Transcriptomic and phenotypic analysis of the effects of T-2 toxin on Saccharomyces cerevisiae: evidence of mitochondrial involvement

Authors


  • Editor: Lubomir Tomaska

Correspondence: Ivor H. Evans, School of Science, University of Greenwich at Medway, Central Avenue, Chatham Maritime, Kent, ME4 4TB, UK. Tel.: +44 208 331 8000; fax: +44 208 331 9805; e-mail: i.h.evans@gre.ac.uk

Abstract

At 5 μg mL−1, T-2 toxin significantly upregulated the transcription of 281 genes and downregulated 86. Strongly upregulated genes included those involved in redox activity, mitochondrial functions, the response to oxidative stress, and cytoplasmic rRNA transcription and processing. Highly repressed genes have roles in mitochondrial biogenesis, and the expression and stability of cytoplasmic rRNAs. T-2 toxin inhibition of growth was greater in a medium requiring respiration, and was antagonized by antioxidants. T-2 toxin treatment induced reactive oxygen species, caused nucleolytic damage to DNA, probably mitochondrial, and externalization of phosphatidylserine. Deletion mutations causing respiratory deficiency substantially increased toxin tolerance, and deletion of some TOR (target of rapamycin) pathway genes altered T-2 toxin sensitivity. Deletion of FMS1, which plays an indirect role in cytoplasmic protein synthesis, markedly increased toxin tolerance. Overall, the findings suggest that T-2 toxin targets mitochondria, generating oxy-radicals and repressing mitochondrial biogenesis genes, thus inducing oxidative stress and redox enzyme genes, and triggering changes associated with apoptosis. The large transcriptional changes in genes needed for rRNA transcription and expression and the effects of deletion of FMS1 are also consistent with T-2 toxin damage to the cytoplasmic translational mechanism, although it is unclear how this relates to the mitochondrial effects.

Introduction

T-2 toxin is a trichothecene mycotoxin produced by phytopathogenic fungi of the Fusarium genus, which can infect crop plants such as wheat, barley, and rice in the field or during storage (Nelson et al., 1994). Trichothecenes are a large group of toxic sesquiterpenoid secondary fungal metabolites, classified into four types, A–D, but all containing a double bond between C9 and 10 and an epoxide at C12 and 13 (Desjardins et al., 1993). T-2 toxin (see Fig. 1), a type A trichothecene, widely detected as a contaminant of human foodstuffs and animal feeds, has been implicated in serious poisoning episodes affecting humans and farm animals, and is considered a significant toxicological hazard (Mirocha, 1983; Sudakin, 2003). The effects of T-2 toxin administration have been studied in a number of animal species, and a wide range of pathophysiological effects of varying severity have been recorded, affecting many organ systems, and rapidly proliferating tissues in particular (Beasley, 1989).

Figure 1.

 T-2 toxin, a type A trichothecene containing a double bond between C9 and 10 and an epoxide at C12 and 13.

At the cellular and subcellular levels, trichothecenes such as T-2 toxin are known to have multiple effects on eukaryotes, including the inhibition of DNA, RNA and protein synthesis, inhibition of cell division, perturbation of plasma membranes, and inhibition of mitochondrial function (Rocha et al., 2005). Earlier in vivo and in vitro work, with both mammalian and yeast cells and subcellular fractions, highlighted protein synthesis as a prime target: trichothecenes were found to inhibit peptidyl transferase and inhibition of DNA and RNA synthesis was presumed to be a secondary effect (Beasley, 1989; Ueno, 1991). Consistent with this role in inhibiting protein synthesis, direct evidence of the binding of trichothecenes to eukaryotic ribosomes was reported (Barbacid & Vasquez, 1974; Wei et al., 1974; McLaughlin et al., 1977). Evidence that trichothecenes differ in their modes of inhibiting protein synthesis was provided by the finding that some, including trichodermin, stabilize polyribosome profiles, suggestive of inhibition of elongation or termination, while others, including T-2 toxin, cause ‘run-off’ and disaggregation of polysomes, indicative of the inhibition of initiation (Schindler et al., 1974; Cannon et al., 1976; Carter & Cannon, 1977). However, Smith et al. (1975) found that T-2 toxin inhibits the formation of the first peptide bond, rather than the initiation complex. The finding that the protein synthesis inhibitor anisomycin caused not only translational arrest but also activated protein kinases involved in stress responses indicated a possible link between the translation mechanism and cell regulatory systems (Schindler et al., 1974; Cannon et al., 1976; Cano et al., 1994). An analysis of the effects on the stress-activated protein kinase (SAPK)/c-Jun NH2-terminal kinase 1 (JNK1) of a range of structurally varied protein synthesis inhibitors, including T-2 toxin (Iordanov et al., 1997), substantiated the connection between inhibition of translation and protein kinase activation, but showed that the relationship was not straightforward: while anisomycin, emetine, cycloheximide, and T-2 toxin all inhibited protein synthesis (>95%), anisomycin activated SAPK/JNK1 over 20-fold, emetine produced no activation, and cycloheximide and T-2 toxin were only weak activators. The activation of kinases involved in cellular stress responses, such as JNK1, was termed a ribotoxic stress response, and was also shown to be triggered by agents damaging 28S rRNA (Iordanov et al., 1997). Anisomycin was found to induce rapid apoptosis in human lymphoid cells (Polverino & Patterson, 1997), and a study using Jurkat T (also human lymphoid) cells and a wide range of trichothecenes showed that many trichothecenes induce apoptosis (Shifrin & Anderson, 1999); however, there was no completely clear correlation between inhibition of protein synthesis, activation of JNK, and activation of caspase-3 (an apoptotic marker), and in the case of T-2, there was a very strong inhibition of protein synthesis, negligible activation of JNK, and only a moderate activation of caspase-3. Nevertheless, Doi and colleagues were able to adduce strong evidence that T-2 toxin can indeed trigger apoptosis in vivo, in the mouse, demonstrating changes in the expression of a number of apoptosis-related genes (Shinozuka et al., 1999; Doi et al., 2006, 2008).

In order to assess the response of a large sample of genes to T-2 intoxication, the Doi group used DNA microarray technology to estimate changes in the transcription levels in various rat tissues exposed in vivo to T-2 toxin (Sehata et al., 2004a, b, 2005). One general finding from these large data sets was an increased expression of oxidative stress- and apoptosis-related genes, as well as mitogen-activated protein kinase pathway genes. Microarray technology was also used in recent studies of the impact of T-2 toxin on Arabidopsis thaliana seedlings. Trichothecenes were found to be phytotoxic, and different trichothecene species could differ markedly in their effects; it was noteworthy that while T-2 toxin treatment caused dwarfism and other aberrations, cycloheximide did not induce these changes (Masuda et al., 2007). In the same study, it was found that out of 8100 genes, T-2 toxin upregulated 35, including AtNFXL1, a gene encoding a member of an evolutionarily conserved family of putative transcription factors including the yeast homologue FAP1 protein, with a role in rapamycin toxicity (Kunz et al., 2000). A subsequent investigation, using an atnfxl1 mutant and further transcriptomics, indicated that AtNFXL1 plays an important role in the response to T-2 toxin, as well as the general defence response in Arabidopsis (Asano et al., 2008).

Yeast (Saccharomyces cerevisiae) can be used as a model system to explore the responses of a whole eukaryotic genome to environmental stresses, including toxic chemicals (Gasch et al., 2000). Toxicants looked at in this way include the pesticide lindane (Parveen et al., 2003), the mycotoxin aflatoxin B1 (Keller-Seitz et al., 2004), and antimicrobial peptides (Morton et al., 2007). Yeast also allows the mechanisms of action of toxicants to be examined through the use of a variety of gene deletion libraries (Holland et al., 2007; Morton et al., 2007; Xia et al., 2007). We have previously reported the use of yeasts, including engineered strains of S. cerevisiae, to detect mycotoxins, such as T-2 toxin (Engler et al., 1999; Li et al., 2006, 2009). In this paper, we report the use of transcriptomics, deletion mutants, and other experimental approaches, in an investigation of the interaction between T-2 toxin and S. cerevisiae [a preliminary report of our transcriptomic data was given by Josséet al. (2005)].

