Journal of Neurochemistry, 2006, 97, 69–78
doi:10.1111/j.1471-4159.2006.03718.x
High susceptibility of neural stem cells to methylmercury toxicity:
effects on cell survival and neuronal differentiation
Christoffer Tamm,* Joshua Duckworth, Ola Hermanson and Sandra Ceccatelli*
*Institute of Environmental Medicine, Division of Toxicology and Neurotoxicology, and Department of Cell and Molecular Biology,
Karolinska Institutet, Stockholm, Sweden
Abstract
Neural stem cells (NSCs) play an essential role in both the
developing embryonic nervous system through to adulthood
where the capacity for self-renewal may be important for
normal function of the CNS, such as in learning, memory and
response to injury. There has been much excitement about
the possibility of transplantation of NSCs to replace damaged
or lost neurones, or by recruitment of endogenous precursors.
However, before the full potential of NSCs can be realized, it is
essential to understand the physiological pathways that control their proliferation and differentiation, as well as the influence of extrinsic factors on these processes. In the present
study we used the NSC line C17.2 and primary embryonic
cortical NSCs (cNSCs) to investigate the effects of the environmental contaminant methylmercury (MeHg) on survival
and differentiation of NSCs. The results show that NSCs, in
particular cNSCs, are highly sensitive to MeHg. MeHg induced
apoptosis in both models via Bax activation, cytochrome c
translocation, and caspase and calpain activation. Remarkably, exposure to MeHg at concentrations comparable to the
current developmental exposure (via cord blood) of the general population in many countries inhibited spontaneous
neuronal differentiation of NSCs. Our studies also identified
the intracellular pathway leading to MeHg-induced apoptosis,
and indicate that NSCs are more sensitive than differentiated
neurones or glia to MeHg-induced cytotoxicity. The observed
effects of MeHg on NSC differentiation offer new perspectives
for evaluating the biological significance of MeHg exposure at
low levels.
Keywords: apoptosis, calpain, caspase, cell death.
J. Neurochem. (2006) 97, 69–78.
Neural stem cells (NSCs), defined by their ability to selfrenew and differentiate into the three major cell types,
neurones, astrocytes and oligodendrocytes, play an essential role in the development and maturation of the nervous
system. They are present not only in the developing brain
but also in the adult brain in different areas with
neurogenic potential (Gage 2000). The importance of
NSCs in the adult CNS is uncertain, but there is evidence
suggesting that capability for self-renewal may be important for normal brain functions, including learning, memory
and emotional responses (Santarelli et al. 2003; Schaffer
and Gage 2004).
Recently there has been much excitement about the
possibility of replacing damaged or lost neural cells by
transplantation of NSCs, or by recruitment of endogenous
precursors to repair adult brain. However, before the full
potential of NSCs can be realized, it is essential to
understand the physiological pathways that control their
proliferation and differentiation, as well as the influence that
extrinsic factors may have on these processes.
The developing nervous system is especially vulnerable to
damage by toxic agents, but so far little attention has been
focused on the effects of environmental neurotoxicants on
NSCs. A relevant environmental contaminant that is
extremely toxic to the developing nervous system is
methylmercury (MeHg). Despite major actions taken to
reduce the use and emission of mercury in the environment,
Received September 30, 2005; revised manuscript received November 7,
2005; accepted November 29, 2005.
Address correspondence and reprint requests to Sandra Ceccatelli,
Institute of Environmental Medicine, Division of Toxicology and
Neurotoxicology, Karolinska Institutet, 171 77 Stockholm, Sweden.
E-mail: [email protected]
Abbreviations used: bFGF, basic fibroblast growth factor; BSA,
bovine serum albumin; cNSC, cortical neural stem cell; Cyt c, cytochrome c; cyto, cytosolic; DEVD-AMC, Ac-Asp-Glu-Val-Asp-7-amino4-methylcoumarin; DTT, dithiothreitol; HBSS, Hanks’ Balanced Salt
Solution; MeHg, methyl mercury; NSC, neural stem cell; PBS, phosphate-buffered saline; pel, membrane-enclosed organelles; PLSD,
protected least significant difference; z-VAD-fmk, benzyloxycarbonylVal-Ala-Asp (OMe) fluoromethylketone.
