PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of ijmsMDPIhomeThis articleThis journalInstructions for authorsSubscribeIJMS
 
Int J Mol Sci. 2011; 12(8): 5213–5237.
Published online 2011 August 15. doi:  10.3390/ijms12085213
PMCID: PMC3179161

Mechanisms of Mycotoxin-Induced Neurotoxicity through Oxidative Stress-Associated Pathways

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

1. Introduction

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 [1], 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. [2] described 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 [6] and has been implicated in additional mycotoxicoses such as red mold disease in humans and animals [7] and beanhull poisoning in horses [8].

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 [911]. 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) [11]. 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 [1417]. T-2 toxin can induce apoptosis in many types of cells bearing rapid rates of proliferation [1822]. 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) [23]. 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 [24]. 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 [27], and therefore, there are only a small number of reports of T-2 toxin-induced neurotoxicity [2830]. Boyd et al. [28] reported that low levels of T-2 toxin were responsible for the changes in the metabolism of brain biogenic monoamines, and Wang et al. [31] showed that ingestion of T-2 toxin leads to changes in amino acid permeability across the blood-brain barrier, which could 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 [3235].

Sehata et al. [36] 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 [36].

T-2 toxin suppresses drug metabolizing enzymes such as glutathione S-transferases (GSTs) [3638]. In addition, a decreased expression in mitochondria-related genes, such as mitochondrial NADH-dehydrogenase and cytochrome oxidase, has been reported in the fetal brain [36], 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. [36] 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 [39]. 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 [40], and JNK and c-jun are important regulators of apoptosis in the nervous system [41].

Differing from the results in the maternal liver, placenta and fetal liver [24], 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 [36], 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 [4244]. 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 [24], 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 [4549].

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 [36]. VEGF is expressed in neurons and may play a role in the maintenance of neurons and endothelial cells in the CNS [50]. 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.

Recently, Chaudhary and Rao [27] 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 [51]. 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 [27] 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 [27], 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 oxidative stress.

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 [5457]. 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 [56,6062]. 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 [56].

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 [63]. 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. [56] have reviewed the relationship between S. chartarum, trichothecene mycotoxins, and DBRI and proposed new insights into a public health enigma.

Besides inhibiting translation, macrocyclic trichothecenes as well as other trichothecenes can simultaneously activate p38, JNK and ERK and MAPKs in vivo and in vitro [6668] via a process referred to as “ribotoxic stress” [69]. 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 [11,7075]. The common ability of macrocyclic trichothecenes to cause protein synthesis inhibition via binding to the 18S rRNA of the ribosomal large subunit [76] 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 [75] 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 [77]. RL2 is said to be nontoxic [78]. 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 [79].

Murine alveolar type II cells and alveolar macrophages are extremely sensitive to intratracheally instilled S. chartarum spores [80]. 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 [81]. In a follow-up study, extract-induced apoptosis in MH-S cells was observed to precede DNA damage [82]. 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. [83] 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 [84]. 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 [83] or RA [84], 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 OSNs [8587]. PKR associates with the ribosome [88] and can selectively shut down translation via phosphorylation of eukaryolic initiation factor 2α (eIF2α) as well as activate NF-κB [89].

Both TNF-α and Fas directly induce apoptosis in OE [90] 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. [93] 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 [93].

In the study by Islam et al. [93], 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 [94]. Moreover, Nusuetrong et al. [79] 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 [96]. 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 [97]. 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].

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 [104]. 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 [106].

FB1 is well known to cause equine leukoencephalomalacia (ELEM) [107,108]. This disease is associated primarily with FB1 and is characterized by high mortality [108]. In histopathological examinations, pathognomonic focal necrotic lesions, located primarily in the subcortical white matter are apparent [102]. In addition, the elevation of free sphingoid bases after FB1 treatment has been demonstrated in the brain of ELEM-diagnosed horses [109], 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 [110]. Treatment with FB1 causes NTDs in ex vivo neurulating mouse embryos [111] and this effect is related to the folic acid receptor deficiency as a result of the FB1-dependent lipid rafts depletion [112].

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 [113], increases levels of sphinganine concentration in the forebrain and brain stem of rats accompanying a concomitant demyelination in the forebrain [114], and disrupts myelination in glial cells but not neurons in aggregating brain cell culture [115] and in developing rats [116]. FB1-dependent changes in neurotransmitter metabolite levels in different brain regions of BALB/c mice [117] and in rat brain [118], and alteration of electrophysiological activity in rat neocortex [199] are also reported.

Osuchowski et al. [120] 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 [123] and that FB1 may disrupt central nervous system homeostasis when brain tissue is directly exposed to this mycotoxin.

All cytokines analyzed in the study of Osuchowski et al. [120] 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) [124]. 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 [120].

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. [127] 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 FB1 [104].

