Int. J. Mol. Sci. 2011, 12, 5213-5237; doi:10.3390/ijms12085213
International Journal of
Mechanisms of Mycotoxin-Induced Neurotoxicity through
Oxidative Stress-Associated Pathways
Kunio Doi 1,2,* and Koji Uetsuka 1
1 Nippon Institute for Biological Science, 9-2221-1, Shin-Machi, Ome, Tokyo 198-0024, Japan
2 Graduate School of Agricultural and Life Sciences, The University of Tokyo, 1-1-1, Yayoi, Bunkyo,
Tokyo 113-8657, Japan
* Author to whom correspondence should be addressed; E-Mail: email@example.com;
Tel.: +81-428-33-1086; Fax: +81-428-31-6166.
Received: 7 June 2011; in revised form: 21 July 2011 / Accepted: 4 August 2011 /
Published: 15 August 2011
Abstract: Among many mycotoxins, T-2 toxin, macrocyclic trichothecenes, fumonisin B1
(FB1) and ochratochin A (OTA) are known to have the potential to induce neurotoxicity in
rodent models. T-2 toxin induces neuronal cell apoptosis in the fetal and adult brain.
Macrocyclic trichothecenes bring about neuronal cell apoptosis and inflammation in the
olfactory epithelium and olfactory bulb. FB1 induces neuronal degeneration in the cerebral
cortex, concurrent with disruption of de novo ceramide synthesis. OTA causes acute
depletion of striatal dopamine and its metabolites, accompanying evidence of neuronal cell
apoptosis in the substantia nigra, striatum and hippocampus. This paper reviews the
mechanisms of neurotoxicity induced by these mycotoxins especially from the viewpoint
of oxidative stress-associated pathways.
Keywords: neurotoxicity; T-2 toxin; macrocyclic trichothecenes; fumonisin B1; ochratoxin A
Mycotoxins are fungal metabolites known to be harmful toward human and animal health. To
date, disorders caused by mycotoxins have been reported in digestive, urinary, immune and
reproduction systems , and many in vivo and in vitro studies have been performed in order to clarify
the mechanisms of mycotoxin-induced toxicity in these systems. Recently, Surai et al.  described
Int. J. Mol. Sci. 2011, 12 5214
that, in many cases, membrane-active properties of various mycotoxins determine their toxicity and
incorporation of mycotoxins into membrane structures causes various detrimental changes, resulting in
alterations in second messenger systems through damaging membrane receptors. In addition,
detrimental effects of mycotoxins on DNA and RNA and protein synthesis together with proapoptotic
action further compromise important metabolic pathways and consequently changes in physiological
functions including growth, development and reproduction occur. During the last decades, the
importance of oxidative stress and lipid peroxidation in all these processes have been pointed out by
many researchers [2,3].
Compared with the amount of research on digestive, urinary, immune and reproduction systems,
there are few reports of the effects of mycotoxins on neuronal tissues. This paper reviews the
mechanisms of neurotoxicity experimentally induced in rats and mice by T-2 toxin, macrocyclic
trichothecenes, fumonisin B1 (FB1) and ochratoxin A (OTA) especially from the viewpoint of
oxidative stress-associated pathways.
2. T-2 Toxin
T-2 toxin is a cytotoxic secondary fungal metabolite that belongs to thetrichothecene mycotoxin
family. They are produced by various species of Fusarium (F. sporotichioides, F. poae, F. equiseti,
and F. acuminatum), which can infect corn, wheat, barley and rice crops in the field or during
storage [4,5]. T-2 toxin is conjectured to be a major factor in alimentary toxic aleukia in humans 
and has been implicated in additional mycotoxicoses such as red mold disease in humans and
animals  and beanhull poisoning in horses .
T-2 toxin is a well-known inhibitor of protein synthesis through its high binding affinity to peptidyl
transferase which is an integral part of the 60 s ribosomal subunit [9–11]. Subsequent inhibition of the
peptidyl transferase reaction can trigger a ribotoxic stress response that activates c-Jun N-terminal
kinase (JNK)/p38 mitogen-activated protein kinases (MAPKs) . Moreover, T-2 toxin interferes
with the metabolism of membrane phospholipids and increases liver lipid peroxides [12,13].
Oral, parenteral and cutaneous exposures to T-2 toxin induce lesions in hematopoietic, lymphoid
and gastrointestinal tissues and suppress reproductive functions in domestic and laboratory
animals [14–17]. T-2 toxin can induce apoptosis in many types of cells bearing rapid rates of
proliferation [18–22]. T-2 toxin also induces apoptosis and fatty change in hepatocytes of mice
following the increased expression of both oxidative stress- and apoptosis-related genes (c-fos and
c-jun) . Moreover, prenatal exposure of rats to T-2 toxin induces apoptosis in maternal liver,
placenta and fetal liver following the increased expression of oxidative stress- and apoptosis-related
genes and decreased expression of lipid metabolism- and drug-metabolizing enzyme-related genes in
these tissues . Doi et al. [25,26] have reviewed the mode of occurrence and mechanisms of T-2
toxin-induced apoptosis in mice and rats.
To date, the effects of T-2 toxin on the central nervous system (CNS) have received limited
attention , and therefore, there are only a small number of reports of T-2 toxin-induced
neurotoxicity [28–30]. Boyd et al.  reported that low levels of T-2 toxin were responsible for the
changes in the metabolism of brain biogenic monoamines, and Wang et al.  showed that ingestion
of T-2 toxin leads to changes in amino acid permeability across the blood-brain barrier, which could
Int. J. Mol. Sci. 2011, 12 5215
lead to neurological effects observed in animals exposed to trichothecenes. T-2 toxin may be easily
distributed to the fetal brain, and induce fetal death and fetotoxicity mainly in the CNS and skeletal
system in addition to maternal toxicity [32–35].
Sehata et al.  have investigated the mechanisms of apoptosis induction in the fetal brain by oral
administration of T-2 toxin (2 mg/kg b.w.) to pregnant rats on day 13 of gestation. In their study, the
number of apoptotic neural progenitor cells in the telencephalon increased from 1 h and peaked at
12 h after T-2 toxin treatment. Microarray analysis revealed that the expression of heat shock protein
70 (HSP70), metallothionein (MT)-2 and 1, and heme oxygenase-1 (HO-1) was strongly elevated by
T-2 toxin at 12 h, and the expression of the Cu, Zn-superoxide dismutase (Cu, Zn-SOD) gene also
increased at 24 h after T-2 toxin treatment. This suggests that oxidative stress might be the main factor
behind the T-2 toxin-induced changes in the fetal brain. In addition, the gene expression of
liver stearyl-CoA desaturase and farnesyl diphosphate synthase genes which are involved in lipid
metabolism was suppressed by T-2 toxin in the fetal brain .
T-2 toxin suppresses drug metabolizing enzymes such as glutathione S-transferases (GSTs) [36–38].
In addition, a decreased expression in mitochondria-related genes, such as mitochondrial
NADH-dehydrogenase and cytochrome oxidase, has been reported in the fetal brain , suggesting a
dysfunction of the mitochondria. Since mitochondria play an important role in cell survival, these
changes in metabolism-related genes may also have a relationship to the induction of apoptosis.
In the study by Sehata et al.  on the fetal brain, the expression of MEKK1 gene increased at 12
and 24 h, and the expression of c-jun gene at 24 h after T-2 toxin treatment. These findings suggest
that the MAPK-JNK-c-jun pathway might be involved in T-2 toxin-induced apoptosis in the fetal
brain. Extracellular signal-related protein kinase (ERK) mediates cell growth and protects cells
from apoptosis, whereas stress-activated protein kinase (SAPK)/JNK and p38 MAPK inhibit cell
proliferation and may promote apoptosis . Each MAPK is activated by an upstream MAPK kinase,
including MEKK1, and JNK activates transcription factors such as c-fos and c-jun. MEKK1 may
induce apoptosis by causing a general deregulation of MAPK signaling , and JNK and c-jun are
important regulators of apoptosis in the nervous system .
Differing from the results in the maternal liver, placenta and fetal liver , the increase in
caspase-2 gene expression with no changes in caspase-9 and Bax-α gene expression was detected in
the fetal brain at 24 h after T-2 toxin treatment , suggesting an involvement of caspase-2 activation
in T-2 toxin-induced apoptosis in the fetal brain. Activation of caspase-2 is induced by reactive oxygen
species (ROS), and caspase-2 is said to play a crucial role in the control of apoptosis [42–44].
Although it is suggested that the p53-related mitochondrial pathway is involved in the T-2
toxin-induced apoptosis in the maternal and fetal livers , apoptosis induction in the fetal brain by
T-2 toxin seems to be independent of the p53-related pathway which is the most important pathway in
DNA-damaging agent-induced apoptosis of neural progenitor cells in the developing brain [45–49].
In microarray analysis on the fetal rat brain from dams exposed to T-2 toxin, the expression of
vascular endothelial growth factor (VEGF) gene increased at 12 and 24 h after T-2 toxin
treatment . VEGF is expressed in neurons and may play a role in the maintenance of neurons and
endothelial cells in the CNS . Therefore, the observed VEGF induction in the fetal brain might
indicate a protective reaction to the apoptotic changes in the fetal brain induced by T-2 toxin.
Int. J. Mol. Sci. 2011, 12 5216
Recently, Chaudhary and Rao  evaluated acute toxicity of dermal and subcutaneous exposure of
T-2 toxin on brain oxidative stress in adult mice. Mice were exposed to LD50 of T-2 toxin either by the
dermal (5.94 mg/kg b.w.) or subcutaneous (1.54 mg/kg b.w.) route and sacrificed at 1, 3 and 7 days
post-exposure. They reported that T-2 toxin-treated animals showed a time-dependent increase in ROS
generation, glutathione (GSH) depletion, lipid peroxidation and protein carbonyl content in the brain in
both routes of exposure. The gene expression profile of antioxidant enzymes showed a significant
increase in SOD and catalase via the percutaneous route and glutathione reductase (GR) and
glutathione peroxidase (GPx) via the subcutaneous route. This indicates that T-2 toxin induces
oxidative damage in adult mouse brain as well as in fetal rat brain. Lipid peroxidation may bring about
protein damage and inactivation of membrane-bound enzyme either through direct attack by free
radicals or through chemical modification by its end products . Indeed, as mentioned above,
protein carbonylation, a sign of oxidative damage, significantly increased in the mouse brain after
exposure to T-2 toxin.
Chaudhary and Roa  also investigated the role of nuclear factor erythroid 2-related factor (Nrf2)
and its downstream targets of phase II antioxidant/detoxifying enzymes in the mouse brain exposed to
T-2 toxin. Upon activation, Nrf2 binds to antioxidant responsive element sites in the promoter regions
of many detoxification and antioxidant genes, leading to coordinate regulation of downstream targets
that boost the cellular detoxification process and antioxidant potential [52,53]. In the study of
Chaudhary and Roa , however, Nrf2 and its downstream target genes were down-regulated, and
the involvement of Nrf2 in augmenting oxidative potential is not significant, although there is
3. Macrocyclic Trichothecenes
The fungus Stachybotrys chartarum, a saprophyte that grows on wet cellulose-containing building
materials including wallboard, ceiling tiles and cardboard is often found in low concentrations among
the mycoflora identified in water-damaged buildings [54–57]. Chronic indoor exposures to S.
chartarum and its products or components have been postulated to etiologically contribute to damp
building-related illnesses (DBRI) such as debilitating respiratory [58,59] and nonrespiratory symptoms
involving immune and neurological impairment [56,60–62]. Experimental rodent studies revealed
that, while this fungus is not infectious, airway exposure to spores of S. chartarum and its components
have the potential to evoke toxicity, inflammation and allergic sensitization in the upper and lower
respiratory tracts .
