehyde (Sigma) for
up to 24 h, washed in cacodylate buffer and mixed with
4 % low-gelling agarose (Sigma). Samples were post-fixed in
1.3 % osmium tetroxide in 0.2 M cacodylate buffer, pH 7.4,
for 1 h and dehydrated in graded ethanol, then in propylene
oxide and embedded in 50 % (w/w) epoxy embedding medium,
26 % (w/w) DDSA (dodecenylsuccinic anhydride), 23 % (w/w)
MNA (methyl nadic anhydride), 1 % (w/w) DMP-30 [2,4,6tris(dimethylaminomethyl)phenol] (all from Fluka and Sigma–
Aldrich). Blocks were cured 48 h at 60 ◦ C, thin sections (70–
80 nm) were cut using an ultramicrotome (Ultracut E, ReichertJung Optische Werke AG) and mounted on 3-mm 200-mesh
copper grids. Grids were stained for 75 min in saturated uranyl
acetate solution (Fluka), then for 100 s in lead citrate (Ultrostain
2, Laurylab), examined and photographed with a Philips CM10
transmission electron microscope combined with a MegaView III
Soft Imaging system to evaluate USPIO NPs associated to cells.
Calculation of results
Each experiment was repeated in three wells at least twice.
Means +
− S.D. were calculated and statistical significance was
evaluated using a Student’s t test.
USPIO NPs, but neither released nor transported them [28],
suggesting an intracellular processing of such NPs and their
retention in the cells. Previous studies using cells from a
different origin and iron oxide-based NPs have demonstrated
lysosomal localization, reorganization of cell cytoskeleton and
intracellular iron-mediated increase in the production of ROS
[29,30]. Therefore the present study focused on the mechanisms
of interactions between NPs with different physicochemical
properties and human cells representative of the BBB, analysing
the cellular consequences of these interactions, focusing on the
generation of oxidative stress and cellular degradative pathways.
TiO2 NPs and uncoated USPIO NPs very rapidly agglomerate
in culture medium containing FBS, resulting in agglomerates
of large dispersion size, average 850 nm [31] and 910 nm
respectively as determined by DLS (M. Dusinska and the
NanoTEST Consortium) and electron microscopy (results not
shown), providing information for agglomerated NPs with two
different physicochemical characteristics. Uncoated and oleic
acid-coated USPIO NPs with similar cores allowed comparison
of NPs with different surface properties. Silica NPs with different
sizes, 25 and 50 nm, allowed evaluation of the size effects of
NPs with similar composition, whereas polymeric NPs were
considered as control NPs not inducing a cellular reaction.
Internalization of NPs by HCECs
After 24 h incubation of the cells with the NPs their uptake
was studied with methods adapted to the NP type: fluorescence
microscopy for fluorescent silica NPs, the histological Prussian
Blue [28] reaction for USPIO NPs and TEM for inorganic
solid core (iron oxide, TiO2 and silica) NPs to analyse the
intracellular localization of solid core NPs inside cell organelles.
The cell uptake of silica NPs was observed only at the highest
concentration tested (75 μg/cm2 ) and the uptake of 25 nm NPs
was slightly higher than the uptake of 50 nm NPs (Figure 1A).
Uncoated USPIO NPs agglomerated in the cellular environment
which resulted in their rapid deposition on the cell surface. The
internalization of these NPs as determined by Prussian Blue
reaction (results not shown) was much higher than the uptake of
oleic-acid-coated USPIO NPs, which were stable in cell culture
medium and were only poorly internalized by endothelial cells,
as previously shown [28]. The intracellular presence of the solid
cores of iron oxide, TiO2 and silica NPs was confirmed by
TEM (Figures 1Ba–1Bf). The NPs were localized in small or
large endosomes in the cytoplasm and in lysosomes. Therefore
TEM and fluorescence microscopy demonstrated the intracellular
localization of solid core NPs in cell organelles, most probably
the endosomal–lysosomal compartment.