Materials and methods

Saccharomyces cerevisiae strains and media

The strain used for transcriptomic analysis was YB110 (MATa/MATα; ura3-52/ura3-52; his3200/his3200; ade2 101/ade2-101; trp11/trp11; gal3-/gal3-; lys2-801/lys2-(leaky); LEU2/leu2, 3-112; gal1/gal1his35′; trp1/trp1his33′∷HOcs; Fasullo et al., 1998). All other strains were either the haploid BY4741 (MATa; his3Δ1; leu2Δ0; met15Δ0; ura3Δ0), or BY4743, the homozygous diploid derived from it, or deletion mutants of BY4741 or BY4743 [created using the KanMX cassette (http://web.uni-frankfurt.de/fb15/mikro//euroscarf/)], all from the Euroscarf collection. Strains were grown in either YPD (1% yeast extract, 1% peptone, 2% glucose) or YPG (1% yeast extract, 1% peptone, 2% glycerol) at 25 or 30 °C, unless otherwise specified.

Online monitoring of yeast growth in microplates

Yeast cells were grown to an A600 nm of 0.4–0.7 (early log stage) in YPD or YPG medium and then diluted to an A600 nm of 0.002. Aliquots (25 μL) of the diluted cells were added to the wells of a 384-well microplate (Grenier, UK), followed by 25 μL of a T-2 toxin/methanol/YPD or YPG mixture, so that on completion of additions, each well contained early log-stage cells with a final A600 nm of 0.001, 0.2% methanol, an appropriate concentration of T-2 toxin, and YPD or YPG medium. The 384-well plate was then covered by Breathe-Easy sealing membrane (Sigma-Aldrich, UK) and placed in a M5 microplate reader (Molecular Devices) for an online kinetic growth assay under the following conditions : temperature 25 °C, wavelength 600 nm, reading interval 30 min, and monitoring duration 48 h (up to 96 h for slow-growing strains).

Antioxidant effects on growth

Glutathione (GSH) and N-acetyl-l-cysteine (NAC) were dissolved in sterile YPD to a final concentration of 50 mM, and the two modified media were resterilized by ultrafiltration (0.2 μm). The YPD–GSH and YPD–NAC solutions were then used to prepare growth test solutions (T-2 toxin/methanol/YPD or YPD–GSH or YPD–NAC), so that on setting up microplate growth assays, as described above, all wells contained early log-stage cells with a final A600 nm of 0.001% and 0.2% methanol, lacking or containing T-2 toxin (final concentration 200 μM), GSH (final concentration 25 mM), and NAC (final concentration 25 mM).

T-2 toxin treatment for transcriptional analysis

A fresh yeast culture [strain YB110 for microarray analysis or strain BY4741 for quantitative real-time PCR (qRT-PCR)] was inoculated into 50 mL YPD in six different shaker flasks and incubated at 30 °C until the cultures reached an A600 nm of 0.5–0.6. Three flasks (solvent controls) were treated with 2 mL 75% methanol (final concentration 3%) and the other three were treated with 2 mL of T-2 toxin at 125 μg mL−1 in 75% methanol (final concentrations 5 μg mL−1 toxin and 3% methanol), and left for 2 h at 30 °C, with continued shaking. Then the cells were harvested from each flask by centrifugation at room temperature for 5 min, the supernatants were discarded, and the cell pellets were frozen immediately on dry ice for microarray analysis, or treated as described below for qRT-PCR.

Microarray hybridization and analysis

Each cell pellet – three controls and three toxin treated – was independently processed and analysed. RNA preparation, hybridization, and data validation were outsourced (Transcriptome Service Facility, Manchester). Total RNA isolation was based on the TRIzol protocol and hybridizations were performed using the Affymetrix GeneChip system (Hayes et al., 2002; Wishart et al., 2005). r/bioconductor packages affy, gcrma, and limma were used for all gene and probe computations.

qRT-PCR

Cell pellet preparation was as described above. The cells were lysed by spheroplasting in 50 U of zymolyase (Sigma-Aldrich), followed by the addition of fresh lysis buffer RLT (Qiagen). The lysates were processed according to the RNeasy Mini Kit protocol (Qiagen). The RNA yield from each replicate was calculated based on the A260 nm and the total elution volume and the quality of RNA was determined by running 1 μg of total RNA on a formaldehyde–agarose gel (the intensity of the 28S rRNA gene band was about twice that of the 18S rRNA gene band).

Fifty nanograms of total RNA were used for transcriptional analysis using the iScript One-Step RT PCR kit with SYBR Green (Bio-Rad) and gene-specific primers (see Table 1).

Table 1.   qRT-PCR primers and results for five genes upregulated in the transcriptomic analysis
Systematic namePrimer sequence (5′–3′)G/HKG ratioStandard name and putative function
  1. G/HGK ratio: change in expression of gene of interest relative to change in expression of house-keeping gene (HKG) PGK1.

YNL335W_FATGGGACTTGGATCAGGAAG1.8DDI3, DNA damage response
YNL335W_RTTTCCACCTGTCGCATTAAA
YHR015W_FTCCGCACACAAGAGTTATCAG1.5MIP6, mRNA export
YHR015W_RAAATGCTGGATGAGGTTTCA
YJR078W_FAAATGGGAAGAAATCGTTGC5.0BNA2, NAD biosynthesis
YJR078W_RTCGTCTCATCCAAGTCCAAG
YMR020W_FGATGGGAGAACGAGGTTTGT3.7FMS1, amino oxidase activity, translation
YMR020W_RTCTGCGAATTTGCTCATTTC
YMR118C_FAGCATCTGTATTCGTGCCAA2.6Unknown
YMR118C_FTCCTTGTTGCTTGACCAAAG

Detection of reactive oxygen species (ROS) via microscopy

A yeast culture was grown to A600 nm 0.3 in 10 mL YPD medium, and then divided into two – A and B. Dimethyl sulphoxide (DMSO) was added to A to a final concentration of 2% v/v, and T-2 toxin/DMSO was added to B, yielding final concentrations of 200 μM for T-2 and 2% v/v for DMSO. After 16 h of incubation at 30 °C, with shaking (200 r.p.m.), volumes yielding c. 1.0 A600 nm for culture A (usually 250 μL), and for B (2.5–3.0 mL), were transferred into new tubes for centrifugation. The cells were collected and washed three times in HEPES buffer (2% glucose, 10 mM Na-HEPES, pH 7.2), and suspended in 50 μL of the same HEPES buffer.

Intracellular ROS were detected by dihydrorhodamine 123 (DHR123, Invitrogen) staining, combined with the counter-stain calcofluor white M2R (Invitrogen).

One microlitre of 100 μM DHR123 in ethanol and 2 μL of 500 μM of calcofluor white M2R in DMSO were added to the above cell suspensions and incubated at 30 °C for 30 min. The cells were then applied to microscopic slides for ROS detection using a fluorescence microscope (Nikon Eclipse 90i).

Quantitative analysis of ROS production using fluorescence-activated cell sorting (FACS)

BY4743 was inoculated into YPD broth and grown for 24 h before being diluted 1/1000 in 5 mL YPD. Cells were then grown further overnight until they reached an A600 nm of 0.2–0.3 on the day of the experiment. T-2 toxin was added at 50 or 200 μM (zero time, T=0) and the presence of free intracellular radicals was assessed using DHR123 or dihydroethidium (DHE) at 0, 2, and 4 h after the addition of the T-2 toxin. DHR123 was added 2 h before the experiment was initiated at 5 μg mL−1 of cell culture from a 5 mg mL−1 stock solution in ethanol. DHE was added 10 min before measurement by resuspending cells in 250 μL of 2.5 μg mL−1 DHE in phosphate-buffered saline (PBS). Detection of necrotic cells was carried out by the addition of propidium iodide (PI) at 4 μg mL−1.

Fluorescence data from 10 000 cells per sample were collected using a Becton-Dickinson FACScalibur Flow Cytometer and analysed using cell quest pro software. Excitation was achieved using a 488 nm laser and emission data for DHR123 were collected using the FL1 detector (515–545 nm), while DHE and PI emissions were detected using FL2 (564–606 nm). For dual detection of DHR123 and PI, 50% FL2/FL1 compensation was used to eliminate bleed-through.

Fluorescein isothiocyanate (FITC) Annexin V- and PI-based apoptotic and dead cell detection via microscopy

FITC Annexin V- and PI-based apoptotic and dead cell detection was performed using an FITC Annexin V kit (Invitrogen).

