Mitochondrial dysfunction reduces yeast replicative lifespan by elevating RAS-dependent ROS production by the ER-localized NADPH oxidase Yno1
Mitochondrial dysfunction reduces yeast replicative lifespan by elevating RAS- dependent ROS production by the ER- localized NADPH oxidase Yno1
Dae-Gwan Yi 0 1
Sujin Hong 0 1
Won-Ki Huh 0 1
☯ These authors contributed equally to this work. 1
0 Department of Biological Sciences, Seoul National University , Seoul , Republic of Korea, 2 Institute of Microbiology, Seoul National University , Seoul , Republic of Korea
1 Editor: Mary Bryk, Texas A&M University , UNITED STATES
Mitochondrial dysfunction leads to the accumulation of reactive oxygen species (ROS) which is associated with cellular dysfunction, disease etiology, and senescence. Here, we used the eukaryotic model Saccharomyces cerevisiae, commonly studied for cellular aging, to demonstrate how defective mitochondrial function affects yeast replicative lifespan (RLS). We show that RLS of respiratory-deficient cells decreases significantly, indicating that the maintenance of RLS requires active respiration. The shortening of RLS due to mitochondrial dysfunction was not related to the accumulation of extrachromosomal ribosomal DNA circles, a well-known cause of aging in yeast. Instead, intracellular ROS and oxidatively damaged proteins increased in respiratory-deficient mutants. We show that, while the protein kinase A activity is not elevated, ROS generation in respiratory-deficient cells depends on RAS signaling pathway. The ER-localized NADPH oxidase Yno1 also played a role in producing ROS. Our data suggest that a severe defect in mitochondrial respiration accelerates cellular aging by disturbing protein homeostasis in yeast.
Data Availability Statement: All relevant data are
within the paper and its Supporting Information
Funding: This work was supported by the National
Research Foundation of Korea grant
(2015R1A2A1A01007871) funded by the Ministry
of Education, Science and Technology, Republic of
Korea (www.nrf.re.kr) to Won-Ki Huh. The funders
had no role in study design, data collection and
analysis, decision to publish, or preparation of the
Over the past few decades, the budding yeast Saccharomyces cerevisiae has contributed to the
search for conserved elements in cellular aging [
]. In budding yeast, two distinct lifespan
paradigms have been proposed. Chronological lifespan (CLS) is a model for the aging process
of post-mitotic cells and measures the amount of time a cell can remain viable in a
non-dividing state . Replicative lifespan (RLS) is useful for understanding aging of dividing cells and
defined as the number of mitotic divisions that each mother cell can undergo before
In replicative aging, three candidates are considered as important senescence factors. One
of them is extrachromosomal ribosomal DNA circles (ERCs) formed by homologous
recombination between ribosomal DNA repeats [
]. Based on many studies showing the effects of
ERCs level on the replicative age [
], it is believed that the accumulation of ERCs to toxic
levels in mother cells leads to senescence. The other candidates of aging factors are oxidatively
damaged proteins and protein aggregates . Levels of carbonylated proteins produced by
oxidative damage increase with the replicative age of the mother cell [
]. Also, aggregates
composed of oxidatively damaged and misfolded proteins are potentially cytotoxic and associated
with age-related phenotypes . Given that heavily oxidized proteins tend to form protein
], these two aging factors are closely connected with the accumulation of
oxidative damage caused by reactive oxygen species (ROS).
A hallmark of aging is the decline in mitochondrial function [
]. One of the
well-characterized phenomena associated with mitochondrial dysfunction is the buildup of ROS. Even
though some enzymes and processes including membrane-associated NADPH oxidases ,
fatty acid β-oxidation in peroxisomes [
], and the ER protein disulfide resolution system [
contribute to ROS generation, it has been known that mitochondria are the main cellular
source of ROS [
]. Because the accumulation of ROS also leads to the damage to
mitochondrial DNA [
], this vicious cycle has been the basis of the mitochondrial free radical theory
for several decades [
]. However, a recent research reported that the increase in free radical
generation is attributed not to the mitochondrial electron transport chain (ETC) but to the
endoplasmic reticulum (ER)-localized NADPH oxidase Yno1 [
], indicating that the issue of
ROS accumulation in living cells remains complex and multifactorial.
In this study, we found that severe respiratory disturbance shortens yeast RLS by using
several respiratory-deficient mutants. Our results show that the accumulation of ERCs is not the
leading cause of reduced RLS in these mutants. On the other hand, respiratory malfunction
disrupted the maintenance of RLS by inducing an increase in intracellular ROS and oxidized
protein level. Well-known signaling pathways involved in the generation of ROS such as the
protein kinase A (PKA) and target of rapamycin (TOR) pathway were not related to ROS
accumulation in these mutant cells. Instead, the suppression of RAS signaling reduced ROS
production and significantly restored RLS of respiratory-deficient cells. In addition, the majority
of detectable ROS was attributed to the ER-localized NADPH oxidase, Yno1. Based on our
results, we suggest that the reduced yeast RLS due to mitochondrial dysfunction is caused by
the failure to maintain proteostasis.