Two toxic “chemotypes” of S. chartarum exist. One chemotype elaborates highly toxic macrocyclic
trichotecene mycotoxins whereas a second chemotype produces less toxic atranones and simple
trichothecenes but no macrocyclic trichothecenes . The former mycotoxins are potent translational
inhibitors and stress kinase activators that appear to be a critical underlying cause for a number of
adverse effects. Notably, these toxins form covalent protein adducts in vitro and in vivo and,
furthermore, cause neurotoxicity and inflammation in the nose and brain of the mouse [64,65]. On the
other hand, the latter mycotoxins can induce pulmonary inflammation. Pestka et al.  have reviewed
the relationship between S. chartarum, trichothecene mycotoxins, and DBRI and proposed new
insights into a public health enigma.
Int. J. Mol. Sci. 2011, 12 5217
Besides inhibiting translation, macrocyclic trichothecenes as well as other trichothecenes can
simultaneously activate p38, JNK and ERK and MAPKs in vivo and in vitro [66–68] via a process
referred to as “ribotoxic stress” . In cell cultures, macrocyclic trichothecenes (Type D
trichotecenes) are 10–100 times more potent than Type A (e.g., T-2 toxin) or Type B (e.g.,
deoxynivalenol) trichothecenes at activating MAPKs, impairing leukocyte proliferation, or inducing
apoptosis [11,70–75]. The common ability of macrocyclic trichothecenes to cause protein synthesis
inhibition via binding to the 18S rRNA of the ribosomal large subunit  has been speculated to be a
major mechanism underlying induction of cell apoptosis by this group of trichothecenes. Moreover, the
potential of macrocyclic trichothecenes to covalently bind to proteins and possibly other
macromolecules has major implications relative to their absorption, metabolism, distribution, toxicity,
and potential allergenicity [64,65].
Satratoxin G (SG) is one of the most potent macrocyclic trichothecenes produced by S. chartarum 
and contributes to the above-mentioned DBRI. Roridin A (RA) is a commercially available
macrocyclic trichithecene used as a SG surrogate, and roridin L2 (RL2) is a putative biosynthetic
precursor of SG. While SG contains an intact macrocyclic ring linking C-4 to C-15, the precursor RL2
contains only an extended carbon chain linked at C-4 . RL2 is said to be nontoxic . Satratoxin
H (SH) is another macrocyclic trichothecene mycotoxin derived from the fungus S. chartaum. This
mycotoxin is one of the toxic constituents of the toxic mushroom, Podostoma cornu-damae .
Murine alveolar type II cells and alveolar macrophages are extremely sensitive to intratracheally
instilled S. chartarum spores . The methanol extract of a trichothecene-producing strain of
S. chartarum particularly up-regulates DNA damage-responsive and DNA repair genes in the murine
alveolar macrophage cell line MH-S early in the treatment, which are suggestive of genotoxic
stress . In a follow-up study, extract-induced apoptosis in MH-S cells was observed to precede
DNA damage . Moreover, both p38- and p53-mediated signaling events seem to occur in
S. chartarum-induced apoptosis of alveolar macrophages.
Macrocyclic tricothecenes also affect the upper respiratory tract (e.g., nasal airway). Using an
intranasal instillation model in adult C57BL/6J mice, Islam et al.  showed that SG exposure
specifically induced apoptosis of the olfactory sensory neurons (OSNs) and subsequent atrophy of the
olfactory epithelium (OE). Concurrently, there was bilateral atrophy of the olfactory nerve layer of the
olfactory bulbs (OBs) of the brain. In addition, SG induced an acute, neutrophilic rhinitis
and encephalitis. Similar findings have also been reported in mice intranasally instilled with RA .
In the ethmoid turbinates and OBs in the frontal brain in mice treated with SG, elevated
mRNA expression for the proinflammatory cytokines, TNF-α, IL-6 and IL-1, the chemokine
macrophage-inflammatory protein-2 (MIP-2), and the proapoptotic genes, Fas, FasL, p75NGFR, p53,
Bax, caspase-3 and CAD, was detected at 24 h post instillation (PI). In the same regions of mice
treated with RA, up-regulated mRNA expression of Fas, TNF-α, IL-6 and IL-1 and MIP-2 was
observed from 6 to 24 h PI, whereas expression of several other proapoptotic genes (p53, Bax, and
caspase-activated DNase) was detectable only at 24 h PI.
Following intranasal instillation of mice to SG  or RA , double-stranded RNA-activated
protein kinase (PKR) mRNA concentrations in the nasal turbinates were up-regulated in parallel with
OSN apoptosis. PKR, Bax and p53 have been previously reported to mediate apoptosis in murine
Int. J. Mol. Sci. 2011, 12 5218
OSNs [85–87]. PKR associates with the ribosome  and can selectively shut down translation
via phosphorylation of eukaryolic initiation factor 2α (eIF2α) as well as activate NF-κB .
Both TNF-α and Fas directly induce apoptosis in OE  and in OE organ cultures [91,92], and
SG- and RA-induced TNF-α and Fas mRNA expression precedes or is concurrent with OSN apoptosis,
induction of caspase-3 mRNA and caspase-3 activation [83,84]. The origins of induced TNF-α are
considered to be OSNs and adjacent cells in the OE which would promote autocrine or paracrine
responses, respectively. SG and RA also directly induces apoptosis in the OSNs by initiating
mitochondrial cell death via an intrinsic pathway involving p53 and Bax. Islam et al.  used the
PC12 rat pheochromocytoma cell models to elucidate potential mechanisms of SG-induced neuronal
cell death. In their experiment, SG-induced apoptosis occurred at 48 h after SG treatment, and the
expression of p53, and PKR, Bax and caspase-activated DNase mRNAs was significantly elevated
from 6 to 48 h after SG treatment. SG-induced p53 and Bax gene expression is known to drive nuclear
translocation of apoptosis-inducing factor (AIF), mitochondrial flavoprotein, in PC-12 cells .
In the study by Islam et al. , SG-induced apoptosis was not affected by inhibitors of oxidative
stress or MAPKs but was suppressed by the PKR inhibitor C16 and by PKR siRNA transfection. PKR
inhibition also blocked SG-induced apoptotic gene expression and AIF translocation but not caspase-3
activation. These results indicate that SG-induced apoptosis in PC12 neural cells is mediated by PKR
via a caspase-independent pathway possibly involving AIF translocation.
SH is thought to induce caspase-3 activation and apoptosis of PC12 cells through the activation of
p38 MAPK and JNK in a GSH-sensitive manner . Moreover, Nusuetrong et al.  carried out the
study to further elucidate the mechanisms by which SH induces cell death in PC12 cells. They reported
that SH causes apoptosis of serum-deprived PC12 cells within 24 h and that SH increases ROS
production and lipid peroxidation which are attenuated by incubation of cells with GSH. They
suggested that SH-induced increase in apoptosis of serum-deprived PC12 cells, may be partially
mediated through the generation of ROS. GSH, the most abundant intracellular thiol, plays an
important role in controlling the redox state of cells, and GSH is thought to play a role in apoptotic cell
death following its efflux through the GSH-specific membrane channels, carrier-mediated GSH
extrusion and oxidative stress [79,95].
The constant activation of inflammatory and apoptotic pathways at low levels of exposure in human
neurological system cells may amplify devastation to neurological tissues and lead to neurological
system cell damage from indirect events triggered by the presence of SH . This suggests that
individuals exposed to SH and microbial organisms resulting in a chronic immune response
(inflammation and oxidative stress) could have increased sensitivity to these agents, leading to neural
damage, further supporting previous in vivo studies demonstrating CNS tissue damage via inhalation
of fungal toxins . The process of inflammation is intended to repair injured tissues; however, this
mechanism tends to induce damage to nervous tissues when activated [98,99]. In this context, studies
on canines by Caldeón-Garcidueñas et al. [100,101] demonstrated increased iNOS, NF-κB, and TNF-α
production among other inflammatory and oxidative stress agents—leading to permanent damage of
DNA and CNS tissues due to passage of small particles via olfactory epithelium and lung tissue. The
fine particles reach CNS tissue via the olfactory bulb and into brain tissue (frontal cortex and cortical
tissues) demonstrating increases in β-amyloid plaques suggestive of pathogenesis similar to
Alzheimer’s disease [100,101].
Int. J. Mol. Sci. 2011, 12 5219
4. Fumonisin B1
Fumonisins belong to the relatively recently discovered group of mycotoxins produced by the
fungus Fusarium verticillioides (formerly F. moniliforme), a widespread fungal concomitant of various
cereals, predominantly corn [2,102]. In this case, FB1 is the most abundant and toxic; it has been linked
to a number of diseases in humans and animals [102,103].
The structures of FB1 and sphingolipids show marked similarities, which may be the reason why
FB1 drastically disrupts the normal sphingolipid metabolism . FB1 inhibits ceramide synthase, a
key enzyme in de novo sphingolipid biosynthesis and sphingolipid turnover, causing elevated levels of
free sphingolipid bases and sphingolipid base metabolites and lowered levels of ceramide [105,106].
FB1-induced inhibition of ceramide synthesis can result in a wide spectrum of changes in lipid
metabolism and associated lipid-depending signaling pathways, and it appears to be a major
contributor to the carcinogenic and other deleterious effects of FB1 .
FB1 is well known to cause equine leukoencephalomalacia (ELEM) [107,108]. This disease is
associated primarily with FB1 and is characterized by high mortality . In histopathological
examinations, pathognomonic focal necrotic lesions, located primarily in the subcortical white matter
are apparent . In addition, the elevation of free sphingoid bases after FB1 treatment has been
demonstrated in the brain of ELEM-diagnosed horses , suggesting that free sphingoid bases may
be important in FB1-related neurotoxicity.
Another emerging neurodevelopmental aspect of FB1 toxicity that implicates the consumption of
fumonisins in the etiology of neuronal tubule defects (NTD) in children has recently been
suggested . Treatment with FB1 causes NTDs in ex vivo neurulating mouse embryos  and
this effect is related to the folic acid receptor deficiency as a result of the FB1-dependent lipid rafts
The detrimental effects of FB1 on neuronal tissue have been shown in a number of reports indicating
its potential for direct neurotoxicity. For example, FB1 drastically inhibits axonal growth in cultured
hippocampal neurons , increases levels of sphinganine concentration in the forebrain and brain
stem of rats accompanying a concomitant demyelination in the forebrain , and disrupts
myelination in glial cells but not neurons in aggregating brain cell culture  and in developing
rats . FB1-dependent changes in neurotransmitter metabolite levels in different brain regions
of BALB/c mice  and in rat brain , and alteration of electrophysiological activity in rat
neocortex  are also reported.
Osuchowski et al.  carried out a study to compare the toxicity of FB1 in mouse brain after an
intracerebroventricular (icv) or subcutaneous (sc) infusion with total doses of 0, 10 or 100 μg/kg of
FB1. The icv infusion of FB1 led to neuronal degeneration in the cortex, concurrent with disruption of
sphingolipid metabolism, i.e., inhibition of de novo ceramide synthesis, stimulation of astrocytes, and
activation of proinflammatory cytokine signaling while the sc infusion of FB1 brought about partial
inhibition of sphingolipid metabolism in the cortex. From these results and the reports showing that
FB1 compromises the endothelial barrier function [121,122], it is suggested that there may be limited
blood-brain barrier transfer of FB1  and that FB1 may disrupt central nervous system homeostasis
when brain tissue is directly exposed to this mycotoxin.
Int. J. Mol. Sci. 2011, 12 5220
All cytokines analyzed in the study of Osuchowski et al.  are CNS-borne and are expressed
on-site by neurons (IL-1β and IL-6), astrocytes (IL-1β, IL-6, TNF-α and interferon-γ; IFN-γ) and
microglia (IL-1β, IL-6 and TNF-α) . IL-1β, IL-6 and TNF-α are primarily associated with
neuronal injury; thus neuronal damage will be accompanied by their elevated expression. In addition,
the increased expression of TNF-α and IL-1β also indirectly enhance neuronal damage via
ceramide-mediated signaling, since both cytokines activate brain neutral sphingomyelinase (nSMase),
and the role of ceramide-dependent neurodegeneration mediated by nSMase is reported [125,126]. The
immunocompetent cells and proinflammatory signaling are first being activated (i.e., astrocytes and
probably microglia) and then neurodegeneration follows in mouse brain after FB1 infusion .