RESULTS AND DISCUSSION
Evaluation of cell viability and DNA synthesis by HCECs
exposed to the NPs
Previously published studies have suggested that following
intravenous injection NPs can be trapped in brain vessels, possibly
being biodegraded inside brain endothelial cells [17,21–24]. We
have previously shown that biocompatible cationic USPIO NPs
were internalized by cells [18], even when functionalized with
a fluorescent reporter [25] or with anti-cancer drugs [26,27].
Their final cellular localization in human melanoma cells was
the lysosomes, where they started to dissolve, inhibiting the
expression of the transferrin receptor and activating the lysosomal
protease cathepsin D [3]. We have also previously shown that
human brain-derived endothelial cells internalized these cationic
To evaluate for potential cytotoxicity, the metabolic activity of
HCECs after 72 h exposure to the NPs was evaluated using the
MTT assay (Figure 2). At the highest concentration, oleic acidcoated USPIO NPs and 25 nm fluorescent silica NPs (Figures 2A
and 2B) induced a decrease in the metabolic activity of HCECs,
whereas uncoated USPIO NPs and 50 nm silica NPs were less
cytotoxic. Acute cytotoxicity was already observed after 24 h
exposure to high doses (75 μg/cm2 ) of oleic acid-coated USPIO
NPs (results not shown). Exposure of cells to TiO2 NPs induced a
significant cytotoxicity for HCECs (Figure 2C). PLGA-PEO NPs
were not cytotoxic at any of the concentrations tested.
c The Authors Journal compilation c 2012 Biochemical Society
NP-induced stress in cerebral endothelial cells
Figure 1
817
Uptake of the NPs by HCECs
(A) Cells were grown to 75 % confluence, then were exposed for 24 h in complete culture
medium to 25 or 50 nm rhodamine-labelled fluorescent silica NPs (75 μg of NPs/cm2 ). The
uptake by the cells of the NPs was observed by fluorescence microscopy. Nuclei are stained blue
with DAPI and silica NPs fluoresce red. (B) Cells were grown to 75 % confluence, then were
exposed for 24 h in complete culture medium to 15 μg/cm2 USPIO NPs, 75 μg/cm2 silica NPs
or 15 μg/cm2 TiO2 NPs. The presence inside HCECs of the solid cores of the NPs was observed
by TEM. (a) Uncoated USPIO NPs; (b) oleic acid-coated USPIO NPs; (c) 25 nm fluorescent
silica NPs; (d) 50 nm fluorescent silica NPs; (e and f) TiO2 NPs. Clusters of NPs in endosomes
(white arrows) or in lysosomes (black arrow) are indicated.
The synthesis of DNA by HCECs was also determined after
48 h exposure of the cells to NPs. A significant decrease in
DNA synthesis was observed after cell exposure to high doses
of oleic acid-coated USPIO NPs and 25 nm silica NPs (Figure 3).
Exposure to uncoated USPIO NPs and TiO2 NPs caused a
slight decrease in DNA synthesis in a dose-dependent manner.
PLGA-PEO NPs and 50 nm silica NPs had no effect on DNA
synthesis by exposed cells (results not shown).
Effect of NPs on oxidative stress generation and DNA damage
in HCECs exposed to the NPs
The mitochondria content of endothelial cells at the BBB is higher
than for endothelial cells of another location, rendering them
more susceptible to generation of oxidative stress. Several studies
have demonstrated that the generation of ROS and oxidative
stress are the key mechanism by which NPs exert deleterious
effects [6], including oxidative-stress-mediated entry of NPs into
oxidation-damaged endothelial cells of cerebral microvessels
[32]. NPs may induce oxidative stress and/or directly interact
with the endosomal–lysosomal pathway, two possibilities that
we evaluated in the experiments reported in the present paper.