Yeast cell growth and T-2 toxin treatment were the same as described in ROS detection. The cells were washed in cold PBS (pH 7.4) and cell pellets were resuspended in 200 μL annexin-binding buffer. Five microlitres of annexin V conjugate and 1 μL of 100 μg mL−1 of PI were added and the cells were incubated at room temperature for 15 min. The cells were washed with annexin-binding buffer and then examined using fluorescence microscopy (Nikon Eclipse 90i).

Terminal deoxynucleotide transferase-mediated dUTP-biotin nick end labelling (TUNEL) analysis

TUNEL analysis was performed using the Roche In Situ Cell Death Detection Kit, TMR Red, for the detection and quantification of apoptosis at the single cell level, based on labelling of DNA strand breaks. Yeast cell growth and T-2 toxin treatment were the same as described for ROS detection, but cells were collected, washed, and resuspended in 300 μL PBS instead of HEPES buffer. Three hundred microlitres of fixation solution (freshly prepared 4% paraformaldehyde in PBS) were then added to the above cell suspension (final concentration 2% paraformaldehyde), followed by 60 min of incubation at room temperature. Cells were collected and washed in PBS buffer, then resuspended in 300 μL of 1 M sorbitol in PBS buffer containing 3 U of lyticase, and incubated at 30 °C with shaking for 2 h. After three washes in PBS buffer, the cells were resuspended in 300 μL of freshly prepared permeabilization solution (0.1% Triton X-100 in 0.1% sodium citrate) and left on ice for 2 min. The cells were washed twice with PBS buffer and resuspended in 50 μL of TUNEL reaction mixture (terminal deoxynucleotide transferase, with the appropriate nucleotides, in a reaction buffer) and incubated for 2 h at 30 °C in the dark. After two final washes in PBS, the cells were analysed using fluorescence microscopy (Nikon Eclipse 90i).

Results

Effect of T-2 toxin on growth

Figure 2 shows the growth of yeast strain YB110 in YED medium as a function of T-2 toxin concentration, in the presence of 3% v/v methanol, as a toxin solvent. At 5 μg mL−1, T-2 toxin showed significant inhibition of growth compared with the solvent control, but the growth and therefore the potential transcript yields were not drastically reduced; hence, 5 μg mL−1 was chosen as the test concentration for our transcriptomic analysis.

Figure 2.

 Effect of T-2 toxin on the growth of Saccharomyces cerevisiae (strain YB110). YB110 was grown in YPD medium, in the presence of varying amounts of T-2 toxin dissolved in methanol (final concentration 3% v/v), the control being YPD medium only.

Overview of DNA microarray results

Initially, the most significantly differentially expressed genes were identified on the basis of a P value of<0.05 for the moderated t-statistic or a B value of 4.6 or higher (B%≥99%), and named genes in this set are used in the account below. This criterion yields 281 significantly upregulated genes and 86 significantly downregulated genes. The transcriptomic response for a larger gene set (P<0.1) used in our GO (gene ontology) analyses is illustrated in Fig. 3 (349 upregulated and 136 downregulated). The substantial excess of induced over-repressed genes evident here has been seen in other analyses of mycotoxin-stressed yeast cells: 183 vs. 66 for aflatoxin B1 (Keller-Seitz et al., 2004) and 363 vs. 73 for cells treated with citrinin (Iwahashi et al., 2007). This disparity is more marked for those genes with most significantly changed transcript levels. The top 100 genes (B>7.40) contain only 11 repressed genes. In the 367 gene set, the greatest upregulation is 45.8-fold and the greatest downregulation is 7.9-fold; the smallest change is 2.1-fold (downregulation), and 272 genes show at least a threefold change. In similar analyses, the minimal change taken as significant has been threefold (Keller-Seitz et al., 2004), twofold (Iwahashi et al., 2007), and 1.5-fold (Morton et al., 2007).

Figure 3.

 Response of the yeast transcriptome to T-2 toxin. Black graph: P<0.05, out of a total of 367 genes, 281 genes were upregulated 2.3–45.8-fold and 86 genes were downregulated 2.1–7.9-fold. Grey graph: P<0.1, out of a total of 484 genes, 349 genes were upregulated 2.1–45.8-fold and 136 genes were downregulated 2.0–7.9-fold.

Genes named in the text below are up- or downregulated at least threefold, the actual factor being given to the nearest integer, in parentheses. The details of gene and protein function cited below are taken from the Saccharomyces Genome Database.

Upregulated genes

GO profiling by Molecular Function (MF) seemed the most illuminating GO analysis of the upregulated genes (349 at P<0.1, 281 annotated), and Table 2 shows the top 10 GO MF categories, on the basis of probability. Oxidoreductase activity is strongly over-represented and contains BNA2– the most highly upregulated (× 40) identified gene – encoding putative tryptophan 2,3-dioxygenase needed for the de novo synthesis of NAD from kynurenine, and BNA4 (× 9), specifying kynurenine 3-monooxygenase, also needed for NAD synthesis. ADE2, another purine nucleotide biosynthesis gene, is also strongly upregulated (× 12). FDH1 (× 27) specifies a formate dehydrogenase involved in NADH regeneration, and GND2 (× 6) encoding phosphogluconate dehydrogenase plays a role in NADPH regeneration through the pentose phosphate pathway. Other oxidoreductases include BDH1 (× 3), GCY1 (× 3), AAD4 (× 6), AAD14 (× 8), and AAD16 (× 11), most of which are known to respond to oxidative stress. ADH2 (× 6) also encodes a dehydrogenase with a possible stress-protection role, and FRE2 (× 11), encoding a ferric reductase, is an important provider of surface reductase activity. FMS1 (× 9) encodes polyamine oxidase, an FAD-coupled enzyme producing spermidine needed for the activation of the translation factor eIF-5A.

Table 2.   The top 10 GO MF categories of upregulated genes based on their probability
GO term
Molecular function
Frequency (%)ProbabilityGenes (fold upregulated)
Upregulated (281)Genome (7146)
  • *

    These gene products have carboxy-lyase activity.

  • Among 349 upregulated genes (P<0.1), 281 genes were annotated and used for GO analysis.

Oxidoreductase activity8.53.30.000032BNA2 (40.61)FDH1 (26.74)
FRE2 (11.23)AAD16 (10.74)
BNA4 (9.19)FMS1 (9.02)
AAD14 (8.07)GND2 (6.03)
AAD4 (5.69)ADH2 (5.76)
GCY1 (5.31)YPL113C (3.86)
TSA2 (3.85)MCR1 (3.85)
GRX4 (3.55)GCV1 (3.10)
MXR1 (3.07)YJR096W (2.99)
JLP1 (2.90)BDH1 (2.69)
YJL045W (2.62)HMX1 (2.45)
MTD1 (2.44)TSC10 (2.25)
Pyruvate decarboxylase activity1.00.040.00054PDC6 (8.32)PDC5 (3.35)
ARO10 (3.23) 
Aryl-alcohol dehydrogenase activity1.00.10.00387AAD16 (10.74)AAD14 (8.07)
AAD4 (5.69) 
Oxidoreductase activity, acting on paired donors, with incorporation or reduction of molecular oxygen1.00.10.00387BNA4 (9.19)JLP1 (2.90)
HMX1 (2.45) 
Electrochemical potential-driven transporter activity2.10.50.0039PUT4 (13.16)PIC2 (5.11)
VBA2 (3.79)FCY21 (2.601)
AAC1 (2.41)CTP1 (2.38)
Oxidoreductase activity, acting on the CH-OH group of donors, NAD or NADP as acceptor2.80.90.0054AAD16 (10.74)AAD14 (8.07)
GND2 (6.03)ADH2 (5.76)
AAD4 (5.69)GCY1 (5.31)
YJR096W (2.99)BDH1 (2.96)
Multidrug transporter activity1.00.10.01169VBA2 (3.79)FLR1 (2.87)
QDR3 (2.79) 
Lyase activity2.81.10.01747ADE2 (12.16)*PDC6 (8.32)*
STR3 (5.33)GAD1 (4.09)*
PDC5 (3.35)*ARO10 (3.23)*
PHA2 (2.88)PAD1 (2.76)*
Protein methyltransferase activity1.00.20.02099SWD3 (3.89)STE14 (3.35)
SPP1 (3.12) 
Carboxylic acid transporter activity1.70.50.02679PUT4 (13.16)DAL5 (5.52)
VBA2 (3.79)YHC3 (3.46)
MMP1 (3.19) 