During the last decades, studies aimed at clarifying the mechanisms of FB1-induced neurotoxicity in
cultured cells has been done mainly from the viewpoints of oxidative stress and/or apoptosis.
Stockmann-Juvala et al.  tried to characterize oxidative stress-related parameters induced by FB1
in three different neural cell lines, human SH-SY5Y neuroblastoma, rat C6 glioblastoma and mouse
GT1-7 hypothalamic cells. In their study, FB1 caused a dose-dependent increase of ROS production in
C6 and GT1-7 cells but was without an effect in SH-SY5Y cells. Decreased GSH levels, increased
malon dialdehyde (MDA)-formation, indicative of lipid peroxidation, and necrotic cell death were
observed in all cell lines after incubation with FB1. From these results, they concluded that FB1
induces oxidative stress in human, rat and mouse neural cell cultures. They also suggested that FB1 is
cytotoxic to neural cells only at high concentrations in vitro, although systemic toxicity, which may be
caused by the inhibition of ceramide synthase, takes place already at very low concentrations of
Mobio et al. [128–130] reported that in rat C6 glioma cells, FB1 inhibits protein synthesis, causes
DNA fragmentation and cell death, increases 8-hydroxy-2’-deoxyguanosine (8-OH-dG), and induces
lipid peroxidation, and that cytotoxic concentrations of FB1 induce cell cycle arrest in C6 cells (in the
G2/M phase after 24 h and in G0/G1 after 48 h incubation with FB1), possibly associated with
genotoxic event. On the other hand, Galvano et al. [131,132] reported that FB1 does not increase ROS
production or cell death in rat astrocytes although DNA-damage and caspase-3 activation take place.
Based on these results, the authors suggested that effects of FB1 are not a result of oxidative injury, but
are instead a response that may occur after modulation of protective genes. These theories support the
above-mentioned observations in FB1-treated SH-SY5Y cells. In SH-SY5Y cells, lipid peroxidation
took place without an increase in ROS production, and was associated with delayed cell death.
Moreover, low expression of the anti-apoptotic Bcl-2 protein in GT1-7 and C6 cells can be linked
to low basal GSH levels in these cell lines , and may increase their susceptibility to radical
attack [133–135]. SH-SY5Y cells, on the other hand, express higher levels of Bcl-2 [133,136], which
may explain why GSH levels in these cells decreased later than in GT1-7 and C6 cells exposed to FB1.
Stockmann-Juvala et al.  have also investigated the effects of FB1 on human U-118MG
glioblastoma cells. In their study, FB1 increased lipid peroxidation and the production of ROS in
U-118 MG cells dose- and time-dependently and these effects were accompanied by decreases in the
GSH levels and cell viability. In addition, signs of apoptosis were indicated by increased caspase-3-
like protease activity and internucleosomal DNA fragmentation. Based on these results, they
concluded that oxidative stress and apoptosis may be involved in the neurotoxicity induced by FB1.
There are a number of studies showing FB1-induced apoptosis in different cell types [129–131,138–140].
Int. J. Mol. Sci. 2011, 12 5221
Apoptosis is considered to be a common result of oxidative stress caused by ROS production,
disturbance of GSH generation and lipid peroxidation [141,142]. In addition, activation of caspase-3
may be one of the events causing an increase in ROS production, and subsequent lipid peroxidation
and reduction of intracellular GSH levels.
5. Ochratoxin A
Ochratoxin A (OTA) is a fungal metabolite produced by Aspergillus ochraceus and Penicillium
verrucosum. OTA is found in a variety of plant food products such as cereals. Because of its long half-
life, it accumulates in the food chain [143,144] and is frequently detected in human plasma at
nanomolar concentrations [145,146]. OTA has been found to be involved in the developments of
certain kidney diseases  and enzymuria  similar to Balkan endemic nephropathy found in
humans [147,149]. In addition to nephrotoxicity, the main OTA-induced toxicity, OTA has also been
reported to have immunotoxic [150,151], teratogenic [152,153], genotoxic  and neurotoxic 
effects. Although there is still insufficient evidence in humans, there is sufficient evidence in
experimental animals for the carcinogenicity of OTA .
OTA has complex mechanisms of action that include evocation of oxidative stress, bio-energetic
compromise, mitochondrial impairment, inhibition of protein synthesis, production of DNA single-
strand breaks and formation of OTA-DNA adducts [155,157–160]. The bio-energetic compromise
induced by OTA may be responsible for the generation of free radicals and ROS that results in global
oxidative damage to DNA and lipids and damage to proteins through generation of oxygen free
radicals and nitric oxide [161,162]. OTA renal toxicity and carcinogenicity may be, at least partly,
mediated by an Nrf2-dependent signal transduction pathway . Effect of OTA on potential
signaling molecules (growth factors, fatty acids, and/or Ca2+) could disrupt PKC-regulated pathways
downstream, and down-regulation of genes under transcriptional control of Nrf2 may lead to a reduced
oxidative stress response. In addition, OTA-induced down-regulation of genes under hepatocyte nuclear
factor 4α(HNF4α)-control may affect key metabolic processes. This could make kidney cells more
vulnerable to OTA-induced toxicity leading to tumor development . OTA has also been reported to
inhibit succinate-dependent electron transfer in the electron transport chain, but at higher concentrations
will also inhibit electron transport at Complex I [164,165], suggesting mitochondrial toxicity.
The developing brain appears to be very susceptible to the deleterious effects of OTA [166–169].
OTA has been shown to affect proliferation and migration of neurons  and reduce DNA
content  in the developing rodent brain. OTA is also reported to be neurotoxic to adult male
rats . Neurotoxicity is more pronounced in the ventral mesencephalon, hippocampus (HP), and
striatum than in the cerebellum (CB) . Recent animal and cellular studies have suggested that
OTA may contribute to the development of human and animal systemic problems, including
neurodegenerative diseases and brain dysfunction .
Sava et al.  investigated the time course of acute effects of OTA in the context of DNA
damage, DNA repair and global oxidative stress across six brain regions, CB, cortex (CX), HP,
midbrain (MB), caudate/putamn (CP) and pons/medulla (PM), in male mice, and they showed that
OTA causes acute depletion of striatal dopamine (DA) and its metabolites as well as decreased
tyrosine hydroxylase immunoreactivity in the corpus striatum on a background of globally increased
Int. J. Mol. Sci. 2011, 12 5222
oxidative stress evidenced by significant increases in lipid peroxidation and oxidative DNA damage,
and transient inhibition of oxidative DNA repair activity (oxyguanosine glyosylase, OGG1) across six
brain regions, accompanying evidence for apoptosis in the substantia nigra (SN), striatum and HP or
other regions. Unlike the monophasic SOD activation, the oxidative DNA repair response exhibited a
Sava et al.  also examined the possibility that OTA can cause parkinsonism in male mice
focusing on the effects of chronic low doses of OTA exposure on regional brain oxidative stress and
striatal DA metabolism. They reported that continuous administration of low doses of OTA with
implanted subcutaneous Alzet minipumps caused a small but significant decrease in striatal DA levels
and an up-regulation of anti-oxidant systems and DNA repair. These data suggest the possibility that
low dose exposure to OTA will result in an earlier onset of parkinsonism when normal age-dependent
decline in striatal DA levels is superimposed on the mycotoxin-induced lesion. In their study, since the
CP and MB showed a relatively diminished OGG1 activity and increased oxidative DNA damage, it
was postulated that the DA terminals of the striatum would suffer damage. This concept is supported
by an earlier report of increased oxidative DNA damage in SN and striatum in post-mortem brain from
Parkinson’s disease cases  and by the above-mentioned report that acute doses of OTA caused a
dose-dependent decrease of striatal DA and a decrease in DA turnover . The regional
vulnerability to the toxin was not directly related to the concentration of the toxin in each region.
Moreover, as mentioned above, not all regions were equally sensitive to the toxin, even though all
brain regions were capable of marked increases in OGG1 activity. Namely, the CP was most sensitive
to the toxin while the CB was the least sensitive. In addition, the HP, a primary site of neurodegeneration
in Alzheimer’s disease, turns out to exhibit relatively high OTA levels with concurrently pronounced
OTA neurotxicity . OTA may also be toxic through other mechanisms. For example, due to its
chemical structure, OTA inhibits protein synthesis by competition with phenylalanine in the
aminoacylation reaction of phenylalanine-tRNA [174,175] and phenylalanine hydroxylase
activity , leading to the impairment of the synthesis of DOPA, dopamine and catecholamines or
enzymes involved in the metabolism of DNA.
The adult brain retains a reservoir of stem-progenitor cells in the hippocampal “neurogenic zone”
capable of proliferative activity throughout life [176,177], and it is known that injury, irradiation,
drugs and endogenous factors such as hormones and trophic factors impact neurogenesis [178–181].
Therefore, OTA may also impact neurogenesis in adult HP. In addition, subchronic administration of
OTA is demonstrated to affect cognitive functions by reducing hippocampal N-methyl-D-aspartate
(NMDA) receptor subunits 2A and 2B concentrations in rats .
Sava et al.  tested neural stem/progenitor cells (NSCs) prepared from HP of adult mouse
brain for their vulnerability to OTA in vitro. OTA, added to the cultures in concentrations of
0.01–100 mg/mL, caused a dose- and time-dependent (6–72 h) decrease in viability of both
proliferating and differentiating NSC. Along with decreased viability, OTA elicited a pronounced
oxidative stress evidenced by a robust increase in total and mitochondrial SOD activity, and OTA also
significantly increased OGG1 activity. Proliferating NSC exhibited a greater vulnerability to the toxin
than differentiated neurons despite robust DNA repair and antioxidant responses. Such a result is
unexpected since DNA repair systems are typically more active and efficient in proliferating cells than
in post-mitotic differentiated cells [184,185]. It suggests that OTA’s mechanism of action as an
Int. J. Mol. Sci. 2011, 12 5223
inhibitor of mitochondrial oxidative metabolism may be less critical than its interference with DNA
synthesis and mitotic competence. Overall, Sava et al.  speculated that OTA exposure may
contribute to impaired hippocampal neurogenesis in vivo, resulting in depression and cognitive
deficits, conditions reported to be linked to mycotoxin exposure in humans [186–188].
Yoon et al.  investigated the potential harm caused by environmental exposure to OTA in
terms of its effects on neuronal cell viability and proteome profiles using mouse hippocampal HT22
and human neuroblastoma SH-SY5Y cells. Generation of ROS was detected in OTA-treated SH-SY5Y
and HT22 cells, however, caspase activation and an increase in p53 phosphorylation were only
detected in HT22 cells, even though OTA treatment caused oxidative stress in both two cell lines. The
expressions of several proteins (valosin containing protein, propyl 4-hydroxylase, Atp5b protein,
nucleophosmin 1, eukaryotic translation elongation factor 1 delta isoform, ornithine aminotransferase,
prohibitin, and peroxiredoxin 6), which have been suggested to be implicated in the pathogenesis of
neurodegenerative disorders, were up-regulated only in HT22 cells after treatment with OTA, which
was interesting because OTA induced the apoptosis of HT22 cells but not of SH-SY5Y cells.
Involvement of the mitochondrial dysfunction-related apoptotic process in OTA toxicity in HT22 cells
was corroborated by the results showing significant decline in mitochondrial activity in OTA-treated
HT22 cells, not in SH-SY5Y cells. Such differences between cell lines might be due in part to the
complex natures of protein expression and functional regulation required during the intracellular
signaling of apoptosis , and the above-mentioned alteration of protein expression profile in HT22
cells after OTA treatment is considered to be related to ROS generation. Because inhibition of
expression of propyl 4-hydoxylase is known to attenuate neuronal death associated with oxidative
stress , ornithine has been shown to be up-regulated during ROS-related apoptotic cell death ,
and valosin containing protein has been proposed to contribute to the conversion of oxidative stress
to an endoplasmic reticulum stress response during the pathological processes of a number of
neurodegenerative disorders . Contrary to the above-mentioned report by Sava et al. ,
Zhang et al.  reported that caspase-9 and caspase-3 were activated in response to OTA treatment
and caspase inhibitors were effective in partly counteracting OTA-induced apoptosis-related
neurocytotoxicity not only in primary rat cortical neuronal cells but also in human neuroblastoma SH-
SY5Y cells and that such OTA-induced apoptosis was accompanied by a loss of mitochondria
membrane potential. The authors suggest that OTA may contribute to the pathogenesis of
neurodegenerative diseases (e.g., Alzheimer’s and Parkinson’s disease) in which apoptotic processes
are centrally involved. The reason for the difference in apoptosis-inducing ability of OTA in human
neuroblastoma SH-SY5Y cells between the two reports still remains unclear.