Mobio et al. [128130] 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 [127], and may increase their susceptibility to radical attack [133135]. 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. [137] 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 [129131,138140]. 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 [147] and enzymuria [148] 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 [154] and neurotoxic [155] effects. Although there is still insufficient evidence in humans, there is sufficient evidence in experimental animals for the carcinogenicity of OTA [156].

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 [155,157160]. 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 [53]. 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 [163]. 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 [166169]. OTA has been shown to affect proliferation and migration of neurons [153] and reduce DNA content [170] in the developing rodent brain. OTA is also reported to be neurotoxic to adult male rats [166]. Neurotoxicity is more pronounced in the ventral mesencephalon, hippocampus (HP), and striatum than in the cerebellum (CB) [171]. 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 [155].

Sava et al. [155] 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 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 biphasic response.

Sava et al. [172] 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 [173] 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 [155]. 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 [166]. 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 [157], 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 [178181]. 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 [182].

Sava et al. [183] 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 inhibitor of mitochondrial oxidative metabolism may be less critical than its interference with DNA synthesis and mitotic competence. Overall, Sava et al. [183] 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 [186188].

Yoon et al. [189] 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 [190], 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 [191], ornithine has been shown to be up-regulated during ROS-related apoptotic cell death [192], 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 [193]. Contrary to the above-mentioned report by Sava et al. [183], Zhang et al. [194] 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 SHSY5Y 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. [195] 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. [196] 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 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.

6. Conclusions

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 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 essentially involved.

Acknowledgments

The authors would like to thank Pete Aughton, D.A.B.T., ITR Laboratories Canada Inc., for proofreading.

