skn-1/nrf2 inhibits dopamine neuron degeneration in a...

TOXICOLOGICAL SCIENCES 118(2), 613–624 (2010)

doi:10.1093/toxsci/kfq285

Advance Access publication September 20, 2010

SKN-1/Nrf2 Inhibits Dopamine Neuron Degeneration in a Caenorhabditiselegans Model of Methylmercury Toxicity

Natalia VanDuyn,*,† Raja Settivari,*,† Garry Wong,‡,§ and Richard Nass*,†,1

*Department of Pharmacology and Toxicology and †Stark Neuroscience Research Institute, Indiana University School of Medicine, Indianapolis, Indiana

46202; and ‡Department of Biosciences, and §Department of Neurobiology, A.I. Virtanen Institute, University of Eastern Finland, Kuopio, Finland 70211

1To whom correspondence should be addressed at Department of Pharmacology and Toxicology, Indiana University School of Medicine, 635 Barnhill Drive,

MS 549, Indianapolis, IN 46202. Fax: (317) 274-7868. E-mail: [email protected].

Received July 25, 2010; accepted September 7, 2010

Methylmercury (MeHg) exposure from occupational, environ-

mental, and food sources is a significant threat to public health.

MeHg poisonings in adults may result in severe psychological and

neurological deficits, and in utero exposures can confer embryonic

defects and developmental delays. Recent epidemiological and

vertebrate studies suggest that MeHg exposure may also

contribute to dopamine (DA) neuron vulnerability and the

propensity to develop Parkinson’s disease (PD). In this study, we

describe a Caenorhabditis elegans model of MeHg toxicity that

shows that low, chronic exposure confers embryonic defects,

developmental delays, decreases in brood size and animal viability,

and DA neuron degeneration. Toxicant exposure results in the

robust induction of the glutathione-S-transferases (GSTs) gst-4 and

gst-38 that are largely dependent on the PD-associated phase II

antioxidant transcription factor SKN-1/Nrf2. We also demonstrate

that the expression of SKN-1, a protein previously localized to

a small subset of chemosensory neurons and intestinal cells in the

nematode, is also expressed in the DA neurons, and a reduction in

SKN-1 gene expression increases MeHg-induced animal vulnerabil-

ity and DA neuron degeneration. These studies recapitulate

fundamental hallmarks of MeHg-induced mammalian toxicity,

identify a key molecular regulator of toxicant-associated whole-

animal and DA neuron vulnerability, and suggest that the nematode

will be a useful in vivo tool to identify and characterize mediators of

MeHg-induced developmental and DA neuron pathologies.

Key Words: methylmercury; Parkinson’s disease; oxidative

stress; C. elegans; nematode; genetics; Nrf2; SKN-1; dopamine

neurodegeneration.

Methylmercury (MeHg) is a pervasive environmental

toxicant that primarily targets the central nervous system

(Aschner and Aschner, 1990; Friberg and Mottet, 1989).

Adults exposed to toxic levels of MeHg can display a range of

symptoms that includes a loss of physical coordination,

abnormal speech, and death. The developing fetus is

particularly vulnerable to MeHg because of the toxicant’s ease

in crossing the placenta and several fold greater toxicant

accumulation relative to the adult (Bakir et al., 1973; Clarkson

and Magos, 2006; Myers and Davidson, 1998; Takeuchi,

1982). Children exposed to toxic levels either pre- or

postnatally display developmental defects that are associated

with motor and sensory deficits and mental retardation

(Clarkson and Magos, 2006).

Although MeHg poisonings have been studied for decades,

the molecular bases for the toxicities are largely ill defined.

MeHg diffuses across most cellular membranes and intracel-

lular compartments and disrupts a wide array of cellular

functions including cell migration, neuronal axon guidance,

microtubule integrity, and cell signaling (Bridges and Zalups,

2005; Kunimoto and Suzuki, 1997; Limke et al., 2004).

Dopamine (DA) neurons are particularly vulnerable to the

toxicant as exposure to MeHg increases DA release and

decreases DA reuptake and catabolism (Dreiem et al., 2009;

Newland et al., 2008). Intriguingly, anecdotal evidence

suggests that MeHg may be an environmental toxicant that

contributes to the development of Parkinson’s disease (PD),

a progressive neurodegenerative disorder that results in motor

deficits, mitochondria dysfunction, and loss of DA neurons in

the substantia nigra (Nass and Przedborski, 2008; Petersen

et al., 2008). The mitochondria are also likely an intracellular

target of MeHg as the toxicant has been shown to disrupt

respiration, oxidative phosphorylation, and calcium regulation,

as well as causing an increase in reactive oxygen species (ROS)

and oxidative stress (Limke et al., 2004; Yu et al., 2010).

Vertebrate studies indicate that MeHg induces the phase II

class of detoxification enzymes glutathione-S-transferases

(GSTs) (Di Simplicio et al., 1993; Goering et al., 2000;

Samali and Orrenius, 1998; Yu et al., 2010). GSTs catalyze the

conjugation of a broad range of electrophilic substrates to the

thiol group of reduced glutathione molecules which typically

inactivates the electrophile and prepares it for excretion (Wilce

and Parker, 1994). Nrf2, a leucine zipper class transcription

factor, regulates a number of detoxifying enzymes including

PD-associated proteins and GSTs (Kensler et al., 2007; Taylor

� The Author 2010. Published by Oxford University Press on behalf of the Society of Toxicology. All rights reserved.For permissions, please email: [email protected]

at Indiana University S

chool of Medicine Libraries on M

ay 6, 2011toxsci.oxfordjournals.org

Dow

nloaded from

http://toxsci.oxfordjournals.org/

et al., 2008; von Otter et al., 2010). MeHg also induces Nrf2

expression, overexpression of the transcription factor inhibits

MeHg-induced cell death, and Nrf2-deficient hepatocytes and

primary cortical neurons are vulnerable to MeHg and PD-

associated toxicants (Lee et al., 2003; Toyama et al., 2007).

The nematode Caenorhabditis elegans is a powerful model

system to explore the molecular basis of neurotoxicity and

human disease (Nass and Hamza, 2007; Nass et al., 2008;

Settivari et al., 2009). The molecular pathways and mechanisms

underlying basic biological processes such as cell development

and migration are highly conserved between the nematode and

humans. Both organisms have virtually an identical number of

genes, and almost all the known signaling and neurotransmitter

systems, including the dopaminergic (DAergic) system, identi-

fied in humans have counterparts in the worm (Nass et al.,2008). The similarities in the DAergic system between the worm

and humans has allowed for the successful recapitulation of both

the familial PD-associated genetic as well as toxicant-induced

DA neuron pathology found in vertebrate models of PD (Nass

and Przedborski, 2008; Nass et al., 2008). Molecular pathways

and components involved in cellular detoxification and cellular

stress in eukaryotes are also conserved in C. elegans including

GSTs, Heat Shock Proteins, cytochrome P450s, and the Nrf2

transcription factor orthologue SKN-1 (Oliveira et al., 2009;

Park et al., 2009).

In this study, we describe a novel C. elegans model for

MeHg toxicity and DA neuron degeneration. We show that that

chronic low exposure to MeHg confers developmental defects,

decrease in brood size, concentration-dependent animal death,

and DA neurodegeneration. Exposure to the toxicant increases

cellular ROS levels, induces the expression of specific GSTs,

and whole-animal vulnerability to the toxicant is mitigated by

the expression of the phase II transcription factor SKN-1.

Furthermore, we show for the first time that SKN-1 is

expressed in C. elegans DA neurons, and genetic knockdown

renders the neurons vulnerable to MeHg-induced cell death.

These studies indicate that the cellular and molecular response

to MeHg-induced toxicity is likely conserved between

the nematode and vertebrates and suggests that C. eleganswill be a powerful in vivo model to explore the molecular basis

of MeHg-induced developmental and DA neuron pathologies.

MATERIAL AND METHODS

Caenorhabditis elegans strains and maintenance. The following strains

were obtained from the C. elegans Genetics Center: wild-type (WT) Bristol N2;

NL2099 rrf-3(pk1426); OD70 unc-119(ed3) III; ItIs44 [pie-1p-

mCherry::PH(PLC1delta) þ unc119 (þ)]; RJ928, a genetic cross between

BY250 (Pdat-1::GFP) and NL2099 (Settivari et al., 2009). The C. elegans

strains were cultured on bacterial lawns containing either OP50 or NA22

bacteria on nematode growth medium (NGM) or 8P plates, respectively, at

20�C according to standard methods (Brenner, 1974; Hope, 1999).