Materials and methods
Yeast strains and growth media
Yeast strains used in this study are listed in S1 Table. Yeast cells were grown in YPD medium
(1% yeast extract, 2% peptone, and 2% glucose) or synthetic complete (SC) medium (0.67%
yeast nitrogen base without amino acids, 2% glucose, and nutritional supplements) lacking
appropriate amino acids for selection [
]. All cultures were incubated at 30ÊC. Gene
disruption was carried out using the one-step PCR-based gene targeting procedure [
lacking mitochondrial DNA (rho0) were generated by growth in YPD medium supplemented with
ethidium bromide (25 μg/ml) [
]. The respiratory deficiency of the strains was confirmed by
growth on YPG (1% yeast extract, 2% peptone, and 2% glycerol) medium.
Cloning and plasmids
Primer sequences used for plasmid construction are shown in S2 Table. The constitutively
active Ras2 mutant, Ras219V, was obtained by the QuickChange multisite-directed mutagenesis
protocol (Stratagene) and cloned into the XbaI and SalI sites of pRS415GPD vector [
overexpression plasmid of C-terminally TAP-tagged SOD1, pRS426ADH-SOD1-TAP, was
generated as described previously [
]. pRS423CUP1-6xMYC-cki12-200(S125/130A) was kindly
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provided by Dr. Jodi Nunnari [
]. pJU676 (pRS416-SCH9-5HA) has been described
Analysis of RLS was carried out by micromanipulation as described previously [
], using a
Zeiss Tetrad Microscope. All measurements of lifespan were performed on YPD plates. For
the effect of respiratory inhibition, antimycin A (3 μg/ml) or oligomycin (10 μg/ml) was added
to the plate. Lifespan was determined from five independent experiments (approximately 100
cells per strain in total). Cells that never budded were excluded from the calculation. For
statistical analysis, lifespan data sets were compared by one-way ANOVA.
rDNA silencing assay
Silencing at the rDNA region was tested as described previously [
]. Yeast cells were
grown to an OD600 of 0.8, and 2.5 μl of 10-fold serial dilutions of the cell suspensions was
spotted on the appropriate media. Plates were incubated at 30ÊC for two days before visualization.
Quantification of mURA3 mRNA
Total RNA was extracted from yeast cells using the RNeasy Mini Kit (Qiagen). 1 μg of total
RNA was reverse transcribed in a 20 μl reaction mixture containing MLV-reverse transcriptase
(M-biotech) and 0.1 μg of oligo-dT (M-biotech) at 42ÊC for 60 min. The mURA3 silencing
reporter gene harboring the TRP1 promoter instead of the URA3 promoter has been described
]. The amount of mRNA was analyzed by quantitative PCR using the Applied
Biosystems 7300 Real-Time PCR system (Applied Biosystems). Gene expression was
quantified by the 2-ΔΔCT method [
] and ACT1 transcript level was used for normalization of
mURA3 mRNA levels. Primers used for amplification of mURA3 and ACT1 are shown in S3
rDNA recombination assay
The rDNA recombination rate was determined by measuring the frequency of the loss of
ADE2 integrated at the rDNA locus of strain DMY3010 as described previously [
cells grown to an OD600 of ~1.0 in SC medium were spread on SC plates. Colonies were
allowed to grow for two days at 30ÊC and then placed at 4ÊC for two days to enhance color
development. The rDNA recombination rate was calculated by dividing the number of
halfred/half-white colonies by the total number of colonies. Entirely red colonies were excluded
from all calculations. Three independent experiments were performed, and more than 10,000
colonies were examined for each assay. For statistical analysis, data sets were compared by
Measurement of intracellular ROS level
Intracellular ROS levels were detected with H2DCFDA (2',7'-dichlorodihydrofluorescein
diacetate, ThermoFisher Scientific) as described previously [
]. Yeast cells were grown to
saturation in YPD medium and diluted to one-hundredth. Then, H2DCFDA was added to a final
concentration of 10 μg/ml followed by incubation with shaking for one day at 30ÊC.
Intracellular ROS levels were also measured with DHE (dihydroethidium, Sigma-Aldrich) as described
previously with some modifications [
]. Yeast cells were grown to saturation and diluted to
one-hundredth. Cells were incubated with shaking for 22 h at 30ÊC, and then DHE was added
to a final concentration of 2.5 μg/ml, followed by incubation with shaking for 2 h at 30ÊC.
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Fluorescence was analyzed using a BD FACS Canto II flow cytometer (Becton Dickinson). A
baseline of zero (background level of fluorescence) was set based on the maximum value of
control sample without the ROS indicator. Cells with higher ROS level than background were
counted and converted into a percentage.