Besides the toxic effects of OTA on neuronal cells, Zurich et al.  investigated the relationship
between OTA toxicity and glial reactivity in serum-free aggregating rat brain cell cultures, and they
showed that OTA affects the cytoskeletal integrity of astrocytes as well as the expression of genes
pertaining to the brain inflammatory response system, and suggested that a relationship exists between
the inflammatory events and the cytoskeletal changes induced by OTA. Furthermore, they also
suggested that, by inducing an atypical glial reactivity, OTA may severely affect the neuroprotective
capacity of glial cells. Moreover, Hong et al.  reported that OTA caused concentration-dependent
reductions in neurite outgrowth and cell number, and induced the activation of transcription factors
activator protein-1 (AP-1) and NF-κB activation in cultured rat embryonic midbrain cells, and that
Int. J. Mol. Sci. 2011, 12 5224
15-deoxy-delta 12, 14-prostaglandin J2 (15-deoxy PGJ2), a peroxisome proliferator-activated receptor
gamma (PPAR-γ) agonist, blocked OTA-induced neurotoxicity by inhibiting AP and NF-κB activation
in cultured rat embryonic midbrain cells.
This paper has reviewed the mechanisms of neurotoxicity induced in rodents and neuronal cell lines
by T-2 toxin, macrocyclic trichothecenes, FB1 and OTA.
T-2 toxin, one of the Type A trichothecene mycotoxins, triggers a ribotoxic response through its
high binding affinity to peptidyl transferase which is an integral part of the 60 s ribosomal subunit,
resulting in activation of JNK/p38 MAPKs. T-2 toxin also interferes with the metabolism of membrane
phospholipids and increases liver lipid peroxides. In the fetal brain, oxidative stress is considered to be
the main factor behind the T-2 toxin-induced changes, and the MAPK-JNK-c-jun pathway is thought
to be involved in T-2 toxin-induced neuronal cell apoptosis. Moreover, activation of caspase-2 is
essential to T-2 toxin-induced apoptosis in the fetal brain. T-2 toxin also induces oxidative damage in
the adult mouse brain as well as in the fetal rat brain.
Macrocyclic trichothecenes have been postulated to etiologically contribute to DBRI such as
debilitating respiratory and nonrespiratory symptoms. The common ability of macrocyclic trichothecenes
to cause protein synthesis inhibition via binding to the 18s rRNA of the ribosomal large subunit has
been speculated to be a major mechanism underlying induction of cell apoptosis by this group of
trichothecenes. In mice, SG or RA exposure specifically induces apoptosis of OSNs and subsequent
atrophy of OE. SG or RA also induces apoptosis in, and atrophy of, the olfactory nerve layer of OBs of
the brain. Moreover, in the ethmoid turbinates and OBs in the frontal brain in mice treated with SG or
RA, elevated mRNA expression for the inflammatory cytokines, chemokine, and proapoptotic genes
and increased mRNA concentrations for PKR are detected. In PC12 neural cells, SG-induced apoptosis
is mediated by PKR via a caspase-independent pathway possibly involving AIF translocation from
mitochondria to the nucleus. On the other hand, SH is thought to induce caspase-3 activation and
apoptosis of PC12 cells through the activation of MAPK and JNK in a GSH-sensitive manner.
FB1 is well known to cause ELEM and may be implicated in the etiology of NTD in children.
FB1-induced inhibition of ceramide synthesis can result in a wide spectrum of changes in lipid
metabolism and associated lipid-dependent pathways. FB1 may disrupt central nervous system
homeostasis when brain tissue is directly exposed to this mycotoxin. Namely, the icv infusion of FB1
leads to neuronal degeneration in the cortex, concurrent with disruption of sphingolipid metabolism,
i.e., inhibition of de novo ceramide synthesis, stimulation of astrocytes, and activation of
proinflammatory cytokine signaling. In in vitro studies, FB1 inhibits protein synthesis, causes DNA
fragmentation and cell death, increases 8-OH-dG, and induces lipid peroxidation in rat C6 glioma
cells, and oxidative stress and apoptosis may be involved in the neurotoxicity induced in human
U-118MG glioblastoma cells by FB1. On the other hand, effects of FB1 are not a result of oxidative
injury but are instead a response that may occur after modulation of protective genes in rat astrocyte
and SH-SY5Y human neuroblastoma cell cultures.
OTA has complex mechanisms of action that include evocation of oxidative stress, bio-energetic
compromise, mitochondrial impairment, inhibition of protein synthesis, production of DNA
Int. J. Mol. Sci. 2011, 12 5225
single-strand breaks and formation of OTA-DNA adducts. OTA causes acute depletion of striatal DA
and its metabolites in the corpus striatum on a background of globally increased oxidative stress across
six brain regions of rats examined, accompanying evidence for apoptosis in the SN, striatum and HP or
other regions. The CP is most sensitive to the toxin while the CB is the least sensitive, and the HP,
primary site of neurodegeneration in Alzheimer’s disease, turns out to exhibit relatively high OTA
levels with concurrently pronounced OTA neurotoxicity. OTA exposure may contribute to impaired
hippocampal neurogenesis in vivo, resulting in depression and cognitive deficits. OTA also induces
oxidative stress and apoptosis through caspase activation and increase in p53 phosphorylation in
various neural cell cultures. OTA seems to contribute to the pathogenesis of neurodegenerative
diseases in humans (e.g., Alzheimer’s and Parkinson’s disease), in which apoptotic processes are
The authors would like to thank Pete Aughton, D.A.B.T., ITR Laboratories Canada Inc.,
1. Haschek, W.M.; Voss, K.A.; Beasley, V.R. Selected Mycotoxins Affecting Animal and Human
Health. In Handbook of Toxicologic Pathology; Haschek, W.M., Rousseaux, C.G.,
Walling, M.A., Eds.; Academic Press: San Diego, CA, USA, 2002; pp. 645–699.
2. Surai, P.F.; Mezes, M.; Melnichuk, S.D.; Fotina, T.I. Mycotoxins and animal health: From
oxidative stress to gene expression. Krmiva 2008, 50, 35–43.
3. Chandra, J.; Samali, A.; Orrenius, S. Triggering and modulation of apoptosis by oxidative stress.
Free Radic. Biol. Med. 2000, 29, 323–333.
4. Desjardins, A.E.; Hohn, T.M.; McComic, S.P. Trichothecene biosynthesis in Fusarium species:
chemistry, genetics, and significance. Microbiol. Mol. Biol. Rev. 1993, 57, 595–604.
5. Nelson, P.E.; Dignani, M.C.; Anaissie, E.J. Taxonomy, biology, and clinical aspects of Fusarium
species. Clin. Microbiol. Rev.1994, 7, 479–504.
6. Joffe, A.Z. Foodborne Diseases: Alimentary Toxic Aleukia. In Handbook of Foodborne Diseases
of Biological Origin; Rochcigle, M., Ed.; CRC Press: Boca Raton, FL, USA, 1983; pp. 353–495.
7. Saito, M.; Ohtsubo, K. Trichothecene Toxins of Fusarium Species. In Mycotoxins;
Purchase, I.F.H., Ed.; Elsevier Scientific Publication: New York, NY, USA, 1977; pp. 264–280.
8. Ueno, Y.; Ishii, K.; Saki, K.; Kanadera, K.; Tsunoda, S.; Tanoka, H.; Enomoto, M. Toxicological
approaches to the metabolites of fusaria. IV. Microbial survey on “bean-hulls poisoning of
horses” with the isolation of toxic trichothecenes, neosonaniol and T-2 toxin of Fusarium solani
M-1-1. Jpn. J. Exp. Med.1972, 42, 187–203.
9. Bennet, J.W.; Klich, M. Mycotoxins. Clin. Microbiol. Rev. 2003, 16, 497–516.
10. Eriksen, G.S.; Petterson, H. Toxicological evaluation of trichothecenes in animal feed. Anim.
Feed Sci. Technol. 2004, 114, 205–239.
Int. J. Mol. Sci. 2011, 12 5226
11. Shifrin, V.I.; Anderson, P. Trichothecene mycotoxins trigger a ribotoxic stress response that
activates c-jun N-terminal kinase and p38 mitogen-activated protein kinase and induces
apoptosis. J. Biol. Chem.1999, 274, 13985–13992.
12. Chang, I.M.; Mar, W.C. Effect of T-2 toxin on lipid peroxidation in rats: Elevation of conjugated
diene formation. Toxicol. Lett. 1988, 40, 275–280.
13. Eriksen, G.S.; Petterson, H.; Lund, H. Comparative cytotoxicity of deoxynivalenol, nivalenol,
triacetylated derivatives and de-epoxy metabolites. Food Chem. Toxicol. 2004, 42, 619–624.
14. IARC. Toxins derived from Fusarium sporotrichoides: T-2 toxin. In IARC Monographson the
Evaluation of Carcinogenic Risks to Humans; IARC: Lyon, France, 1993; pp. 467–488.
15. Sharma, R.P. Immunotoxicity of mycotoxins. J. Dairy Sci. 1993, 76, 892–897.
16. Stanford, G.K.; Hood, R.D.; Haynes, A.W. Effects of prenatal administration of T-2 toxin to
mice. Res. Commu. Chem. Pathol. Pharmacol.1975, 10, 743–746.
17. Williams, P.P. Effects of T-2 mycotoxin on gastrointestinal tissues: A review of in vivo and
in vitro models. Arch. Environ. Contam. Toxicol.1989, 18, 374–387.
18. Shinozuka, J.; Li, G.; Kiatipattanasakul, W.; Uetsuka, K.; Nakayama, H.; Doi, K. T-2
toxin-induced apoptosis in lymphoid organs of mice. Exp. Toxicol. Pathol. 1997, 49, 387–392.
19. Li, G.; Shinozuka, J.; Uetsuka, K.; Nakayama, H.; Doi, K. T-2 toxin-induced apoptosis in
Peyer’s patches of mice. J. Toxicol. Pathol. 1997, 10, 59–61.
20. Shinozuka, J.; Suzuki, M.; Noguchi, N.; Sugimoto, T.; Uetsuka, K.; Nakayama, H.; Doi, K. T-2
toxin-induced apoptosis in hematopoietic tissues of mice. Toxicol. Pathol. 1998, 26, 674–681.
21. Li, G.; Shinozuka, J.; Uetsuka, K.; Nakayama, H.; Doi, K. T-2 toxin-induced apoptosis in
intestinal crypt epithelial cells of mice. Exp. Toxicol. Pathol. 1997, 49, 447–450.
22. Albarenque, S.M.; Shinozuka, J.; Iwamoto, S.; Nakayama, H.; Doi, K. T-2 toxin-induced acute
skin lesions in Wistar-derived hypotrichotic WBN/ILA-Ht rats. Histol. Histopathol. 1999, 14,
23. Shinozuka, J.; Miwa, S.; Fujimura, H.; Toriumi, W.; Doi, K. Hepatotoxicity of T-2 Toxin,
Trichothecene Mycotoxin. In New Strategies for Mycotoxin Research in Asia (Proceedings of
ISMYCO Bangkok ‘06); Kumagai, S., Ed.; Japanese Association of Mycotoxicology: Tokyo,
Japan, 2007; pp. 62–66.