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Journal Compilation 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78
69
70 C. Tamm et al.
MeHg contamination remains a persistent problem, and is of
major concern to developing organisms. Emission of
mercury occurs naturally mainly from the earth’s crust, but
also from anthropogenic sources, including mining, chloroalkali manufacturing and combustion of fossil fuels. Once in
aquatic environments, mercury is methylated by widespread
sulphate-reducing bacteria into MeHg. MeHg then enters the
aquatic food chain, and accumulates in fish and large sea
mammals (Morel et al. 1998). Dietary MeHg is almost
totally absorbed by the gastrointestinal tract and rapidly
enters the bloodstream, easily crossing the blood–brain
barrier and the placenta (Clarkson 1997). The levels of
MeHg in fetal blood are about 25% higher than those of the
mother (Amin-Zaki et al. 1976). It has also been shown that
fetuses exposed in utero can be affected in the absence of
maternal toxicity (Matsumoto et al. 1965; Choi et al. 1978;
Takeuchi 1982). The pattern of damage depends on the stage
of development when the exposure occurs, as well as
duration and concentration (Rodier 1995). At the cellular
level the cytotoxicity of MeHg has been ascribed to three
major mechanisms: perturbation of intracellular Ca2+ levels
(Sarafian 1993; Atchison and Hare 1994; Graff et al. 1997),
induction of oxidative stress by either overproduction of
reactive oxygen species (LeBel et al. 1990; Sarafian and
Verity 1991) or by reduced antioxidant defences (Sarafian
and Verity 1991; Yee and Choi 1994), and interactions with
sulphydryl groups (Clarkson 1972).
Exposure to MeHg can result in necrotic or apoptotic cell
death. Necrosis is characterized by ATP-independent cell and
organelle swelling, loss of plasma membrane integrity and
cell lysis. In contrast, apoptosis is an ATP-driven process
with cell shrinkage, chromatin condensation, plasma membrane blebbing, activation of specific proteases (e.g. caspases
and calpains), and DNA fragmentation at specific sites
(Orrenius et al. 2003). Several studies have shown that high
concentrations of MeHg induce necrosis, whereas lower
levels induce apoptosis in neuronal and glial cells (Miura and
Imura 1987; Nagashima et al. 1996; Castoldi et al. 2000;
Daré et al. 2000, 2001a).
In the present study we investigated the effects of MeHg
on NSCs by using the murine-derived multipotent NSC
line, C17.2, and primary cultures of cortical NSCs (cNSCs)
from E15 rat embryos. We focused our attention on the
mechanisms of MeHg-induced cell death, as well as on the
effects of non-cytotoxic levels of MeHg on differentiation
of NSCs.
Materials and methods
Cell culture procedures and experimental treatments
The murine-derived multipotent neural stem cell line C17.2 and
primary embryonic cNSCs obtained from E15 rat embryos were used
as experimental models. Both cell types have previously been shown
to maintain an undifferentiated state, or to differentiate into neurones,
astrocytes and oligodendrocytes (Snyder et al. 1992, 1997; Johe et al.
1996; Hermanson et al. 2002). The C17.2 cell line was maintained in
cell culture dishes (Corning Inc., Corning, NY, USA) in Dulbecco’s
modified Eagle’s medium (Life Technologies, Gibco BRL, Grand
Island, NY, USA) containing supplementary 10% fetal bovine serum
and 5% horse serum (HS) (Life Technologies) in a humidified
atmosphere of 5% CO2 and 95% air at 37C. For experimental
analyses, cells were grown in either cell culture dishes or on glass
coverslips coated with poly-L-lysine (Sigma, St Louis, MO, USA)
(50 lg/mL). At the time of the experiments all cells were nestin
positive, confirming their proliferative and undifferentiated status.
Primary cultures of NSCs were obtained from embryonic cortices
dissected in Hanks’ Balanced Salt Solution (HBSS) (Life Technologies) from timed-pregnant Sprague–Dawley rats (B & K, Sollentuna,
Sweden) at E15 (E1 was defined as the day of copulatory plug). The
tissue was gently mechanically dispersed, and meninges and larger
cell clumps were allowed to sediment for 10 min. The cells were
plated at a density of 0.6 · 106 cells per 100-mm cell culture dish
precoated with poly-L-ornithine and fibronectin (both from Sigma).