The production of ROS by HCECs exposed to the NPs was first
measured using carboxy-H2 DCFDA, detecting the intracellular
presence of H2 O2 (Figure 4). After 4 h of cell exposure [which we
had determined as the optimal time for the determination of ROS
Figure 2
Cytotoxicity of the NPs for HCECs
The MTT assay was performed after 72 h exposure of HCECs to increasing concentrations of
(A) 䉬 uncoated USPIO NPs and 䊐 oleic acid-coated USPIO NPs; (B) 䉬 25 nm and 䊐 50 nm
silica NPs; (C) 䉬 PLGA-PEO NPs and 䊐 TiO2 NPs.
production (results not shown)], to uncoated USPIO NPs, oleic
acid-coated USPIO NPs and TiO2 NPs, a significant increase
in ROS production was detected by quantitative fluorescence
measurements (Figure 4A), which was confirmed by fluorescence
microscopy (Figure 4B).
Then, the content in thiols of HCECs was determined after 4,
24 or 48 h exposure of the cells to the NPs using the bromobimane
assay (Figure 5). A more pronounced dose-dependent decrease in
thiol levels was observed after cell exposure to uncoated USPIO
c The Authors Journal compilation c 2012 Biochemical Society
818
Figure 3
B. Halamoda Kenzaoui and others
Effect of NPs on DNA synthesis by HCECs
DNA synthesis by HCECs was measured using thymidine incorporation after 48 h exposure of
the cells to either uncoated USPIO NPs (䉬), oleic acid-coated USPIO NPs (䊐), 25 nm silica
NPs () and TiO2 NPs (䉱), then the incorporation of radioactive thymidine was performed for
the last 2 h.
Figure 5
Determination of the thiol content of HCECs exposed to NPs
HCECs were exposed to the NPs for either 4 h (white bars), 24 h (light grey bars) or 48 h
(black bars) exposure, then the bromobimane assay was performed to determine thiol levels in
the cells. Cells were treated with 100 μM NEM (c + ) to deplete the cells of thiols. Statistical
significance was assessed by a Student’s t test; *P 0.05, **P 0.01, ***P 0.001, exposed
cells compared with control cells.
Figure 4
ROS production by HCECs exposed to NPs
Figure 6
Evaluation of DNA-damage induced by NPs in HCECs
(A) The formation of free radical was determined by carboxy-H2 DCFDA in HCECs exposed for 4 h
to either uncoated USPIO NPs (left) or TiO2 (right) NPs. TBHP (t-butyl-hydroperoxide) (5 mM)
was used as a positive control (c + ). Statistical significance was assessed by a Student’s t test;
*P 0.05, exposed cells compared with control cells. (B) HCECs were exposed for 4 h to NPs
(15 μg of NPs/cm2 ) or TBHP (200 μM) as a positive control, then fixed and stained with DAPI
(blue) and carboxy-H2 DCFDA (green) and photographed. (a) Non-treated cells, (b) uncoated
USPIO NPs, (c) oleic acid-coated USPIO NPs, (d) PLGA-PEO NPs, (e) TiO2 NPs and (f) TBHP.
HCECs were exposed to increasing non-cytotoxic concentrations of either uncoated iron oxide
NPs, oleic acid-coated USPIO NPs or TiO2 NPs for 24 h (grey bars) or 48 h (black bars), then
the Comet assay was performed to evaluate DNA strand breaks. Cells treated with 100 μM H2 O2
during 5 min were used as a positive control (c + ). Statistical significance was assessed by a
Student’s t test; *P 0.05, exposed cells compared with control cells.
NPs, already apparent after 4 h, than to oleic acid-coated USPIO
NPs (results not shown). As exposure of cells to 75 μg/cm2 oleic
acid-coated USPIO NPs induced almost complete cytotoxicity
(Figure 2A) we limited evaluation of this particular NP to the
highest non-cytotoxic concentration of 30 μg/cm2 . PLGA-PEO
and 50 nm silica NPs did not affect cellular thiol content after 4 h
(results not shown), 24 or 48 h, whereas TiO2 and 25 nm silica
NPs induced a slight decrease in cellular thiols after 24 and 48 h
at the highest concentration tested (Figure 5).