References

1. Haschek WM, Voss KA, Beasley VR. Selected Mycotoxins Affecting Animal and Human Health. In: Haschek WM, Rousseaux CG, Walling MA, editors. Handbook of Toxicologic Pathology. Academic Press; San Diego, CA, USA: 2002. pp. 645–699.
2. Surai PF, Mezes M, Melnichuk SD, Fotina TI. 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. [PubMed]
4. Desjardins AE, Hohn TM, McComic SP. Trichothecene biosynthesis in Fusarium species: chemistry, genetics, and significance. Microbiol. Mol. Biol. Rev. 1993;57:595–604. [PMC free article] [PubMed]
5. Nelson PE, Dignani MC, Anaissie EJ. Taxonomy, biology, and clinical aspects of Fusarium species. Clin Microbiol Rev. 1994;7:479–504. [PMC free article] [PubMed]
6. Joffe AZ. Foodborne Diseases: Alimentary Toxic Aleukia. In: Rochcigle M, editor. Handbook of Foodborne Diseases of Biological Origin. CRC Press; Boca Raton, FL, USA: 1983. pp. 353–495.
7. Saito M, Ohtsubo K. Trichothecene Toxins of Fusarium Species. In: Purchase IFH, editor. Mycotoxins. 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. [PubMed]
9. Bennet JW, Klich M. Mycotoxins. Clin. Microbiol. Rev. 2003;16:497–516. [PMC free article] [PubMed]
10. Eriksen GS, Petterson H. Toxicological evaluation of trichothecenes in animal feed. Anim. Feed Sci. Technol. 2004;114:205–239.
11. Shifrin VI, 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. [PubMed]
12. Chang IM, Mar WC. Effect of T-2 toxin on lipid peroxidation in rats: Elevation of conjugated diene formation. Toxicol. Lett. 1988;40:275–280. [PubMed]
13. Eriksen GS, Petterson H, Lund H. Comparative cytotoxicity of deoxynivalenol, nivalenol, triacetylated derivatives and de-epoxy metabolites. Food Chem. Toxicol. 2004;42:619–624. [PubMed]
14. IARC IARC Monographson the Evaluation of Carcinogenic Risks to Humans. IARC; Lyon, France: 1993. Toxins derived from Fusarium sporotrichoides: T-2 toxin; pp. 467–488. [PubMed]
15. Sharma RP. Immunotoxicity of mycotoxins. J. Dairy Sci. 1993;76:892–897. [PubMed]
16. Stanford GK, Hood RD, Haynes AW. Effects of prenatal administration of T-2 toxin to mice. Res Commu Chem Pathol Pharmacol. 1975;10:743–746. [PubMed]
17. Williams PP. 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. [PubMed]
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. [PubMed]
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. [PubMed]
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. [PubMed]
22. Albarenque SM, 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;14:337–342. [PubMed]
23. Shinozuka J, Miwa S, Fujimura H, Toriumi W, Doi K. Hepatotoxicity of T-2 Toxin, Trichothecene Mycotoxin. In: Kumagai S, editor. New Strategies for Mycotoxin Research in Asia (Proceedings of ISMYCO Bangkok ‘06) 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. [PubMed]
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. [PMC free article] [PubMed]
27. Chaudhary M, Rao PV. Brain oxidative stress after dermal and subcutaneous exposure of T-2 toxin in mice. Food Chem. Toxicol. 2010;48:3436–3442. [PubMed]
28. Boyd KE, Fitzpatrick DW, Wilson JR, Wilson LM. Effect of T-2 toxin on brain biogenic monoamines in rats and chickens. Can. J. Vet. Res. 1988;52:181–185. [PMC free article] [PubMed]
29. Martin LJ, Morse JD, Anthony A. Quantitative cytophotometric analysis of brain neuronal RNA and protein changes in acute T-2 mycotoxin poisoned rats. Toxicon. 1986;24:933–941. [PubMed]
30. Wang J, Fitzpatrick DW, Wilson JR. Effects of the trichothecene mycotoxin T-2 toxin on neurotransmitters and metabolites in discrete areas of the rat brain. Food Chem. Toxicol. 1998;36:947–953. [PubMed]
31. Wang J, Fitzpatrick DW, Wilson JR. 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. [PubMed]
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. [PubMed]
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. [PubMed]
34. Rousseaux CG, Schiefer HB. Maternal toxicity, embryolethality and abnormal fetal development in CD-1 mice following one oral dose of T-2 toxin. J. Appl. Toxicol. 1987;7:281–288. [PubMed]
35. Stanford GK, Hood RD, Hayes AW. Effect of prenatal administration of T-2 toxin to mice. Res. Commun. Chem. Path. Pharmacol. 1975;10:743–746. [PubMed]
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. [PubMed]
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;27:215–220. [PubMed]
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. [PubMed]
39. Jarpe MB, Widmann C, Knall C, Schlesinger TK, Gibson S, Yujiri T, Fanger GR, Gelfand EW, Johnson GL. Anti-apoptotic versus pro-apoptotic signal transduction: checkpoints and stop signs along the roard to death. Oncogene. 1998;17:1475–1582. [PubMed]