Genes involved in mitochondrial biogenesis, function, and maintenance occur in several of the GO MF categories – the mitochondrial role in redox biochemistry is evident. Among these are MCR1 (× 5 – cytochrome b5 reductase), YLR164W (× 5 – an inner-membrane protein), YJR079W (× 20), and YER187W (× 37 – induced in respiratory-deficient cells). SUE1 (× 5) is a mitochondrial protein required for the degradation of unstable forms of cytochrome c, and THI4 (× 7 – required in thiamine synthesis), PET18 (× 8), and SML1 (× 7) all play roles in mitochondrial genome stability, while three other thiamine biosynthesis genes are also substantially upregulated –THI11 (× 8), THI21 (× 6), and THI74 (× 3). In the case of SML1, an inhibitor of ribonucleotide reductase, upregulation could reduce dNTP levels, and thus increase mitochondrial genome instability – although T-2 toxin appears not to induce significant petite formation (results not shown). The response to stress, including oxidative stress, can involve redox enzymes such as MXR1 (× 3), TSA2 (× 4 – a thioredoxin peroxidase) and GRX4 (× 4 – a superoxide-radical-responsive GSH-dependent oxidoreductase). Interestingly, GTT2 (× 7) specifies a GSH-S-transferase involved in GSH–glutaredoxin redox reactions, and a mitochondrial location has been inferred from high-throughput studies (Reinders et al., 2006). Also, it can be noted that the HOG-regulated GRE1 (× 12) gene encodes a hydrophilin induced by varied, including oxidative, stresses.

Not unexpectedly, a number of genes encoding drug efflux pumps are upregulated, including FLR1 (× 3) and QDR3 (× 3) of the major facilitator superfamily.

Unexpectedly, four genes normally only expressed anaerobically are strongly upregulated –TIR4 (× 8), DAN2 (× 5) and PAU5 (× 5), and PAU2 (× 5), as are meiosis- and sporulation-specific genes including SGA1 (× 7), ISC10 (× 6), SLZ1 (× 8), SPO21 (× 4), DMC1 (× 6), GIP1 (× 5), TEP1 (× 6), and WTM2 (× 5) – a transcriptional modulator involved in the regulation of meiosis, and these genes are not highlighted by GO analyses; as YB110 is a diploid, transient induction of meiosis is a possibility. A clear category highlighted by GO Cellular Component profiling (not shown here) is the spliceosome complex, including CWC23 (× 3), PRP42 (× 4), SMD1 (× 4), PRP9 (× 3), CUS1 (× 6), LEA1 (× 3), MUD1 (× 4), PRP42 (× 4), and PRP4 (× 4); MIP6 (× 11) specifies a pore component involved in the nuclear export of mRNA. There is also very strong upregulation of rRNA-related genes, including SNR11 (× 8), RRT13 (× 23) – which regulates rRNA transcription, and RRN10 (× 11) – involved in promoting high-level transcription of rDNA.

Although there is little upregulation of well-known DNA repair genes, one of the most highly induced genes in the data set, DDI3 (× 32, P=0.00002), currently has an unknown gene product, but it is known to be inducible over 100-fold by DNA damage; YGR066c (× 23, P=0.00010) is another gene of unknown protein product, but the null mutant increases MMS (methyl methanesulfonate) sensitivity and the gene is induced over 64-fold by DNA damage (Gasch et al., 2000).

Interestingly, a number of Ty elements (four Ty1 LTR, one Ty3 LTR, 13 Ty4 LTR, and one full-length Ty4) are significantly upregulated, the highest increase being 15-fold. It can also be noted that YNR064c (× 7) encodes an epoxide hydrolase, and that T-2 toxin contains an epoxide that is essential for activity in several biological systems (Desjardins et al., 1993).

Downregulated genes

GO profiling of the downregulated genes (136 at P<0.1, 124 recognized by GO) by Biological Process showed that of the top 10 downregulated categories (Table 3), ‘Mitochondrial organization’ has the lowest probability (1.42E-6) and, with 22 out of 124 genes, it is 4.5-fold over-represented. Detailed inspection of all repressed genes shows that genes involved in respiration and mitochondrial biogenesis constitute 22 out of the 66 repressed threefold or more. Examples of these are exons of the mitochondrially encoded cytochrome b subunit [(× 8, P=0.002 – the most highly repressed transcript in the data set), (× 6), (× 4)] and of the mitochondrially encoded subunit 1 of cytochrome c oxidase [(× 6), (× 4), (× 3)], PET309 (× 5 – translational activator for COX1 mRNA), and CYT2 (× 4 – lyase needed for the maturation of cytochrome c1). Strongly repressed mitochondrial transcription and translation genes include RPO41 (× 4 – mitochondrial RNA polymerase), IFM1 (× 4 – mitochondrial translation initiation factor), MEF1 (× 4 – mitochondrial translation elongation factor), RSM19 (× 4 – mitochondrial ribosomal small subunit protein), MRP13 (× 4 – mitochondrial ribosomal small subunit protein), RML2 (× 3 – mitochondrial ribosomal large subunit protein), MRPL13 (× 4 – mitochondrial ribosomal large subunit protein), ISM1 (× 4 – mitochondrial isoleucyl tRNA synthetase), VAS1 (× 3 – mitochondrial and cytoplasmic valyl-tRNA synthetase), and MTO1 (× 5 – needed for the modification of mitochondrial tRNAs).

Table 3.   The top 10 GO Biological Process categories of downregulated genes based on their probability
GO term
Biological Process
Frequency (%)ProbabilityGenes (fold downregulated)
Downregulated (124)Genome (7146)Top 10, if>10 genes
  1. 124 out of 136 downregulated genes/exons (P<0.1) were GO recognized; the three exons of COB, and four exons of COX1, were counted as two genes.

Mitochondrion organization17.74.01.42E-06PET309 (5.21)SSC1 (4.40)
MEF1 (4.32)RSM19 (4.32)
ISM1 (3.90)COX17 (3.80)
IMF1 (3.77)RPO41 (3.69)
MRPL13 (3.59)MYO2 (3.29)
Cellular component organization44.422.08.29E-06PET309 (5.21)TLC1 (4.512)
SSC1 (4.40)TRA1 (4.39)
RAX2 (4.38)MEF1 (4.32)
RSM19 (4.32)JEM1 (3.96)
IMF1 (3.77)COX17 (3.80)
Cellular process87.966.81.83E-05SNR38 (7.89)COB (7.59)
SNR75 (6.11)COX1 (6.10)
ROT2 (6.07)SNR77 (5.75)
SNR40 (3.39)SNR13 (5.37)
PET309 (5.21)SNR61 (5.20)
Organelle organization32.314.80.00023PET309 (5.21)SSC1 (4.40)
TRA1 (4.39)MEF1 (4.32)
RSM19 (4.32)JEM1 (3.96)
IMF1 (3.77)COX17 (3.80)
KCC (3.79)MRPL13 (3.59)
Gene expression42.724.80.0033SNR38 (7.89)SNR75 (6.11)
CAF120 (6.01)SNR77 (5.75)
SNR40 (3.39)SNR13 (5.37)
PET309 (5.21)SNR61 (5.20)
MOT1 (4.93)SNR76 (4.60)
rRNA methylation5.60.60.00426SNR38 (7.89)SNR75 (6.11)
SNR77 (5.75)SNR40 (3.39)
SNR13 (5.37)SNR61 (5.20)
SNR76 (4.60) 
RNA metabolic process31.516.10.00546SNR38 (7.89)SNR75 (6.11)
CAF120 (6.01)SNR77 (5.75)
SNR40 (3.39)SNR13 (5.37)
PET309 (5.21)SNR61 (5.20)
SNR76 (4.60)CDC39 (4.49)
Mitochondrial translation8.11.50.00816PET309 (5.21)RSM19 (4.32)
MEF1 (4.32)ISM1 (3.90)
MRPL13 (3.59)MRPL44 (3.17)
IMF1 (3.77)MRP1 (2.42)
MRPS18 (2.41)VAR1 (2.41)
Cell cycle17.77.30.03462CDC39 (4.49)RAX2 (4.38)
KCC4 (3.79)PIN4 (3.68)
RAD9 (3.57)SMC3 (3.57)
SAP155 (3.35)MYO2 (3.29)
SGS1 (3.17)VHS3 (3.06)
Biological regulation29.015.90.0565CAF120 (6.01)PET309 (5.21)
CDC39 (4.49)SSC1 (4.40)
TRA1 (4.39)KCC4 (3.79)
PIN4 (3.68)RAD9 (3.57)
GPR1 (3.40)MYO2 (3.29)