24. Sehata, S.; Kiyosawa, N.; Atsumi, F.; Ito, K.; Yamoto, T.; Teranishi, M.; Uetsuka, K.;
Nakayama, H.; Doi, K. Microarray analysis of T-2 toxin-induced liver, placenta and fetal liver
lesions in pregnant rats. Exp. Toxicol. Pathol. 2005, 57, 15–28.
25. Doi, K.; Shinozuka, J.; Sehata, S. T-2 toxin and apoptosis. J. Toxicol. Pathol. 2006, 19, 15–27.
26. Doi, K.; Ishigami, N.; Sehata, S. T-2 toxin-induced toxicity in pregnant mice and rats. Int. J. Mol.
Sci. 2008, 9, 2146–2158.
27. Chaudhary, M.; Rao, P.V. Brain oxidative stress after dermal and subcutaneous exposure of T-2
toxin in mice. Food Chem. Toxicol. 2010, 48, 3436–3442.
28. Boyd, K.E.; Fitzpatrick, D.W.; Wilson, J.R.; Wilson, L.M. Effect of T-2 toxin on brain biogenic
monoamines in rats and chickens. Can. J. Vet. Res. 1988, 52, 181–185.
29. Martin, L.J.; Morse, J.D.; Anthony, A. Quantitative cytophotometric analysis of brain neuronal
RNA and protein changes in acute T-2 mycotoxin poisoned rats. Toxicon 1986, 24, 933–941.
Int. J. Mol. Sci. 2011, 12 5227
30. Wang, J.; Fitzpatrick, D.W.; Wilson, J.R. Effects of the trichothecene mycotoxin T-2 toxin on
neurotransmitters and metabolites in discrete areas of the rat brain. Food Chem. Toxicol. 1998,
31. Wang, J.; Fitzpatrick, D.W.; Wilson, J.R. Effect of T-2 toxin on blood-brain barrier permeability
monoamine oxidase activity and protein synthesis in rats. Food Chem. Toxicol. 1998, 36, 955–961.
32. Ishigami, N.; Shinozuka, J.; Katayama, K.; Uetsuka, K.; Nakayama, H.; Doi, K. Apoptosis in the
developing mouse embryos from T-2 toxin-inoculated dams. Histol. Histopathol. 1999, 14, 729–733.
33. Ishigami, N.; Shinozuka, J.; Katayama, K.; Uetsuka, K.; Nakayama, H.; Doi, K. Apoptosis in
mouse fetuses from dams exposed to T-2 toxin at different days of gestation. Exp. Toxicol.
Pathol. 2001, 52, 493–501.
34. Rousseaux, C.G.; Schiefer, H.B. Maternal toxicity, embryolethality and abnormal fetal
development in CD-1 mice following one oral dose of T-2 toxin. J. Appl. Toxicol. 1987, 7,
35. Stanford, G.K.; Hood, R.D.; Hayes, A.W. Effect of prenatal administration of T-2 toxin to mice.
Res. Commun. Chem. Path. Pharmacol. 1975, 10, 743–746.
36. Sehata, S.; Kiyosawa, N.; Makino, T.; Atsumi, F.; Ito, K.; Yamoto, T.; Teranishi, M., Baba, Y.;
Uetauka, K.; Nakayama, H.; Doi, K. Morphological and microarray analysis of T-2
toxin-induced rat fetal brain lesion. Food Chem. Toxicol. 2004, 42, 1727–1736.
37. Galtier, P.; Paulin, F.; Eeckhoutte, C.; Larrieu, G. Comparative effects of T-2 toxin and
diacetoxyscirpenol on drug metabolizing enzymes in rat tissues. Food Chem. Toxicol. 1989, 27,
38. Guerre, P.; Eeckhoutte, C.; Burgat, V.; Galtier, P. The effects of T-2 toxin exposure on liver drug
metabolizing enzymes in rabbit. Food Add. Contam. 2000, 17, 1019–1026.
39. Jarpe, M.B.; Widmann, C.; Knall, C.; Schlesinger, T.K.; Gibson, S.; Yujiri, T.; Fanger, G.R.;
Gelfand, E.W.; Johnson, G.L. Anti-apoptotic versus pro-apoptotic signal transduction:
checkpoints and stop signs along the roard to death. Oncogene1998, 17, 1475–1582.
40. Bold, S.; Weidle, U.H.; Kolch, W. The kinase domain of MEKK1 induces apoptosis by
dysregulation of MAP kinase pathways. Exp. Cell Res. 2003, 283, 80–90.
41. Ham, J.; Eilers, A.; Whitfield, J.; Neame, S.J.; Shah, B. c-JUN and the transcriptional control of
neuronal apoptosis. Biochem. Pharmacol. 2000, 60, 1015–1021.
42. Annunziato, L.; Amoroso, S.; Pannaccione, A.; Cataldi, M.; Pignataro, G.; D’Alessio, S.;
Sirabella, R.; Second, A.; Sibaud, L.; DiRenzo, G.F. Apoptosis induced in neuronal cells by
oxidative stress: role played by caspases and intracellular calcium ions. Toxicol. Lett. 2003, 139,
43. Troy, C.M.; Shelanski, M.L. Caspase-2 redux. Cell Death Differ. 2003, 10, 101–107.
44. Huang, P.; Akagawa, K.; Yokoyama, Y.; Nohara, K.; Kano, K.; Morimoto, K. T-2 toxin initially
activates caspase-2 and induces apoptosis in U937 cells. Toxicol. Lett. 2007, 170, 1–10.
45. Kanemitsu, H.; Yamauchi, H.; Komatsu, M.; Yamamoto, S.; Okazaki, S.; Uchida, K.; Nakayama, H.
6-Mercaptopurine (6-MP) induces cell cycle arrest and apoptosis of neural progenitor cells in the
developing rat brain. Neurotox. Teratol. 2009, 31, 104–109.
Int. J. Mol. Sci. 2011, 12 5228
46. Katayama, K.; Ueno, M.; Yamauchi, H.; Nakayama, H.; Doi, K. Microarray analysis of genes in
fetal central nervous system after ethylnitrosourea administration. Birth Defects Res. Part B
2005, 74, 255–260.
47. Nam, C.; Yamauchi, H.; Nakayama, H.; Doi, K. Etoposide induces apoptosis and cell cycle arrest
of neuroepithelial cells in a p53-related manner. Neurotox. Teratol. 2009, 28, 664–672.
48. Ueno, M.; Katayama, K.; Yamauchi, H.; Nakayama, H.; Doi, K. Cell cycle and cell death
regulation of neural progenitor cells in the 5-azacytidine (5AzC)-treated developing fetal brain.
Exp. Neurol .2006, 198, 154–166.
49. Woo, G.H.; Bak, E.J.; Nakayama, H.; Doi, K. Molecular mechanisms of hydroxyurea
(HU)-induced apoptosis in the mouse fetal brain. Neurotox. Teratol. 2006, 28, 125–134.
50. Ogunshola, O.O.; Antic, A.; Donoghue, M.J.; Fan, S.-Y.; Kim, H.; Stewart, W.B.; Madri, J.A.;
Ment, L.R. Paracrine and autocrine function of neuronal vascular endothelial growth factor
(VEGF) in the central nervous system. J. Biol. Chem. 2002, 277, 11410–11415.
51. Halliwell, B.; Gutteridge, J.M.C. Free Radicals in Biology and Medicine, 3rd ed.; Oxford
University Press: New York, NY, USA, 1999.
52. Lee, J.-M.; Jiang, L.; Johnson, D.A.; Stein, T.D.; Kraft, A.D.; Calkins, M.J.; Jakel, R.J.;
Jofnson, J.A. Nrf2, a multiorgan protector? FASEB J. 2005, 19, 1061–1066.
53. Boesch-Saadatmandi, C.; Wagner, A.E.; Graeser, A.C.; Hundhausen, C.; Wollram, S.;
Rimbach, G. Ochratoxin A impairs Nrf2-dependent gene expression in porcine kidney tubulus
cells. J. Anim. Phys. Anim. Nutr. 2009, 93, 547–555.
54. Boutin-Forzano, S.; Charpin-Kadouch, C.; Chabbi, S.; Bennedjai, N.; Dumon, H.; Charpin, D.
Wall relative humidity: A simple and reliable index for predicting Stachybotrys chartarum
infestation in dwellings. Indoor Air 2004, 14, 196–199.
55. Tsumori, T.; Reijula, K.; Johnsson, T.; Hemminki, K.; Hintikka, E.L.; Lindroos, O.; Kalso, S.;
Koukila-Kahkola, P.; Mussalo-Rauhamaa, H.; Haahtela, T. Mycotoxins in crude building
materials from water-damaged buildings. Appl. Environ. Microbiol. 2000, 66, 1899–1904.
56. Pestka, J.J.; Yike, I.; Dearborn, D.G.; Ward, M.D.W.; Harkema, J.R. Stachybotrys chartarum,
trichothecene mycotoxins, and damp building-related illness: New insights into a public health
enigma. Toxicol. Sci. 2008, 104, 4–26.
57. Shelton, B.G.; Kirkland, K.H.; Flanders, W.D.; Morris, G.K. Profiles of airborne fungi in
buildings and outdoor environments in the United States. Appl. Environ. Microbiol. 2002, 68,
58. Hodgson, M.J.; Morey, P.; Leung, W.Y.; Morrow, L.; Miller, D.; Jarvis, B.B.; Robbins, H.;
Halsey, J.F.; Storey, E. Building-associated pulmonary disease from exposure to Stachybotrys
chartarum and Aspergillus versicolor. J. Occup. Environ. Med. 1998, 40, 241–249.
59. Johanning, E.; Biagini, R.; Hull, D.; Morey, P.; Jarvis, B.; Landsbergis, P. Health and
immunology study following exposure to toxigenic fungi (Stachybotrys chartarum) in a
water-damaged office environment. Int. Arch. Occup. Environ. Health 1996, 68, 207–218.
60. Gordon, W.A.; Cantor, J.B.; Johanning, E.; Charatz, H.J.; Ashman, T.A.; Breeze, J.L.;
Haddad, L.; Abramowitz, S. Cognitive impairment associated with toxigenic fungal exposure:
A replication and extension of previous findings. Appl. Neuropsychol. 2004, 11, 65–74.
Int. J. Mol. Sci. 2011, 12 5229
61. Hossain, M.A.; Ahmed, M.S.; Ghannoum, M.A. Attributes of Stachybotrys chartarum and its
association with human disease. J. Allergy Clin. Immunol. 2004, 113, 200–208.
62. Kirburn, K.H. Role of molds and myxotoxins in being sick in buildings: Neurobehavioral and
pulmonary impairment. Adv. Appl. Microbiol. 2004, 55, 339–359.
63. Andersen, B.; Nielsen, K.F.; Jarvis, B.B. Characterization of Stachybotrys from water-damaged
buildings based on morphology, growth, and metabolite production. Mycologia 2002, 94, 392–403.
64. Gregory, L.; Pestka, J.J.; Dearborn, D.G.; Rand, T.G. Localization of satratoxin-G in
Stachybotrys chartarum spores and spore-impacted mouse lung using immunocytochemistry.
Toxicol. Pathol. 2004, 32, 26–34.
65. Yike, I.; Distler, A.M.; Ziady, A.G.; Dearborn, D.G. Mycotoxin adducts on human serum
albumin: Biomerkers of exposure to Stachybotrys chartarum. Environ. Health Perspect. 2006,
66. Chung, Y.J.; Zhou, H.R.; Pestka, J.J. Transcriptional and posttranscriptional roles for p38
mitogen-activated protein kinase in upregulation of TNF-α expression by deoxynivalenol
(vomitoxin). Toxicol. Appl. Pharmacol. 2003, 193, 188–201.
67. Moon, Y.; Pestka, J.J. Deoxynivalenol-induced mitogen-activated protein kinase phosphorylation
and IL-6 expression in mice suppressed by fish oil. J. Nutr. Biochem. 2003, 14, 717–726.
68. Zhou, H.R.; Lau, A.S.; Pestka, J.J. Role of double-stranded RNA-activated protein kinase R
(PKR) in deoxynivalenol-induced ribotoxic stress response. Toxicol. Sci. 2003, 74, 335–344.