40. Bold S, Weidle UH, Kolch W. The kinase domain of MEKK1 induces apoptosis by dysregulation of MAP kinase pathways. Exp. Cell Res. 2003;283:80–90. [PubMed]
41. Ham J, Eilers A, Whitfield J, Neame SJ, Shah B. c-JUN and the transcriptional control of neuronal apoptosis. Biochem. Pharmacol. 2000;60:1015–1021. [PubMed]
42. Annunziato L, Amoroso S, Pannaccione A, Cataldi M, Pignataro G, D’Alessio S, Sirabella R, Second A, Sibaud L, DiRenzo GF. Apoptosis induced in neuronal cells by oxidative stress: role played by caspases and intracellular calcium ions. Toxicol. Lett. 2003;139:125–133. [PubMed]
43. Troy CM, Shelanski ML. Caspase-2 redux. Cell Death Differ. 2003;10:101–107. [PubMed]
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. [PubMed]
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. [PubMed]
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. [PubMed]
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. [PubMed]
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. [PubMed]
49. Woo GH, Bak EJ, Nakayama H, Doi K. Molecular mechanisms of hydroxyurea (HU)-induced apoptosis in the mouse fetal brain. Neurotox. Teratol. 2006;28:125–134. [PubMed]
50. Ogunshola OO, Antic A, Donoghue MJ, Fan S-Y, Kim H, Stewart WB, Madri JA, Ment LR. Paracrine and autocrine function of neuronal vascular endothelial growth factor (VEGF) in the central nervous system. J. Biol. Chem. 2002;277:11410–11415. [PubMed]
51. Halliwell B, Gutteridge JMC. Free Radicals in Biology and Medicine. 3rd ed. Oxford University Press; New York, NY, USA: 1999.
52. Lee J-M, Jiang L, Johnson DA, Stein TD, Kraft AD, Calkins MJ, Jakel RJ, Jofnson JA. Nrf2, a multiorgan protector? FASEB J. 2005;19:1061–1066. [PubMed]
53. Boesch-Saadatmandi C, Wagner AE, Graeser AC, 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. [PubMed]
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. [PubMed]
55. Tsumori T, Reijula K, Johnsson T, Hemminki K, Hintikka EL, 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. [PMC free article] [PubMed]
56. Pestka JJ, Yike I, Dearborn DG, Ward MDW, Harkema JR. Stachybotrys chartarum, trichothecene mycotoxins, and damp building-related illness: New insights into a public health enigma. Toxicol. Sci. 2008;104:4–26. [PubMed]
57. Shelton BG, Kirkland KH, Flanders WD, Morris GK. Profiles of airborne fungi in buildings and outdoor environments in the United States. Appl. Environ. Microbiol. 2002;68:1743–1753. [PMC free article] [PubMed]
58. Hodgson MJ, Morey P, Leung WY, Morrow L, Miller D, Jarvis BB, Robbins H, Halsey JF, Storey E. Building-associated pulmonary disease from exposure to Stachybotrys chartarum and Aspergillus versicolor. J. Occup. Environ. Med. 1998;40:241–249. [PubMed]
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. [PubMed]
60. Gordon WA, Cantor JB, Johanning E, Charatz HJ, Ashman TA, Breeze JL, Haddad L, Abramowitz S. Cognitive impairment associated with toxigenic fungal exposure: A replication and extension of previous findings. Appl. Neuropsychol. 2004;11:65–74. [PubMed]
61. Hossain MA, Ahmed MS, Ghannoum MA. Attributes of Stachybotrys chartarum and its association with human disease. J. Allergy Clin. Immunol. 2004;113:200–208. [PubMed]
62. Kirburn KH. Role of molds and myxotoxins in being sick in buildings: Neurobehavioral and pulmonary impairment. Adv. Appl. Microbiol. 2004;55:339–359. [PubMed]
63. Andersen B, Nielsen KF, Jarvis BB. Characterization of Stachybotrys from water-damaged buildings based on morphology, growth, and metabolite production. Mycologia. 2002;94:392–403. [PubMed]
64. Gregory L, Pestka JJ, Dearborn DG, Rand TG. Localization of satratoxin-G in Stachybotrys chartarum spores and spore-impacted mouse lung using immunocytochemistry. Toxicol. Pathol. 2004;32:26–34. [PubMed]
65. Yike I, Distler AM, Ziady AG, Dearborn DG. Mycotoxin adducts on human serum albumin: Biomerkers of exposure to Stachybotrys chartarum. Environ. Health Perspect. 2006;114:1221–1226. [PMC free article] [PubMed]
66. Chung YJ, Zhou HR, Pestka JJ. 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. [PubMed]
67. Moon Y, Pestka JJ. Deoxynivalenol-induced mitogen-activated protein kinase phosphorylation and IL-6 expression in mice suppressed by fish oil. J. Nutr. Biochem. 2003;14:717–726. [PubMed]
68. Zhou HR, Lau AS, Pestka JJ. Role of double-stranded RNA-activated protein kinase R (PKR) in deoxynivalenol-induced ribotoxic stress response. Toxicol. Sci. 2003;74:335–344. [PubMed]
69. Iordanov MS, Pribnow D, Magun JL, Dinh TH, Pearson JA, Chen SL, Magun BE. 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. [PMC free article] [PubMed]
70. Chung YJ, Jarvis B, Pestka JJ. 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. [PubMed]