Cells were maintained in enriched N2 medium (Bottenstein and Sato
1979) with 10 ng/mL basic fibroblast growth factor (bFGF) (R & D
Systems, Minneapolis, MN, USA) added every 24 h and the medium
changed every other day to keep cells in an undifferentiated and
proliferative state. When still subconfluent, cells were passaged by
detaching by incubation with HBSS and subsequent scraping.
Afterwards, the cells were gently mixed in N2 medium, counted,
and plated at the desired density. The cells were used for experiments
48 h after the first and second passage. When used, the general caspase
inhibitor z-VAD-fmk [benzyloxycarbonyl-Val-Ala-Asp (OMe)
fluoromethylketone] (20 lM) (Peptide Institute, Osaka, Japan) and/
or the calpain inhibitor E64d (Sigma) was added 60 min before
exposure to MeHg (0.05–2 lM).
Trypan blue exclusion test
Cells were harvested with trypsin and a small aliquot of the cell
suspension was diluted with an equal volume of 0.4% Trypan blue
solution (Sigma). Cells were counted under a phase-contrast
microscope using a Neubauer improved counting chamber. Cells
with a damaged cell membrane (necrotic cells) stained blue, whereas
cells with an intact plasma membrane (healthy or apoptotic cells)
remained unstained. All experiments were performed in triplicate
and repeated at least three times.
Immunocytochemistry
Cells were fixed with cold 4% paraformaldehyde (Sigma) for 60 min
or ice-cold 80% methanol for 30 min and then washed with
phosphate-buffered saline (PBS). Primary antibodies were diluted in
PBS with 0.3% Triton X-100 and 0.5% bovine serum albumin (BSA)
(Boehringer Mannheim, Bromma, Sweden). The following primary
antibodies were used: rabbit anti-nestin (1 : 1000; Karolinska
Institute, Stockholm, Sweden) (Dahlstrand et al. 1992), mouse antiBax (1 : 400, clone 6A7; BD PharMingen) and mouse anti-cytochrome c (1 : 100, BD PharMingen). Cells were incubated in a humid
chamber at 4C overnight, rinsed with PBS and incubated with
secondary FITC-conjugated antibodies (Jackson ImmunoResearch,
West Grove, PA, USA) for 60 min at 24C. After rinsing with PBS,
coverslips were mounted in glycerol–PBS containing 0.1% phenyl-
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Journal Compilation 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78
Susceptibility of neural stem cells to methylmercury toxicity 71
enediamine. All experiments were performed in triplicate and
repeated at least three times.
Nuclear staining with Hoechst 33342 and propidium iodide
To evaluate the nuclear morphology, C17.2 cells grown on
coverslips and fixed with 80% methanol for 30 min at ) 20C.
They were then washed with PBS, and stained with propidium
iodide (1.0 lg/mL) or Hoechst 33342 (30 lg/mL) (both from
Molecular Probes, Eugene, OR, USA) for 5 min at room temperature. After rinsing in PBS, coverslips were mounted with glycerol–
PBS containing 0.1% phenylenediamine and examined by fluorescence microscopy. Cells were counted, scoring at least 300 cells in
five microscopic fields randomly selected on each coverslip. The
experiments were performed three times in triplicate.
Microscopy and photography
Cells were examined using an Olympus BX60 fluorescence
microscope (Olympus, Tokyo, Japan) equipped with a C4742-9510sc digital camera (Hamamatsu Photomics Norden AB, Solna,
Sweden), or a Zeiss LSM 510 Meta confocal microscope (Zeiss,
Jena, Germany).
Measurement of caspase activity
To evaluate the activity of class II caspases (2, 3 and 7) we measured
Ac-Asp-Glu-Val-Asp-7-amino-4-methylcoumarin (DEVD-AMC)
(Peptide Institute) cleavage using a fluorometric assay, as described
previously (Gorman et al. 1999). Some 200 000 cells/lL were
pelleted and washed once with ice-cold PBS. Cells were resuspended in 25 lL PBS, added to a microtitre plate, and combined
with substrate dissolved in a standard reaction buffer [100 mM
HEPES, pH 7.25, 10% sucrose, 10 mM dithiothreitol (DTT), 0.1%
CHAPS]. Cleavage of the fluorogenic peptide substrates was
monitored by AMC liberation in a Fluoroscan II plate reader
(Labsystems, Stockholm, Sweden) using 355 nm excitation and
460 nm emission wavelengths. Fluorescence units were converted
to pmoles of AMC released using a standard curve generated with
free AMC and subsequently related to amount of protein in each
sample. The experiments were performed three times in triplicate.