A second distinct group of repressed genes have roles in rDNA stability and expression: six of the most highly repressed genes are C/D box small nucleolar RNAs, which guide methylation of rRNAs –SNR38 (× 8), SNR75 (× 6), SNR77 (× 6), SNR13 (× 5), SNR61 (× 5), and SNR76 (× 5). RDN37-1 (× 4), RDN5-1 (× 4), jointly specify the cytoplasmic rRNAs and SGS1 (× 3) encodes a nucleolar DNA helicase. Repressed genes involved in rRNA processing include UTP20 (× 4), RAT1 (× 3), and BMS1 (× 3). Among genes with more general transcriptional roles are CAF120 (× 6) and CDC39 (× 5) – both part of the CCR4–NOT transcriptional regulatory complex, and TRA1 (× 4), which contributes to histone acetyl transferase activity, thus being involved in transcriptional activation.

A number of repressed genes are involved in growth and cell division. Examples of the more highly repressed genes are TLC1 (× 5 – the RNA template component of telomerase), RAD9 (× 4 – DNA damage checkpoint protein), RAX2 (× 4 – bud site selection), KCC4 (× 4 – bud neck protein kinase involved in the septin checkpoint), PIN4 (× 4 – G2/M phase progression), and VHS3 (× 3 – G1/S transition of mitosis).

qRT-PCR

To substantiate evidence of altered gene expression as detected by the microarray analysis, qRT-PCR analysis was performed on transcripts of five upregulated genes, and the results are given in Table 1. All five genes upregulated in the microarray data were significantly upregulated in this test, but the factors were lower, and the relativities were altered.

Sensitivity of growth to the T-2 toxin in fermentable and nonfermentable media

Preferential toxic activity affecting the mitochondrial/respiratory system can be indicated by greater toxin sensitivity of growth on a medium with a nonfermentable carbon source compared with a fermentable medium, as S. cerevisiae is a facultative anaerobe (Egilsson et al., 1979). Strain BY4743 was grown in glucose or glycerol media, in microwell plates, with and without 200 μM T-2 toxin; three replicate wells were used for each condition. The growth curves in Fig. 4 show that growth in glycerol is nearly completely inhibited by the T-2 toxin, whereas growth on glucose in the presence of the same toxin concentration, although reduced, is quite vigorous: this is consistent with T-2 toxin selectively inhibiting the respiratory activity of the mitochondrion, or a function essential for growth that depends on respiratory activity.

Figure 4.

 T-2 toxin effects on the growth of Saccharomyces cerevisiae (strain BY4743) on different carbon sources. The yeast extract/peptone-based media were as described in the text.

Induction of ROS

Figure 5 shows that treatment with 200 μM T-2 toxin generates ROS in strain BY4743, as detected by DHR123: the colourless DHR123 reacts with oxidants to form rhodamine 123, which fluoresces green, while calcofluor white binds to cellulose and chitin in the cell walls of fungi, and emits bright blue fluorescence. Green fluorescence is seen in 56% of T-2 toxin-treated cells (the sample scored was 538), whereas green fluorescence is not observed in controls. There is no evidence of a particular subset (defined by size/morphology/position in the cell cycle) of cells being especially prone to ROS formation in the presence of T-2 toxin. DHR123-detected ROS-induction analysed using FACS (Fig. 6) showed clear concentration dependence, with<0.5% ROS-positive cells at 0 μM T-2 toxin, and up to 64% at 200 μM (percentages of positive cells cited here refer to the M2 figures i.e. the percentages of cells emitting above the background levels of fluorescence). With respect to time, ROS-positive cells induced by 200 μM T-2 toxin increased from 55.2% at 2 h to 64% at 4 h, although at 50 μM, the levels at 2 h (13.9%) and 4 h (10.1%) were similar, perhaps because the intracellular reduction of ROS was more effective than at higher toxin concentration. PI fluorescence monitored in the above FACS analysis showed no significant formation of necrotic or dead cells (2% maximum), while a heat-treated positive control (2 min at 80 °C) yielded 97% positive cells (data not shown).

Figure 5.

 Induction of ROS by T-2 toxin treatment. Saccharomyces cerevisiae (strain BY4743) was treated as described in the text. (a) Control cells (16-h incubation at 30°C with shaking, no T-2 toxin treatment) showed no ROS production. (b) Cells incubated as in (a), but treated with 200 μM T-2 toxin, showed substantial levels of ROS production (green fluorescence) in the buds, mother cells, and nonbudding cells.

Figure 6.

 Quantitative analysis of ROS production (DHR123 staining) by different concentrations of T-2 toxin. y-Axis – Counts: cell numbers (strain BY4743); x-axis – FL1-H: fluorescence intensity, >10 considered as positive. M2 shows the proportion of the population that has a fluorescence level above the background. Without T-2 toxin in the medium, there is almost no ROS production, at times 0, 2, and 4 h; however, 50 μM T-2 toxin caused 13.9% of ROS at 2 h and 10.1% of ROS at 4 h, and 200 μM T-2 toxin generated 55.2% of ROS positives at 2 h and 64.2% ROS at 4 h.

DHE staining was then used in a FACS investigation of superoxide induction (Fig. 7): three independent clones of strain BY4743 treated with 200 μM T-2 toxin yielded high levels of induction (a mean of 78.8%), compared with the level at 0 h (mean of 0.8%). Parallel FACS analysis of PI fluorescence showed no significant necrosis in the three clones at 0 h (mean of 0.6%), while after 4 h of treatment with 200 μM T-2 toxin, the frequencies of dead or necrotic cells were 0.9%, 0.3%, and 22.5% (perhaps in the case of the third clone, cell breakdown following ROS-induced apoptosis was particularly far advanced).

Figure 7.

 Superoxide induction by T-2 toxin detected by DHE staining. y-Axis – Counts: cell numbers (strain BY4743); x-axis – FL2-H: fluorescence intensity, >10 considered as positive. M2 designates the proportion of the population that has a fluorescence level above the background. All three randomly selected clones contained high frequencies of superoxide-positive cells after 4 h of T-2 toxin treatment: clone 1 – 79.7%, clone 2 – 87.4%, and clone 3 – 69.4%.

Effects of antioxidants

Figure 8 shows that the antioxidants NAC and GSH have no significant effect on the log-phase growth of a glucose culture of strain BY4743, although NAC mildly increases the A600 nm of the stationary phase of growth. Inhibition of growth by T-2 toxin (200 μM) was substantially although incompletely relieved by NAC and GSH, with NAC being marginally more effective in enhancing log-phase growth in the presence of the toxin. Three replicate wells were used for each condition.

Figure 8.

 Effects of the antioxidants NAC and GSH on the growth of Saccharomyces cerevisiae (strain BY4743) with or without T-2 toxin. The basic growth medium was YPD (see text).

Annexin V staining of externalized phosphatidylserine

Externalization of phosphatidylserine is considered to be an early marker of apoptosis (Fadok et al., 1998), and can be detected by Annexin V-FITC staining, with PI double staining. In normal live cells, phosphatidylserine is located on the cytoplasmic surface of the plasma membrane. However, in apoptotic cells, phosphatidylserine is translocated from the inner to the outer leaflet of the plasma membrane, thus exposing phosphatidylserine to the external cellular environment. Annexin V is a 35–36 kDa Ca2+-dependent phospholipid-binding protein that has a high affinity for phosphatidylserine. Annexin V conjugated to fluorescein (Annexin V-FITC) can identify apoptotic cells by binding to phosphatidylserine exposed on the outer leaflet. PI is excluded by live and apoptotic cells, but enters dead cells, binding tightly to nucleic acids, and generating red fluorescence in the cell. Combining Annexin V-FITC and PI with calcofluor white, cell walls and bud scars of all cells display blue fluorescence, apoptotic cells show green fluorescence, dead cells show red fluorescence and occasionally some green fluorescence, and living cells show neither green nor red fluorescence.