71. Chung YJ, Yang GH, Islam Z, Pestka JJ. Up-regulation of macrophage inflammatory protein-2 and complement 3A receptor by the trichothecenes deoxynivalenol and satratoxin G. Toxicology. 2003;186:51–65. [PubMed]
72. Hughes BJ, Hsieh GC, Jarvis BB, Sharma RP. Effects of macrocyclic trichothecene mycotoxins on the murine immune system. Arh. Environ. Contam. Toxicol. 1989;18:388–395. [PubMed]
73. Hughes BJ, Jarvis BB, Sharma RP. Effects of macrocyclictrichothecene congeners on the viability and mitogenesis of mirine splenic lymphocytes. Toxicol Lett. 1990;50:57–67. [PubMed]
74. Pestka JJ, Forsell JH. Inhibition of human lymphocyte transformation by the macrocyclic trichothecene roridin A and verrucarin A. Toxicol Lett. 1988;41:215–222. [PubMed]
75. Yang G-H, Jarvis BB, Chung Y-J, Pestka JJ. 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. [PubMed]
76. Cundliffe E, Davies JE. Inhibition of initiation, elongation, and termination of eukaryotic protein synthesis by trichothecene fungal toxins. Antimicrob. Agents Chemother. 1977;11:491–499. [PMC free article] [PubMed]
77. Nielsen KF, Huttunen K, Hyvarinen A, Andersen B, Jarvis BB, Hirvonen MR. Metabolite profiles of Stachybotrys isolates from water-damaged buildings and their induction of inflammatory mediators and cytotoxicity in macrophages. Mycopathologia. 2002;154:201–205. [PubMed]
78. Islam Z, Shinozuka J, Harkema JR, Pestka JJ. Purification and comparative neurotoxicity of the trichothecenes satratoxin G and roridin L2 from Stachybotrys chartarum. J. Toxicol. Environ. Health A. 2009;72:1242–1251. [PMC free article] [PubMed]
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. [PubMed]
80. Rand TG, 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. [PubMed]
81. Wang H, Yadav JS. 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. [PubMed]
82. Wang H, Yadav JS. 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;214:297–308. [PubMed]
83. Islam Z, Harkema JR, Pestka JJ. 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. [PMC free article] [PubMed]
84. Islam Z, Amuzie CJ, Harkema JR, Pestka JJ. 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. [PubMed]
85. Chang RC, Suen KC, Ma CH, Elyaman W, Ng HK, 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. [PubMed]
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;27:521–527. [PubMed]
87. Huang CC, Chen K, Huang TY. Immunohistochemical studies of sensory neurons in rat olfactory epithelium. Eur. Arch. Otorhinolaryngol. 1995;252:86–91. [PubMed]
88. Wu S, Kumar KU, Kaufmam RJ. Identification and requirement of three ribosome binding domains in dsRNA-dependent protein kinase (PKR) Biochemistry. 1998;37:13816–13826. [PubMed]
89. Garcia MA, Meurs EF, Esteban M. The dsRNA protein kinase PKR: Virus and cell control. Biochemie. 2007;89:799–811. [PubMed]
90. Cowan CM, Roskams AJ. Apoptosis in the mature and developing olfactory neuroepithelium. Microsc. Res. Technol. 2002;58:204–215. [PubMed]
91. Farbman AI, Buchholz JA, Suzuki Y, Coines A, Speert D. A molecular basis of cell death in olfactory epithelium. J. Comp. Neurol. 1999;414:306–314. [PubMed]
92. Suzuki Y, Farbman AI. 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. [PubMed]
93. Islam Z, Hegg CC, Bae HY, Pestka JJ. Satratoxin G-induced apoptosis in PC-12 neuronal cells is mediated by PKR and caspase independent. Toxicol. Sci. 2008;105:142–152. [PMC free article] [PubMed]
94. Nusuetrong P, Yoshida M, Tanitsu MA, 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;507:239–246. [PubMed]
95. Chandra J, Samali A, Orrenius S. Triggering and modulation of apoptosis by oxidative stress. Free Radic. Biol. Med. 2000;29:323–333. [PubMed]
96. Karunasena E, Larrañaga MD, Simoni JS, Douglas DR, Straus DC. 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. [PubMed]
97. Thrasher JD, Crawley S. The biocontaminants and complexity of damp indoor spaces; more than what meets the eyes. Toxicol. Ind. Health. 2009;25:583–615. [PubMed]
98. Campbell IL. Neuropathogenic acions of cytokines assessed in transgenic mice. Int. J. Dev. Neurosci. 1995;13:275–284. [PubMed]
99. Peters A, Vweronesi B, Calderon-Garciduenas J, Gehr P, Chen LC, 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;3:1–13. [PMC free article] [PubMed]
100. Calderón-Garcidueñas L, Azzarelli B, Acuna H, Garcia R, Gambling TM, Osnaya N, Monroy S, Tizapantzi MDR, Carson JL, Villarreal-Calderon A, et al. Air pollution and brain damage. Toxicol. Pathol. 2002;30:373–389. [PubMed]
101. Calderón-Garcidueñas L, Maronpot RR, 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. [PubMed]
102. Dutton MF. Fumonisins, mycotoxins of increasing importance: their nature and their effects. Pharmacol. Ther. 1996;70:137–161. [PubMed]
