High susceptibility of neural stem cells to methylmercury ... · High susceptibility of neural stem...

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 con-

trol their proliferation and differentiation, as well as the influ-

ence 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 envi-

ronmental 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. Remark-

ably, exposure to MeHg at concentrations comparable to the

current developmental exposure (via cord blood) of the gen-

eral 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 self-renew and differentiate into the three major cell types,neurones, astrocytes and oligodendrocytes, play an essen-tial role in the development and maturation of the nervoussystem. They are present not only in the developing brainbut also in the adult brain in different areas withneurogenic potential (Gage 2000). The importance ofNSCs in the adult CNS is uncertain, but there is evidencesuggesting that capability for self-renewal may be import-ant for normal brain functions, including learning, memoryand emotional responses (Santarelli et al. 2003; Schafferand Gage 2004).

Recently there has been much excitement about thepossibility of replacing damaged or lost neural cells bytransplantation of NSCs, or by recruitment of endogenousprecursors to repair adult brain. However, before the fullpotential of NSCs can be realized, it is essential tounderstand the physiological pathways that control theirproliferation and differentiation, as well as the influence thatextrinsic factors may have on these processes.

The developing nervous system is especially vulnerable todamage by toxic agents, but so far little attention has beenfocused on the effects of environmental neurotoxicants onNSCs. A relevant environmental contaminant that isextremely toxic to the developing nervous system ismethylmercury (MeHg). Despite major actions taken toreduce 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 andNeurotoxicology, Karolinska Institutet, 171 77 Stockholm, Sweden.E-mail: [email protected] used: bFGF, basic fibroblast growth factor; BSA,

bovine serum albumin; cNSC, cortical neural stem cell; Cyt c, cyto-chrome c; cyto, cytosolic; DEVD-AMC, Ac-Asp-Glu-Val-Asp-7-amino-4-methylcoumarin; DTT, dithiothreitol; HBSS, Hanks’ Balanced SaltSolution; MeHg, methyl mercury; NSC, neural stem cell; PBS, phos-phate-buffered saline; pel, membrane-enclosed organelles; PLSD,protected least significant difference; z-VAD-fmk, benzyloxycarbonyl-Val-Ala-Asp (OMe) fluoromethylketone.

Journal of Neurochemistry, 2006, 97, 69–78 doi:10.1111/j.1471-4159.2006.03718.x

� 2006 The AuthorsJournal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78 69

MeHg contamination remains a persistent problem, and is ofmajor concern to developing organisms. Emission ofmercury occurs naturally mainly from the earth’s crust, butalso from anthropogenic sources, including mining, chloro-alkali manufacturing and combustion of fossil fuels. Once inaquatic environments, mercury is methylated by widespreadsulphate-reducing bacteria into MeHg. MeHg then enters theaquatic food chain, and accumulates in fish and large seamammals (Morel et al. 1998). Dietary MeHg is almosttotally absorbed by the gastrointestinal tract and rapidlyenters the bloodstream, easily crossing the blood–brainbarrier and the placenta (Clarkson 1997). The levels ofMeHg in fetal blood are about 25% higher than those of themother (Amin-Zaki et al. 1976). It has also been shown thatfetuses exposed in utero can be affected in the absence ofmaternal toxicity (Matsumoto et al. 1965; Choi et al. 1978;Takeuchi 1982). The pattern of damage depends on the stageof development when the exposure occurs, as well asduration and concentration (Rodier 1995). At the cellularlevel the cytotoxicity of MeHg has been ascribed to threemajor mechanisms: perturbation of intracellular Ca2+ levels(Sarafian 1993; Atchison and Hare 1994; Graff et al. 1997),induction of oxidative stress by either overproduction ofreactive oxygen species (LeBel et al. 1990; Sarafian andVerity 1991) or by reduced antioxidant defences (Sarafianand Verity 1991; Yee and Choi 1994), and interactions withsulphydryl groups (Clarkson 1972).

Exposure to MeHg can result in necrotic or apoptotic celldeath. Necrosis is characterized by ATP-independent cell andorganelle swelling, loss of plasma membrane integrity andcell lysis. In contrast, apoptosis is an ATP-driven processwith cell shrinkage, chromatin condensation, plasma mem-brane blebbing, activation of specific proteases (e.g. caspasesand calpains), and DNA fragmentation at specific sites(Orrenius et al. 2003). Several studies have shown that highconcentrations of MeHg induce necrosis, whereas lowerlevels induce apoptosis in neuronal and glial cells (Miura andImura 1987; Nagashima et al. 1996; Castoldi et al. 2000;Dare et al. 2000, 2001a).