Figure 9 shows that 21% (the total sample included 585 cells) of the cells of strain BY4743 treated with 200 μM T-2 toxin for 16 h fluoresced green, indicating externalization of phosphatidylserine. Dead cells (fluorescing red) were also seen, at a frequency of 19%. About 5% of the cells exhibited both green and red fluorescence.

Figure 9.

 T-2 toxin-induced apoptosis in Saccharomyces cerevisiae (strain BY4743) – externalization of phosphatidylserine. Annexin V-FITC and PI were used to detect apoptotic and dead cells. (a) Control, without T-2 toxin treatment: no evidence of apoptotic or dead cells. (b) Treated with 200 μM of T-2 toxin for 16 h: shows some dead cells (red fluorescence), some apoptotic cells (green fluorescence), and some cells with both fluorescent signals (red and green).

TUNEL analysis

TUNEL analysis was introduced by Gavrieli et al. (1992) to detect 3′OH ends (i.e. cleaved phosphodiester bonds or ‘nicks’) in DNA, derived from endonucleolytic cleavage and deemed to be indicative of apoptosis. As Fig. 10 shows, 8.3% of the T-2 toxin-treated BY4743 cells (200 μM for 16 h) were labelled by the TUNEL procedure; no fluorescence is seen in the control sample, not exposed to the T-2 toxin. A striking feature is that in all cases (1900 cells screened), label is confined to very young buds, no larger than 25% of the length (ellipsoidal major axis) of the mother cell.

Figure 10.

 TUNEL staining of T-2 toxin-treated Saccharomyces cerevisiae cells indicating DNA damage. Cells of strain BY4743 were treated with DMSO only (a1–a3) or 200 μM T-2 toxin for 16 h (b1–b3) and then TUNEL-stained. For control cells (DMSO treatment only), no TUNEL-staining positive cells were found; however, for T-2 toxin-treated cells, in a sample of 1900 cells, 158 (8.3%) were TUNEL-positive (red fluorescence, mother cell plus bud counted as one cell), and in all cases, fluorescence was restricted to the bud. Magnifications are × 100 (a1 and b1), and × 600 (a2 and b2 – fluorescence, a3 and b3 – bright field), with a2 and a3 and b2 and b3 being identical fields.

Effects of selected deletion mutations on T-2 toxin sensitivity

Seven deletion mutants of the haploid parent strain BY4741 were selected for study on the basis that the genes were one of the top seven most up- or downregulated annotated genes, in the transcriptomic experiment. Growth was monitored in the presence and absence of T-2 toxin (200 μM) for 56–72 h using the microplate method described above, each datum point being the average of three (minimum) replicate wells, and indices comparing mutant and wild-type responses to the T-2 toxin were calculated; indices>1.0 indicate that the mutant is more sensitive than the parent strain, and <1.0 indicates greater tolerance (see Table 4). Deletion of MIP6 showed no significant change in tolerance, deletions of BNA2, YMR118C, COX17, and DDI3 exhibited marginally enhanced tolerance, and deletion of PET309 conferred significant tolerance. The deletion mutant showing most difference from the parent strain was FMS1, encoding polyamine oxidase: as the growth curves in Fig. 11 indicate, it is markedly more tolerant to the T-2 toxin than the parent strain.

Table 4.   T-2 toxin effects on selected yeast deletion strains
Mutant nameORFGene functionTranscriptomic
changes (fold)
Index of effect
on growth
  • All yeast strains listed in the table are derived from BY4741 except the last two. Index of the effect on growth calculated as follows: (Tmut_T2/200 μM_A 0.1/Tmut_T2/0_A 0.1)/(Twt_T2/200 μM_A 0.1/Twt_T−2/0_A 0.1). Tmut_T2/200 μM represents the time needed by mutant strains to reach A600 nm 0.1 in the presence of 200 μM T-2 toxin; Tmut_T2/0 represents the time needed by mutant strains to reach A600 nm 0.1 without T-2 toxin treatment; Twt_T2/200 μM_A 0.1 represents the time needed by parent strains to reach A600 nm 0.1 in the presence of 200 μM T-2 toxin; and Twt_T2/0_A 0.1 represents the time needed by parent strains to reach A600 nm 0.1 without T-2 toxin treatment. If a mutant strain is completely tolerant to T-2 toxin, the index would be 0.45 for BY4741-derived strains, and 0.5 for BY4743-derived strains.

  • *

    P>0.1. The transcriptomic changes cited in the table are those seen in our transcriptomic analysis of diploid strain YB110 (see also Tables 2 and 3).

PET309YLR067CSpecific translational activator for the COX1 mRNA, also influences stability of intron-containing COX1 primary transcripts↓5.210.81 ± 0.10
FMS1YMR020WPolyamine oxidase, converts spermine to spermidine, also involved in pantothenic acid biosynthesis↑9.020.61 ± 0.08
BNA2YJR078WPutative tryptophan 2,3-dioxygenase or indoleamine 2,3-dioxygenase↑40.610.86 ± 0.12
YMR118CYMR118CSimilarity to succinate dehydrogenase cytochrome b subunit↑12.120.86 ± 0.12
COX17YLL009CCopper metallochaperone↓3.800.87 ± 0.12
MIP6YHR015WPutative RNA-binding protein, interacts with Mex67p↑11.281.04 ± 0.09
DDI3YNL335WExpression is induced over 100-fold by DNA damage↑32.760.84 ± 0.14
FAP1YNL023CProtein that binds to Fpr1p, conferring rapamycin resistance by competing with rapamycin for Fpr1p binding, similarity to human NFX1↑1.23*0.80 ± 0.12
FPR1YNL135CPeptidyl-prolyl cistrans isomerase↓1.39*0.78 ± 0.14
TOR1YJR066WPIK-related protein kinase and rapamycin target; subunit of TORC1↓2.781.05 ± 0.09
GLN3YER040WTranscriptional activator of genes regulated by nitrogen catabolite repression↓ 2.323.03 ± 0.03
VPS27YNR006WEndosomal protein that forms a complex with Hse1p↑1.28*3.70 ± 0.04
RIP1YEL024WUbiquinol-cytochrome-c reductase↓1.13*0.63 ± 0.08
RSM28YDR494WMitochondrial ribosomal protein of the small subunit↓1.17*0.89 ± 0.10
RMD9YGL107CMitochondrial protein required for respiratory growth, may play a role in delivering mitochondrial mRNAs to ribosomes↓2.280.54 ± 0.05
PET122YER153CMitochondrial translational activator specific for the COX3 mRNA↓1.02*0.65 ± 0.07
OXA1YER154WMitochondrial inner membrane insertase↓2.10*0.56 ± 0.07
MEF1YLR069CMitochondrial elongation factor↓4.320.61 ± 0.09
IFM1YOL023WMitochondrial translation initiation factor 2↓3.770.92 ± 0.04
TUF1YOR187WMitochondrial translation elongation factor Tu↓2.45*0.57 ± 0.07
ISM1YPL040CMitochondrial isoleucyl-tRNA synthetase↓3.900.61 ± 0.09
MSS116YDR194CDEAD-box protein required for efficient splicing of mitochondrial Group I and II introns↓2.230.62 ± 0.09
BY4743_Pet1NARandom spontaneous petite 1NA0.54 ± 0.10
BY4743_Pet2NARandom spontaneous petite 2NA0.63 ± 0.08
Figure 11.

 T-2 toxin effects on BY4741 parental and FMS1 deletion strains. The growth medium was YPD (see text). Triplicates were used for each condition/treatment; hence, each datum point shown is the average of the absorbances of three wells. Reading intervals were 30 min, but for clarity, only absorbances at 2-h intervals are shown.

In view of the recent findings in Arabidopsis that AtNFXL1 is involved in the action of T-2 toxin, AtNFXL being a homologue of the yeast TORC1 (target of rapamycin complex 1) pathway-related gene FAP1 (Asano et al., 2008), we also looked at deletions of genes involved in TOR function. Deletion mutants of five genes involved in TOR function were assessed: deletion of TOR1 caused no change in tolerance, whereas deletions of FAP1 and FPR1 significantly increased resistance, and deletions of GLN3 and VPS27 markedly increased sensitivity (Table 4).