Whole-animal vulnerability assay. Synchronized L1 stage worms were

obtained by hypochlorite treatment of gravid adults followed by incubation of

the embryos in M9 buffer for 18 h and washed at least 33 in dH2O as described

previously (Nass et al., 2002; Nass and Hamza, 2007). L1 staged worms were

incubated on 8P/NA22 plates at 20�C for 48 h until they reached L4 stage.

They were then transferred to 8P/NA22 plates containing various concen-

trations of MeHg (0–125lM) (CH3HgCl, 442534-5G-A; Sigma-Aldrich,

St Louis, MO) and grown for 48 h at 20�C. Worms were assayed for viability

by touching the nose of the animal with a metal pick. Worms that moved were

considered alive. Each of the experiments was performed at least in triplicate,

and the results were reported as mean ± SE.

Brood-size assay. Single synchronized L4 stage N2 worms were each

placed onto an 8P/NA22 plate containing various concentrations of MeHg

(0–15lM). Each worm was transferred to a fresh plate after 24 h and then twice

daily for 5 days. The total number of live worms on each plate was counted

following a 48- to 72-h incubation.

Larvae development rate assay. L1 synchronized N2 nematodes were

incubated on 8P/NA22 plates containing 0 or 10lM MeHg at 20�C. The plates

were examined every 12 h, and the time was recorded when the worms reached

the L4 and adult stages. Each of the experiments was repeated at least four times.

Embryonic development assay. Synchronized L1 stage OD70 worms were

incubated on 8P/NA22 growth plates containing ± 10lM MeHg for 72 h.

OD70 worms contain a fusion of the pleckstrin homology domain derived from

mammalian PLC1d1 to the red fluorescent protein mCherry, which binds to

a phosphoinositide lipid on the plasma membrane. Use of the pie-1 promoter

drives expression specifically in germ cells, allowing them to be visualized

under an inverted microscope (Zeiss Axioplan 2, Jena, Germany) (Axiovision

Release 4.5, Jena, Germany) (Audhya et al., 2005). Following the incubation

period, at least 50 worms were immobilized on 2% agarose pads with 2%

sodium azide, and the image of the embryos was captured within the young

adult using an inverted fluorescent microscope. Each of the experiments was

performed at least in triplicate.

RNA extraction and complementary DNA synthesis. Synchronized L4

stage worms were incubated on 8P/NA22 media plates containing 25lM MeHg

and collected by washing with water, Trizol added to pellet, and frozen

at �80�C until RNA purification. Total RNA was isolated from a synchronized

C. elegans population using Trizol reagent as described previously with minor

modifications (Novillo et al., 2005). Briefly, the worm pellets were resuspended

in Trizol (1 ml/100 ll compact worm pellet), the protein and other impurities

were separated from nucleic acids using chloroform, and the RNA was

precipitated with isopropyl alcohol. The RNA pellet was washed with 75%

ethanol, air-dried and dissolved in RNase-free water, and stored at �80�C until

used for further analysis. The RNA concentrations were measured using an

ND-1000 spectrophotometer (Nanodrop Technology, Wilmington, DE). One

microgram of total RNA was reverse transcribed to complementary DNA

(cDNA) using the iScript cDNA Synthesis Kit and following the manufac-

turer’s instructions (Bio-Rad, Hercules, CA). The cDNA was further purified

using Microcon YM30 filters (Millipore Corp., Bedford, MA) and measured

using an ND-1000 spectrophotometer.

qPCR measurements. Gene-specific primers were designed using Primer3

software, and primers were designed to be exon spanning to avoid amplification

of contaminating genomic DNA. Glyceraldehyde-3-dehydrogenase (GAPDH)

was selected as the housekeeping gene as its expression does not change as

a result of MeHg treatment. Primers used to determine changes in gene

expression are found in Supplementary figure 1. Real-time PCR was performed

using 2x SYBR Green PCR master mix and the ABI Prism 7500 sequence

detection system (Applied Biosystems, Warrington, UK). Gene expression

studies were performed in triplicate and the formation of a single PCR product

was confirmed using dissociation curves. Negative controls with the primers

consisted of all the components of PCR mix except cDNA. Relative fold

change in gene expression for each gene was calculated using normalized

CT values (the cycle number at which the fluorescence passes the threshold).

ROS analysis. Total ROS formation was evaluated in the whole worm

using 2,7-dichlorodihydro-fluorescein-diacetate (H2-DCF-DA). ROS oxidize

the dye to the fluorescent 2,7-dichlorofluorescein, and the level of fluorescence

614 VANDUYN ET AL.




Dow

nloaded from

Supplementary figure 1


corresponds to cellular ROS levels. ROS levels were determined following

previously established protocols with slight modifications (Kampkotter et al.,

2007; Schulz et al., 2007). Briefly, synchronized L4 worms were exposed to

25lM MeHg as described above for 8 h and washed 33 with M9 buffer. The

animals were suspended in M9, and 50 ll of the suspension was incubated in

50 ll freshly prepared 100lM H2-DCF-DA (for a final concentration of 50lM)

in four replicates in a 96-well plate. Control wells included worms from each

treatment without H2-DCF-DA or wells containing H2-DCF-DA without

worms. The basal fluorescent signal from each well was measured at excitation

and emission wavelengths of 485 and 520 nm, respectively, using a Tecan

spectrophotometer (Tecan Spectrafluor Plus, Durham, NC). Following the

initial reading, plates were incubated on a shaker at room temperature for an

hour and were then measured following earlier conditions. Fluorescence values

from the control wells were subtracted from the corresponding signal value of

each well after the final measurement and the change in fluorescence was

determined by subtracting the initial value from the final value for each well.

All assays were performed in triplicate.

Antibodies and Western analysis. Antibodies to amino acids 6–92 from

the putative C. elegans GST-38 protein (WP:CE15958) were generated using

Genomic Antibody Technology at Strategic Diagnostics, Inc. (SDI) (Newark,

DE). Rabbit polyclonal antibodies were further purified at SDI. GAPDH

(ab36840 Abcam, Cambridge, MA) was used as a loading control for Western

analysis. To prepare protein for Western blot analysis, synchronized L4 stage

worms exposed to MeHg were pelleted from media plates as described above:

150 ll of mito buffer (20mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic

acid [HEPES], pH 7.5, 250mM sucrose, 1mM ethylene diamine tetraacetic

acid, 1mM ethylene glycol tetraacetic acid, 10mM KCl, 1.5mM MgCl2,

1mM dithiothreitol, 0.1mM PMSF, 2 lg/ml leupeptin, 2 lg/ml pepstatin, 2 lg/

ml aprotinin) was added to 300–400 ll of pelleted worms and the tubes frozen

at �20�C until protein purification. Worm samples stored at �20�C were

thawed and homogenized on ice with 50–60 strokes with a 2-ml glass

homogenizer. The lysate was spun at 400 g at 4�C for 4 min, the supernatant

collected in a sterile tube, and protein concentration determined using the

Bradford assay with bovine gamma globulin as the standard. The samples were

diluted in NuPAGE LDS buffer (Invitrogen, Carlsbad, CA), heated at 95�C for

20 min, and total cell lysates (50 lg protein) were separated by SDS-

polyacrylamide gel electrophoresis (PAGE) and transferred to PVDF

membranes (Bio-Rad). Membranes were blocked with 5% nonfat dry milk

dissolved in TBST (Tris-buffered saline, 0.1% Tween-20) for 2 h at RT,

followed by incubation with the appropriate primary antibody dilution (anti-

GST-38 at 1:20,000; anti-GAPDH at 1:5,000) at 4�C overnight. The

membranes were washed three times at RT for 15 min and incubated with

horseradish peroxidase-conjugated secondary anti-rabbit IgG (611-1302;

Rockland, Gilbertsville, PA). The membrane was developed using enhanced

chemiluminescence (ECL) (Amersham Biosciences, Pittsburgh, PA), image

captured using Bio-Rad ChemiDoc XRS, and total-protein intensities were

measured using QuantityOne software (Bio-Rad).