Measurement of Sch9 phosphorylation
Analysis of phosphorylated Sch9 was conducted by Western blotting as described previously
]. Cells were grown to log phase and trichloroacetic acid was added up to 6%. Samples were
put on ice for at least 5 min, spun down, washed twice with cold acetone, and dried. Cells were
bead-beaten in 100 μl of urea buffer (6 M urea, 50 mM Tris-HCl, pH 7.5, 5 mM EDTA, 1%
SDS, 1 mM phenylmethylsulfonyl fluoride, 5 mM NaF, 5 mM NaN3, 5 mM p-nitrophenyl
phosphate, 5 mM Na2P2O4, and 5 mM β-glycerophosphate) followed by heating for 10 min to
65ÊC. For 2-nitro-5-thiocyanobenzoic acid cleavage, 30 μl of 0.5 M CHES (pH 10.5) and 20 μl
of 2-nitro-5-thiocyanobenzoic acid (7.5 mM in H2O) were added, and samples were incubated
overnight at room temperature before adding 6× sample buffer. Sch9 phosphorylation was
detected by SDS-PAGE and immunoblotting using HRP-conjugated mouse anti-HA antibody
(sc-7392 HRP, Santa Cruz Biotechnology).
Western blot analysis and determination of PKA activity
Cell extracts were prepared by suspending cells in lysis buffer (50 mM Tris-HCl, pH 7.5, 150
mM NaCl, 0.01% NP-40, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 1 mM
benzamidine, 1 μg/ml leupeptin, and 1 μg/ml pepstatin), followed by bead-beating. Extracts were spun
by centrifugation at 1600 g for 10 min at 4ÊC and the supernatant was subjected to SDS-PAGE.
Western blot analysis was performed by standard methods using HRP-conjugated mouse
antiMyc antibody (sc-40 HRP, Santa Cruz Biotechnology) for the detection of Myc-tagged
proteins. Images were captured using a luminescent image analyzer, AE-9150 Ez-Capture II
(ATTO), and the quantification of phosphorylated protein was performed using CS analyzer 3
Measurement of oxidized proteins
The detection of oxidized proteins (protein carbonyls) was performed using Oxidized Protein
Detection Kit (ab178020, Abcam) as described previously [
] with some modifications. Cell
extracts were prepared by suspending cells in lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM
NaCl, 0.01% NP-40, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 1 mM benzamidine,
1 μg/ml leupeptin, and 1 μg/ml pepstatin), followed by bead-beating. Cell lysates were
incubated on ice for 20 min and quantified by Bradford assay. About 30 μg/μl of protein was
derivatized per sample and mixed with 6× sample buffer. Further procedures for detecting oxidized
protein were done by SDS-PAGE and immunoblotting using rabbit anti-DNP antibody and
HRP-conjugated goat anti-rabbit antibody (supplied in Oxidized Protein Detection Kit).
Mitochondrial respiration is required for the maintenance of RLS
To confirm whether mitochondrial respiration regulates replicative aging, we first investigated
RLS of rho0 cells that lack both mitochondrial genome and respiration. Although the loss of
mitochondrial DNA is known to impact the longevity of cells in a strain-specific manner [
RLS of BY4741 rho0 cells decreased by about 40% compared to that of wild-type cells (Fig 1A
and 1B). Since cells that do not carry cytochrome c heme lyase (Cyc3) or cytochrome c oxidase
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AA 3 μg/ml
OM 10 μg/ml
Fig 1. Mitochondrial respiration defect impairs the maintenance of RLS. (A) RLS analysis was performed with wild-type (WT), rho0, cyc3Δ, and shy1Δ cells.
(B) The relative changes in RLS were calculated as the ratio of the mean RLS to that of WT cells in (A). (C) RLS analysis was performed with WT, rho0, and WT
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cells on media containing 3 μg/ml antimycin A (AA) or 10 μg/ml oligomycin (OM). (D) The relative changes in RLS of indicated strains were calculated as the
ratio of the mean RLS to that of WT cells in (C). (E) RLS analysis was performed with WT, rho0, cox5aΔ, and cyc1Δ cells. (F) The relative changes in RLS of
indicated strains were calculated as the ratio of the mean RLS to that of WT cells in (E). Mean RLS values are shown in parentheses. All asterisks indicate
P<0.01, compared with WT cells (one-way ANOVA).
(COX) assembly chaperone (Shy1) have been reported to retain no detectable respiration [
we investigated RLS of cyc3Δ or shy1Δ mutants in order to exclude the possibility that RLS
reduction in rho0 cells might be caused by unknown factors other than respiratory
malfunction. We confirmed respiratory deficiency of these mutants by the severe growth defect on
media containing glycerol, a nonfermentable carbon source (S1 Fig). RLS of each mutant was
similar to that of rho0 cells (Fig 1A and 1B). Interestingly, RLS of wild-type cells was
significantly reduced by the addition of inhibitors such as antimycin A and oligomycin that
specifically block mitochondrial respiration (Fig 1C and 1D). These results suggest that cellular
respiration is important for the maintenance of RLS in yeast.