A potential DNA-damaging effect of NPs in HCECs was
measured by the alkaline single-cell gel electrophoresis (Comet
assay) [20] detecting single and double DNA strand breaks and
alkali-labile sites (Figure 6). Significant DNA damage was found
after 24 and 48 h of cell exposure to both uncoated and oleic acidcoated USPIO NPs and this effect was even more pronounced
for TiO2 NPs. Thus we show that defined NPs induced ROS
production by brain-derived endothelial cells, decreasing the
levels of thiols in the cells and increasing DNA damage.
c The Authors Journal compilation c 2012 Biochemical Society
NP-induced stress in cerebral endothelial cells
Figure 7
TEM images of autophagic vacuoles in HCECs exposed to NPs
After 48 h exposure of HCECs to uncoated USPIO NPs (a, b), TiO2 NPs (c, d) and 50 nm silica
NPs (e, f), the cells were examined by TEM. Black arrows, NPs clusters. *Denotes autophagic
vacuoles with double-layered membranes containing cellular debris; @ denotes autolysosomes
with partly degraded content, some of them possibly containing NPs.
Autophagy induction and lysosomal activation by the NPs
in HCECs
Autophagy is a process by which cells consume their own
components in lysosomes and is the only cell process able to
degrade large components, therefore it can be postulated that
NPs may be processed by autophagy. There are several forms
of autophagy, but all involve the delivery of cell components to
lysosomes in response to sublethal cell stress, such as oxidative
stress or DNA damage, protein aggregates, particulate structures,
mitochondrial damage or pathogens [33,34]. ROS activation
of autophagy may represent a feedback mechanism to limit
ROS-mediated cell activation by removing oxidatively damaged
molecules and cell structure [7]. NP-mediated autophagy may
be an adaptive cellular response, aiding in the degradation
and clearance of nanomaterials, but may also cause harmful
cellular dysfunction. Some previous publications have described
NP-related autophagy processes [8,14,15,34–36], but none has
compared the autophagy-inducing properties with the oxidative
stress-inducing properties of several NPs. For example, gold
NPs induced oxidative stress and autophagy in human fibroblasts
[35]. The reports of NP-mediated induction of autophagy in the
endothelium, in particular in cerebral endothelial cells, are very
scarce. Pollution NPs have been associated with disruption of the
BBB [37] and fullerene NPs were shown to induce autophagic
vacuolization of HUVECs (human umbilical vein endothelial
cells) [36].
During autophagy, a small flattened vesicle forms in the
cytoplasm and grows to a cup-shaped isolation membrane,
engulfing the materials to be degraded, then closing to form
a double-membrane autophagosome that fuses with lysosomes.
Cellular structures presenting the characteristics of an autophagic
response of the cells to the NPs as well as lysosomal involvement
were observed by TEM in HCECs following exposure to the
NPs (Figure 7). The presence of characteristic double membrane
vacuoles containing cellular material indicated the formation of
autophagosomes. After the fusion with lysosomes they become
autolysosomes able to degrade cellular material. Autolysosomes
were observed in NP-exposed cells (Figures 7a–7d and 7f),
some of them containing NPs (Figure 7f). Generalized high
vacuolization of cells was another autophagy characteristic that
could be observed in cells exposed to the NPs (Figure 7e),
in particular in cells exposed to 50 nm silica NPs. Thus we
819
demonstrated in cells exposed to NPs the appearance of cellular
structures presenting the characteristics of an autophagic response
of the cells as well as lysosomal involvement.
The formation of autophagosomes involves complex protein
pathways, one of which includes the conjugation of an ubiquitinlike protein, LC3, that is cleaved at its C-terminal by a ROSsensitive thiol protease to release LC3-I exposing a glycine residue
that is then conjugated to the lipid PE (phosphatidylethanolamine)
in an ubiquitin-like reaction to produce LC3-II. LC3-II is then
attached to the double membranes of mature autophagosomes
and maintained after their fusion with lysosomes. LC3-I into
LC3-II conversion is commonly used to monitor autophagy [38].