RNA interference. RNA-mediated interference (RNAi) using the sensitive

strain NL2099 rrf-3(pk1426) were carried out on NGM plates containing 1mM

isopropyl b-D-thiogalactoside and 100 lg/ml ampicillin and seeded with HT115

(DE3), an RNase III-deficient Escherichia coli strain carrying L4440 vector

with the gene fragment (skn-1) (GeneService, Source BioScience, PLC,

Nottingham, UK) or empty vector (Addgene, Cambridge, MA) (Timmons and

Fire, 1998). Synchronized L1 stage NL2099 worms were transferred onto

RNAi plates and the feeding protocol was followed with slight modifications

(Kamath and Ahringer, 2003). As knockdown of skn-1 is embryonic lethal,

assays were performed with first-generation synchronized L1 larvae incubated

on RNAi plates at 20�C for 48 h. The L4 staged worms were then transferred

onto RNAi plates containing various [MeHg]’s (1–25lM), and the number of

live worms were counted 24 h later as described above. For DA neuron

degeneration assays, animals were incubated on plates containing MeHg

(0, 0.5, 1.0, and 2.0lM) at 20�C for 96 h, 50–60 animals at each concentration

were immobilized on 2% agarose pads with 2% sodium azide, and were scored

for DA neurodegeneration under fluorescent microscope (Leica MZ 16FA,

Switzerland). Worms were scored positive for DA neuron degeneration when

GFP in any part of the four cephalic dendrites (CEP; which run from nerve ring

to tip of the nose) was absent (Nass et al., 2002). For mRNA and protein

analysis, worms that had been growing on RNAi bacteria for 48 h were

transferred to RNAi plates containing 25lM MeHg for 4 h. Worms were

collected, and mRNA or protein was isolated for use in quantitative Polymerase

Chain Reaction (qPCR) or Western blotting as described above.

Immunohistochemistry. Primary C. elegans cultures were prepared as

previously described with slight modifications (Bianchi and Driscoll, 2006;

Settivari et al., 2009). Briefly, RNAi-sensitive (RJ928) L1 stage worms

expressing GFP in the DA neurons were grown on RNAi plates containing

control (HT115) or skn-1 (RNAi) bacteria until the gravid adult stage. The

gravid adults were then lysed with the synchronization solution, and the egg

pellet was washed using egg buffer (118mM NaCl, 48mM KCl, 2mM CaCl2,

2mM MgCl2, 25mM HEPES) (Settivari et al., 2009). The eggs were then

separated from the debris using a 60% sucrose solution, digested using 4 mg/ml

chitinase (Sigma), and the embryonic cells were dissociated using a syringe.

The embryonic cells were then resuspended in L-15 medium (containing 10%

fetal bovine serum and 1% pen/strep) and grown on polylysine-coated cover

slips at 20�C. Following growth for 72 h, cells were fixed in 4%

paraformaldehyde, permeabilized in 0.5% Triton X-100, and 1% normal

donkey serum was used as blocking buffer. The cells were then incubated with

either SKN-1 primary antibodies (1:1000) (sc-9244; Santa Cruz Biotechnology,

Santa Cruz, CA) at 4�C overnight (14 h), followed by incubation with a Cy5-

conjugated donkey anti-goat secondary antibodies (Millipore) (1:1000) at room

temperature for 1 h. Images were captured using confocal microscopy (Zeiss

LSM 510 microscope). To further confirm the specificity of SKN-1

immunoreactivity, the antibody and its complementary antigenic peptide (sc-

9244p; Santa Cruz Biotechnology) were mixed in a ratio of 1:5 and incubated

at 4�C for 14 h. Following incubation, primary cultures were stained with the

SKN-1 antibody-complementary peptide mixture for 4�C for 14 h and analyzed

as described above.

RESULTS

MeHg Confers Concentration-Dependent Animal Death

Vertebrate studies indicate that chronic exposure to MeHg

confers whole-animal and cellular death. In order to determine

whether chronic exposure to low concentrations of MeHg

affects C. elegans viability, we added synchronized WT N2 L4

larvae to agar media plates containing between 0 and 125lM

MeHg and determined the number of live animals 48 h later.

We find that MeHg causes a concentration-dependent loss of

viability, with an LC50 of approximately 95lM (Fig. 1). MeHg

exposure also appears to significantly reduce the number of

viable eggs on the plate once the animals reach adulthood (data

not shown). These results indicate that C. elegans are sensitive

to low concentrations of MeHg in a concentration-dependent

manner and suggest that the toxicant may affect embryo

viability and development.

MeHg Exposure Decreases Brood Size

Mammalian studies indicate that MeHg exposures can cause

a reduction in the number of offspring. Studies with rodents

have shown a decrease in progeny relatively to nonexposed

animals (Beyrouty et al., 2006; Hughes and Annau, 1976). To

determine whether exposure to MeHg can cause a decrease

in the number of progeny, we incubated L4 animals, the

INHIBITION OF DOPAMINE NEURON DEGENERATION BY SKN-1/NRF2 615




Dow

nloaded from


developmental stage immediately prior to self-fertilization, on

media plates containing low concentrations of MeHg. Every

24 h, the adult animals were moved to a new plate until the

adults ceased producing embryos, and the total number of

progeny on each plate was counted. As can be seen in

Figure 2A, exposure to low concentrations of MeHg

dramatically reduced the brood size. Growth on media plates

containing 2.5lM MeHg reduces the number of progeny by

almost 20%, whereas growth on agar containing 10lM reduces

the number of progeny by over 90%. These results indicate that

exposure to MeHg decreases the number of viable progeny

and, as in vertebrate studies, may affect maturation and cell

division.

MeHg Confers Developmental Delays and EmbryonicDefects

Mammalian and invertebrate studies indicate that exposure

to MeHg can cause delays in animal development and confer

embryonic defects (Carvalho et al., 2008; Curle et al., 1987;

Weis, 2009). In order to determine if MeHg can affect normal

development, we examined the rate of development of animals

grown on a low concentration of MeHg from L1 to the L4

stage. L1 animals exposed to 10lM MeHg took approximately

30% longer to reach adulthood at 20�C relative to non-MeHg–

exposed animals (56 vs. 78 h) (Fig. 2B). These results indicate

that MeHg confer delays in morphogenesis and gonadogenesis

and suggest that the toxicant may have a deleterious effect on

embryogenesis and cellular division. To determine whether

chronic MeHg exposure may affect early embryonic de-

velopment, we examined embryos during the early stages of

cell division in the adult hermaphrodite in vivo. Exposure of L1

animals to 10lM of MeHg until they reached adulthood results

in the generation of embryos with significant developmental

defects relative to control (Figs. 2C–F). Embryonic defects

were seen in nearly all the animals evaluated; some embryos

contain three nuclei within a single cell, suggesting defects in

mitosis (Figs. 2D–F). Taken together, these results indicate that

exposure of C. elegans to MeHg has deleterious effects on

animal growth and development.

MeHg Exposure Increases ROS Levels

Although the origin of the cellular stress induced by MeHg

is not well defined, the toxicant is believed to generate an

increase in cellular ROS (InSug et al., 1997; Limke et al.,

2004). To determine if MeHg may also increase ROS levels in

C. elegans, we compared total ROS levels in control and

MeHg-exposed animals. L4 animals were incubated on plates

containing 25lM MeHg for 8 h, and ROS levels were

determined using the ROS-dependent dye DCF as previously

described (Settivari et al., 2009). We find that a brief exposure

to MeHg confers over a twofold increase in cellular ROS

relative to non–MeHg-exposed animals (Fig. 3A). These

results suggest that oxidative stress may contribute to

C. elegans cellular sensitivity to MeHg.

MeHg Induces GST Expression

An increase in cellular ROS levels and oxidative stress can

induce GST expression in an attempt to maintain normal

cellular and metabolic processes. In order to determine if

C. elegans GSTs are induced following exposure to MeHg, we

utilized qPCR and evaluated mRNA levels of a number of

GSTs, some of which have been associated with toxicant-

induced gene expression changes, following sublethal expo-

sures to MeHg. L4 animals exposed to 25lM MeHg for 2 h

display a significant increase in a number of GSTs expression

including gst-4, gst-5, gst-12, gst-21, and gst-38 (Fig. 3B).

Animals exposed under these conditions and then incubated on

non-MeHg media do not demonstrate reduced viability relative

to nonexposed animals, indicating that these growth conditions

are not lethal (data not shown). mRNA levels for gst-5 and

gst-38 also increased dramatically, up to 10-fold (over 50-fold

relative to nonexposed animals), when exposed for an

additional 6 h (Fig. 3B). Taken together, these results indicate

that C. elegans GSTs are induced following MeHg exposure.

MeHg-Associated Induction of gst-4 and gst-38 Is Dependenton skn-1

Caenorhabditis elegans hyperbaric stress microarray studies

indicate that gst-4 and gst-38 mRNA inductions are largely

FIG. 1. MeHg exposure confers concentration-dependent animal death.

WT (N2) L4 nematodes (� 30 worms/condition, six replicates) were grown on

8P plates with various concentrations of MeHg, and the number of live animals

was determined 48 h later. Data were analyzed using one-way ANOVA

followed by Dunnett’s multiple comparison test. The number of live animals

was significantly different from control (0lM MeHg) at 100 and 125lM

(*p < 0.01).