103. Howard PC, Eppley RM, Stack ME, Warbritton A, Voss KA, Lorentzen RJ, Kovach RM, Bucci TJ. Fumonisin B1 carcinogenicity in a 2-year feeding study using F344 rats and B6C3 F1 mice. Environ. Health Perspect. 2001;109:277–282. [PMC free article] [PubMed]
104. Wang E, Norred WP, Bacon CW, Riley RT, Merrill AH., Jr Inhibition of sphingolipid biosynthesis by fumonisins. J. Biol. Chem. 1991;22:14486–14490. [PubMed]
105. Merrill AH, Jr, Sullards MC, Wang E, Voss KA, Riley RT. Sphingolipid metabolism: Role in signal transduction and disruption by fumonisins. Environ. Health Perspect. 2001;109:283–289. [PMC free article] [PubMed]
106. Riley RT, Enongene E, Voss KA, Norred WP, Meredith FI, Sharma RP, Spitsbergen J, Williams DE, Carlson DB, Merrill AH., Jr Sphingolipid perturbations as mechanisms for fumonisin carcinogenesis. Environ. Health Perspect. 2001;109:301–308. [PMC free article] [PubMed]
107. Ross PF, Rice LG, Reagor JC, Osweiler GD, Wilson TM, Nelson HA, Owens DL, Plattner RD, Harlin KA, Richard JL, et al. Fumonisin B1 concentrations in feeds from 45 confirmed equine leukoencephalomalacia cases. J. Vet. Diagn. Invest. 1991;3:238–241. [PubMed]
108. Wilson TM, Ross PF, Rice LG, Osweiler GD, Nelson HA, Owen DL, Plattner RD, Reggiardo C, Noon TH, Pickrell JW. Fumonisin B1 levels associated with an epizootics of equine leukoencephalomalacia. J Vet Diagn Invest. 1990;2:213–216. [PubMed]
109. Goel S, Schumacher J, Lenz SD, Kemppanien BW. Effects of fusarium moniliforme isolates on tissue and serum sphingolipid concentrations in horses. Vet. Hum. Toxicol. 1996;38:265–270. [PubMed]
110. Marasas WF, Riley RT, Hendricks KA, Stevens VL, Sadler TW, 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. [PubMed]
111. Sadler TW, Merrill AH, Stevens VL, Sullards MC, Wang E, Wang P. Prevention of fumonisin B1-induced neural tube defects by folic acid. Teratology. 2002;66:169–176. [PubMed]
112. Stevens VL, Tang J. Fumonisin B1-induced sphingolipid depletion inhibits vitamin uptake via the glycosylphosphatidylinositol-anchored folate receptor. J. Biol. Chem. 1997;272:18020–18025. [PubMed]
113. Harel R, Futerman AH. Inhibition of sphingolipid synthesis affects axonal outgrowth in cultured hippocampal neurons. J. Biol. Chem. 1993;268:14476–14481. [PubMed]
114. Kwon OS, Slikker W, Jr, Davies DL. Biochemical and morphological effects of fumonisin B1 on primary cultures of rat cerebrum. Neurotoxicol. Teratol. 2000;22:565–572. [PubMed]
115. Monnet-Tschudi F, Zurich MG, Sorg O, Matthieu JM, Honegger P, Schilter B. The naturally occurring food mycotoxin fumonisin B1 impairs myelin formation in aggregating brain cell culture. Neurotoxicology. 1999;20:41–48. [PubMed]
116. Kwon OS, Schmued LC, Slikker W., Jr Fumonisin B1 in developing rats alter brain sphinganine levels and myelination. Neurotoxicolog. 1997;18:571–580. [PubMed]
117. Tsunoda M, Dugyala RR, Sharma RP. 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. [PubMed]
118. Porter JK, Voss KA, Chamberlain WJ, Bacon CW, Norred WP. Neurotransmitters in rats fed fumonisin B1. Proc. Soc. Exp. Biol. Med. 1993;202:360–364. [PubMed]
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. [PubMed]
120. Osuchowski MF, Edwards GL, Sharma RP. 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. [PubMed]
121. Bouhet S, Hourcade E, Loiseau N, Fikry A, Martinez S, Roselli M, Galtier P, Mengheri E, Oswald IP. The mycotoxin fumonisin B1 alters the proliferation and the barrier function of porcine intestinal epithelial cells. Toxicol. Sci. 2004;77:165–171. [PubMed]
122. Ramasamy S, Wang E, Hennig B, Merrill AH., Jr Fumonisin B1 alters sphingolipid metabolism and disrupt the barrier function of endothelial cells in culture. Toxicol. Appl. Pharmacol. 1995;133:343–348. [PubMed]
123. Osuchowski MF, He Q, Sharma RP. 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. [PubMed]
125. Buccoliero R, Futerman AH. The roles of ceramide and complex sphingolipids in neuronal cell function. Pharmacol. Res. 2003;47:409–419. [PubMed]
126. Pettus BJ, Chalfant CE, Hannun YA. Ceramide in apoptosis: An overview and current perspectives. Biochem. Biophys. Acta. 2002;1585:114–125. [PubMed]
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. [PubMed]
128. Mobio TA, Anane R, Baudrimont I, Carratū MR, Shier TW, Dano SD, Ueno Y, Creppy EE. Epigenetic properties of fumonisin B1: cell cycle arrest and DNA base modification in C6 glioma cells. Toxicol. Appl. Pharmacol. 2000;164:91–96. [PubMed]
129. Mobio TA, Baudrimont I, Sanni A, Shier TW, Saboureau D, Dano SD, Ueno Y, Steyn PS, Creppy EE. Prevention by vitamin E of DNA fragmentation and apoptosis induced by fumonisin B1 in C6 glioma cells. Arch. Toxicol. 2000;74:112–119. [PubMed]