Because of our transcriptomic and other evidence that the mitochondrial system is targeted by the T-2 toxin, we also assessed the effect on toxin sensitivity of deletion mutations in mtDNA (two so-called ρ, spontaneous mutations) and of the deletion of nuclear genes (10) encoding important elements of the mitochondrial system (see Table 4 for descriptions): all these mutations cause respiratory deficiency. Table 4 shows that for all these mutants, the tolerance of the T-2 toxin is increased, in most cases substantially. PET309, mentioned above, also falls into this category (Table 4). It is noteworthy that the two mtDNA petites tested show very high levels of tolerance to the T-2 toxin: we have observed that T-2 toxin can select strongly for cytoplasmic petites (results not shown).

Discussion

Transcriptomics

Transcriptomic analyses of the T-2 toxin effects have been undertaken recently in two very different higher eukaryotes. In A. thaliana, 35 genes were highly upregulated (× 3 or greater), including a WRKY transcription factor and three peroxidase genes – consistent with ROS generation by the toxin, while other induced genes suggested the induction of plant defence responses and inactivation of the plant steroid hormone brassinosteroid (Masuda et al., 2007); hydrogen peroxide (H2O2) generation was shown to occur in T-2 toxin-treated Arabidopsis leaves (Nishiuchi et al., 2006). Unlike T-2 toxin, the well-known protein synthesis inhibitor cycloheximide did not induce lesions in treated leaves (Nishiuchi et al., 2006), indicating a different mechanism of action. In a microarray study of T-2 toxin-induced lesions on foetal rat brains (Sehata et al., 2004a), only four genes were upregulated more than threefold, three being oxidative stress related – HSP70 (× 145), metallothionein (MT)-2 and -1 (× 16), heme oxygenase (HO –× 7), and mitochondrion-related genes such as cytochrome oxidase were substantially downregulated; significant, although low (5–6%) frequencies of TUNEL-positive cells were also seen in brain tissue, indicative of apoptosis. This animal work shows clear parallels with our yeast transcriptomic results, the authors concluding that T-2 toxin induced oxidative stress, triggering apoptosis – increased expression of caspase 2 (inducible by ROS) being seen 24 h after treatment (Sehata et al., 2004a). Sehata et al. (2004b, 2005) reported similar transcriptional changes and induction of apoptosis in T-2 toxin-treated pregnant rats.

In a recent transcriptomic study using S. cerevisiae, published after our transcriptomic experiment was undertaken, T-2 toxin was used at 108 μg mL−1 for 2 h, RNA levels corresponding to 6131 ORFs were analysed, and, using a P<0.05 criterion, 515 genes were≥2 upregulated, and 490 genes were≥2 downregulated (Iwahashi et al., 2008). The top 45 named upregulated (between × 15.7 and × 5.0) genes are largely different from those identified as most upregulated in our study, but a few genes appear in both lists: BNA2 (up to 40.6-fold in our study, up to 6.8-fold in the Iwahashi study), THI11, and FDH1. Differences in the strain and the precise treatment conditions will contribute to differences in transcript profiles; however, the oxidative stress response genes AAD4, AAD16, and AAD14, as in our study, were significantly upregulated (between × 2 and × 5) – indeed, both yeast studies show substantial numbers of upregulated redox function genes. Although Iwahashi et al. (2008) do not list downregulated genes, another similarity to our findings that they report is the suppression of many genes concerned with the electron transport chain (especially cytochrome c-related) and ribosome biogenesis and rRNA processing. Because they found that 11 of their top 45 upregulated genes were transporter genes, Iwahashi and colleagues concluded that the T-2 toxin attacks the plasma membrane, although no direct evidence of this was given.

Upregulation of Ty retrotranposons is a feature of the results reported here, not reported by Iwahashi et al. (2008); Ansari and colleagues (unpublished data quoted in Rocha et al., 2005) found that retrotransposon transcripts were overexpressed in wheat roots treated with the trichothecene mycotoxin deoxynivalenol, and there is a recent suggestion that Ty1-driven transcription could regulate yeast gene expression in response to stress (Servant et al., 2008).

The qRT-PCR results broadly support the microarray evidence of transcriptional change, with five of the most upregulated genes, according to microarray hybridization, also showing significant upregulation by qRT-PCR (see Table 1). It should be noted that this positive correlation is seen, although different strains were used and despite the different techniques used for RNA isolation.

Patterns of transcriptional change

A major theme in the transcriptional response described above is strong upregulation of many genes concerned with redox activity, as well as of a number of mitochondrial maintenance genes, while the most notable group of strongly repressed genes are concerned with mitochondrial biogenesis and the electron transport chain. There is also substantial upregulation of genes that respond to stress, including oxidative stress, a noteworthy example being GTT2 (× 7), which encodes an apparently mitochondrial GSH-S-transferase. This response pattern is consistent with T-2 toxin triggering the formation of ROS and the consequent localized damage in the mitochondrial system, impairing the mitochondrial transcription and translation apparatus and reducing the transcript levels of key genes such as those needed for mitoribosome production and activity, and for mitochondrially encoded inner-membrane proteins such as cytochromes b and c1. Damage to mtDNA may also occur. A cellular response to this toxic stress could be upregulation of genes needed to generate higher levels of reduced coenzymes, through biosynthesis, for example BNA2 (× 40), and dehydrogenase activity, for example FDH2 (× 27), and of genes, for example PET18 (× 8), involved in mitochondrial genome stability. Depression of mitochondrial activity and elevated reducing levels in the cell might prompt induction of genes, such as TIR4 and DAN5, normally only expressed anaerobically, and lowered ATP levels consequent on mitochondrial dysfunction could repress genes involved in growth and cell division, for example CDC6 (× 3) and RAX2 (× 4). Induction of meiosis and sporulation genes such as DMC1 (× 6), SGA1 (× 7) and SLZ1 (× 8) could be related to growth inhibition and/or stress. Apparently, contradictory responses also occur, such as upregulated rRNA transcription (e.g RRN10, × 11) and downregulation of rRNA processing genes (e.g. UTP20, × 4). Also, while there is upregulation of genes such as PET18 (× 8), THI4 (× 7), and SML1 (× 7), concerned with mitochondrial genome stability and response to DNA damage, there is no extensive induction of DNA repair genes, although the poorly understood gene DDI3, known to respond to DNA damage, is highly upregulated (× 32).