616 VANDUYN ET AL.




Dow

nloaded from


dependent on the transcription factor skn-1 (Park et al., 2009).

In order to determine whether MeHg-induced gst-4 and gst-38expression is skn-1 dependent, mRNA was isolated from

RNAi-sensitive nematodes incubated on media plates contain-

ing bacteria expressing skn-1 double-stranded RNA (dsRNA)

following exposure to 25lM MeHg, and gst-4 and gst-38expression levels were determined by RT-PCR. A brief 4-h

incubation in the presence of MeHg results in an approximate

15-fold and 55-fold decrease in gst-4 and gst-38 mRNA levels

relative to RNA levels from worms grown on control bacteria,

respectively (Fig. 4A). These results indicate that MeHg-

associated induction of gst-4 and gst-38 mRNAs is largely

dependent on the expression of skn-1 and suggests that the

corresponding proteins levels may also be skn-1 dependent. To

determine if GST-38 protein expression is also dependent on

skn-1, we generated antibodies to GST-38 and determined

protein levels following exposure to 25lM MeHg for 4 h.

Consistent with the induction of gst-38 mRNA expression,

exposure to the toxicant results in an approximate 12-fold

increase in GST-38 protein levels (Fig. 4B; data not shown).

FIG. 2. Chronic exposure to MeHg confers reproductive and developmental defects in Caenorhabditis elegans. MeHg exposure results in decreases in

C. elegans brood size (A). L4 animals (� 15 worms/condition) were placed on 8P/NA22 plates and allowed to lay eggs, and every 24 h, the parent worm was

transferred to a new plate until the animal terminated laying eggs (approximately 5 days). The total number of progeny from each plate was counted for each animal

and combined to determine the animal’s brood size. MeHg delays animal development (B). L1 nematodes (� 100 worms/condition) were placed on 8P/NA22

plates containing 10lM MeHg, and the time to reach adulthood was determined. Exposure to MeHg causes embryonic defects (D–F). L1 nematodes expressing the

mCherry fluorophore behind the pie-1 promoter were grown to adulthood in the presence of water (C) or 10lM MeHg (D–F), and fluorescence (C, D, and F) or

differential interference contrast (E) image was evaluated. Relative to control (C), MeHg causes abnormal cell division (D), and some cells contain several nuclei

(E and F). Scale bar ¼ 50 lm. Data were analyzed using one-way ANOVA followed by Dunnett’s multiple comparison test. All individual treatments were

statistically significant (*p < 0.01) compared with control group (0lM MeHg).





Dow

nloaded from


Furthermore, MeHg-associated induction of the GST-38 pro-

tein appears to be highly dependent on skn-1 as genetic

knockdown of the transcription factor does not result in an

increase in GST-38 following exposure to MeHg (Fig. 4B).

Taken together, these results indicate that MeHg-associated

induction of gst-4 and gst-38 is largely skn-1 dependent and

suggests that skn-1 may inhibit cellular toxicity following

exposure to the toxicant.

skn-1 Inhibits MeHg-Induced Animal Death

Nrf2/SKN-1 has been shown to play a significant role in

inhibiting oxidative stress and cellular toxicity following

MeHg exposure in vertebrates. Considering that at least two

of the skn-1–dependent C. elegans GSTs are highly induced

following exposure to MeHg, we asked whether a genetic

knockdown of skn-1 may affect toxicant-induced animal

vulnerability. To determine whether a reduction in skn-1 may

increase sensitivity to MeHg, we grew animals on media plates

on bacteria containing either an empty vector or a vector

expressing skn-1 dsRNA to reduce skn-1 mRNA expression.

A reduction in skn-1 mRNA levels was confirmed by qPCR

and likely translates to lower skn-1 protein levels because

growth on bacteria expressing the dsRNA also results in a

skn-1 deletion phenotype of eggs containing dead embryos

(Simmer et al., 2003). Consistent with prior skn-1 studies,

genetic knockdown of the transcription factor at this animal’s

age does not reduce animal viability under nonoxidative stress

conditions (An and Blackwell, 2003) (Fig. 4C). Conversely,

skn-1 animals exposed to MeHg are up to 53 more sensitive to

the toxicant relative to the controls (Fig. 4C). These results

indicate that skn-1 expression inhibits MeHg-induced toxicity

in C. elegans and further suggests that the signaling cascades

involved in MeHg-induced cytotoxicity are conserved between

nematodes and humans.

SKN-1 Is Expressed in DA Neurons

The mammalian SKN-1 orthologue NRF2 is expressed in

DA neurons, and a decrease in gene expression in vitro and

in vivo renders the cells vulnerable to DA neurotoxins (Jakel

et al., 2007; Siebert et al., 2009). Detailed analysis of

endogenous SKN-1 expression and localization has not been

performed in the whole worm, but transgenic animals

overexpressing the three isoforms of SKN-1 fused to the

green fluorescent protein (SKN-1::GFP) display protein

expression in the intestines and the chemosensory

ASI neurons (Bishop and Guarente, 2007; Tullet et al.,2008). However, translational reporter fusions may not give a

complete representation of endogenous protein expression

levels and cellular localization (Boulin et al., 2006).

To determine whether the C. elegans DA neurons express

*

0

10

20

30

40

50

60

70

Control MeHg

Cha

nge

in fl

uore

scen

ce(p

er m

g pr

otei

n)

A

0

10

20

30

40

50

60

70

gst-1 gst-4

gst-5 gst-10

gst-12 gst-21

gst-38 Fo

ld c

hang

e(re

lativ

e to

con

trol)

*

* *

*

*

*

*

*

#

B

#

#

FIG. 3. MeHg exposure increases ROS levels and GST mRNA expression. MeHg exposure increases the levels of ROS (A). Synchronized L4 animals were

grown on 8P/NA22 plates containing 25lM MeHg for 8 h and then incubated with DCF-DA for 60 min. Change in fluorescence was normalized to protein

concentration. Shown are mean values of ± SE of four individual replicates. Mean ROS levels between MeHg and control groups were analyzed using a paired t-test,

and the difference was statistically significant (*p ¼ 0.02). MeHg exposure induces the expression of GST mRNAs (B). Caenorhabditis elegans were exposed to

25lM for 2 h (black bars) or 8 h (gray bars), mRNA extracted, and reverse-transcribed to cDNA. Relative gene expression changes of the GSTs were quantitated

using real-time PCR. The fold change in gene expression relative to GAPDH was calculated for each gene following the DDCT method. Shown are mean values

for ± SE of at least three individual replicates. The level of significance between the groups was calculated using paired t-test. MeHg induced (p < 0.03) gst-4,

gst-12, and gst-21 gene expression at 2- and at 8-h time point, gene expression of gst-4, gst-5, gst-12, gst-21, and gst-38 were increased (indicated by ‘‘*,’’

p < 0.05). To determine gene expression changes between 2- and 8-h time point, log fold-change values were analyzed using one-way ANOVA followed by

unpaired t-test. Transcript levels of gst-5, gst-12, and gst-38 were increased (indicated by #, p � 0.05) in worms exposed to MeHg for 8 h compared with worms

exposed for 2 h. Exposure to MeHg did not change GAPDH gene expression (p ¼ 0.76) in worms at either time point.

618 VANDUYN ET AL.




Dow

nloaded from


SKN-1, we generated C. elegans primary cultures from RJ928

animals that have robust expression of GFP in DA

neurons both in vivo and in vitro (Carvelli et al., 2004; Nass

et al., 2002). We used affinity-purified anti-SKN-1 to evaluate

cellular SKN-1 expression levels. SKN-1 immunoreactivity is

observed in all DA neurons (Figs. 5A–D). No specific

staining was observed in primary DA neurons or other

cells from animals in which skn-1 mRNA levels were reduced

using RNAi (Figs. 5E–H; data not shown). Furthermore,

SKN-1–associated DA neuron immunoreactivity was not

observed following coincubation of the cultures with

the SKN-1 antibody and the complementary antigenic

peptide, providing further evidence that the antibody is

binding to the SKN-1 protein in the DA neurons (Supple-

mentary fig. 2). Taken together, these results indicate that

SKN-1 is expressed in DA neurons and that a reduction in the

transcription factor mRNA levels by RNAi results in

a significant loss of SKN-1–immunoreactive protein expres-

sion in the DAergic cells.

SKN-1 Inhibits MeHg-Induced DA Neurodegeneration

The activation of Nrf2-dependent stress response pathways

have been shown to inhibit MeHg- and PD-associated

toxicant pathologies (Jakel et al., 2007; Taylor et al., 2008).