Western blot analysis
After treatment, cells were harvested by trypsinization, washed with
cold PBS and incubated in lysis buffer (10 mM Tris, 10 mM NaCl,
3 mM MgCl2, 0.1% Nonidet P-40, 0.1 mM phenylmethylsulphonyl
fluoride, 1 mM DTT, 2 lg/mL aprotinin) for 30 min on ice. For
fractionation of organelles and cytosol, cells were harvested by
trypsinization and washed once with cold PBS. The cells were then
resuspended at 1 million cells per 50 lL 0.005% digitonin (Sigma)
in a lysis buffer (250 mM sucrose, 20 mM HEPES, pH 7.4, 5 mM
MgCl2, 10 mM KCl, 1 mM EDTA, 1 mM EGTA). After 5 min, the
cells were centrifuged at 16 000 g for 5 min; 80 lL of the cytosolic
fraction-containing supernatant was removed and kept at ) 20C.
The residual pellet was resuspended in 100 lL of the digitonin lysis
buffer described above, snap-frozen in liquid nitrogen and quickthawed to complete the lysis. Some 80 lL of the membrane fraction
was saved and stored at ) 20C. The protein content of samples was
determined with a Micro BSA protein assay (Pierce, Rockford, IL,
USA) to ensure equal protein loading in each well during gel
electrophoresis. All samples were mixed with Laemmli loading
buffer, boiled for 5 min, and subjected to sodium dodecyl sulphate–
polyacrylamide gel electrophoresis (12%) at 100 V followed by
electroblotting to nitrocellulose for 2 h at 110 V. Membranes were
blocked for 1 h with 5% non-fat milk in PBS at room temperature
and subsequently probed overnight with mouse anti-fodrin
(1 : 1000; Chemicon, Temecula, CA, USA) or rabbit anti-cytochrome c (1 : 2500; BD PharMingen). The membranes were rinsed
and incubated with a horseradish peroxidase-conjugated secondary
antibody (1 : 10 000; Pierce). Following incubation with secondary
antibody, the membranes were rinsed, developed with enhanced
chemiluminescence reagents (Amersham, Little Chalfont, UK) and
exposed to autoradiography films (Fuji). All experiments were
performed in triplicate and repeated at least three times.
Differentiation assay
Primary cultures of NSCs were plated at low density (500 cells/cm2)
on coverslips coated with poly-L-ornithine and fibronectin, and
grown in the presence of bFGF. Forty-eight hours after first passage,
the medium was changed and no further bFGF was added during the
course of the experiment to promote spontaneous differentiation;
meanwhile medium was changed every second day. Cells were
exposed once to doses of MeHg ranging from 2.5 to 5 nM for
7 days. Subsequently cells were fixed with cold 4% paraformaldehyde for 60 min as described above. Primary antibodies were
diluted in PBS supplemented with 0.3% Triton X-100 and 0.5%
BSA. To determine the differentiation state of NSCs, cells were
stained for the neuronal marker Tuj1, using mouse anti-Tuj1
(1 : 1000; Nordic Biosite), rabbit anti-nestin (1 : 1000; Pharmigen,
Täby, Sweden) and Hoechst 33342. After incubation with the
appropriate secondary antibody (Alexa; Molecular Probes) for 1 h at
room temperature, coverslips were rinsed and mounted in glycerol–
PBS containing 0.1% phenylenediamine. Stained cells were examined by fluorescence microscopy; differentiated cells were counted
in five microscopic fields randomly selected on each coverslip and
then related to the total number of cells in each field assessed by
counting of Hoechst 33342-stained nuclei. The experiments were
performed twice in triplicate.
Results
NSCs are highly susceptible to MeHg toxicity
To investigate the cytotoxic effect of MeHg on NSCs, we
exposed C17.2 cells and primary cNSCs to concentrations of
MeHg that have been used previously to induce apoptotic cell
death in neuronal or glial cells (Daré et al. 2000, 2001a). In
C17.2 cells, the Trypan blue exclusion test showed a significant level of cell membrane damage at concentrations ranging
from 0.5 to 2 lM, in a dose-dependent manner, with a peak of
45% at 2 lM (Fig. 1). In cNSCs the same concentrations
damaged the plasma membrane of almost all exposed cells,
with 90% of the cells already showing signs of cytotoxicity at
0.5 lM MeHg (Fig. 1). To keep the focus of our studies on the
identification of intracellular pathways leading to apoptotic
cell death, we lowered the concentration of MeHg to
0.1–0.5 lM for C17.2 cells and to 0.025–0.05 lM for cNSCs,
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72 C. Tamm et al.