We also examined RLS of cells with mutations that are associated with mitochondrial
respiration but do not cause severe respiratory failure. These mutants include cells lacking the
subunit Va of COX (Cox5a) or iso-1-cytochrome c (Cyc1) [
]. Although the viability of cox5aΔ
or cyc1Δ cells is reduced in the medium containing glycerol (S1 Fig), both strains showed no
noticeable change in RLS compared to wild-type cells (Fig 1E and 1F). This result suggests that
respiratory failure above a certain threshold value is required to induce a reduction in
rDNA silencing is not a major cause for RLS reduction in
One of the factors that influence yeast longevity is rDNA silencing [
]. To examine whether
respiratory deficiency worsens the stability of rDNA, we carried out an rDNA silencing assay
using the mURA3 silencing reporter gene. For this test, we used yeast strains harboring the
mURA3 silencing reporter gene integrated either into the non-transcribed spacer regions
(NTS1 and NTS2) of the rDNA locus or outside the rDNA array [
]. CYC1, CYC3, COX5A,
or SHY1 genes were deleted from each strain, and cells of each strain were 10-fold serially
diluted and spotted on SC medium in which uracil was omitted or 5-fluoroorotic acid (FOA)
was added. In wild-type cells, the mURA3 reporter gene was effectively silenced at both the
NTS1 and NTS2 regions, as indicated by decreased growth on uracil-deficient medium and
increased growth on medium containing FOA (Fig 2A). Compared to wild-type cells, rho0
cells did not exhibit significant changes in growth on medium lacking uracil or containing
FOA (Fig 2A), suggesting that rDNA silencing has a low correlation with RLS reduction due to
mitochondrial respiratory failure. Likewise, no significant changes in rDNA silencing were
observed in respiratory-deficient cyc3Δ and shy1Δ cells (Fig 2B) or wild-type cells treated with
respiratory inhibitors (Fig 2C). cox5aΔ and cyc1Δ cells with little respiratory defect also did not
show a remarkable difference in rDNA silencing from wild-type cells, except that rDNA
silencing was slightly increased at the NTS1 region in cyc1Δ cells (Fig 2D). Given that RLS of cox5aΔ
and cyc1Δ cells did not differ significantly from that of wild-type cells (Fig 1E and 1F), these
mutations do not seem to be related to rDNA silencing.
To more directly examine rDNA silencing, we measured the transcript levels of the mURA3
gene by using a real-time reverse transcription-PCR analysis as described previously [
wildtype cells, the transcription of the mURA3 gene at the NTS1 and NTS2 regions was effectively
silenced (>50%) compared to that outside rDNA (Fig 2E). The relative transcript levels of mURA3
were not significantly changed in mitochondrial respiratory-deficient rho0, cyc3Δ, and shy1Δ cells
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Fig 2. Transcriptional silencing is not a major cause for shortening of RLS in respiratory-deficient cells. Silencing at the rDNA region was assessed by monitoring
the growth of 10-fold serial dilution of cells on SC media lacking uracil or supplemented with FOA. SC medium was used as a control. (A) 10-fold serial dilutions of
wild-type (WT) and rho0 cells were spotted on SC media without uracil or with FOA. (B) 10-fold serial dilutions of WT, cyc3Δ, and shy1Δ cells were spotted on SC media
without uracil or with FOA. (C) 10-fold serial dilutions of WT cells were spotted on SC media without uracil containing 3 μg/ml antimycin A (AA) or 10 μg/ml
oligomycin (OM). (D) 10-fold serial dilutions of WT, cox5aΔ, and cyc1Δ cells were spotted on SC media without uracil or with FOA. (E) Total RNA was extracted from
WT, rho0, cyc3Δ, shy1Δ, cox5aΔ, cyc1Δ, and sir2Δ cells. Quantitative real-time reverse transcription-PCR analysis was performed to measure the mURA3 transcript level.
Amplification efficiencies were validated and normalized against ACT1. The relative transcript levels of the mURA3 gene were calculated as the ratio of the normalized
transcript levels of the mURA3 gene inside the rDNA array (NTS1::mURA3 and NTS2::mURA3) to that outside the rDNA array (leu2::mURA3). Values represent the
average of three independent experiments, and error bars indicate the standard deviation. (F) rDNA recombination assay was performed to check rDNA stability of the
indicated cells. rDNA recombination is represented by the frequency of loss of the ADE2 marker gene integrated at the rDNA locus in the corresponding cells. Values
represent the average of three independent experiments, and error bars indicate the standard deviation. Asterisks indicate P<0.01, compared with WT cells (one-way
showing reduced RLS, compared to that of wild-type cells. The transcription of mURA3 was also
not significantly changed in cox5aΔ and cyc1Δ cells that do not show RLS reduction.
To further test rDNA stability in respiratory-deficient cells, the frequency of loss of the
ADE2 marker gene integrated at the rDNA locus was monitored. As reported previously [
], mutant cells without Sir2, a major rDNA silencing factor, exhibited a considerable
increase in the frequency of ADE2 marker loss compared to wild-type cells (Fig 2F). However,
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in agreement with the above observations that rDNA silencing is unrelated to mitochondrial
respiratory defect, the frequency of ADE2 marker loss in respiratory-deficient cells was not
significantly different from that of wild-type cells. Taken together, these results suggest that
rDNA silencing, a well-known longevity factor in yeast, is not related to RLS reduction caused
by mitochondrial respiratory failure.
