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Report |
Address correspondence to Kathryn R. Ayscough, Department of Molecular Biology and Biotechnology, University of Sheffield, Western Bank, Sheffield S10 2TN, UK. Tel.: 44 114 222 2332. Fax: 44 114 272 8697. email: k.ayscough{at}sheffield.ac.uk
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Abstract |
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Key Words: actin; senescence; Scp1; apoptosis; ROS
Abbreviations used in this paper: ROS, reactive oxygen species; Tpm, tropomyosin.
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Introduction |
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Coupled to the process of aging is the presence of tightly regulated mechanisms, which ensure that aged cells undergo cell death. Such pathways are critical in multicellular organisms to allow maintenance of tissues and organ structure. However, it is clear that apoptotic pathways that culminate in cell death also occur in single cell organisms. One possible reason for this in yeast is that old cells are less robust and their presence may become deleterious to the well being of a colony. The presence of apoptotic pathways in yeast that resemble those in higher organisms has been well demonstrated (Frohlich and Madeo, 2001; Tissenbaum and Guarente, 2002). As observed in mammalian cells, apoptotic yeast cells show mitochondrial depolarization, high levels of reactive oxygen species (ROS), DNA fragmentation, and phosphoserine exposure at the plasma membrane. There is also good evidence that these apoptotic pathways are part of the normal aging process (Laun et al., 2001; Jazwinski, 2002). The role of the caspase-like protein Yca1p in yeast is less well understood, though there is evidence that caspase-dependent cell death pathways are present and contribute to the process of programmed cell death (Madeo et al., 2002). However, even yca1 cells show rapid death in elevated H2O2 levels, demonstrating the presence of other mechanisms to induce cell death.
Several recent studies have reported induction of apoptosis in mammalian cells associated with changes in the actin cytoskeleton (Kim et al., 2003; Morley et al., 2003). To further our understanding of this phenomenon, we have undertaken studies in the budding yeast Saccharomyces cerevisiae. Previous studies in yeast have generated much information on fundamental roles of the actin cytoskeleton because this organism expresses a single form of actin and there is ready availability of mutant actin alleles. Here we demonstrate that a decrease in actin dynamics can induce cell death through the production of ROS from the mitochondria. Conversely, increasing actin dynamics reduces production of ROS and increases cell viability. Finally, we determine that deletion of a single gene, encoding the actin bundling protein Scp1, leads to a reduced production of ROS and a highly significant increase in longevity.
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Results and discussion |
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An important consideration is whether the changes to actin are caused by, or are a consequence of, elevated ROS. Actin staining of the act1-159 strain revealed some disorganization of the actin cytoskeleton with fewer, larger chunks of cortical actin and aberrant actin cables (Fig. 1 e). This phenotype was not rescued in the rho° strain that has lower levels of ROS, demonstrating that actin disorganization occurs upstream of the production of ROS from the mitochondria. We also observed that the actin phenotype was exacerbated as act1-159 cells entered stationary phase and that it correlated with an increased incidence of DNA fragmentation (unpublished data). To determine whether this actin "chunk" phenotype correlated with the ROS production, we also investigated actin organization of wild-type and act1-157 cells after 28 d in stationary phase. Here we found a clear difference between the strains, with 45% of wild-type cells containing large actin chunks compared with only 16% of act1-157 cells (Fig. 1 f), demonstrating a role for actin dynamics within stationary phase cells.
To ascertain whether the changes in ROS levels correlated with longevity, we compared viability of the yeast strains over a period of 28 d after entry into stationary phase. This approach allows assessment of chronological lifespan of yeast, which mimics aging of post-mitotic tissues in higher organisms (MacLean et al., 2001). The strain with reduced actin dynamics had all died by 10 d. After 28 d, wild-type cells showed 38.5% (±2.1%) viability, and the strain with increased dynamics showed 51% (±3.3%) viability, an increase of 32%, suggesting a link between actin dynamics, ROS production, and longevity. To demonstrate that the ROS responsible for cell death are derived from mitochondrial activity, viability of the act1-159 strain and act1-159 rho° strain were compared. This measurement was performed on log phase cells because the lack of mitochondria precludes the rho° strains from entering stationary phase. Under these conditions, viability of act1-159 cells was 50.1% (±5.8), and viability of the act1-159 rho° cells was 93.7% (±2.2), demonstrating the importance of mitochondria in production of ROS, which results in cell death.