(a)
(b)
Fig. 1 Altered membrane permeability in C17.2 cells and cNSCs
exposed to different doses of MeHg (0.5–2 lM) for 24 h. Following
treatment, cells were harvested and stained with Trypan blue and
analysed under light microscopy. Values are mean ± SD (n ¼ 3).
*p < 0.05 [ANOVA; Fisher’s protected least significant difference (PLSD)
test].
(a)
(c)
Fig. 3 (a) Chromatin condensation (apoptotic cells) and alterations in
permeability (necrotic cells) in cNSC cells after treatment with MeHg
(0.025–0.1 lM) for 24 h. Values are mean ± SD (n ¼ 3). *p < 0.05
(ANOVA; Fisher’s PLSD test). (b) Untreated cNSCs stained with
Hoechst 33342. (c) cNSCs exposed to 0.05 lM MeHg stained with
Hoechst 33342 showing apoptotic chromatin condensation (arrows).
Scale bar 10 lm.
condensed chromatin visualized with the DNA stain Hoechst
33342. Remarkably, in cNSCs similar percentages of apoptotic
cells were induced by 10-fold lower doses of MeHg (Fig. 3),
indicating that embryonic NSCs were extremely sensitive to
the toxic effects of MeHg.
(b)
(c)
Fig. 2 (a) Chromatin condensation (apoptotic cells) and alterations in
membrane permeability (necrotic cells) in C17.2 cells after treatment
with MeHg (0.1–0.5 lM) for 24 h. Values are mean ± SD (n ¼ 3).
*p < 0.05 (ANOVA; Fisher’s PLSD test). (b) C17.2 control cells stained
with Hoechst 33342. (c) C17.2 cells exposed to 0.5 lM MeHg stained
with Hoechst 33342 showed apoptotic chromatin condensation
(arrows). Scale bar 20 lm.
at which concentrations only a negligible amount (< 5%) of
cells had a damaged plasma membrane. In C17.2 cells, 0.25 or
0.5 lM MeHg induced apoptosis in approximately 15–20% of
the exposed cells (Fig. 2), as estimated by quantifying
MeHg induces apoptosis in NSCs via Bax activation,
cytochrome c release, and caspase and calpain activation
After treatment for 24 h with the selected doses of MeHg,
both the C17.2 cells (Fig. 4a) and the cNSCs (Fig. 4b)
showed oligomerized and activated Bax, which may form a
channel or a membrane pore and allow the release of
apoptogenic factors (Antonsson et al. 2000). The exposed
cells also showed clear cytochrome c release from the
intramembrane space of the mitochondria into the cytosol, as
detected by immunocytochemistry (Figs 4c–f) and by western blotting (Fig. 5).
The activation of caspase 3 was examined by spectrofluorometric analysis of DEVD-AMC cleavage and detection
of the active fragment of caspase 3, p17, by immunocytochemistry. In C17.2 cells, MeHg induced a significant
increase in caspase 3-like cleavage of this synthetic substrate
in a dose-dependent manner, compared with levels in
untreated cells (Fig. 6a). Activation of caspase 3 was also
observed in cNSCs exposed to 0.05 lM MeHg, as detected
by the presence of the active fragment p17 in cells with a
condensed nucleus (Fig. 6b).
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Susceptibility of neural stem cells to methylmercury toxicity 73
To gain further evidence that caspase 3 is involved in
MeHg-induced apoptosis in NSCs, we investigated the
specific cleavage of the endogenous substrate a-fodrin in
the C17.2 cell line. Western blot analysis with an antibody
against the cytoskeletal protein a-fodrin showed a clear
increase in the 120-kDa fragment that is a cleavage product
of activated caspase 3 (Fig. 6c). In addition, there was a
significant increase in the 150-kDa fodrin breakdown product
(Fig. 6c), indicating that calpains are also activated in NSCs
exposed to MeHg.