130. Mobio TA, Tavan E, Baudrimont I, Anane R, Carratū MR, Sanni A, Gbeassor MF, Shier TW, Narbonne J-F, Creppy EE. 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. [PubMed]
131. Galvano F, Campisi A, Russo A, Galvano G, Palumbo M, Renis M, Barcellona ML, Perez-Polo JR, Vanella A. DNA damage in astrocytes exposed to fumonisin B1. Neurochem. Res. 2002;27:345–351. [PubMed]
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. [PubMed]
133. Ellerby LM, Ellerby HM, Park SM, Holleran AL, Murphy AN, Fiskum G, Kane DJ, Testa MP, Kayalar C, Bredesen DE. Shift of cellular oxidation-reduction potential in neural cells expressing Bcl-2. J Neurochem. 1996;67:1259–1267. [PubMed]
134. Kane DJ, Sarafian TA, Anton R, Hahn H, Butler GE, Selverstone VJ, Ord T, Bredesen DE. Bcl-2 inhibition of neural death: decreased generation of reactive oxygen species. Science. 1993;262:1274–1277. [PubMed]
135. Tjalkens RB, Ewing MM, Philbert MA. 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. [PubMed]
136. Reed JC, 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. [PubMed]
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. [PubMed]
138. Schmelz EM, Dombrink-Kurzman MA, Roberts PC, Kozutsumi Y, Kawasaki T, Merrill AH., 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. [PubMed]
139. Tolleson WH, Dooley KL, Sheldon WG, Thurman JD, Bucci TJ, Howard PC. The Mycotoxin Fumonisin Induces Apoptosis in Cultured Human Cells and in Livers and Kidneys of Rats. In: Jackson LS, DeVries JW, Bullerman LB, editors. Advances in Experimental and Medical Biology Fumonisins in Food. Plenum Press; New York, NY, USA: 1996. pp. 237–250. [PubMed]
140. Tolleson WH, Melchior WB, Morris SM, McGarrity LJ, Domon OE, Muskhelishvili L, James SJ, Howard PC. Apoptotic and anti-proliferaive effects of fuminisin B1 in human keratinocytes, fibroblasts, esophageal epithelial cells and hepatoma cells. Carcinogenesis. 1996;17:239–249. [PubMed]
141. Higuchi Y. Chromosomal DNA fragmentation in apoptosis and necrosis induced by oxidative stress. Biochem. Pharmacol. 2003;66:1527–1535. [PubMed]
142. Slater AFG, Nobel CSI, van den Dobbelsteen DJ, Orrenius S. Signaling mechanisms and oxidative stress in apoptosis. Toxicol Lett. 1995;82/83:149–153. [PubMed]
143. Galtier P. Pharmacokinetics of ochratoxin A in animals. IARC Sci Publ. 1991:187–200. [PubMed]
144. Pfohl-Leszkowicz A, Manderville RA. Ochratoxin A: An overview on toxicity and carcinogenicity in animals and humans. Mol. Nutr. Food Res. 2007;51:61–99. [PubMed]
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. [PubMed]
146. Mally A, Hard GC, 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. [PubMed]
147. Krogh P. Role of ochratoxin in disease causation. Food Chem. Toxicol. 1992;30:213–224. [PubMed]
148. Kane A, Creppy EE, Roschenthaler R, Dirheimer G. Changes in urinary and renal tubular enzymes caused by subchronic administration of ochratoxin A in rats. Toxicology. 1986;42:233–243. [PubMed]
149. Petkova-Bocharova T, Chernozemsky IN, 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. [PubMed]
150. Lea T, Steinen K, Stormer FC. Mechanism of ochratoxin A-induced immunosuppression. Mycopathologia. 1989;107:153–159. [PubMed]
151. Stromer FC, Lea T. Effects of ochratoxin A upon early and late events in human T-cell proliferation. Toxicology. 1995;95:45–50. [PubMed]
152. Arora RG, Frolen H, Fellner-Feldegg H. Inhibition of ochratoxin A teratogenesis by zearalenone and diethylstilbesterol. Food Chem. Toxicol. 1983;21:779–783. [PubMed]
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. [PubMed]
154. Pfohl-Leszkowicz A, Chakor K, Creppy EE, Dirheimer G. DNA adduct formation in mice treated with ochratoxin A. IARC Sci Publ. 1991:245–253. [PubMed]
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. [PubMed]
156. Kuiper-Goodman T, Hilts C, Billiard SM, Kiparissis Y, Richard ID, 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. [PMC free article] [PubMed]
157. Creppy EE, Chakor K, Fisher MJ, Dirheimer G. The mycotoxin ochratoxin A is a substrate for phenylalanine hydroxylase in isolated rat hepatocytes and in vivo. Arch. Toxicol. 1990;64:279–284. [PubMed]
158. Creppy EE, 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. [PubMed]
159. Dirheimer G, Creppy EE. Mechanism of action of ochratoxin A. IARC Sci Publ. 1991:171–186. [PubMed]
160. Gautier JC, Holzhaeuser D, Markovic J, Gremaud E, Schilter B, Turesky RJ. Oxidative damage and stress response from ochratoxin exposure in rats. Free Radic. Biol. Med. 2001;30:1089–1098. [PubMed]
161. Bryan NS, Rassaf T, Maloney RE, Rodriguez CM, Saijo F, Rodriguez JR, 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. [PubMed]
162. Thomas JA, Mallis RJ. Aging and oxidation of reactive protein sulfhydryls. Exp. Gerontol. 2001;36:1519–1526. [PubMed]