In the present study we investigated the effects of MeHgon NSCs by using the murine-derived multipotent NSCline, C17.2, and primary cultures of cortical NSCs (cNSCs)from E15 rat embryos. We focused our attention on themechanisms of MeHg-induced cell death, as well as on theeffects of non-cytotoxic levels of MeHg on differentiationof 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 37�C. 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 Technol-

ogies) 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 anti-

Bax (1 : 400, clone 6A7; BD PharMingen) and mouse anti-cyto-

chrome c (1 : 100, BD PharMingen). Cells were incubated in a humid

chamber at 4�C overnight, rinsed with PBS and incubated with

secondary FITC-conjugated antibodies (Jackson ImmunoResearch,

West Grove, PA, USA) for 60 min at 24�C. After rinsing with PBS,

coverslips were mounted in glycerol–PBS containing 0.1% phenyl-

70 C. Tamm et al.

Journal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78� 2006 The Authors

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 ) 20�C.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 tempera-

ture. After rinsing in PBS, coverslips were mounted with glycerol–

PBS containing 0.1% phenylenediamine and examined by fluores-

cence 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 C4742-95-

10sc 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 resus-

pended 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 ) 20�C.The residual pellet was resuspended in 100 lL of the digitonin lysis

buffer described above, snap-frozen in liquid nitrogen and quick-

thawed to complete the lysis. Some 80 lL of the membrane fraction

was saved and stored at ) 20�C. 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-cyto-

chrome 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% paraformalde-

hyde 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,

Taby, 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 exam-

ined 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, weexposed C17.2 cells and primary cNSCs to concentrations ofMeHg that have been used previously to induce apoptotic celldeath in neuronal or glial cells (Dare et al. 2000, 2001a). InC17.2 cells, the Trypan blue exclusion test showed a signifi-cant level of cell membrane damage at concentrations rangingfrom 0.5 to 2 lM, in a dose-dependent manner, with a peak of45% at 2 lM (Fig. 1). In cNSCs the same concentrationsdamaged the plasma membrane of almost all exposed cells,with 90% of the cells already showing signs of cytotoxicity at0.5 lMMeHg (Fig. 1). To keep the focus of our studies on theidentification of intracellular pathways leading to apoptoticcell death, we lowered the concentration of MeHg to0.1–0.5 lM for C17.2 cells and to 0.025–0.05 lM for cNSCs,

Susceptibility of neural stem cells to methylmercury toxicity 71

� 2006 The AuthorsJournal Compilation � 2006 International Society for Neurochemistry, J. Neurochem. (2006) 97, 69–78

at which concentrations only a negligible amount (< 5%) ofcells had a damaged plasma membrane. In C17.2 cells, 0.25 or0.5 lMMeHg induced apoptosis in approximately 15–20% ofthe exposed cells (Fig. 2), as estimated by quantifying

condensed chromatin visualized with the DNA stain Hoechst33342.Remarkably, in cNSCs similar percentages of apoptoticcells were induced by 10-fold lower doses of MeHg (Fig. 3),indicating that embryonic NSCs were extremely sensitive tothe toxic effects of MeHg.

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 achannel or a membrane pore and allow the release ofapoptogenic factors (Antonsson et al. 2000). The exposedcells also showed clear cytochrome c release from theintramembrane space of the mitochondria into the cytosol, asdetected by immunocytochemistry (Figs 4c–f) and by west-ern blotting (Fig. 5).

The activation of caspase 3 was examined by spectro-fluorometric analysis of DEVD-AMC cleavage and detectionof the active fragment of caspase 3, p17, by immunocyto-chemistry. In C17.2 cells, MeHg induced a significantincrease in caspase 3-like cleavage of this synthetic substratein a dose-dependent manner, compared with levels inuntreated cells (Fig. 6a). Activation of caspase 3 was alsoobserved in cNSCs exposed to 0.05 lM MeHg, as detectedby the presence of the active fragment p17 in cells with acondensed nucleus (Fig. 6b).

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)

(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.

(a)

(b) (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.

72 C. Tamm et al.


To gain further evidence that caspase 3 is involved inMeHg-induced apoptosis in NSCs, we investigated thespecific cleavage of the endogenous substrate a-fodrin inthe C17.2 cell line. Western blot analysis with an antibodyagainst the cytoskeletal protein a-fodrin showed a clearincrease in the 120-kDa fragment that is a cleavage productof activated caspase 3 (Fig. 6c). In addition, there was asignificant increase in the 150-kDa fodrin breakdown product(Fig. 6c), indicating that calpains are also activated in NSCsexposed to MeHg.