Considering SKN-1 inhibits MeHg-induced animal death and

is expressed in DA neurons, we asked whether the

transcription factor may also mitigate toxicant-associated

DA neuron vulnerability. Nematodes were grown on RNAi

bacteria to reduce expression of skn-1; reduction in mRNA

levels of skn-1 was confirmed by quantitative PCR (data not

shown). Animals were then exposed to various concentrations

of toxicant ranging from 0 to 2lM MeHg for 96 h, and DA

neuron integrity was assessed in vivo as previously described

(Settivari et al., 2009). We found that low chronic exposures

to MeHg caused a significant loss of DA neurons in animals

(up to 30% of the animals exposed to 1lM MeHg) with

a reduction of skn-1 mRNA within 96 h at all concentrations

tested (Figs. 6A and 6B). The DA neuron degeneration appear

similar to our prior PD-associated toxicant studies in which

we characterized the cellular pathology by loss of DA neuron

GFP expression in the CEP dendrites (since our prior studies

have correlated a loss of GFP in the dendrites with DA

neuronal death), DA levels, and loss of DA neuron integrity

by electron microscopy (Nass et al., 2002; Settivari et al.,2009). Animals exposed at these concentrations for up to

120 h did not display a decrease in animal viability, and

approximately a 10% decrease in viability following exposure

for 144 h (consistent with the decrease in longevity of the

skn-1 mutants [Oliveira et al., 2009]), suggesting that there is

not large-scale MeHg-associated cellular death following the

96 h exposure (Supplementary fig. 3). Taken together, these

results indicate that the DA neurons are vulnerable to MeHg,

and the expression of SKN-1 inhibits the toxicant-induced

DA neuron degeneration.

FIG. 4. SKN-1 induces MeHg-associated gst-4 and gst-38 expression and

decreases whole-animal toxicity. MeHg induces skn-1–dependent mRNA

expression (A). RNAi-sensitive nematodes (NL2099) were grown on bacteria

on NGM plates containing either an empty vector (black bars) or a vector

expressing dsRNA to skn-1 (gray bars) and were exposed to 25lM MeHg for

4 h, mRNA extracted, and reverse transcribed to cDNA. Relative gene

expression changes of the gst-4 and gst-38 were quantitated utilizing RT-PCR.

Shown are mean values for ± SE of three individual replicates. The fold change

in gene expression relative to GAPDH was calculated for each gene following

the DDCT method. Paired t-test was performed on log-transformed fold-change

values to determine changes in gene expression between worms exposed to

MeHg versus controls. Genetic knockdown of skn-1 decreased (p � 0.01) gst-4and gst-38 gene expression following MeHg exposure. MeHg-induced

expression of GST-38 protein is dependent on skn-1 (B). Animals were

exposed as in (A), and protein lysates were prepared and isolated as described

in the ‘‘Materials and Methods’’ section; 50 lg of the cell lysate protein was

resolved by SDS-PAGE, transferred to PVDF membranes, and probed with

antibodies specific for GST-38. Western blots were performed in triplicate, and

a representative image is shown. skn-1 mRNA knockdown increases whole-

animal vulnerability to MeHg (C). RNAi-sensitive L4 nematodes (NL2099)

(� 50 worms/condition) were grown on bacteria on media plates containing

either an empty vector (black bars) or a vector expressing dsRNA to skn-1 (gray

bars) for 48 h and then exposed to various concentrations of MeHg for 24 h,

and the number of live animals was determined as described in the ‘‘Materials

and Methods’’ section. Data were analyzed using one-way ANOVA followed

by Dunnett’s multiple comparison test. MeHg significantly induced animal

death at 25lM in the HT115-fed animals (p < 0.05) and at 10 and 25lM in the

skn-1 RNAi-fed animals (p < 0.01). At 10 and 25lM MeHg, animal death was

significantly increased in skn-1 RNAi worms compared with HT115 worms as

determined by Tukey’s test following one-way ANOVA (*p < 0.01).





Dow

nloaded from

Supplementary fig. 3




DISCUSSION

MeHg is a potent xenobiotic that is associated with human

developmental abnormalities, neurological dysfunction, and

embryonic defects (Budtz-Jorgensen et al., 2007; Grandjean

et al., 1997; Murata et al., 2004; Rodier, 1995). Vertebrate

studies indicate that low chronic exposure to MeHg confers

animal and cellular developmental delays and embryonic

defects, impairs cellular proliferation and migration, and toxic

exposures decrease the number of offspring and animal death

(Glover et al., 2009; Limke et al., 2004). Recent Drosophilastudies involving relatively low MeHg exposures also show

toxicant-induced inhibition of embryonic development and

delays, suggesting conservation of the molecular targets

across phyla (Rand et al., 2009). Our studies recapitulate key

hallmarks of MeHg-induced mammalian toxicity as low and

chronic exposure of the toxicant cause embryonic defects,

postembryonic developmental delays, a reduction in brood

size, and concentration-dependent animal death (Figs. 1

and 2).

FIG. 5. SKN-1 is expressed in DA neurons. Primary Caenorhabditis elegans cultures expressing GFP in the DA neurons were generated with WT (A–D) or

skn-1 knockdown animals (E–H), and fixation was performed as described under the ‘‘Materials and Methods’’ section. Primary cultures were incubated with

SKN-1 primary antibody followed by incubation with Texas Red–conjugated donkey anti-goat secondary antibodies. DIC images of (A) and (E) of WT and

skn-1RNAi cultures, respectively. DA neurons from WT and skn-1 knockdown animals expressing GFP driven by the dat-1 promotor (B) and (F), respectively.

SKN-1 expressed in DA neurons in WT animals (C) but not in skn-1 knockdown animals (G). (D) Overlay of (B–C) and (H) overlay of (F–G). Images were

observed under a Zeiss confocal microscope (Zeiss LSM 510). Scale bar represents 45 lm.

FIG. 6. SKN-1 inhibits MeHg-induced DA neuron degeneration. Synchronized L1 nematodes (RJ928) (� 50 worms/condition) were grown on 1.0lM MeHg

media plates containing either control (empty vector) or skn-1 RNAi bacteria (vector expressing dsRNA for skn-1) for 96 h, and worms were scored for DA neuron

degeneration as described in the ‘‘Materials and Methods’’ section. (A) GFP-expressing DA neurons (CEPs) within the head of a control worm exposed to MeHg.

(B) DIC image of the corresponding control animal in (A). (C) GFP-expressing DA neurons within the head of an skn-1 knockdown animal exposed to MeHg;

image chosen emphasizes significant loss of CEP cell bodies and dendrites. (D) DIC image of the corresponding control animal (C). (E) Quantification of DA

neuron integrity in control animals or skn-1 knockdown animals exposed to various concentrations MeHg. Shown are mean values of ± SE of three individual

replicates. Data were analyzed using two-way ANOVA. All comparisons between control and skn-1 knockdown animals were significant (p � 0.01) at 0.5, 1, and

2lM MeHg concentrations. Within the skn-1 RNAi group, all the MeHg concentration exposure groups were significantly different (p < 0.03) from the 0lM

MeHg group.

620 VANDUYN ET AL.




Dow

nloaded from


In vertebrate systems, glutathione (GSH) and the phase II

detoxification enzymes play significant roles in inhibiting

MeHg-induced cellular pathology. MeHg exposure has been

shown to induce gene expression of the rate-limiting enzyme

for GSH synthesis, c-glutamylcysteine synthetase, which

results in an increase in cellular GSH levels and cytoprotection

(Thompson et al., 2000). The toxicant also induces the

expression of GSTs that conjugate the compound to GSH,

allowing for cellular excretion through phase III detoxification

proteins. Caenorhabditis elegans contains over 40 putative

GSTs, and previous studies demonstrate that there are

significant gene expression changes for several of the GSTs

following exposure to neurotoxicant and oxidative stress

(Hasegawa et al., 2008; Liao and Freedman, 1998; Oliveira

et al., 2009; Park et al., 2009). Consistent with vertebrate

studies, exposure of C. elegans to MeHg results in a significant

and robust increase in mRNA levels of a number of GSTs, with

the highest induction in gst-4, gst-5, gst-12, and gst-38 (Fig. 3B).

Among the GSTs examined, gst-38, a GST that is expressed in

the intestines and nervous system, shows the largest change in

gene expression relative to non–toxicant-exposed animals, with

a 50-fold increase in mRNA levels, as well as a 12-fold increase

in GST-38 protein levels (Fig. 3B) (Hasegawa et al., 2008).