69. Iordanov, M.S.; Pribnow, D.; Magun, J.L.; Dinh, T.H.; Pearson, J.A.; Chen, S.L.; Magun, B.E.
Ribotoxic stress response: Activation of the stress-activated protein kinase JNK1 by inhibitors of
the peptidyl transferase reaction and by sequence-specific RNA damage to the alphasarcin/ricin
loop in the 28S rRNA. Mol. Cell Biol. 1997, 17, 3373–3381.
70. Chung, Y.J.; Jarvis, B.; Pestka, J.J. Modulation of lipopolysaccharide-induced proinflammatory
cytokine production by satratoxins and other macrocyclic trichothecenes in the murine
macrophage. J. Toxicol. Environ. Health A 2003, 66, 379–391.
71. Chung, Y.J.; Yang, G.H.; Islam, Z.; Pestka, J.J. Up-regulation of macrophage inflammatory
protein-2 and complement 3A receptor by the trichothecenes deoxynivalenol and satratoxin G.
Toxicology 2003, 186, 51–65.
72. Hughes, B.J.; Hsieh, G.C.; Jarvis, B.B.; Sharma, R.P. Effects of macrocyclic trichothecene
mycotoxins on the murine immune system. Arh. Environ. Contam. Toxicol. 1989, 18, 388–395.
73. Hughes, B.J.; Jarvis, B.B.; Sharma, R.P. Effects of macrocyclictrichothecene congeners on the
viability and mitogenesis of mirine splenic lymphocytes. Toxicol. Lett.1990, 50, 57–67.
74. Pestka, J.J.; Forsell, J.H. Inhibition of human lymphocyte transformation by the macrocyclic
trichothecene roridin A and verrucarin A. Toxicol. Lett.1988, 41, 215–222.
75. Yang, G.-H.; Jarvis, B.B.; Chung, Y.-J.; Pestka, J.J. Apoptosis induction by the satratoxins and
other trichothecene mycotoxins: relationship to ERK, p38 MAPK, and SAPK/JNK activation.
Toxicol. Appl. Pharmacol. 2000, 164, 149–160.
76. Cundliffe, E.; Davies, J.E. Inhibition of initiation, elongation, and termination of eukaryotic protein
synthesis by trichothecene fungal toxins. Antimicrob. Agents Chemother. 1977, 11, 491–499.
Int. J. Mol. Sci. 2011, 12 5230
77. Nielsen, K.F.; Huttunen, K.; Hyvarinen, A.; Andersen, B.; Jarvis, B.B.; Hirvonen, M.R.
Metabolite profiles of Stachybotrys isolates from water-damaged buildings and their induction of
inflammatory mediators and cytotoxicity in macrophages. Mycopathologia 2002, 154, 201–205.
78. Islam, Z.; Shinozuka, J.; Harkema, J.R.; Pestka, J.J. Purification and comparative neurotoxicity
of the trichothecenes satratoxin G and roridin L2 from Stachybotrys chartarum. J. Toxicol.
Environ. Health A 2009, 72, 1242–1251.
79. Nusuetrong, P.; Pengsuparp, T.; Meksuriyen, D.; Tanitsu, M.; Kikuchi, H.; Muzugaki, M.;
Shimazu, K; Oshima, Y.; Nakahata, N.; Yoshida, M. Satratoxin H generates reactive oxygen
species and lipid peroxides in PC12 cell. Biol. Pharm. Bull. 2008, 31, 1115–1120.
80. Rand, T.G.; Mahoney, M.; White, K.; Oulton, M. Microanatomical changes in alveolar type II
cells in juvenile mice intratracheally exposed to Stachybotrys chartarum spores and toxin.
Toxicol. Sci. 2002, 65, 239–245.
81. Wang, H.; Yadav, J.S. Global gene expression changes underlying Stachybotrys chartarum
toxin-induced apoptosis in murine alveolar macrophages: Evidence of multiple signal
transduction pathways. Apoptosis 2007, 12, 535–548.
82. Wang, H.; Yadav, J.S. DNA damage, redox changes, and associated stress-inducible signaling
events underlying the apoptosis and cytotocity in murine alveolar macrophage cell line MH-S by
methanol-extracted Stachybotrys chartarum toxins. Toxicol. Appl. Pharmacol. 2006, 214,
83. Islam, Z.; Harkema, J.R.; Pestka, J.J. Satratoxin G from the black mold Stachybotrys chartarum
evokes olfactory sensory neuron loss and inflammation in the murine nose and brain. Environ.
Health Perspect. 2006, 114, 1099–1107.
84. Islam, Z.; Amuzie, C.J.; Harkema, J.R.; Pestka, J.J. Neurotoxicity and inflammation in the nasal
airways of mice exposed to the macrocyclic trichothecene mycotoxin roridin A: Kinetic and
potentiation by bacterial lipipolysaccharide coexposure. Oxford J. Life Sci. Med. Toxicol. Sci.
2007, 98, 526–541.
85. Chang, R.C.; Suen, K.C.; Ma, C.H.; Elyaman, W.; Ng, H.K.; Hugon, J. Involvement of
double-stranded RNA-dependent protein kinase and phosphorylation of eukaryotic initiation
factor-2alpha in neuronal degeneration. J. Neurochem. 2002, 83, 1215–1225.
86. Ge, Y.; Tsukatani, T.; Nishimura, T.; Furukawa, M.; Miwa, T. Cell death of olfactory receptor
neurons in a rat with nasosinusitis infected artificially with Staphylococcus. Chem. Senses 2002,
87. Huang, C.C.; Chen, K.; Huang, T.Y. Immunohistochemical studies of sensory neurons in rat
olfactory epithelium. Eur. Arch. Otorhinolaryngol. 1995, 252, 86–91.
88. Wu, S.; Kumar, K.U.; Kaufmam, R.J. Identification and requirement of three ribosome binding
domains in dsRNA-dependent protein kinase (PKR). Biochemistry 1998, 37, 13816–13826.
89. Garcia, M.A.; Meurs, E.F.; Esteban, M. The dsRNA protein kinase PKR: Virus and cell control.
Biochemie 2007, 89, 799–811.
90. Cowan, C.M.; Roskams, A.J. Apoptosis in the mature and developing olfactory neuroepithelium.
Microsc. Res. Technol. 2002, 58, 204–215.
91. Farbman, A.I.; Buchholz, J.A.; Suzuki, Y.; Coines, A.; Speert, D. A molecular basis of cell death
in olfactory epithelium. J. Comp. Neurol. 1999, 414, 306–314.
Int. J. Mol. Sci. 2011, 12 5231
92. Suzuki, Y.; Farbman, A.I. Tumor necrosis factor-alpha-induced apoptosis in olfactory epithelium
in vitro: Possible roles of caspase 1 (ICE), caspase-2 (ICH-1), and caspase-3 (CPP32). Exp.
Neurol. 2000, 165, 35–45.
93. Islam, Z.; Hegg, C.C.; Bae, H.Y.; Pestka, J.J. Satratoxin G-induced apoptosis in PC-12 neuronal
cells is mediated by PKR and caspase independent. Toxicol. Sci. 2008, 105, 142–152.
94. Nusuetrong, P.; Yoshida, M.; Tanitsu, M.A.; Kikuchi, H.; Mizugaki, M.; Shimazu, K.;
Pengsuparp, T.; Meksuriyen, D.; Oshima, Y.; Nakahata, N. Involvementof reactive oxygen
species and stress activated MAPKs in satoratoxin H-induced apoptosis. Eur. J. Pharmacol. 2005,
95. Chandra, J.; Samali, A.; Orrenius, S. Triggering and modulation of apoptosis by oxidative
stress. Free Radic. Biol. Med. 2000, 29, 323–333.
96. Karunasena, E.; Larrañaga, M.D.; Simoni, J.S.; Douglas, D.R.; Straus, D.C. Building-associated
neurological damage modeled in human cells: A mechanism of neurotoxic effects by exposure to
mycotoxins in the indoor environment. Mycopathologia 2010, 170, 377–390.
97. Thrasher, J.D.; Crawley, S. The biocontaminants and complexity of damp indoor spaces; more
than what meets the eyes. Toxicol. Ind. Health 2009, 25, 583–615.
98. Campbell, I.L. Neuropathogenic acions of cytokines assessed in transgenic mice. Int. J.
Dev. Neurosci. 1995, 13, 275–284.
99. Peters, A.; Vweronesi, B.; Calderon-Garciduenas, J.; Gehr, P.; Chen, L.C.; Greiser, M.;
Reed, W.; Rothen-Rutishauser, B.; Schurch, S.; Schulz, H. Translocation and potential
neurological effects of fine and ultrafine particles a critical update. Part. Fibre Toxicol. 2006, 3,
100. Calderón-Garcidueñas, L.; Azzarelli, B.; Acuna, H.; Garcia, R.; Gambling, T.M.; Osnaya, N.;
Monroy, S.; Tizapantzi, M.D.R.; Carson, J.L.; Villarreal-Calderon, A.; et al. Air pollution and
brain damage. Toxicol. Pathol. 2002, 30, 373–389.
101. Calderón-Garcidueñas, L.; Maronpot, R.R.; Torres-Jardon, R.; Henríquez-Roldán, C.;
Schoonhoven, R.; Acuña-Ayala, H.; Villarreal-Carderón, A.; Nakamura, J.; Fernando, R.;
Reed, W.; et al. DNA damage in nasal and brain tissues of canines exposed to air pollutants is
associated with evidence of chronic brain inflammation and neurodegeneration. Toxico. Pathol.
2003, 31, 524–538.
102. Dutton, M.F. Fumonisins, mycotoxins of increasing importance: their nature and their effects.
Pharmacol. Ther. 1996, 70, 137–161.
103. Howard, P.C.; Eppley, R.M.; Stack, M.E.; Warbritton, A.; Voss, K.A.; Lorentzen, R.J.;
Kovach, R.M.; Bucci, T.J.; Fumonisin B1 carcinogenicity in a 2-year feeding study using F344
rats and B6C3 F1 mice. Environ. Health Perspect. 2001, 109, 277–282.
104. Wang, E.; Norred, W.P.; Bacon, C.W.; Riley, R.T.; Merrill, A.H., Jr. Inhibition of sphingolipid
biosynthesis by fumonisins. J. Biol. Chem. 1991, 22, 14486–14490.
105. Merrill, A.H., Jr.; Sullards, M.C.; Wang, E.; Voss, K.A.; Riley, R.T. Sphingolipid metabolism:
Role in signal transduction and disruption by fumonisins. Environ. Health Perspect. 2001, 109,
Int. J. Mol. Sci. 2011, 12 5232
106. Riley, R.T.; Enongene, E.; Voss, K.A.; Norred, W.P.; Meredith, F.I.; Sharma, R.P.; Spitsbergen, J.;
Williams, D.E.; Carlson, D.B.; Merrill, A.H., Jr. Sphingolipid perturbations as mechanisms for
fumonisin carcinogenesis. Environ. Health Perspect. 2001, 109, 301–308.
107. Ross, P.F.; Rice, L.G.; Reagor, J.C.; Osweiler, G.D.; Wilson, T.M.; Nelson, H.A. Owens, D.L.;
Plattner, R.D.; Harlin, K.A.; Richard, J.L.; et al. Fumonisin B1 concentrations in feeds from 45
confirmed equine leukoencephalomalacia cases. J. Vet. Diagn. Invest. 1991, 3, 238–241.
108. Wilson, T.M.; Ross, P.F.; Rice, L.G.; Osweiler, G.D.; Nelson, H.A.; Owen, D.L.; Plattner, R.D.;
Reggiardo, C.; Noon, T.H.; Pickrell, J.W. Fumonisin B1 levels associated with an epizootics of
equine leukoencephalomalacia. J. Vet. Diagn. Invest.1990, 2, 213–216.