To determine whether the cell death induced by ROS in response to the actin mutation is due to caspase activation, we incubated yeast with FITC-labeled VAD-fmk. This molecule binds specifically to the catalytic site of metazoan caspases and has been used as a method to detect yeast cells with active caspases using flow cytometry (Madeo et al., 2002). As shown in Fig. 1 f, caspase levels are elevated in strains expressing the act1-159 mutation, correlating with increased cell death of this strain. Interestingly however, the rho° strain lacking mitochondria, which has restored viability under these conditions, still has elevated caspase levels, indicating that high caspase levels alone may not trigger cell death.
One caveat of such studies is that the presence of a mutant allele in cells may for a variety of reasons initiate changes that induce ROS production. To determine whether a reduction in actin dynamics in an otherwise wild-type cell also causes an increase in ROS, we used a yeast strain that is sensitive to the actin stabilizing drug jasplakinolide (KAY339). Previously we have shown that jasplakinolide addition to this strain causes a block in actin dynamics and the actin to accumulate as a single clump (Ayscough, 2000). To analyze the effect of jasplakinolide in this study, the drug was added at low levels overnight to allow concomitant uptake of the dye used for analysis of ROS production. After this time, with 0 and 1 µM jasplakinolide addition, actin was still punctate and polarized, and mitochondrial and nuclear DNA was normal in appearance. However, 2 µM jasplakinolide addition resulted in clumped actin and fragmentation of nuclear DNA, indicative of apoptosis (Fig. 2 a). In addition, a dramatic increase in ROS was measured, indicating that the high ROS levels are a consequence of the changes in the actin cytoskeleton (Fig. 2 b).
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In summary, we have demonstrated the importance of actin cytoskeleton dynamics in regulating the production of ROS in yeast and have determined that coupling of actin and the mitochondria is central to this mechanism. Importantly, we also show that deletion of the SCP1 gene encoding an actin bundling protein significantly increases the lifespan of yeast, indicating that this pathway is also part of a pathway regulating aging processes.
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Materials and methods |
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Yeast were grown with shaking at 30°C in liquid YPD (1% yeast extract, 2% peptone, 2% glucose) or synthetic medium (lacking uracil; Sigma-Aldrich). Solid media was made by addition of 2% agar. For glycerol plates, the glucose was replaced with 3% glycerol. Overexpression of SCP1 was induced by switching cells onto media lacking methionine (Winder et al., 2003). The rho° strain of KAY375 was made as previously described (Guthrie and Fink, 1991). Jasplakinolide was added to amounts indicated from a 5 mM stock in DMSO.
Fluorescence procedures
Rhodamine-phalloidin and DAPI staining was performed as previously described (Hagan and Ayscough, 2000). Staining of mitochondria with DiOC6 was performed as published (Pringle et al., 1989). Visualization of mitochondria using GFP on a mitochondrial localization peptide was achieved by transformation of cells with PVTU100U-mtGFP (Westermann and Neupert, 2000). Cell images were recorded from a BX60 Olympus microscope with a Roper Scientific Micromax camera and captured on a Macintosh 7300 computer. Images were processed using Adobe Photoshop 7.0® software.
ROS production.
Cells were incubated overnight in the presence of 5 µg/ml 2',7'-dichloro dihydrofluorescein diacetate (H2-DCFDA; Molecular Probes). Cells were sonicated and analyzed using a Becton Dickinson flow cytometer. Parameters were set at excitation and emission settings of 304 and 551 nm (filter FL-1), respectively. For each experiment, two peaks of fluorescence were observed. The second peak represents cells with high levels of ROS. Experiments were repeated at least three times independently. Representative datasets are depicted. Errors shown are standard deviations. In vivo caspase levels were measured as previously described (Madeo et al., 2002) using a staining solution containing FITC-VAD-fmk (CaspACETM; Promega).
Aging assays
Chronological aging assay.
Cells were grown in YPD medium on a shaker with high aeration (220 rpm) at 30°C. Samples were sonicated before cell number determination, in triplicate, with a Schärfe Systems TT cell counter. Serial dilutions were plated on YPD plates, and numbers of growing colonies were counted in triplicate platings.
Reproductive capacity was assessed using the methods previously described (Lin et al., 2000). In brief, cells from log phase cultures were plated at low density. Daughter cells were isolated as buds that emerged from mother cells and removed using a Singer Instruments MSM manual micromanipulator. Life span was determined by removing all subsequent daughter cells generated. The average reproductive lifespan was calculated from the dissection of budding cells from 32 starting cells for each strain tested.
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Acknowledgments |
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This work was supported by a Medical Research Council (MRC) senior research fellowship to K.R. Ayscough (G117/394), MRC career establishment grant to S.J. Winder (G104/51660), a Wellcome Trust studentship to L.N. Carpp, and a Biotechnology and Biological Sciences Research Council grant to K.R. Ayscough and S.J. Winder (17/C12769).
Submitted: 31 October 2003
Accepted: 30 January 2004
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