Mitochondrial respiratory deficiency induces an increase in the amount of
intracellular ROS regardless of TOR and PKA pathways
Given that the accumulation of ROS contributes to the buildup of other aging factors such as
oxidatively damaged proteins and protein aggregates [
], we examined whether defective
respiration induces intracellular ROS production. To detect ROS accumulation, we employed
flow cytometry using H2DCFDA, a fluorescent probe that reacts with several ROS such as
hydroxyl radicals and H2O2 [
]. Cell population (P2) showing higher fluorescence than
background level was converted into a percentage (S2A Fig). In wild-type cells, about 14% of cell
population emitted higher fluorescence than background level (Fig 3A and 3B). Remarkably,
respiratory-deficient rho0, cyc3Δ, and shy1Δ cells showed 4~5 fold increase in P2 percentage. It
was also found that the level of intracellular ROS increased significantly when wild-type cells
were treated with antimycin A or oligomycin. In contrast, cox5aΔ and cyc1Δ cells, which
showed no decrease in RLS, did not exhibit a significant change in P2 percentage. In addition,
we employed another fluorescent probe DHE to detect intracellular superoxide radical. Similar
to the above results with H2DCFDA, cells with a severe defect in respiration showed increased
percentages of the high-ROS cell population, whereas a significant change was not observed in
cox5aΔ and cyc1Δ cells (S2B±S2D Fig). These observations suggest that the reduction of RLS in
respiratory-deficient mutants may rely on the elevated level of intracellular ROS.
To test a possibility that elevated ROS production might increase the level of oxidatively
damaged proteins in cells with respiratory failure, we analyzed protein oxidation by using the
oxyblot assay. When proteins are oxidized by ROS, carbonyl groups such as aldehydes and
ketones are produced on protein side chains [
]. These groups can be derivatized with
2,4-dinitrophenylhydrazine to form a stable 2,4-dinitrophenyl (DNP) hydrazone product and
be measured with the anti-DNP antibody by western blot immunoassay. Consistent with the
above results that a severe defect in mitochondrial respiration induces ROS accumulation, the
levels of protein oxidation in respiratory-deficient rho0, cyc3Δ, and shy1Δ cells were about two
times higher than that in wild-type cells (Fig 3C and S3A Fig). Treatment of antimycin A and
oligomycin mimicking respiratory failure also increased intracellular protein oxidation. In
case of cox5aΔ and cyc1Δ cells, however, the oxidized protein level was not significantly
different from that of wild-type cells. These results support our hypothesis that increased ROS by
mitochondrial respiratory deficiency induces protein oxidation and thus affects RLS.
It is known that the activation of TOR and PKA pathways leads to the accumulation of ROS
in yeast [
]. We examined whether these two pathways are involved in the elevation of
ROS level due to respiratory defect. Given that Sch9 is a major substrate of TOR kinase, Sch9
phosphorylation was quantitatively analyzed using SDS-PAGE mobility shift assay in order to
measure the activity of TOR kinase . As expected, the up-shifted, phosphorylated forms of
Sch9 were observed in wild-type cells (Fig 3D and S3B Fig). Consistent with previous reports
that nitrogen starvation inhibits the activity of TOR kinase [
], the level of phosphorylated
Sch9 was reduced under nitrogen starvation. Notably, a significant change in Sch9
phosphorylation was not observed in respiratory-deficient rho0, cyc3Δ, and shy1Δ cells or in cells treated
with respiratory inhibitors. This observation suggests that TOR pathway is not involved in the
elevation of ROS level caused by respiratory failure.
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DCF fluorescence intensity
WT rho0 cyc3∆ shy1∆
rho0 cyc3∆ shy1∆
OM cox5a∆ cyc1∆
OM cox5a∆ cyc1∆
Fig 3. Increased ROS level decreases RLS in respiratory-deficient cells. (A) Intracellular ROS levels in wild-type (WT),
rho0, cyc3Δ, shy1Δ, cox5aΔ, cyc1Δ, and WT cells treated with 3 μg/ml antimycin A (AA) or 10 μg/ml oligomycin (OM) were
detected with H2DCFDA. Fluorescence was analyzed using a BD FACS Canto II flow cytometer. (B) Cells with high ROS
were calculated as a percentage of cells with higher fluorescence intensity than the maximum fluorescence intensity of
control sample without the ROS indicator. Values represent the average of three independent experiments, and error bars
indicate the standard deviation. All asterisks indicate P<0.01, compared with WT cells (one-way ANOVA). (C)
Carbonylated proteins in WT, rho0, cyc3Δ, shy1Δ, cox5aΔ, cyc1Δ, and WT cells treated with 3 μg/ml AA or 10 μg/ml OM
were detected using Oxidized Protein Detection kit. Hexokinase was used as a loading control. The relative ratio of
carbonylated proteins in the indicated strain to those of WT cells is shown below each lane. Data are representative of at
least three independent experiments. (D) Total protein was extracted from WT, rho0, cyc3Δ, shy1Δ, and WT cells treated
with 3 μg/ml AA or 10 μg/ml OM, and WT cells under nitrogen starvation. All cells harbor pRS416-SCH9T570A-5HA.
Immunoblotting was performed using a mouse anti-HA antibody. Data are representative of at least three independent
experiments. (E) Total protein was extracted from WT, rho0, cyc3Δ, shy1Δ, WT cells treated with 3 μg/ml AA or 10 μg/ml
OM, WT cells expressing constitutively active RAS2val19 (19V), and WT cells under glucose starvation. All cells harbor
pRS423-CUP1-6xMYC-cki12-200(S125/130A). Immunoblotting was performed using a mouse anti-Myc antibody. The relative
ratio of phosphorylated to unphosphorylated forms of Cki1 is shown below each lane. Data are representative of at least
three independent experiments.