109. Goel, S.; Schumacher, J.; Lenz, S.D.; Kemppanien, B.W. Effects of fusarium moniliforme
isolates on tissue and serum sphingolipid concentrations in horses. Vet. Hum. Toxicol. 1996, 38,
110. Marasas, W.F.; Riley, R.T.; Hendricks, K.A.; Stevens, V.L.; Sadler, T.W.; Gelineau-van Wanes, J.
Fumonisins disrupt sphingolipid metabolism, folate transport, and neural tube development in
embryo culture and in vivo: A potential risk factor for human neural tube defects among
populations consuming fumonisin-contaminated maize. J. Nutr. 2004, 134, 711–716.
111. Sadler, T.W.; Merrill, A.H.; Stevens, V.L.; Sullards, M.C.; Wang, E.; Wang, P. Prevention of
fumonisin B1-induced neural tube defects by folic acid. Teratology 2002, 66, 169–176.
112. Stevens, V.L.; Tang, J. Fumonisin B1-induced sphingolipid depletion inhibits vitamin uptake via
the glycosylphosphatidylinositol-anchored folate receptor. J. Biol. Chem. 1997, 272, 18020–18025.
113. Harel, R.; Futerman, A.H. Inhibition of sphingolipid synthesis affects axonal outgrowth in
cultured hippocampal neurons. J. Biol. Chem. 1993, 268, 14476–14481.
114. Kwon, O.S.; Slikker, W., Jr.; Davies, D.L. Biochemical and morphological effects of fumonisin
B1 on primary cultures of rat cerebrum. Neurotoxicol. Teratol. 2000, 22, 565–572.
115. Monnet-Tschudi, F.; Zurich, M.G.; Sorg, O.; Matthieu, J.M.; Honegger, P.; Schilter, B. The
naturally occurring food mycotoxin fumonisin B1 impairs myelin formation in aggregating brain
cell culture. Neurotoxicology 1999, 20, 41–48.
116. Kwon, O.S.; Schmued, L.C.; Slikker, W., Jr. Fumonisin B1 in developing rats alter brain
sphinganine levels and myelination. Neurotoxicolog 1997, 18, 571–580.
117. Tsunoda, M.; Dugyala, R.R.; Sharma, R.P. Fumonisin B1-induced increases in neurotransmitter
metabolite levels in different brain regions of BALB/c mice. Comp. Biochem. Physiol. C.
Pharmacol. Toxicol. Endocrnol. 1998, 120, 457–465.
118. Porter, J.K.; Voss, K.A.; Chamberlain, W.J.; Bacon, C.W.; Norred, W.P. Neurotransmitters in
rats fed fumonisin B1. Proc. Soc. Exp. Biol. Med. 1993, 202, 360–364.
119. Banczerowski-Pelyhe, I.; Vilagi, I.; Detri, L.; Doczi, J.; Kovacs, F.; Kukorelli, T. In vivo and in
vitro electrophysiological monitoring of rat neocortical activity after dietary fumonisin exposure.
Mycopathologia 2002, 153, 149–156.
120. Osuchowski, M.F.; Edwards, G.L.; Sharma, R.P. Fumonisin B1-induced neurodegeneration in
mice after intracerebroventricular infusion is concurrent with disruption of sphingolipid
metabolism and activation of proinflammatory signaling. Neurotoxicology 2005, 26, 211–221.
Int. J. Mol. Sci. 2011, 12 5233
121. Bouhet, S.; Hourcade, E.; Loiseau, N.; Fikry, A.; Martinez, S.; Roselli, M.; Galtier, P.;
Mengheri, E.; Oswald, I.P. The mycotoxin fumonisin B1 alters the proliferation and the barrier
function of porcine intestinal epithelial cells. Toxicol. Sci. 2004, 77, 165–171.
122. Ramasamy, S.; Wang, E.; Hennig, B.; Merrill, A.H., Jr. Fumonisin B1 alters sphingolipid
metabolism and disrupt the barrier function of endothelial cells in culture. Toxicol. Appl.
Pharmacol. 1995, 133, 343–348.
123. Osuchowski, M.F.; He, Q.; Sharma, R.P. Fumoniin B1 toxicity in the brain during coexisting
lipopolysaccharide-related endotoxemia in BALB/c mice. Toxicol. Sci. 2003, 72, 252–253.
124. Szelenyi, J. Cytokines and the central nervous system. Brain Res. Bull. 2001, 54, 329–338.
125. Buccoliero, R.; Futerman, A.H. The roles of ceramide and complex sphingolipids in neuronal
cell function. Pharmacol. Res. 2003, 47, 409–419.
126. Pettus, B.J.; Chalfant, C.E.; Hannun, Y.A. Ceramide in apoptosis: An overview and current
perspectives. Biochem. Biophys. Acta 2002, 1585, 114–125.
127. Stockmann-Juvalla, H.; Mikkola, J.; Naarala, J.; Loikkanen, J.; Elovaara, E.; Savolainen, K.
Oxidative stress induced by fumonisin B1 in continuous human and rodent neural cell cultures.
Free Radic. Res. 2004, 38, 933–942.
128. Mobio, T.A.; Anane, R.; Baudrimont, I.; Carratū, M.R.; Shier, T.W.; Dano, S.D.; Ueno, Y.;
Creppy, E.E. Epigenetic properties of fumonisin B1: cell cycle arrest and DNA base modification
in C6 glioma cells. Toxicol. Appl. Pharmacol. 2000, 164, 91–96.
129. Mobio, T.A.; Baudrimont, I.; Sanni, A.; Shier, T.W.; Saboureau, D.; Dano, S.D.; Ueno, Y.;
Steyn, P.S.; Creppy, E.E. Prevention by vitamin E of DNA fragmentation and apoptosis induced
by fumonisin B1 in C6 glioma cells. Arch. Toxicol. 2000, 74, 112–119.
130. Mobio, T.A.; Tavan, E.; Baudrimont, I.; Anane, R.; Carratū, M.R.; Sanni, A.; Gbeassor, M.F.;
Shier, T.W.; Narbonne, J.-F.; Creppy, E.E. Comparative study of the toxic effects of fumonisin
B1 in rat C6 glioma cells and p53-null mouse embryo fibroblasts. Toxicology 2003, 183, 65–75.
131. Galvano, F.; Campisi, A.; Russo, A.; Galvano, G.; Palumbo, M.; Renis, M.; Barcellona, M.L.;
Perez-Polo, J.R.; Vanella, A. DNA damage in astrocytes exposed to fumonisin B1. Neurochem.
Res. 2002, 27, 345–351.
132. Galvano, F.; Russo, A.; Cardile, V.; Galvano, G.; Vanella, A.; Renis, M. DNA damage in human
fibroblasts exposed to fumonisin B1. Food Chem. Toxicol. 2002, 40, 25–31.
133. Ellerby, L.M.; Ellerby, H.M.; Park, S.M.; Holleran, A.L.; Murphy, A.N.; Fiskum, G.; Kane, D.J.;
Testa, M.P.; Kayalar, C.; Bredesen, D.E. Shift of cellular oxidation-reduction potential in neural
cells expressing Bcl-2. J. Neurochem.1996, 67, 1259–1267.
134. Kane, D.J.; Sarafian, T.A.; Anton, R.; Hahn, H.; Butler, G.E.; Selverstone, V.J.; Ord, T.;
Bredesen, D.E. Bcl-2 inhibition of neural death: decreased generation of reactive oxygen species.
Science 1993, 262, 1274–1277.
135. Tjalkens, R.B.; Ewing, M.M.; Philbert, M.A. Differential cellular regulation of the mitochondrial
permeability transition in an in vitro model of 1,3-dinitrobenzene-induced encephalopathy. Brain
Res. 2000, 874, 165–177.
136. Reed, J.C.; Meister, L.; Tanaka, S.; Cuddy, M.; Yum, S.; Geyer, C.; Pleasure, D. Differential
expression of bcl-2 protooncogene in neuroblastoma and other human tumor cell lines of
neuronal origin. Cancer Res. 1991, 51, 6529–6538.
Int. J. Mol. Sci. 2011, 12 5234
137. Stockmann-Juvala, H.; Mikkola, J.; Naarala, J.; Loikkanen, J.; Elovaara, E.; Savolainen, K.
Fuminisin B1-induced toxicity and oxidative damage in U-118MG glioblastoma cells. Toxicology
2004, 202, 173–183.
138. Schmelz, E.M.; Dombrink-Kurzman, M.A.Roberts, P.C.; Kozutsumi, Y.; Kawasaki, T.;
Merrill, A.H., Jr. Induction of apoptosis by fumonisin B1 in HT29 cells is mediated by the
accumulation of endogenous free sphingoid bases. Toxicol. Appl. Pharmacol.1998, 148, 252–260.
139. Tolleson, W.H.; Dooley, K.L.; Sheldon, W.G.; Thurman, J.D.; Bucci, T.J.; Howard, P.C. The
Mycotoxin Fumonisin Induces Apoptosis in Cultured Human Cells and in Livers and Kidneys of
Rats. In Advances in Experimental and Medical Biology. Fumonisins in Food; Jackson, L.S.,
DeVries, J.W., Bullerman, L.B., Eds.; Plenum Press: New York, NY, USA, 1996; pp. 237–250.
140. Tolleson, W.H.; Melchior, W.B.; Morris, S.M.; McGarrity, L.J.; Domon, O.E.; Muskhelishvili, L.;
James, S.J.; Howard, P.C. Apoptotic and anti-proliferaive effects of fuminisin B1 in human
keratinocytes, fibroblasts, esophageal epithelial cells and hepatoma cells. Carcinogenesis 1996,
141. Higuchi, Y. Chromosomal DNA fragmentation in apoptosis and necrosis induced by oxidative
stress. Biochem. Pharmacol. 2003, 66, 1527–1535.
142. Slater, A.F.G.; Nobel, C.S.I.; van den Dobbelsteen, D.J.; Orrenius, S. Signaling mechanisms and
oxidative stress in apoptosis. Toxicol. Lett. 1995, 82/83, 149–153.
143. Galtier, P. Pharmacokinetics of ochratoxin A in animals. IARC Sci. Publ. 1991, 187–200.
144. Pfohl-Leszkowicz, A.; Manderville, R.A. Ochratoxin A: An overview on toxicity and
carcinogenicity in animals and humans. Mol. Nutr. Food Res. 2007, 51, 61–99.
145. Garies, M.; Wolff, J. Relevance of mycotoxin contaminated feed for farm animals and carryover
of mycotoxins to food of animal origin. Mycoses 2000, 43, 79–83.
146. Mally, A.; Hard, G.C.; Dekant, W. Ochratoxin A as a potential etiologic factor in endemic
nephropathy: lesions from toxicity studies in rats. Food Chem. Toxicol. 2007, 45, 2254–2260.
147. Krogh, P. Role of ochratoxin in disease causation. Food Chem. Toxicol. 1992, 30, 213–224.
148. Kane, A.; Creppy, E.E.; Roschenthaler, R.; Dirheimer, G. Changes in urinary and renal tubular
enzymes caused by subchronic administration of ochratoxin A in rats. Toxicology 1986, 42,
149. Petkova-Bocharova, T.; Chernozemsky, I.N.; Castegnaro, M. Ochratoxin A in human blood in
relation to Balkan endemic nephropathy and urinary system tumors in Bulgaria. Food Addit.
Contam. 1988, 5, 299–301.
150. Lea, T.; Steinen, K.; Stormer, F.C. Mechanism of ochratoxin A-induced immunosuppression.
Mycopathologia 1989, 107, 153–159.
151. Stromer, F.C.; Lea, T. Effects of ochratoxin A upon early and late events in human T-cell
proliferation. Toxicology 1995, 95, 45–50.
152. Arora, R.G.; Frolen, H.; Fellner-Feldegg, H. Inhibition of ochratoxin A teratogenesis by
zearalenone and diethylstilbesterol. Food Chem. Toxicol. 1983, 21, 779–783.
153. Fukui, Y.; Hayasaka, S.; Itoh, M.; Takeuchi, Y. Development of neurons and synapses in
ochratoxin A-induced microcephalic mice: a quantitative assessment of somatosensory cortex.
Neurotoxicol. Teratol. 1992, 14, 191–196.