T-2 toxin and the mitochondrion

The glucose vs. glycerol growth curve data reported here provide clear quantitative evidence that the mitochondrial system – respiratory function – is a significant target for T-2 toxin: as Fig. 4 shows, growth with glycerol as a carbon source – where respiratory function is essential – is completely inhibited at a concentration that only partially inhibits glucose growth. Higher T-2 toxicity towards the growth of S. cerevisiae on a glycerol medium has also been seen using a filter disc method with a solid medium (Schappert & Khachatourians, 1983). Interestingly, in the recent paper by McLaughlin et al. (2009), preferential toxicity of the type B trichothecene, trichothecin, on glycerol compared with glucose medium was also seen. Direct inhibition of site II of the electron transport chain of isolated yeast mitochondria has been reported (Koshinsky et al., 1988) and in the case of isolated rat liver mitochondria, T-2 toxin was found to inhibit site I of the electron transport chain (Pace, 1983), and also mitochondrial protein synthesis (Pace et al., 1988). Mitochondria are known to be a source of ROS (Poyton et al., 2009), and ROS generation can be enhanced by impairment of the electron transport chain and damage to mtDNA (Indo et al., 2007). As Fig. 5 shows, T-2 toxin indeed induces substantial ROS levels in a large fraction of treated cells – the first such report for yeast. Further evidence of ROS induction is provided by the FACS analysis using the oxidant detector DHR123 reported in Fig. 6, and the data in Fig. 7, using DHE, indicate that superoxide anions are an important component of the ROS generated by T-2 toxin. This induction of ROS explains the efficacy of antioxidants in antagonizing T-2 toxin inhibition of growth (Fig. 8) as well as our transcriptomic evidence of increased expression of genes with roles in combating oxidative stress. Interestingly, GSH and cysteine have recently been reported (Gardiner et al., 2010) to antagonize the toxicity towards yeast of the trichothecene deoxynivalenol. T-2 toxin has been shown to induce antioxidant enzymes in mice (Chaudhari et al., 2009), and induction of mitochondria-dependent apoptosis has been seen in cultured mammalian cells (Bouaziz et al., 2009) – consistent with T-2 toxin provoking oxidative stress and triggering apoptosis in yeast cells. Our observation of externalization of phosphatidylserine (Fig. 9) is strong evidence that T-2 toxin is indeed triggering apoptosis in yeast cells. It can be noted that the FITC Annexin V-staining/phosphatidylserine externalization, an early marker of apoptosis, often appears to be localized in a portion of the cell or in a bud. With respect of cell localization, the TUNEL staining results are particularly striking: 8.3% of 1900 cells scored were TUNEL-positive, and in all cases, the red fluorescence was restricted to young buds (Fig. 10). This is evidence of damaged DNA in this cell compartment, and as it is known that mitochondria enter buds early in the cell cycle (Stevens, 1981; Roeder et al., 1998), but the nucleus does not (Hoepfner et al., 2000), the damaged DNA is highly likely to be mitochondrial: this could suggest that mitochondria and mtDNA in the early bud are particularly vulnerable to the effects of the T-2 toxin, or that damaged mtDNA is somehow segregated into the early bud. As ROS formation can be seen throughout the cell, mother cell, and bud (Fig. 5), perhaps the latter explanation is more likely. TUNEL staining of mtDNA, as opposed to nuclear DNA, may also be an early marker of apoptosis in yeast. There is now considerable evidence for mitochondrial involvement, including mitochondrial fission, in apoptosis in yeast (Gourlay et al., 2006; Cheng et al., 2008), and apoptosis induction in yeast mediated via mitochondrial damage and ROS induction by the fungal toxin bostrycin has been reported recently (Xu et al., 2010).

However, it can be pointed out that antioxidant antagonism of yeast growth inhibition by T-2 toxin is incomplete (Fig. 6), suggesting that a mitochondrial ROS toxicity pathway is not the only mechanism of action by this mycotoxin. Whatever the mechanism, severe T-2 treatment (200 μM for 16 h) causes significant levels of cell death (Fig. 9), although the levels of necrosis are low at 4 h (PI FACS data reported in the text above).

Effects of deletion mutants

Clearly, up- or downregulation of transcription does not mean that a particular gene is a target of the T-2 toxin or even closely connected, functionally or structurally, but it may be significant that six of seven deletion mutants tested, chosen because of large transcriptional changes in response to the toxin, showed enhanced tolerance compared with the parent strain. Deletion of FMS1 caused the greatest change in this group (Table 4 and Fig. 9). The polyamine oxidase encoded by FMS1 converts spermine to spermidine, which is required for the essential hypusination modification of translation factor eIF-5A; hence, the deletion of FMS1 could reduce the functionality of a protein factor now considered to be involved in translation elongation (rather than initiation), but still poorly understood and with roles not directly connected to the translation mechanism (Saini et al., 2009). It can also be noted that polyamine oxidase can generate H2O2– deletion of FMS1 potentially reducing oxidative stress, and that polyamine (spermine) oxidase activity has been found in the mitochondrial fraction of yeast cells (Tanaka et al., 2004). The connection between FMS1 and protein synthesis could be significant, in view of many early findings showing that trichothecenes are potent inhibitors of protein synthesis in vivo and in vitro, with T-2 toxin and others causing rapid disaggregation of polysomes in H-HeLa cells, suggesting the inhibition of initiation of translation (Cundliffe et al., 1974), possibly caused by binding to the peptidyl transferase (McLaughlin et al., 1977). However, the remarkably greater sensitivity of in vivo systems compared with in vitro and the importance of lipophilicity in T-2 toxin's potency (Sato & Ueno, 1977) is consistent with the involvement of a membranous structure or function in the toxic mechanism (Ueno, 1977), before interaction with the ribosome.

Regarding TOR-related genes, FAP1 encodes a transcription factor able to confer rapamycin resistance, when overexpressed, by competing for binding to rapamycin with Fpr1p, the ligand that normally binds to rapamycin, the Fpr1p–rapamycin complex inhibiting TORC1; deletion of FAP1 produced some increase in tolerance to T-2 toxin (Table 4), which is unexpected if TORC1, like rapamycin, is a T-2 toxin target. In Arabidopsis, deletion of the FAP1 analogue AtNFXL1 resulted in hypersensitivity to T-2 toxin, a severe growth defect being caused, but not cell death, the interpretation being that AtNFXL1p negatively regulates defence-related genes (Asano et al., 2008). Deletion of FPR1, encoding the rapamycin-binding peptidyl prolyl isomerase (Fpr1p), causes increased rapamycin resistance and showed very similar, enhanced, T-2 toxin tolerance to the FAP1 deletion mutant. Deletion of TOR1 had no effect on T-2 toxin tolerance in our study (Table 4), again consistent with TORC1 not being a T-2 toxin target. GLN3 encodes a transcriptional factor involved in nitrogen catabolite repression negatively regulated by TORC1: deletion is known to increase resistance to rapamycin, but very significantly reduced tolerance to the T-2 toxin here, again indicating different toxicity pathways. Deletion of VPS27 (an endosomal protein-coding gene), which increases sensitivity to rapamycin, also increased sensitivity to T-2 toxin (Table 4). Overall, our TOR gene-related data certainly indicate that T-2 toxin has a different mechanism of action from rapamycin, but do not exclude the possibility of some involvement of the TOR pathway in the T-2 toxicity mechanism. Another feature of the gene deletion data is the lack of correspondence between transcriptional changes and effects on T-2 toxin tolerance, also seen in the case of rapamycin (Xie et al., 2005).

Table 4 also reports the effects of mitochondrial and nuclear mutations causing a respiratory-deficient phenotype: it is notable that in each case, sensitivity to T-2 toxin inhibition of growth is significantly, often substantially, reduced. The fact that the mutational inactivation of respiratory activity – by a variety of routes – causes enhanced resistance, strongly implicates an active, respiring mitochondrial system in the mechanism of action of the T-2 toxin. McLaughlin et al. (2009) have recently reported that induction of the ρ0 mutation increases resistance to the type B trichothecene, trichothecin, and that in a genome-wide screen of deletion mutations, the largest group of trichothecin-resistant deletion strains were mutants affected in mitochondrial function: they interpreted this as evidence of a critical role for the mitochondrion in trichothecin toxicity. McLaughlin et al. (2009) also observed that trichothecin causes fragmentation of the tubular mitochondrial network, and, as noted above, mitochondrial fission may be a significant step on the road to apoptosis in both yeast and mammalian cells (Cheng et al., 2008).

Concluding remarks

The transcriptomic data reported here are consistent with the mitochondrial system being a significant target for the T-2 toxin, one outcome being the substantial repression of key genes concerned with mitochondrial biogenesis and electron transport. This is supported by growth on a nonfermentable carbon source being more sensitive to T-2 toxin than growth on glucose. It is possible that the lipophilic T-2 toxin molecule interacts with the actively respiring mitochondrial inner membrane to provoke ROS formation, resulting in the induction of genes controlling redox activity and protecting against oxidative stress, but also triggering apoptosis. We have shown that T-2 toxin treatment does cause substantial ROS formation, that inhibition of growth by the T-2 toxin is antagonized by antioxidants, that genetic respiratory deficiency increases tolerance to the T-2 toxin, and that markers indicative of apoptosis are induced by the toxin. However, neither antioxidant treatment nor respiratory deficiency completely suppresses T-2 toxicity, indicating other toxicity targets, and it is well known that trichothecenes including T-2 toxin can inhibit cytoplasmic protein synthesis both in vitro and in vivo. It is possible that T-2 toxin has at least two targets, in the mitochondrion and the cytoplasmic ribosome, or that these two targets are linked in an as yet unknown pathway.

Acknowledgements

The authors would like to thank COGEME for a grant towards the transcriptomic analysis, and the staff of the UMIST microarray facility, especially Dr Michael Wilson, for help with transcriptomic analysis. Thanks are also due to Dr Mariangela Bizzarri for her assistance with the work.

Statement

L.J. and X.L. are joint first authors.

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