For the determination of PKA activity, we used a PKA substrate reporter derived from a
native substrate Cki1 [
]. By analyzing the mobility shift on SDS-PAGE, PKA-dependent
phosphorylation of the Cki1 reporter was detected and the ratio of phosphorylated and
unphosphorylated forms was calculated. As expected, cells with a constitutively active variant of
] exhibited a considerable increase in the phosphorylated form of Cki1, while a
nearly 90% reduction in the phosphorylated form of Cki1 was observed in cells under glucose
starvation (Fig 3E and S3C Fig). However, we could not observe a significant change in Cki1
phosphorylation not only in respiratory-deficient rho0, cyc3Δ, and shy1Δ cells but also in cells
treated with respiratory inhibitors. This result suggests that, like TOR pathway, PKA pathway
is not related to ROS accumulation induced by mitochondrial respiratory deficiency.
RAS signaling and NADPH oxidase Yno1 contribute to intracellular ROS
Above, we have shown that PKA activity does not contribute to the elevation of ROS level in
respiratory-deficient cells. It is well established that Ras2 activates adenylyl cyclase depending
on the levels of glucose and regulates the activation of PKA by controlling cAMP levels in yeast
]. However, a recent study reported that RAS signaling operates independently of PKA to
promote ROS accumulation in cells lacking COX activity [
]. To test whether RAS signaling
is responsible for ROS accumulation in respiratory-deficient cells, we measured intracellular
ROS level in RAS2-deleted rho0 cells. Interestingly, the loss of Ras2 resulted in a significant
reduction of ROS level in rho0 cells (Fig 4A, S4A and S5A and S5B Figs). RAS2 deletion also
led to a significant recovery in RLS of rho0 cells (Fig 4B and 4C). Consistent with our notion
that ROS accumulation impairs the maintenance of RLS by promoting protein oxidation, the
level of oxidized proteins was lowered by about 60% in RAS2-deleted rho0 cells compared to
rho0 cells (Fig 4D and S6A Fig). These results suggest that RAS signaling contributes to ROS
accumulation and concomitant protein oxidation induced by mitochondrial dysfunction.
Intriguingly, although ras2Δ cells have quite low ROS level compared to wild-type cells,
RLS of ras2Δ cells was not increased but even slightly decreased (Fig 4B and 4C). Ras2 has
been known not only as a key component of generating ROS in respiratory-deficient cells but
also as a major regulator responsive to external environment [
]. One of the downstream
effectors of Ras2 is PKA which plays crucial roles in a wide variety of cellular processes [
Moreover, there is a complex network that covers multiple regulatory pathways in the
downstream of Ras2 [
]. Therefore, even though ras2Δ cells have lower ROS level than wild-type
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Fig 4. RAS signaling pathway and Yno1 contribute to the buildup of intracellular ROS in cells lacking mitochondrial respiration. (A, E, and H) Intracellular ROS
levels in the indicated strains were detected with H2DCFDA. Fluorescence was analyzed using a BD FACS Canto II flow cytometer. Cells with high ROS were calculated
as a percentage of cells with higher fluorescence intensity than the maximum fluorescence intensity of control sample without the ROS indicator. Values represent the
average of three independent experiments, and error bars indicate the standard deviation. All asterisks indicate P<0.01, compared with rho0 cells (one-way ANOVA).
(B, F, and I) RLS analysis was performed with the indicated strains. (C, G, and J) The relative changes in RLS were calculated as the ratio of the mean RLS to that of WT
cells. Mean RLS values are shown in parentheses. All asterisks indicate P<0.01, compared with rho0 cells (one-way ANOVA). (D and K) Carbonylated proteins in
indicated strains were detected using Oxidized Protein Detection kit. Hexokinase was used as a loading control. The relative ratio of carbonylated proteins in the
indicated strain to those of WT cells is shown below each lane. Data are representative of at least three independent experiments.
cells, ras2Δ cells seem to exhibit reduced RLS compared to wild-type cells because of defects in
several regulatory pathways important for normal cell function. It is presumable that the effect
of defective cell function overrides the beneficial effect of reduced ROS in ras2Δ cells.
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RAS signaling has been involved in the regulation of the endoplasmic reticulum-associated
degradation (ERAD) pathway which plays an important role in ER quality control
]. According to a previous study, among other ER-associated oxidases under the
control of the ERAD pathway, ER-localized NADPH oxidase Yno1 is required for ROS generation
in COX-deficient cells [
]. Based on these reports, we next examined whether Yno1 plays a
role in the buildup of ROS in cells with respiratory malfunction. Notably, YNO1 deletion led to
about 50% decrease in ROS accumulation in rho0 cells (Fig 4E, S4B, S5A and S5B Figs). RLS of
yno1Δ rho0 cells was also significantly restored compared to that of rho0 cells (Fig 4F and 4G).