To further assess the involvement of caspases and calpains
in MeHg-induced apoptosis, we used the pan-caspase
inhibitor zVAD-fmk to block activation of caspases, and
the general calpain inhibitor E64d. Pretreatment with 20 lM
zVAD-fmk for 1 h fully blocked DEVD-AMC cleavage in
C17.2 cells exposed to 0.5 lM MeHg for 24 h (data not
shown). Staining of the exposed cells with Hoechst 33342 to
allow identification and quantification of apoptotic nuclei
revealed that pretreatment with zVAD-fmk significantly but
only partially protected C17.2 cells and cNSCs from MeHginduced apoptosis (Figs 7a and b). Preincubation with the
calpain inhibitor E64d for 1 h before exposure to MeHg
resulted in partial protection in both cell models (Figs 7c and
d). When the two inhibitors were mixed together and added
to the cell cultures before MeHg, apoptosis was almost
completely prevented in both C17.2 cells and cNSCs
(Figs 7e and f). These data indicate that two independent
pathways, one involving the activation of caspases and the
other the activation of calpains, function in parallel during
MeHg-induced apoptotic cell death.
Fig. 4 Bax oligomerization and activation in C17.2 cells (a) and
cNSCs (b) after exposure to 0.5 and 0.05 lM MeHg respectively for
24 h. Green fluorescent-positive staining for oligomerized Bax (arrow)
is shown with blue Hoechst 33342-stained condensed nuclei. Cytochrome c (Cyt c) immunoreactivity and nuclear staining (Hoechst) in
control (c, e) and MeHg-exposed cells (d, f). In control C17.2 cells (c)
cytochrome c staining appeared as a mitochondrial dot-like pattern,
whereas in control primary cNSCs it appeared as a mitochondrial
network (e). Diffuse cytosolic staining (arrows) was observed in C17.2
cells (d) and cNSCs (e) exposed to MeHg (0.5 and 0.05 lM respectively) for 24 h, suggesting release of cytochrome c into the cytosol.
Scale bar 20 lm (a, c), 10 lm (b, e).
Concentrations of MeHg relevant to human exposure
inhibit neuronal differentiation of primary cNSCs
To test whether MeHg may affect differentiation of NSCs,
we used the cNSC primary culture. We exposed the cells to
concentrations that are not cytotoxic and are relevant to
human exposure. During normal culture conditions bFGF is
added to the cNSC medium to keep the stem cells in a
proliferative and undifferentiated state. When the medium is
replaced and no further bFGF is added, the cells begin to
differentiate spontaneously. This process takes approximately
7 days from the removal of bFGF to the appearance of a
strong b-tubulin III (Tuj1) neuronal staining. We observed
that exposure to 2.5 or 5 nM MeHg significantly impaired the
neuronal differentiating potential, as visualized by a clear
decrease in Tuj1-positive neuronal cells compared with the
control (Fig. 8). Undifferentiated cells in the control and
exposed conditions remained nestin positive (data not
shown).
Discussion
Fig. 5 Western blot of cytosolic (cyto) and membrane-enclosed
organelles (pel) showing mitochondrial cytochrome c release in C17.2
cells exposed to MeHg (0.25–0.5 lM) for 24 h.
In the present study we used the murine NSC cell line C17.2
and primary cultures of NSCs from cortices of E15 rat
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74 C. Tamm et al.
(a)
(b)
Fig. 6 (a) MeHg induces caspase 3-like
activity in C17.2 cells. Control and MeHgexposed cells were harvested, and caspase
3-like activity was measured as DEVD-AMC
cleavage. Values are mean ± SD (n ¼ 3).
*p < 0.05 (ANOVA; Fisher’s PLSD test). (b)
cNSCs exposed to 0.05 lM MeHg for 24 h
showed activated caspase 3 (p17) and
nuclear condensation as detected by
Hoechst 33342 staining. Scale bar 10 lm.
(c) Western blot analysis of C17.2 cells
exposed to 0.5–1 lM MeHg for 24 h showing specific breakdown of a-fodrin by calpain (150 kDa) and caspase (120 kDa).
(c)
embryos to investigate the cytotoxic effects of the environmental organometal MeHg on neurodevelopment. Our results
show that NSCs, especially embryonic NSCs, are highly
sensitive to MeHg, and that cells undergo apoptotic cell
death via activation of two parallel pathways involving
caspases and calpains. In addition, low doses of MeHg,
relevant to the current exposure via cord blood of the general
population, inhibit spontaneous neuronal differentiation of
embryonic NSCs.