GST-38 expression levels have previously been shown to

increase fivefold or less following exposure to cadmium,

acrylamide, arsenite, or hyperoxia, suggesting that gst-38expression is likely a sensitive indicator of tissue-associated

oxidative stress (Hasegawa et al., 2008; Liao and Freedman,

1998; Oliveira et al., 2009; Park et al., 2009). Furthermore,

although care must be taken in comparing mRNA expression

changes across analysis platforms (microarray vs. RT-PCR), as

well as growth conditions (i.e., different medias and bacteria),

age and length of toxicant exposures, the greater than 10-fold

induction of gst-38 relative to those found in previously reported

toxicant exposure studies suggests that the C. elegans neurons

and/or intestinal cells may be particularly sensitive to MeHg.

Numerous phase II detoxification enzymes, including the

GSTs in mammals and C. elegans, are regulated by the

transcription factor Nrf2/SKN-1 (Oliveira et al., 2009; Yu

et al., 2010). In vertebrates and nematodes, oxidative stress can

trigger Nrf2/SKN-1 translocation from the cytoplasm to the

nucleus that results in the induction of cytoprotective proteins

(Sykiotis and Bohmann, 2010). In vertebrates, Nrf2-deficient

animals are also sensitive to MeHg and oxidative stress, and

exposure to MeHg confers an increase in Nrf2 expression

(Toyama et al., 2007). The most highly induced GSTs in our

C. elegans studies (gst-4, gst-5, gst-12, and gst-38) have at

least three canonical SKN-1–binding sites within 1 kb 5# of the

start ATG, and gene expression for each of these GSTs has

previously been shown to be at least partially dependent on

skn-1 (An and Blackwell, 2003; Park et al., 2009). Consistent

with MeHg-induced skn-1–dependent gene expression, knock-

down of skn-1 results in decreases in GST mRNA and protein

levels (Figs. 4A and 4B). Furthermore, a decrease in skn-1

mRNA levels in the nematode results in a dramatic increase in

MeHg sensitivity (Fig. 4C). Taken together, these results

indicate that SKN-1 and its downstream targets play an

essential role in mediating MeHg-induced cellular toxicity.

Developing and adult DA neurons are particularly sensitive to

MeHg, and recent studies suggest that the toxicant may be

a contributing factor in the development of PD (Dreiem et al.,2009; Newland et al., 2008; Petersen et al., 2008). Here we

show that SKN-1 is expressed in C. elegans DA neurons, and

genetic knockdown of skn-1 renders the DA neurons highly

vulnerable to MeHg in vivo, as 1lM of the toxicant causes DA

neuron degeneration in approximately 30% of the animals

examined (Figs. 5 and 6). These results indicate that SKN-1

plays a vital role in limiting MeHg-induced DA neuron toxicity.

It is interesting to speculate that in individuals who have reduced

levels of Nrf2, MeHg exposure could be an environmental risk

factor for the development of PD. Indeed, Nrf2 haplotypes have

been correlated with the propensity to develop PD (von Otter

et al., 2010). Furthermore, DNA polymorphisms in Nrf2 that

result in reduced cellular levels have been associated with

a number of human diseases that could be affected by

environmental toxicant exposures, consistent with Nrf2 expres-

sion levels potentially mediating DA neuron vulnerability and

the propensity to develop PD (Sykiotis and Bohmann, 2010).

SKN-1 likely contributes to limiting MeHg-induced DA

neuron degeneration through the upregulation of phase II

enzymes, although a reduction in SKN-1 may also adversely

affect the regulation of DA synthesis that could contribute to DA

neuron vulnerability. For example, Blackwell and colleagues

have recently shown through microarray expression profiles that

a reduction in skn-1 mRNA levels reduces GTP cyclohydrolase I

(GTPCH) and 6-pyruvoyl tetrahydrobiopterin synthase (PTPS)

gene expression (Oliveira et al., 2009). Both proteins are

involved in the synthesis of tetrahydrobiopterin (BH4) that is

required for the generation of tyrosine hydroxylase, the rate-

limiting step in DA synthesis, and reduced GTPCH and PTPS

expression could theoretically increase toxic metabolite levels

that contribute to DA neuron pathogenesis (Blau et al., 2001).

Consistent with a reduction in GTPCH or PTPS gene expression

and a concomitant increase in DA neuron pathogenesis,

a reduction in gene expression of both proteins is observed

prior to PD-associated behavioral changes and neurodegenera-

tion in a Drosophila PD model (Scherzer et al., 2003).

Furthermore, autosomal dominant human mutations in GTPCH

have been correlated with the development of parkinsonism in

children and adults, and cerebrospinal fluid from PD patients

have been shown to have reduced BH4 levels relative to control

subjects (Blau et al., 2001; Lovenberg et al., 1979; Nygaard

et al., 1992). Taken together, these studies suggest that the SKN-

1–dependent MeHg-induced DA neuron vulnerability may also

be because of dysregulation of enzymes involved in DA

synthesis or metabolism.

The concentrations of MeHg that C. elegans were exposed in

this study are environmentally relevant and similar to those





Dow

nloaded from


incorporated in vertebrate studies (Wang et al., 2009; Yu et al.,2010). Considering low and chronic exposures of 10lM MeHg

reduces whole-animal viability, brood size, and normal

embryonic development within 2–3 days, our studies suggest

that C. elegans is highly sensitive to the toxicant. This is in

sharp contrast to a number of other toxicity studies in

C. elegans (acute and chronic) that often require a 10- to

a 100-fold or greater toxicant concentration exposure relative to

vertebrates to facilitate toxicant transport through the worm’s

orifices or across the moderately impermeable cuticle; toxicant

concentrations within the cytoplasm though are often similar to

vertebrates (Nass and Hamza, 2007; Rand and Johnson, 1995).

It is likely that MeHg’s lipophilic properties facilitate its ease in

crossing the cuticle and cellular membranes and its intracellular

bioaccumulation (Lakowicz and Anderson, 1980; Nakada and

Imura, 1982; Parran et al., 2001). A recent phenomenological

study that describes effects of significantly higher concen-

trations of MeHg on C. elegans (100–1000lM) also reports

a decrease in nematode viability and larvae growth (Helmcke

et al., 2009); embryo or brood size defects were not apparent

possibly because of the length of the exposure period or the

severity of the insult that may reduce embryo viability. The

rapid and robust induction of GST mRNA (within 2 h of MeHg

exposure) in this study also further emphasizes the acute

sensitivity of the nematode to the toxicant. The induction of

these phase II enzymes likely increase animal survival until

other Nrf2/SKN-1–regulated events (i.e., GSH generation,

induction of multidrug resistance efflux proteins, etc.), or other

stress-associated pathways can reverse the cellular redox

imbalance to normal cellular homeostasis.

In summary, our findings describe a novel C. elegans model

for MeHg toxicity that demonstrates that low and chronic

exposure to MeHg confers embryonic defects, developmental

delays, decreases in brood size, and concentration-dependent

whole-animal death. We also show for the first time that the

stress-associated transcription factor SKN-1 is expressed in DA

neurons, specific SKN-1–dependent GSTs are induced follow-

ing exposure to MeHg, and a reduction in SKN-1 gene

expression increases MeHg-induced animal vulnerability and

confers DA neuron degeneration. The development of this

genetic model should facilitate the identification of genes and

molecular pathways in vivo involved in MeHg-induced

developmental defects and cellular dysfunction.

SUPPLEMENTARY DATA

Supplementary data are available online at http://toxsci

.oxfordjournals.org/.

FUNDING

National Institute of Environmental Health (R011ES014459)

(to R.N.); Vanderbilt University Toxicology Program Pilot

Grant (to R.N.); Saastamoinen Foundation (to G.W.); a PhRMA

Foundation Predoctoral Award (to N.V.).

ACKNOWLEDGMENTS

We gratefully appreciate the technical assistance of Ken

Grimes and Randy Hunter. We thank Lihsia Chen for her

invaluable assistance with the embryonic development assays.

We also thank Bill Atchison for valuable discussions. Some of

the strains were also provided by the Caenorhabditis GeneticsCenter, which is supported by the National Institutes of Health

Center for Research Resources.

REFERENCES

An, J. H., and Blackwell, T. K. (2003). SKN-1 links C. elegans mesendodermal

specification to a conserved oxidative stress response. Genes Dev. 17,

1882–1893.

Aschner, M., and Aschner, J. L. (1990). Mercury neurotoxicity: mechanisms of

blood-brain barrier transport. Neurosci. Biobehav. Rev. 14, 169–176.