Both NP exposure [3] and autophagy induction also involves
the activation of the enzymes of lysosomes, including the
cathepsins. The expression of LC3-I and LC3-II, and of
the (pro)cathepsins B and D were determined in extracts
of HCECs exposed to the NPs (Figure 8A). We demonstrated NPspecific induction of LC3-II expression in HCECs, supporting
our TEM observations. NP-specific activation of procathepsins
B and D was also demonstrated (Figures 8A and 8B). In the
HCECs, LC3-II levels together with the activation of procathepsin
D were particularly enhanced in cells exposed to uncoated USPIO
NPs and TiO2 NPs. It has to be emphasized that these two NPs
are particularly prone to agglomeration in cell culture medium.
(Pro)cathepsin B is expressed at only very low levels in HCECs,
and we showed that NP-dependent induction or activation of
procathepsin B was very low for all NPs, possibly being
slightly more marked with the non-agglomerating solid-core NPs
(Figure 8B). In a time-course study to evaluate the autophagic
flux, the rate at which intracellular material is transported to the
lysosomes [39], in HCECs exposed or not exposed for 6–72 h
to uncoated USPIO NPs, we demonstrated that the appearance
of activated cathepsin D and LC3-II conversion were correlated,
starting after 48 h exposure, whereas the iron-mediated repression
of the transferrin receptor/CD71, being apparent already after 24 h
exposure, preceded cathepsin D activation and LC3-II conversions
(Figure 8C). However, NP-mediated autophagy induction seems
to be different from aggregated protein-related degradation by
autophagy, since the expression of sequestosome 1/SQTM1/p62
[38–40] was not decreased in HCECs exposed to uncoated
USPIO NPs (Figure 8C). The polyubiquitin-binding protein
sequestosome SQSTM/p62 has been shown to be degraded in
some autophagic processes, and inhibition of autophagy increased
its expression [41]. Thus we showed that defined NPs have the
potential to induce ROS generation, DNA damage or to selectively
activate procathepsins B and D to their active forms and to induce
autophagy. However, different NPs were responsible for these
effects. These effects were particularly obvious for the two NPs
that agglomerate in the culture conditions and induced oxidative
stress, i.e. uncoated USPIO NPs and TiO2 NPs. Therefore we
hypothesize that NP aggregation may be important in ROS
generation, activation of defined lysosomal cathepsins and the
NP-mediated autophagic cell reaction. However, NP-mediated
DNA damage seems to play a less relevant role, since oleic
acid-coated USPIO NPs that induce DNA damage at the same
level as uncoated USPIO NPs do not induce such an autophagic
reaction and generate much lower oxidative stress. NP size has
also been suggested to be relevant for the induction of the
autophagic process. PEG [poly(ethylene glycol)]-coated quantum
dots induction of autophagy was size-dependent in human
mesenchymal stem cells [12,42]. Rare earth oxide nanocrystals
induced autophagy and vacuolization in human cancer cells.
Vacuolization was size-dependent and actually caused by large
aggregates of NPs [10]. All of these results are in agreement with
these of the present study.
c The Authors Journal compilation c 2012 Biochemical Society
820
Figure 8
B. Halamoda Kenzaoui and others
Expression of autophagy and lysosomal markers by HCECs exposed to NPs
(A) HCECs were exposed to 30 μg/cm2 NPs for 48 h and then the expression of LC3-I/II, cathepsin B or cathepsin D was determined by Western blotting in cell extracts. (B) The ratio of the band
intensities of either cathepsin D or cathepsin B to α-tubulin has been determined. (C) HCECs were either not exposed or exposed to 30 μg/cm2 uncoated USPIO NPs for 6–72 h, then the expression
of either LC3-I/II, cathepsin D, the transferrin receptor/CD71 or SQSTM/p62 was determined by Western blotting in cell extracts.