MeHg has been a threat to public health for more than
50 years because of its neurotoxic effects in adults and
infants, ranging from minor behavioural changes to
morbidity (NRC 2000). The developing nervous system
has a unique susceptibility to MeHg and prenatal exposure
results in wide ranging adverse effects on brain development and organization, compared with the limited damage
that occurs when exposure takes place in adult life
(Matsumoto et al. 1965; Choi et al. 1978). The fetus is
far more susceptible to the toxic effects of MeHg than the
mother, and adverse neurological effects have been
described in the progeny of women who showed little or
no signs of toxicity (Harada 1978; Clarkson et al. 1985;
Marsh et al. 1987). The damage that occurs in cases of
fetal exposure in both animal models and humans is
mostly associated with a decrease in the number of neural
cells and altered cytoarchitecture (Matsumoto et al. 1965;
Mottet and Body 1974; Takeuchi et al. 1977; Choi et al.
1978, 1986, 1989; Geelen et al. 1990; Eto et al. 1992).
This can be ascribed to interference with processes such as
cell division, migration, differentiation and cell death,
which regulate neural development.
The capacity of MeHg to reduce the number of progenitor
cells has been attributed to impaired cell cycle transition and
mitotic inhibition (Matsumoto et al. 1965; Miura et al. 1978;
Rodier et al. 1984; Sager et al. 1984; Howard and Mottet
1986; Sager 1988; Choi 1989, 1991; Ponce et al. 1994). Our
studies indicate that MeHg can also decrease the number of
NSCs by the activation of intracellular pathways that lead to
apoptosis, even at exposure levels that do not cause death of
other neural cell types.
Apoptotic cell death occurs via the activation of well
characterized biochemical pathways (Zimmermann et al.
2001). The release of proteins, including cytochrome c from
the intermembrane space of the mitochondria, initiates the
classical mitochondrial pathway (Liu et al. 1996; Bratton
et al. 2000). Cytochrome c interacts with apoptotic protease-
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Susceptibility of neural stem cells to methylmercury toxicity 75
Fig. 7 Chromatin condensation in apoptotic C17.2 cells and cNSCs
exposed for 24 h to apoptosis-inducing concentrations of 0.5 and
0.05 lM MeHg respectively. Following treatment cells were fixed,
stained with Hoechst 33342 and analysed by fluorescence microscopy.
Values are mean ± SD (n ¼ 3). *p < 0.05 (ANOVA; Fisher’s PLSD test).
Pretreatment for 1 h with the pan-caspase inhibitor zVAD-fmk (20 lM)
partially decreased the amount of both C17.2 cells (a) and cNSCs (b)
with apoptotic morphology after exposure to MeHg. A similar effect was
observed after pretreatment for 1 h with the calpain inhibitor E64d
(10 lM) (c, d). Combination of the two inhibitors almost completely
prevented the formation of nuclei with condensed chromatin after
exposure to MeHg (e, f).
activating factor-1 in the cytosol, leading to the activation of
pro-caspase 9, which in turn cleaves and activates procaspase 3 (Bratton et al. 2000).
Another pathway involves the binding of members of
the death receptor family (e.g. Fas/tumor necrosis factor
receptor 1/tumor necrosis factor-related apoptosis-inducing
ligand receptor) and their cognate ligands (Nagata 1997).
Besides caspases, other cysteine proteases, i.e. calpains, can be
activated in apoptosis (Chan and Mattson 1999). An increase
in intracellular Ca2+ is the main signal for activation of these
proteases (Sorimachi et al. 1997).
In previous studies we investigated the cell death
mechanisms activated when NSCs are exposed to toxic
stimuli, such as oxidative stress, and showed that NSCs
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76 C. Tamm et al.