Audhya, A., Hyndman, F., McLeod, I. X., Maddox, A. S., Yates, J. R., III.,

Desai, A., and Oegema, K. (2005). A complex containing the Sm protein

CAR-1 and the RNA helicase CGH-1 is required for embryonic cytokinesis

in Caenorhabditis elegans. J. Cell. Biol. 171, 267–279.

Bakir, F., Damluji, S. F., Amin-Zaki, L., Murtadha, M., Khalidi, A.,

al-Rawi, N. Y., Tikriti, S., Dahahir, H. I., Clarkson, T. W., Smith, J. C.,

et al. (1973). Methylmercury poisoning in Iraq. Science 181, 230–241.

Beyrouty, P., Stamler, C. J., Liu, J. N., Loua, K. M., Kubow, S., and

Chan, H. M. (2006). Effects of prenatal methylmercury exposure on brain

monoamine oxidase activity and neurobehaviour of rats. Neurotoxicol.

Teratol. 28, 251–259.

Bianchi, L., and Driscoll, M. (2006). Culture of embryonic C. elegans cells for

electrophysiological and pharmacological analyses (September 30, 2006).

WormBook. (The C. elegans Research Community, WormBook Ed.). doi/

10.1895/wormbook.1.122.1. Available at: http://www.wormbook.org.

Bishop, N. A., and Guarente, L. (2007). Genetic links between diet and

lifespan: shared mechanisms from yeast to humans. Nat. Rev. Genet. 8,

835–844.

Blau, N., Bonafe, L., and Thony, B. (2001). Tetrahydrobiopterin deficiencies

without hyperphenylalaninemia: diagnosis and genetics of dopa-responsive

dystonia and sepiapterin reductase deficiency. Mol. Genet. Metab. 74,

172–185.

Boulin, T., Etchberger, J., and Hobert, O. (2006). Reporter gene fusions (April

5, 2006). WormBook. (The C. elegans Research Community, WormBook

Ed.). doi/10.1895/wormbook.1.122.1. Available at: http://www.wormbook

.org.

Brenner, S. (1974). The genetics of Caenorhabditis elegans. Genetics 77,

71–94.

Bridges, C. C., and Zalups, R. K. (2005). Molecular and ionic mimicry and the

transport of toxic metals. Toxicol. Appl. Pharmacol. 204, 274–308.

Budtz-Jorgensen, E., Grandjean, P., and Weihe, P. (2007). Separation of

risks and benefits of seafood intake. Environ. Health Perspect. 115, 323–327.

Carvalho, M. C., Nazari, E. M., Farina, M., and Muller, Y. M. (2008).

Behavioral, morphological, and biochemical changes after in ovo exposure

to methylmercury in chicks. Toxicol. Sci. 106, 180–185.

Carvelli, L., McDonald, P. W., Blakely, R. D., and Defelice, L. J. (2004).

Dopamine transporters depolarize neurons by a channel mechanism. Proc.

Natl. Acad. Sci. U.S.A. 101, 16046–16051.

622 VANDUYN ET AL.




Dow

nloaded from

Supplementary data



http://www.wormbook.org

http://www.wormbook.org


Clarkson, T. W., and Magos, L. (2006). The toxicology of mercury and its

chemical compounds. Crit. Rev. Toxicol. 36, 609–662.

Curle, D. C., Ray, M., and Persaud, T. V. (1987). In vivo evaluation of

teratogenesis and cytogenetic changes following methylmercuric chloride

treatment. Anat. Rec. 219, 286–295.

Di Simplicio, P., Gorelli, M., Vignani, R., and Leonzio, C. (1993). The

differential modulation of the enzymes of glutathione metabolism. Indication

of overlapping effects of toxicity and repair in mouse liver and kidney after

dietary treatment with methylmercury and sodium selenite. Biol. Trace Elem.

Res. 36, 167–181.

Dreiem, A., Shan, M., Okoniewski, R. J., Sanchez-Morrissey, S., and

Seegal, R. F. (2009). Methylmercury inhibits dopaminergic function in rat

pup synaptosomes in an age-dependent manner. Neurotoxicol. Teratol. 31,

312–317.

Friberg, L., and Mottet, N. K. (1989). Accumulation of methylmercury and

inorganic mercury in the brain. Biol. Trace Elem. Res. 21, 201–206.

Glover, C. N., Zheng, D., Jayashankar, S., Sales, G. D., Hogstrand, C., and

Lundebye, A. K. (2009). Methylmercury speciation influences brain gene

expression and behavior in gestationally-exposed mice pups. Toxicol. Sci.

110, 389–400.

Goering, P. L., Fisher, B. R., Noren, B. T., Papaconstantinou, A., Rojko, J. L.,

and Marler, R. J. (2000). Mercury induces regional and cell-specific stress

protein expression in rat kidney. Toxicol. Sci. 53, 447–457.

Grandjean, P., Weihe, P., White, R. F., Debes, F., Araki, S., Yokoyama, K.,

Murata, K., Sorensen, N., Dahl, R., and Jorgensen, P. J. (1997). Cognitive

deficit in 7-year-old children with prenatal exposure to methylmercury.

Neurotoxicol. Teratol. 19, 417–428.

Hasegawa, K., Miwa, S., Isomura, K., Tsutsumiuchi, K., Taniguchi, H., and

Miwa, J. (2008). Acrylamide-responsive genes in the nematode Caeno-

rhabditis elegans. Toxicol. Sci. 101, 215–225.

Helmcke, K. J., Syversen, T., Miller, D. M., III., and Aschner, M. (2009).

Characterization of the effects of methylmercury on Caenorhabditis elegans.

Toxicol. Appl. Pharmacol. 240, 265–272.

Hope, I., Ed. (1999). C. elegans: A Practical Approach. New York, Oxford

University Press.

Hughes, J. A., and Annau, Z. (1976). Postnatal behavioral effects in mice after

prenatal exposure to methylmercury. Pharmacol. Biochem. Behav. 4,

385–391.

InSug, O., Datar, S., Koch, C. J., Shapiro, I. M., and Shenker, B. J. (1997).

Mercuric compounds inhibit human monocyte function by inducing

apoptosis: evidence for formation of reactive oxygen species, development

of mitochondrial membrane permeability transition and loss of reductive

reserve. Toxicology 124, 211–224.

Jakel, R. J., Townsend, J. A., Kraft, A. D., and Johnson, J. A. (2007). Nrf2-

mediated protection against 6-hydroxydopamine. Brain Res. 1144, 192–201.

Kamath, R. S., and Ahringer, J. (2003). Genome-wide RNAi screening in

Caenorhabditis elegans. Methods 30, 313–321.

Kampkotter, A., Pielarski, T., Rohrig, R., Timpel, C., Chovolou, Y.,

Watjen, W., and Kahl, R. (2007). The Ginkgo biloba extract EGb761

reduces stress sensitivity, ROS accumulation and expression of catalase and

glutathione S-transferase 4 in Caenorhabditis elegans. Pharmacol. Res. 55,

139–147.

Kensler, T. W., Wakabayashi, N., and Biswal, S. (2007). Cell survival

responses to environmental stresses via the Keap1-Nrf2-ARE pathway.

Annu. Rev. Pharmacol. Toxicol. 47, 89–116.

Kunimoto, M., and Suzuki, T. (1997). Migration of granule neurons in

cerebellar organotypic cultures is impaired by methylmercury. Neurosci.

Lett. 226, 183–186.

Lakowicz, J. R., and Anderson, C. J. (1980). Permeability of lipid bilayers to

methylmercuric chloride: quantification by fluorescence quenching of

a carbazole-labeled phospholipid. Chem. Biol. Interact. 30, 309–323.

Lee, J. M., Shih, A. Y., Murphy, T. H., and Johnson, J. A. (2003). NF-E2-

related factor-2 mediates neuroprotection against mitochondrial complex I

inhibitors and increased concentrations of intracellular calcium in primary

cortical neurons. J. Biol. Chem. 278, 37948–37956.

Liao, V. H., and Freedman, J. H. (1998). Cadmium-regulated genes from the

nematode Caenorhabditis elegans. Identification and cloning of new

cadmium-responsive genes by differential display. J. Biol. Chem. 273,

31962–31970.

Limke, T. L., Heidemann, S. R., and Atchison, W. D. (2004). Disruption of

intraneuronal divalent cation regulation by methylmercury: are specific

targets involved in altered neuronal development and cytotoxicity in

methylmercury poisoning? Neurotoxicology 25, 741–760.

Lovenberg, W., Levine, R. A., Robinson, D. S., Ebert, M., Williams, A. C., and

Calne, D. B. (1979). Hydroxylase cofactor activity in cerebrospinal fluid of

normal subjects and patients with Parkinson’s disease. Science 204, 624–626.