Cathepsin D (EC 3.4.23.5) is an aspartyl lysosomal protease
expressed in all cells, synthesized in the ER (endoplasmic
reticulum) as an inactive proenzyme of 52 kDa, then activated by
intramolecular autocatalytic splitting in the acidic environment of
lysosomes into active cathepsin D, constituted of two subunits
of 34 and 14 kDa. Cathepsin B (EC 3.4.22.1) is a thiol lysosomal
protease, biosynthesized as an inactive proenzyme of 39 kDa,
which is transported into the lysosomes where acidification
triggers the activation of the enzyme. The 30 kDa single chain
active enzyme can be further processed into an active twochain form (25 and 5 kDa), linked by a disulfide bridge, by
removal of an internal dipeptide. Using a different model of
induction of apoptosis and autophagy by serum deprivation in
PC12 cells, it was previously shown that cell death was mediated
by mitochondria and involved caspase-3 activation. However,
during this process, the functions of the lysosomal cathepsins,
particularly cathepsin B and cathepsin D, were also altered,
cathepsin B decreased whereas cathepsin D increased in serumdeprived PC12 cells [43]. It was also shown that overexpression
of cathepsin D accelerated apoptosis, whereas overexpression of
cathepsin B increased cell viability. In another report of nutrient
deprivation of breast cancer cells, it was shown that inhibition
of autophagy stimulated cathepsin D expression and induced
apoptosis via cathepsin D accumulation; apoptosis was reduced by
knockdown of cathepsin D [41]. It was also previously shown
that increased cathepsin D expression may be responsible for
switching cell pathways from apoptosis to autophagy, possibly by
depleting caspase 3 [44] and salvaging the cells from cell death.
In conclusion, we have explored the mechanisms of interactions
between NPs with different biophysical characteristics and human
brain-derived endothelial cells. Taken together, our previous
results [28] and those of the present study indicate that depending
on their physicochemical characteristics, NPs may be found
in brain vascular cells, where they induce an NP-specific cell
reaction, suggesting that they may modulate brain functions
c The Authors Journal compilation c 2012 Biochemical Society
from this location. We have shown that the NPs induced the
production of ROS, were transported into lysosomes, interfering
with the lysosomal hydrolases, cathepsins D and B, and induced an
autophagy process. The observation that NPs induced autophagy
in these cells probably in response to oxidative stress and as
being recognized as agglomerated materials, suggests a defensive
mechanism of the cerebral endothelial cells to protect the brain.
These observations have important biomedical consequences.
Owing to the suspected risks of distributing deleterious chemicals,
such as metals and cations to the CNS (central nervous system),
the technologies based on NPs are designed in such a way that
they cannot get access to the CNS. Therefore the response of cells
forming the interface between the brain and the blood, the
cells of the BBB, needs to be evaluated in detail for NPs that
will be designed in such a way.
AUTHOR CONTRIBUTION
Blanka Halamoda Kenzaoui and Lucienne Juillerat-Jeanneret designed the experiments,
analysed and interpreted the results and wrote the paper; Blanka Halamoda Kenzaoui,
Catherine Chapuis Bernasconi and Seher Guney-Ayra performed the experiments and
participated in the evaluation of the results.
ACKNOWLEDGEMENTS
We thank P. Bowen and H. Hofmann from EPFL, Lausanne, for providing the
thermogravimetric analyses of the silica NPs; M. Dusinska and her team at NILU, Oslo,
for their help in the Comet assay and helpful comments; and D. Bilanicova, G. Pojana and
A. Marcomini, University of Venice, for providing physicochemical characterization data
of the NPs.
FUNDING
This work was supported by the European Community 7th Framework Programme [project
number 2007-201335 ‘NanoTEST’]; the Switzerland-France InterReg programme, project
‘NAOMI’; and the Swiss Government-University of Bern BNF programme.
NP-induced stress in cerebral endothelial cells
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Received 13 July 2011/21 October 2011; accepted 26 October 2011
Published as BJ Immediate Publication 26 October 2011, doi:10.1042/BJ20111252
c The Authors Journal compilation c 2012 Biochemical Society