Int. J. Mol. Sci. 2011, 12 5235
154. Pfohl-Leszkowicz, A.; Chakor, K.; Creppy, E.E.; Dirheimer, G. DNA adduct formation in mice
treated with ochratoxin A. IARC Sci. Publ. 1991, 245–253.
155. Sava, V.; Reunova, O.; Velasquez, A.; Harbison, R.; Sanchez-Ramos, J. Acute neurotoxic effects
of the fungal netabolite ochratoxin-A. Neurotoxicology 2006, 27, 82–92.
156. Kuiper-Goodman, T.; Hilts, C.; Billiard, S.M.; Kiparissis, Y.; Richard, I.D.; Hayward, S. Health
risk assessment of ochratoxin A for all age-sex strata in a market economy. Food Addit. Contam.
Part A Chem. Anal. Control Expo. Risk Assess. 2010, 27, 212–240.
157. Creppy, E.E.; Chakor, K.; Fisher, M.J.; Dirheimer, G. The mycotoxin ochratoxin A is a substrate
for phenylalanine hydroxylase in isolated rat hepatocytes and in vivo. Arch. Toxicol.1990, 64,
158. Creppy, E.E.; Kane, D.; Dirheimer, G.; Lafarge-Frayssinet, C.; Mousset, S.; Frayssinet, C.
Genotoxicity of ochratoxin A in mice: DNA single-strand break evaluation in spleen, liver and
kidney. Toxicol. Lett. 1985, 28, 29–35.
159. Dirheimer, G.; Creppy, E.E. Mechanism of action of ochratoxin A. IARC Sci. Publ. 1991,
160. Gautier, J.C.; Holzhaeuser, D.; Markovic, J.; Gremaud, E.; Schilter, B.; Turesky, R.J. Oxidative
damage and stress response from ochratoxin exposure in rats. Free Radic. Biol. Med. 2001, 30,
161. Bryan, N.S.; Rassaf, T.; Maloney, R.E.; Rodriguez, C.M.; Saijo, F.; Rodriguez, J.R.; Feelisch, M.
Cellular targets and mechanisms of nitros(yl)ation: An insight into their nature and kinetics in
vivo. Proc. Natl. Acad. Sci. USA 2004, 101, 4308–4313.
162. Thomas, J.A.; Mallis, R.J. Aging and oxidation of reactive protein sulfhydryls. Exp. Gerontol.
2001, 36, 1519–1526.
163. Marin-Kuan, M.; Nestler, S.; Verguet, C.; Bezençon, C.; Piguet, D.; Mansourian, R.; Holzwarth, J.;
Grigorov, M.; Delatour, T.; Mantel, P.; et al. A toxicogenomics approach to identify new
plausible epigenetic mechanisms of ochratoxin A carcinogenicity in rat. Toxicol. Sci. 2006, 89,
164. Aleo, M.D.; Wyatt, R.D.; Schnellmann, R.G. Mitochondrial dysfunction is an early event in
ochratoxin A but not oosporein toxicity to rat renal proximal tubules. Toxicol. Appl. Pharmacol.
1991, 107, 73–80.
165. Wei, Y.H.; Lu, C.Y.; Lin, T.N.; Wei, R.D. Effect of ochratoxin A on rat liver mitochondrial
respiration and oxidative phosphorylation. Toxicology 1985, 36, 119–130.
166. Belmadani, A.; Tramu, G.; Betbeder, A.M.; Creppy, E.E. Subchronic effects of ochratoxin A on
young adult rat brain and partial prevention by aspartate, a sweetener. Hum. Exp. Toxicol. 1998,
167. Hayes, A.W.; Cain, J.A.; Moore, B.G. Effects of aflatoxin B1, ochratoxin A and rubratoxin B on
infant rats. Food Cosmet. Toxicol. 1977, 15, 23–27.
168. Hayes, A.W.; Hood, R.D.; Lee, H.L. Teratogenic effects of ochratoxin A in mice. Teratology
1974, 9, 93–97.
169. Wangikar, P.B.; Dwivedi, P.; Sharma, A.K.; Sinha, N. Effect in rats of simultaneous prenatal
exposure to ochratoxin A and aflatoxin B(1). II. Histopathological features of teratological
anomalies induced in fetuses. Birth Defects Res. B 2004, 71, 352–358.
Int. J. Mol. Sci. 2011, 12 5236
170. Tamura, M.; Hirata, Y.; Matsutani, T. Neurochemical effects of prenatal treatment with
ochtatoxin A on fetal and adult mouse brain. Neurochem. Sci. 1988, 13, 1139–1147.
171. Belmadani, A.; Tramu, G.; Betbeder, A.M.; Steyn, P.S.; Creppy, E.E. Regional selectivity to
ochratoxin A, distribution and cytotoxicity in rat brain. Arch. Toxicol. 1998, 72, 656–662.
172. Sava, V.; Reunova, O.; Velasquez, A.; Sanchez-Ramos, J. Can low level exposure to ochratoxin-
A cause parkinsonism? J. Neurol. Sci. 2006, 249, 68–75.
173. Sanchez-Ramos, J.; Overvik, E.; Ames, B.N. A marker of oxyradical-mediated DNA damage
(oxo8dG) is increased in nigro-striatum of Parkinson’s disease brain. Neurodegeneration
(incorporated into Exp. Neurol.) 1994, 3, 197–204.
174. Bunge, I.; Dirheimer, G.; Roschenthaler, R. In vivo and in vitro inhibition of protein synthesis in
Bacillus stearothermophilus by ochratoxin A. Biochem. Biophys. Res. Commun.1978, 83,
175. Creppy, E.E.; Kern, D.; Steyn, P.S.; Vleggaar, R.; Roschenthaler, R.; Dirheimer, G. Comparative
study of the effect of ochratoxin a analogues on yeast aminoacyl-tRNA synthetases and on the
growth and protein synthesis of hepatoma cells. Toxicol. Lett. 1983, 19, 217–224.
176. Palmer, T.D.; Takahashi, J.; Gage, F.H. The adult rat hippocampus contains primordial neural
stem cells. Mol. Cell. Neurosci.1997, 8, 389–404.
177. Song, H.J.; Stevens, C.F.; Gage, F.H. Neuronal stem cells from adult hippocampus develop
essential properties of functional CNS neurons. Nat. Neurosci. 2002, 5, 438–445.
178. Chen, H.; Tung, Y.C.; Li, B.; Iqbal, K.; Grundke-Iqbal, I. Trophic factors counteract elevated
FGF-2-induced inhibition of adult neurogenesis. Neurobiol. Aging 2006, 28, 1148–1162.
179. Kawai, T.; Takagi, N.; Mochizuki, N.; Besshoh, S.; Sakanishi, K.; Nakahara, M.: Takeo, S.
Inhibitor of vascular endothelial growth factor receptor tyrosine kinase attenuates cellular
proliferation and differentiation to mature neurons in the hippocampal dentate gyrus after
transient forebrain ischemia in the adult rats. Neuroscience 2006, 141, 1209–1216.
180. Lagace, D.C.; Yee, J.K.; Bolanos, C.A.; Eisch, A.J. Juvenile administration of methylphenidate
attenuates adult hippocampal neurogenesis. Biol. Psychiatry 2006, 60, 1121–1130.
181. Rossi, C.; Angelucci, A.; Costantin, L.; Braschi, C.; Mazzantini, M.; Babbini, F.; Fabbri, M.E.;
Tessarollo, L.; Maffei, L.; Berardi, N.; Caleo, M. Brain-derived neurotrophic factor (BDNF) is
required for the enhancement of hippocampal neurogenesis following environmental enrichment.
Eur. J. Neurosci .2006, 24, 1850–1856.
182. Delibas, N.; Altuntas, I.; Yonden, Z.; Ozcelik, N. Ochratoxin A reduces NMDA receptor
subunits 2A and 2B concentrations in rat hippocampus: partial protective effect of melatonin.
Hum. Exp. Toxicol. 2003, 22, 335–339.
183. Sava, V.; Velasquez, A.; Song, S.; Sanchez-Ramos, J. Adult hippocampal neural stem/progenitor
cells in vitro are vulnerable to the mycotoxin ochratoxin A. Toxicol. Sci. 2007, 98, 187–197.
184. Alexander, P. The role of DNA lesions in processes leading to aging in mice. Sym. Soc. Exp.
Biol. 1967, 21, 29–50.
185. Korr, H.; Schultz, B. Unscheduled DNA synthesis in various types of cells of the mouse brain in
vivo. Exp. Brain Res. 1989, 74, 573–578.
Int. J. Mol. Sci. 2011, 12 5237
186. Crago, B.R.; Gray, M.R.; Nelson, L.A.; Davis, M.; Arnold, L.; Thrasher, J.D. Psychological,
neuropsychological, and electrocortical effects of mixed mold exposure. Arch. Environ. Health
2003, 58, 452–563.
187. Gordon, W.A.; Cantor, J.B. The diagnosis of cognitive impairment associated with exposure to
mold. Adv. Appl. Microbiol. 2004, 55, 361–374.
188. Rea, W.J.; Didriksen, N.; Simon, T.R.; Pan, Y.; Fenyves, E.J.; Griffiths, B. Effects of toxic
exposure to molds and mycotoxins in building-related illnesses. Arch. Environ. Health 2003, 58,
189. Yoon, S.; Cong, W.-T.; Bang, Y.; Lee, S.N.; Yoon, C.S.; Kwack, S.J.; Kang, T.S.; Lee, K.Y.;
Choi, J.-K.; Choi, H.J. Proteome response to ochratoxin A-induced apoptotic cell death in mouse
hippocampal HT22 cells. Neurotoxicology 2009, 30, 666–676.
190. Sato, A.; Miyazaki, E.; Satake, A.; Hiramoto, A.; Hiraoka, O.; Miyake, T.; Kim, H.S.; Wataya, Y.
Proteome and transcriptome analysis of cell death induced by 5-fluoro-2’-deoxyuridine. Nucleic
Acids Symp. Ser (Oxf). 2007, 51, 433–434.
191. Siddiq, A.; Ayoub, I.A.; Chavez, J.C.; Aminova, L.; Shah, S.; LaManna, J.C.; Patton, S.M.;
Connor, J.R.; Cherny, R.A.; Volitakis, I.; et al. Hypoxia-indicible factor prolyl 4-hydroxylase
inhibition. A target for neurprotection in the central nervous system. J. Biol. Chem. 2005, 280,
192. Lei, T.; He, Q.; Cai, Z.; Zhou, Y.; Wang, Y.; Si, L.; Cai, Z.; Chiu, J.F. Proteomic analysis of
chromium cytotoxicity in cultured rat lung epithelial cells. Proteomics 2008, 8, 2420–2429.
193. Noguchi, M.; Takata, T.; Kimura, Y.; Manno, A.; Murakami, K.; Koike, M.; Ohizumi, H.;
Hori, S.; Kakizuka, A. ATPase activity of p97/valosin-containing protein is regulated by
oxidative modification of the evolutionally conserved cysteine 522 residue in Waker A motif.
J. Biol. Chem. 2005, 280, 41332–41341.
194. Zhang, X.; Boesch-Saadatmandi, C.; Lou, Y.; Wolffram, S.; Huebbe, P.; Rimbach, G.
Ochratoxin A induces apoptosis in neuronal cells. Genes Nutr. 2009, 4, 41–48.
195. Zurich, M.G.; Lengacher, S.; Braissant, O.; Monnet-Tschudi, F.; Pellerin, L.; Honegger, P.
Unusual astrocyte reactivity caused by the food mycotoxin ochratoxin A in aggregating rat brain
cell cultures. Neuroscience 2005, 134, 771–782.
196. Hong, J.T.; Lee, M.K.; Park, K.S.; Jung, K.M.; Lee, R.D.; Jung, H.K.; Park, K.L.; Yang, K.J.;
Chung, Y.S. Inhibitory effect of peroxisome proliferator-activated receptor gamma agonist on
ochratoxin A-induced cytotoxicity and activation of transcription factors in cultured rat
embryonic midbrain cells. J. Toxicol. Environ. Health A 2002, 65, 407–418.
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