Murata, K., Weihe, P., Budtz-Jorgensen, E., Jorgensen, P. J., and Grandjean, P.

(2004). Delayed brainstem auditory evoked potential latencies in 14-year-old

children exposed to methylmercury. J. Pediatr. 144, 177–183.

Myers, G. J., and Davidson, P. W. (1998). Prenatal methylmercury exposure

and children: neurologic, developmental, and behavioral research. Environ.

Health Perspec.t 106(Suppl. 3), 841–847.

Nakada, S., and Imura, N. (1982). Uptake of methylmercury and inorganic

mercury by mouse glioma and mouse neuroblastoma cells. Neurotoxicology

3, 249–258.

Nass, R., Hall, D. H., Miller, D. M., III., and Blakely, R. D. (2002).

Neurotoxin-induced degeneration of dopamine neurons in Caenorhabditis

elegans. Proc. Natl. Acad. Sci. U.S.A. 99, 3264–3269.

Nass, R., and Hamza, I. (2007). The nematode Caenorhabditis elegans as a model

to explore toxicology in vivo: solid and axenic growth culture conditions and

compound exposure parameters. Curr. Protoc. Toxicol. 1.9.1–1.9.18.

Nass, R., Merchant, K. M., and Ryan, T. (2008). Caenohabditis elegans in

Parkinson’s disease drug discovery: addressing an unmet medical need. Mol.

Interv. 8, 284–293.

Nass, R. and Przedborski, S., Eds. (2008). Parkinson’s Disease: Molecular and

Therapeutic Insights from Model Systems, 1st ed. Elsevier Academic Press.

Newland, M. C., Paletz, E. M., and Reed, M. N. (2008). Methylmercury and

nutrition: adult effects of fetal exposure in experimental models. Neuro-

toxicology 29, 783–801.

Novillo, A., Won, S.-J., Li, C., and Callard, I. P. (2005). Changes in nuclear

receptor and vitellogenin gene expression in response to steroids and heavy

metal in Caenorhabditis elegans. Int. Comp. Biol. 45, 61–71.

Nygaard, T. G., Takahashi, H., Heiman, G. A., Snow, B. J., Fahn, S., and

Calne, D. B. (1992). Long-term treatment response and fluorodopa positron

emission tomographic scanning of parkinsonism in a family with dopa-

responsive dystonia. Annu. Neurol. 32, 603–608.

Oliveira, R. P., Porter Abate, J., Dilks, K., Landis, J., Ashraf, J., Murphy, C. T.,

and Blackwell, T. K. (2009). Condition-adapted stress and longevity gene

regulation by Caenorhabditis elegans SKN-1/Nrf. Aging Cell 8, 524–541.

Park, S. K., Tedesco, P. M., and Johnson, T. E. (2009). Oxidative stress and

longevity in Caenorhabditis elegans as mediated by SKN-1. Aging Cell 8,

258–269.

Parran, D. K., Mundy, W. R., and Barone, S., Jr. (2001). Effects of

methylmercury and mercuric chloride on differentiation and cell viability

in PC12 cells. Toxicol. Sci. 59, 278–290.

Petersen, M. S., Halling, J., Bech, S., Wermuth, L., Weihe, P., Nielsen, F.,

Jorgensen, P. J., Budtz-Jorgensen, E., and Grandjean, P. (2008). Impact of

dietary exposure to food contaminants on the risk of Parkinson’s disease.

Neurotoxicology 29, 584–590.

Rand, J. B., and Johnson, C. D. (1995). Genetic pharmacology: interactions

between drugs and gene products in Caenorhabditis elegans. Methods Cell.

Biol. 48, 187–204.





Dow

nloaded from


Rand, M. D., Dao, J. C., and Clason, T. A. (2009). Methylmercury disruption of

embryonic neural development in Drosophila. Neurotoxicology 30, 794–802.

Rodier, P. M. (1995). Developing brain as a target of toxicity. Environ. Health

Perspect. 103(Suppl. 6), 73–76.

Samali, A., and Orrenius, S. (1998). Heat shock proteins: regulators of stress

response and apoptosis. Cell Stress Chaperones 3, 228–236.

Scherzer, C. R., Jensen, R. V., Gullans, S. R., and Feany, M. B. (2003). Gene

expression changes presage neurodegeneration in a Drosophila model of

Parkinson’s disease. Hum. Mol. Genet. 12, 2457–2466.

Schulz, T. J., Zarse, K., Voigt, A., Urban, N., Birringer, M., and Ristow, M.

(2007). Glucose restriction extends Caenorhabditis elegans life span by

inducing mitochondrial respiration and increasing oxidative stress. Cell

Metab. 6, 280–293.

Settivari, R., Levora, J., and Nass, R. (2009). The divalent metal transporter

homologues SMF-1/2 mediate dopamine neuron sensitivity in Caenorhabdi-

tis elegans models of manganism and parkinson disease. J. Biol. Chem. 284,

35758–35768.

Siebert, A., Desai, V., Chandrasekaran, K., Fiskum, G., and Jafri, M. S. (2009).

Nrf2 activators provide neuroprotection against 6-hydroxydopamine toxicity

in rat organotypic nigrostriatal cocultures. J. Neurosci. Res. 87, 1659–1669.

Simmer, F., Moorman, C., van der Linden, A. M., Kuijk, E., van den

Berghe, P. V., Kamath, R. S., Fraser, A. G., Ahringer, J., and Plasterk, R. H.

(2003). Genome-wide RNAi of C. elegans using the hypersensitive rrf-3

strain reveals novel gene functions. PLoS Biol. 1, E12.

Sykiotis, G. P., and Bohmann, D. (2010). Stress-activated cap#n#collar

transcription factors in aging and human disease. Sci. Signal 3, re3.

Takeuchi, T. (1982). Pathology of Minamata disease. With special reference to

its pathogenesis. Acta Pathol. Jpn 32(Suppl. 1), 73–99.

Taylor, R. C., Acquaah-Mensah, G., Singhal, M., Malhotra, D., and Biswal, S.

(2008). Network inference algorithms elucidate Nrf2 regulation of mouse

lung oxidative stress. PLoS Comput. Biol. 4, e1000166.

Thompson, S. A., White, C. C., Krejsa, C. M., Eaton, D. L., and

Kavanagh, T. J. (2000). Modulation of glutathione and glutamate-L-cysteine

ligase by methylmercury during mouse development. Toxicol. Sci. 57,

141–146.

Timmons, L., and Fire, A. (1998). Specific interference by ingested dsRNA.

Nature 395, 854.

Toyama, T., Sumi, D., Shinkai, Y., Yasutake, A., Taguchi, K., Tong, K. I.,

Yamamoto, M., and Kumagai, Y. (2007). Cytoprotective role of Nrf2/Keap1

system in methylmercury toxicity. Biochem. Biophys. Res. Commun. 363,

645–650.

Tullet, J. M., Hertweck, M., An, J. H., Baker, J., Hwang, J. Y., Liu, S.,

Oliveira, R. P., Baumeister, R., and Blackwell, T. K. (2008). Direct

inhibition of the longevity-promoting factor SKN-1 by insulin-like signaling

in C. elegans. Cell 132, 1025–1038.

von Otter, M., Landgren, S., Nilsson, S., Celojevic, D., Bergstrom, P.,

Hakansson, A., Nissbrandt, H., Drozdzik, M., Bialecka, M., Kurzawski, M.,

et al. (2010). Association of Nrf2-encoding NFE2L2 haplotypes with

Parkinson’s disease. BMC Med. Genet. 11, 36.

Wang, L., Jiang, H., Yin, Z., Aschner, M., and Cai, J. (2009). Methylmercury

toxicity and Nrf2-dependent detoxification in astrocytes. Toxicol. Sci. 107,

135–143.

Weis, J. S. (2009). Reproductive, developmental, and neurobehavioral effects

of methylmercury in fishes. J. Environ. Sci. Health C Environ. Carcinog.

Ecotoxicol. Rev. 27, 212–225.

Wilce, M. C. J., and Parker, M. W. (1994). Structure and function of

glutathione S-transferases. Biochim. Biophys. Acta 1205, 1–18.

Yu, X., Robinson, J. F., Sidhu, J. S., Hong, S., and Faustman, E. M. (2010).

A system-based comparison of gene expression reveals alterations in

oxidative stress, disruption of ubiquitin-proteasome system and altered cell

cycle regulation after exposure to cadmium and methylmercury in mouse

embryonic fibroblast. Toxicol. Sci. 114, 356–377.

624 VANDUYN ET AL.




Dow

nloaded from


skn-1/nrf2 inhibits dopamine neuron degeneration in a...

Documents