Furthermore, we observed a considerable reduction in the level of protein oxidization in
YNO1-deleted rho0 cells compared to rho0 cells (Fig 4D and S6A Fig), suggesting that Yno1 is
involved in ROS accumulation and protein oxidation in respiratory-deficient cells.
Next, we checked whether RAS signaling and Yno1 contribute independently to ROS
accumulation and concomitant RLS reduction in rho0 cells. To test this, we measured intracellular
ROS level and RLS of ras2Δ yno1Δ rho0 cells. With respect to ROS reduction, no synergistic
effect was observed in ras2Δ yno1Δ rho0 cells compared to ras2Δ rho0 or yno1Δ rho0 cells (Fig
4H, S4C, S5A and S5B Figs). In addition, we could not observe the synergistic effect of deletion
of RAS2 and YNO1 on RLS of rho0 cells (Fig 4I and 4J). In accordance with the above
observations, the synergistic effect of deletion of RAS2 and YNO1 on protein oxidation of rho0 cells
was not detected (Fig 4K and S6B Fig). These results suggest that RAS signaling and Yno1 act
on the same pathway in the buildup of ROS and consequent reduction of RLS induced by
mitochondrial dysfunction. These results are also consistent with a recent report that RAS
signaling is associated with degradation of Yno1 through the ERAD pathway [
Although not yet clearly identified, mitochondria are believed to play important roles in
cellular aging. In this study, we used the budding yeast S. cerevisiae to investigate the effects of
mitochondrial respiratory deficiency on replicative aging. Through analysis of RLS in
respiratorydeficient mutants, we found that mitochondrial respiration is required for the maintenance of
RLS (Fig 1). Our data also show that rDNA silencing, one of the major aging factors, is not
relevant to RLS reduction induced by defective mitochondria (Fig 2). Instead, we suggest that the
elevated ROS levels triggered by mitochondrial malfunction are responsible for the reduction
of RLS by impairing proteostasis (Fig 3A, 3B and 3C). Given that mitochondrial respiration is
also essential to sustain yeast CLS [
], our results highlight that the state of mitochondrial
respiration is widely involved in the longevity process.
In contrast to our results, some studies reported that RLS of mitochondrial
respiratory-deficient cells is maintained or even increased depending on laboratory strains used [
37, 53, 54
Nonetheless, the accumulation of ROS is observed in respiratory-deficient cells derived from
several laboratory strains and this phenomenon is also conserved in higher eukaryotes [
]. Previous studies have found that cells lacking mitochondrial respiration generally have
low viability due to hypersensitivity to various stresses such as hydrogen peroxide, ethanol,
and heat [
38, 56, 57
]. Furthermore, the tolerance for intracellular ROS varies depending on the
difference of genetic background among the laboratory strains . Taking all of the above
into consideration, it is plausible that RLS of respiratory-deficient mutants is influenced by the
resistance to accumulated ROS, which is different among the laboratory strains. Meanwhile, it
has been reported that a mild inhibition of mitochondrial respiration prolongs the lifespan of
several organisms [37, 59±62]. Consistent with these reports, although cox5aΔ and cyc1Δ cells
had some defects in respiration (S1 Fig), these cells did not exhibit RLS reduction nor ROS
accumulation (Figs 1C and 3). In addition, thresholds of mitochondrial respiration are
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necessary to regulate yeast CLS [
]. Therefore, the degree of respiratory capacity seems to be
another determining factor for lifespan regulation in cells with respiratory defects.
We observed that mitochondrial dysfunction has no significant effect on the activation of
TOR and PKA kinases (Fig 3D and 3E). On the basis of these results, we suggest that the
buildup of ROS in respiratory-deficient cells is not attributed to TOR and PKA pathways.
Instead, it is plausible that the RAS signaling pathway regulates the activity of NADPH oxidase
Yno1 responsible for ROS overproduction in respiratory-deficient cells (Fig 4). Although
Yno1 is known to be regulated by ERAD-mediated degradation under the control of RAS
], the underlying mechanism is expected to be more complicated. It has been
reported that the activity of RAS is involved in actin dynamics and remodeling upon nutrient
]. In addition, previous genome-wide screening studies reported that yno1Δ
cells show a hypersensitivity to inhibitors of the actin cytoskeleton and that Yno1 functionally
regulates the nucleation and elongation step in the biosynthesis of F-actin cables [
Taken together, these findings suggest that the interactive regulation between RAS signaling
and Yno1 can be linked through actin dynamics. Meanwhile, the mitochondrial retrograde
pathway is known to regulate many cellular activities and aging in yeast . Given that the
retrograde response has crosstalk with Ras2 [
], it will be of interest to investigate whether
the retrograde signaling links RAS signaling and Yno1.
Our findings suggest that a severe defect in mitochondrial respiration impairs the
maintenance of RLS by the accumulation of intracellular ROS rather than the loss of rDNA silencing.
In addition, we suggest that the NADPH oxidase Yno1 significantly contributes to replicative
aging by regulating ROS production in cells lacking respiratory activity along with RAS
signaling. This study highlights the complex and multifaceted effects of ROS accumulation in yeast,
which are relevant to the physiology of aging of higher organisms. Given that dysfunctional
mitochondria are known to be connected with age-related metabolic and degenerative
diseases, our results also provide implications for the mechanisms underlying cancerogenesis,
neurodegenerative disease, diabetes, and obesity.