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;89:120–134. [PubMed]
164. Aleo MD, Wyatt RD, Schnellmann RG. 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. [PubMed]
165. Wei YH, Lu CY, Lin TN, Wei RD. Effect of ochratoxin A on rat liver mitochondrial respiration and oxidative phosphorylation. Toxicology. 1985;36:119–130. [PubMed]
166. Belmadani A, Tramu G, Betbeder AM, Creppy EE. Subchronic effects of ochratoxin A on young adult rat brain and partial prevention by aspartate, a sweetener. Hum. Exp. Toxicol. 1998;17:380–386. [PubMed]
167. Hayes AW, Cain JA, Moore BG. Effects of aflatoxin B1, ochratoxin A and rubratoxin B on infant rats. Food Cosmet. Toxicol. 1977;15:23–27. [PubMed]
168. Hayes AW, Hood RD, Lee HL. Teratogenic effects of ochratoxin A in mice. Teratology. 1974;9:93–97. [PubMed]
169. Wangikar PB, Dwivedi P, Sharma AK, 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. [PubMed]
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. [PubMed]
171. Belmadani A, Tramu G, Betbeder AM, Steyn PS, Creppy EE. Regional selectivity to ochratoxin A, distribution and cytotoxicity in rat brain. Arch. Toxicol. 1998;72:656–662. [PubMed]
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. [PubMed]
173. Sanchez-Ramos J, Overvik E, Ames BN. 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;83:398–405. [PubMed]
175. Creppy EE, Kern D, Steyn PS, 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. [PubMed]
176. Palmer TD, Takahashi J, Gage FH. The adult rat hippocampus contains primordial neural stem cells. Mol Cell Neurosci. 1997;8:389–404. [PubMed]
177. Song HJ, Stevens CF, Gage FH. Neuronal stem cells from adult hippocampus develop essential properties of functional CNS neurons. Nat. Neurosci. 2002;5:438–445. [PubMed]
178. Chen H, Tung YC, Li B, Iqbal K, Grundke-Iqbal I. Trophic factors counteract elevated FGF-2-induced inhibition of adult neurogenesis. Neurobiol. Aging. 2006;28:1148–1162. [PubMed]
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. [PubMed]
180. Lagace DC, Yee JK, Bolanos CA, Eisch AJ. Juvenile administration of methylphenidate attenuates adult hippocampal neurogenesis. Biol. Psychiatry. 2006;60:1121–1130. [PubMed]
181. Rossi C, Angelucci A, Costantin L, Braschi C, Mazzantini M, Babbini F, Fabbri ME, 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. [PubMed]
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. [PubMed]
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. [PubMed]
184. Alexander P. The role of DNA lesions in processes leading to aging in mice. Sym. Soc. Exp. Biol. 1967;21:29–50. [PubMed]
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. [PubMed]
186. Crago BR, Gray MR, Nelson LA, Davis M, Arnold L, Thrasher JD. Psychological, neuropsychological, and electrocortical effects of mixed mold exposure. Arch. Environ. Health. 2003;58:452–563. [PubMed]
187. Gordon WA, Cantor JB. The diagnosis of cognitive impairment associated with exposure to mold. Adv. Appl. Microbiol. 2004;55:361–374. [PubMed]
188. Rea WJ, Didriksen N, Simon TR, Pan Y, Fenyves EJ, Griffiths B. Effects of toxic exposure to molds and mycotoxins in building-related illnesses. Arch. Environ. Health. 2003;58:399–405. [PubMed]
189. Yoon S, Cong W-T, Bang Y, Lee SN, Yoon CS, Kwack SJ, Kang TS, Lee KY, Choi J-K, Choi HJ. Proteome response to ochratoxin A-induced apoptotic cell death in mouse hippocampal HT22 cells. Neurotoxicology. 2009;30:666–676. [PubMed]
190. Sato A, Miyazaki E, Satake A, Hiramoto A, Hiraoka O, Miyake T, Kim HS, Wataya Y. Proteome and transcriptome analysis of cell death induced by 5-fluoro-2′-deoxyuridine. Nucleic Acids Symp. Ser (Oxf) 2007;51:433–434. [PubMed]
191. Siddiq A, Ayoub IA, Chavez JC, Aminova L, Shah S, LaManna JC, Patton SM, Connor JR, Cherny RA, Volitakis I, et al. Hypoxia-indicible factor prolyl 4-hydroxylase inhibition. A target for neurprotection in the central nervous system. J. Biol. Chem. 2005;280:41732–41743. [PMC free article] [PubMed]
192. Lei T, He Q, Cai Z, Zhou Y, Wang Y, Si L, Cai Z, Chiu JF. Proteomic analysis of chromium cytotoxicity in cultured rat lung epithelial cells. Proteomics. 2008;8:2420–2429. [PubMed]
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. [PubMed]
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. [PMC free article] [PubMed]
195. Zurich MG, 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. [PubMed]
196. Hong JT, Lee MK, Park KS, Jung KM, Lee RD, Jung HK, Park KL, Yang KJ, Chung YS. 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. [PubMed]

Articles from International Journal of Molecular Sciences are provided here courtesy of Multidisciplinary Digital Publishing Institute (MDPI)