To further assess the involvement of caspases and calpainsin MeHg-induced apoptosis, we used the pan-caspaseinhibitor zVAD-fmk to block activation of caspases, andthe general calpain inhibitor E64d. Pretreatment with 20 lMzVAD-fmk for 1 h fully blocked DEVD-AMC cleavage inC17.2 cells exposed to 0.5 lM MeHg for 24 h (data notshown). Staining of the exposed cells with Hoechst 33342 toallow identification and quantification of apoptotic nucleirevealed that pretreatment with zVAD-fmk significantly butonly partially protected C17.2 cells and cNSCs from MeHg-induced apoptosis (Figs 7a and b). Preincubation with thecalpain inhibitor E64d for 1 h before exposure to MeHgresulted in partial protection in both cell models (Figs 7c andd). When the two inhibitors were mixed together and addedto the cell cultures before MeHg, apoptosis was almostcompletely prevented in both C17.2 cells and cNSCs(Figs 7e and f). These data indicate that two independentpathways, one involving the activation of caspases and theother the activation of calpains, function in parallel duringMeHg-induced apoptotic cell death.

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 toconcentrations that are not cytotoxic and are relevant tohuman exposure. During normal culture conditions bFGF isadded to the cNSC medium to keep the stem cells in aproliferative and undifferentiated state. When the medium isreplaced and no further bFGF is added, the cells begin todifferentiate spontaneously. This process takes approximately7 days from the removal of bFGF to the appearance of astrong b-tubulin III (Tuj1) neuronal staining. We observedthat exposure to 2.5 or 5 nM MeHg significantly impaired theneuronal differentiating potential, as visualized by a cleardecrease in Tuj1-positive neuronal cells compared with thecontrol (Fig. 8). Undifferentiated cells in the control andexposed conditions remained nestin positive (data notshown).

Discussion

In the present study we used the murine NSC cell line C17.2and primary cultures of NSCs from cortices of E15 rat

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. Cyto-

chrome 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 respect-

ively) for 24 h, suggesting release of cytochrome c into the cytosol.

Scale bar 20 lm (a, c), 10 lm (b, e).

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.



embryos to investigate the cytotoxic effects of the environ-mental organometal MeHg on neurodevelopment. Our resultsshow that NSCs, especially embryonic NSCs, are highlysensitive to MeHg, and that cells undergo apoptotic celldeath via activation of two parallel pathways involvingcaspases and calpains. In addition, low doses of MeHg,relevant to the current exposure via cord blood of the generalpopulation, inhibit spontaneous neuronal differentiation ofembryonic NSCs.

MeHg has been a threat to public health for more than50 years because of its neurotoxic effects in adults andinfants, ranging from minor behavioural changes tomorbidity (NRC 2000). The developing nervous systemhas a unique susceptibility to MeHg and prenatal exposureresults in wide ranging adverse effects on brain develop-ment and organization, compared with the limited damagethat occurs when exposure takes place in adult life(Matsumoto et al. 1965; Choi et al. 1978). The fetus isfar more susceptible to the toxic effects of MeHg than themother, and adverse neurological effects have beendescribed in the progeny of women who showed little orno 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 ismostly associated with a decrease in the number of neuralcells 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 ascell division, migration, differentiation and cell death,which regulate neural development.

The capacity of MeHg to reduce the number of progenitorcells has been attributed to impaired cell cycle transition andmitotic inhibition (Matsumoto et al. 1965; Miura et al. 1978;Rodier et al. 1984; Sager et al. 1984; Howard and Mottet1986; Sager 1988; Choi 1989, 1991; Ponce et al. 1994). Ourstudies indicate that MeHg can also decrease the number ofNSCs by the activation of intracellular pathways that lead toapoptosis, even at exposure levels that do not cause death ofother neural cell types.

Apoptotic cell death occurs via the activation of wellcharacterized biochemical pathways (Zimmermann et al.2001). The release of proteins, including cytochrome c fromthe intermembrane space of the mitochondria, initiates theclassical mitochondrial pathway (Liu et al. 1996; Brattonet al. 2000). Cytochrome c interacts with apoptotic protease-

(a)

(c)

(b)

Fig. 6 (a) MeHg induces caspase 3-like

activity in C17.2 cells. Control and MeHg-

exposed 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 show-

ing specific breakdown of a-fodrin by cal-

pain (150 kDa) and caspase (120 kDa).