S1 Fig. Comparative analysis of the mitochondrial respiration capacity in used strains.
10-fold serial dilutions of wild-type (WT), rho0, cyc3Δ, shy1Δ, cox5aΔ, and cyc1Δ cells were
spotted on YPG medium. The respiratory capacity of the indicated strains was assessed by
monitoring the growth of 10-fold serial dilution of cells on YPG media. YPD medium was
used as a control.
S2 Fig. Determination of intracellular ROS level by flow cytometry. (A and B) Wild-type
(WT) cells were grown and their fluorescence was analyzed without the indicated ROS probes.
Based on this, the fluorescence output was set to zero. Any cells that have a value above zero
were counted as P2. (C) Intracellular ROS levels in the indicated strains were detected with
DHE. (D) Cells with high ROS were calculated as a percentage of cells with higher fluorescence
intensity than the maximum fluorescence intensity of control sample without the ROS
indicator. Values represent the average of three independent experiments, and error bars indicate
the standard deviation. All asterisks indicate P<0.01, compared with WT cells (one-way
S3 Fig. Quantification of protein oxidation, TOR, and PKA activity. (A) Carbonylated
proteins in wild-type (WT), rho0, cyc3Δ, shy1Δ, cox5aΔ, cyc1Δ, and WT cells treated with 3 μg/ml
13 / 18
AA or 10 μg/ml OM were detected using Oxidized Protein Detection kit. The relative change
in protein oxidation was calculated as the ratio of carbonylated proteins in the indicated strain
to those of WT cells. (B) Total protein was extracted from WT, rho0, cyc3Δ, shy1Δ, WT cells
treated with 3 μg/ml AA or 10 μg/ml OM, and WT cells under nitrogen starvation. All cells
harbor pRS416-SCH9T570A-5HA. Immunoblotting was performed using a mouse anti-HA
antibody. Then the relative ratio of phosphorylated to unphosphorylated forms of Sch9 was
calculated. (C) Total protein was extracted from WT, rho0, cyc3Δ, shy1Δ, WT cells treated with
3 μg/ml AA or 10 μg/ml OM, WT cells expressing constitutively active RAS2val19 (19V), and
WT cells under glucose starvation. All cells harbor pRS423-CUP1-6xMYC-cki12-200(S125/130A).
Immunoblotting was performed using a mouse anti-Myc antibody. Then the relative ratio of
phosphorylated to unphosphorylated forms of Cki1 was calculated. All values represent the
average of three independent experiments, and error bars indicate the standard deviation. All
asterisks indicate P<0.01, compared with WT cells (one-way ANOVA).
S4 Fig. Determination of intracellular ROS levels affected by RAS and Yno1 in rho0 cells
using H2DCFDA. (A, B, and C) Intracellular ROS levels in the indicated strains were detected
with H2DCFDA. Fluorescence was analyzed using a BD FACS Canto II flow cytometer.
S5 Fig. Determination of intracellular ROS levels affected by RAS and Yno1 in rho0 cells
using DHE. (A) Intracellular ROS levels in the indicated strains were detected with DHE.
Fluorescence was analyzed using a BD FACS Canto II flow cytometer. (B) Cells with high ROS
were calculated as a percentage of cells with higher fluorescence intensity than the maximum
fluorescence intensity of control sample without the ROS indicator. Values represent the
average of three independent experiments, and error bars indicate the standard deviation. All
asterisks indicate P<0.01, compared with rho0 cells (one-way ANOVA).
S6 Fig. Quantification of protein oxidation affected by RAS and Yno1 in rho0 cells. (A and
B) Carbonylated proteins in the indicated strains were detected using Oxidized Protein
Detection kit. Then the relative change in protein oxidation was calculated as the ratio of
carbonylated proteins in the indicated strain to those of WT cells. Values represent the average of three
independent experiments, and error bars indicate the standard deviation. All asterisks indicate
P<0.01, compared with rho0 cells (one-way ANOVA).
S1 Table. Yeast strains used in this study.
S2 Table. Primers used for plasmid construction.
S3 Table. Primers used for quantitative PCR.
This work was supported by the National Research Foundation of Korea grant
(2015R1A2A1A01007871) funded by the Ministry of Education, Science and Technology,
Republic of Korea.
14 / 18
Conceptualization: Dae-Gwan Yi, Won-Ki Huh.
Data curation: Dae-Gwan Yi, Sujin Hong.
Formal analysis: Dae-Gwan Yi, Sujin Hong.
Funding acquisition: Won-Ki Huh.
Investigation: Dae-Gwan Yi, Sujin Hong, Won-Ki Huh.
Methodology: Dae-Gwan Yi, Sujin Hong.
Project administration: Won-Ki Huh.
Resources: Won-Ki Huh.
Supervision: Won-Ki Huh.
Validation: Dae-Gwan Yi, Sujin Hong.
Writing ± original draft: Dae-Gwan Yi, Sujin Hong, Won-Ki Huh.
Writing ± review & editing: Won-Ki Huh.
15 / 18
16 / 18
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