Fig. 8 Non-cytotoxic levels of MeHg inhibit neuronal differentiation in
cNSCs. In the absence of bFGF, cells were treated with a single dose
of MeHg (2.5–5 nM) for 48 h and subsequently allowed to differentiate
spontaneously for an additional 120 h. Cells were fixed and immunocytochemically stained with the early neuronal marker Tuj1 (arrows)
and analysed by fluorescence microscopy. Compared with untreated
control cells (a, b), single doses of MeHg (2.5 and 5 nM) (c, d)
decreased neuronal differentiation of cNSCs. Semiquantitative analysis showed that the decrease in cNSC neuronal differentiation was
significant (e). Values are mean ± SD (n ¼ 3). *p < 0.05 (ANOVA;
Fisher’s PLSD test). Scale bar 10 lm.
can undergo apoptosis via the mitochondrial caspasemediated pathway, whereas the Fas-mediated cell death
pathway does not seem to be operative (Sleeper et al.
2002; Tamm et al. 2004)
Here we show that Bax is oligomerized in NSCs exposed
to MeHg, and cytochrome c is released from the mitochondria with subsequent activation of the effector caspase 3. In
addition to caspases, NSCs can activate the calpain pathway.
This pathway is activated by the perturbation in intracellular
Ca2+ homeostasis that can occur during exposure to toxic
stimuli such as MeHg. The caspase and calpain pathways are
concomitantly activated in NSCs exposed to concentrations
of MeHg inducing apoptosis, as proven by either the partial
protection exerted by the caspase (zVAD-fmk) or calpain
(E64d) inhibitor alone, or the full protective effect of the two
inhibitors together.
NSCs appear to be much more vulnerable than other
in vitro neural models used previously to investigate the
cytotoxicity of MeHg. In contrast to neuronal and glial cells
(Daré et al. 2000, 2001b), MeHg-exposed NSCs undergoing
apoptosis show activation of different execution pathways,
with caspases playing a critical role. The percentage of
apoptotic cells in primary cNSC cultures induced by 0.05 lM
MeHg was similar to that observed in C17.2 cells after
exposure to a 10-fold higher dose. Several factors involved in
cell signalling may contribute to the observed effects of
MeHg on NSC survival and differentiation, including
alterations in neurotrophic factors, neurotrophic factor
receptors and in the Eph/Ephrin family, which have been
shown to be affected by MeHg (Soderström and Ebendal
1995; Andersson et al. 1997; Parran et al. 2003; Wilson
et al. 2005). Further investigation is needed to clarify this
issue.
A noteworthy finding is that levels of MeHg (2.5–5 nM)
lower than those found in the umbilical cord blood of
pregnant women in the general Swedish population (0.99 lg/
L) (Björnberg et al. 2005) can inhibit spontaneous neuronal
differentiation. Remarkably, a daily exposure to 0.1 lg
MeHg/kg bodyweight, which is considered to be without
risk of deleterious effects (NRC 2000), equates to 5.8 lg/L in
cord blood, which is 10-fold higher than the concentration
used in our studies. Thus, in light of our results, there seems
to be a narrow margin of safety against the risk of
neurodevelopmental effects. Thus dietary advice for pregnant
women is necessary and justified. In addition, considering
that NSCs are also present in the adult nervous system, where
they may have a role in learning, memory and response to
injury, exposure to low levels of MeHg may have negative
consequences in adulthood as well.
By confirming in vivo data on the increased sensitivity of
the developing nervous system to MeHg, our study shows
that cultures of NSCs represent a good in vitro model with
which to identify the neurodevelopmental effects of toxic
substances, and the effects that extrinsic factors play in the
cell biology of NSCs. The difference in sensitivity to MeHg
that we have observed in NSCs further strengthens the need
for multiple cellular models when in vitro studies are used to
identify the toxic effects of a potential neurotoxic substance.
2006 The Authors
Journal Compilation 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78
Susceptibility of neural stem cells to methylmercury toxicity 77
In conclusion, the present data show that NSCs are highly
sensitive to the toxic effects of MeHg, and that cells undergo
apoptosis via activation of two parallel pathways involving
caspases and calpains. The effects of low concentrations of
MeHg, similar to those to which humans are exposed, on
spontaneous neuronal differentiation of NSCs point to the
need for further investigations of NSCs exposed to subtoxic
doses of neurotoxic substances.
Acknowledgements
The authors thank Dr Evan Y. Snyder for providing the C17.2 cells.
This work was supported by grants from the European Commission
(CT 2003-506143), the Swedish Research Council (33X-10815), the
Swedish Research Council for Environment, Agricultural Sciences
and Spatial Planning, the Swedish Animal Welfare Agency, the
Swedish Cancer Society and the Swedish Children’s Cancer
Foundation.
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