74 C. Tamm et al.


activating factor-1 in the cytosol, leading to the activation ofpro-caspase 9, which in turn cleaves and activates pro-caspase 3 (Bratton et al. 2000).

Another pathway involves the binding of members ofthe death receptor family (e.g. Fas/tumor necrosis factorreceptor 1/tumor necrosis factor-related apoptosis-inducingligand receptor) and their cognate ligands (Nagata 1997).

Besides caspases, other cysteine proteases, i.e. calpains, can beactivated in apoptosis (Chan and Mattson 1999). An increasein intracellular Ca2+ is the main signal for activation of theseproteases (Sorimachi et al. 1997).

In previous studies we investigated the cell deathmechanisms activated when NSCs are exposed to toxicstimuli, such as oxidative stress, and showed that NSCs

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).



can undergo apoptosis via the mitochondrial caspase-mediated pathway, whereas the Fas-mediated cell deathpathway does not seem to be operative (Sleeper et al.2002; Tamm et al. 2004)

Here we show that Bax is oligomerized in NSCs exposedto MeHg, and cytochrome c is released from the mitochon-dria with subsequent activation of the effector caspase 3. Inaddition to caspases, NSCs can activate the calpain pathway.This pathway is activated by the perturbation in intracellularCa2+ homeostasis that can occur during exposure to toxicstimuli such as MeHg. The caspase and calpain pathways areconcomitantly activated in NSCs exposed to concentrationsof MeHg inducing apoptosis, as proven by either the partialprotection exerted by the caspase (zVAD-fmk) or calpain(E64d) inhibitor alone, or the full protective effect of the twoinhibitors together.

NSCs appear to be much more vulnerable than otherin vitro neural models used previously to investigate thecytotoxicity of MeHg. In contrast to neuronal and glial cells(Dare et al. 2000, 2001b), MeHg-exposed NSCs undergoingapoptosis show activation of different execution pathways,with caspases playing a critical role. The percentage ofapoptotic cells in primary cNSC cultures induced by 0.05 lMMeHg was similar to that observed in C17.2 cells afterexposure to a 10-fold higher dose. Several factors involved incell signalling may contribute to the observed effects ofMeHg on NSC survival and differentiation, includingalterations in neurotrophic factors, neurotrophic factorreceptors and in the Eph/Ephrin family, which have beenshown to be affected by MeHg (Soderstrom and Ebendal1995; Andersson et al. 1997; Parran et al. 2003; Wilsonet al. 2005). Further investigation is needed to clarify thisissue.

A noteworthy finding is that levels of MeHg (2.5–5 nM)lower than those found in the umbilical cord blood ofpregnant women in the general Swedish population (0.99 lg/L) (Bjornberg et al. 2005) can inhibit spontaneous neuronaldifferentiation. Remarkably, a daily exposure to 0.1 lgMeHg/kg bodyweight, which is considered to be withoutrisk of deleterious effects (NRC 2000), equates to 5.8 lg/L incord blood, which is 10-fold higher than the concentrationused in our studies. Thus, in light of our results, there seemsto be a narrow margin of safety against the risk ofneurodevelopmental effects. Thus dietary advice for pregnantwomen is necessary and justified. In addition, consideringthat NSCs are also present in the adult nervous system, wherethey may have a role in learning, memory and response toinjury, exposure to low levels of MeHg may have negativeconsequences in adulthood as well.

By confirming in vivo data on the increased sensitivity ofthe developing nervous system to MeHg, our study showsthat cultures of NSCs represent a good in vitro model withwhich to identify the neurodevelopmental effects of toxicsubstances, and the effects that extrinsic factors play in thecell biology of NSCs. The difference in sensitivity to MeHgthat we have observed in NSCs further strengthens the needfor multiple cellular models when in vitro studies are used toidentify the toxic effects of a potential neurotoxic substance.

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 immuno-

cytochemically 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 ana-

lysis 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.

76 C. Tamm et al.


In conclusion, the present data show that NSCs are highlysensitive to the toxic effects of MeHg, and that cells undergoapoptosis via activation of two parallel pathways involvingcaspases and calpains. The effects of low concentrations ofMeHg, similar to those to which humans are exposed, onspontaneous neuronal differentiation of NSCs point to theneed for further investigations of NSCs exposed to subtoxicdoses 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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