How does DNA damage cause ageing in yeast?

How does DNA damage cause ageing in yeast?

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As I understand it, in yeast ageing there is daughter cell and mother cell. The daughter cell is has newly "fresh" DNA and mother cell dies after some counts of replication.

What happens to the accumulated DNA damage in the yeast mother cell?

One of theory of human ageing is accumulation of damage in nuclear DNA. The daughter yeast cell is clone of mother, and has same DNA. What happens with this damage?

Edit to simplify: Why is the DNA damage not transferred at all to the daughter cell? The repair mechanism in humans is evidently not perfect, because DNA damage is one of major ageing theories. So if we die from this, how do yeast get around the detrimental affects of DNA damage that cause ageing in humans?

In eukaryotic cells there is no difference between a mother and a daughter cell - the later is an exact copy of the mother cell. This is true for yeasts as well for example for human cells. The only thing that happens over time is that the telomeres at the end of the chromosomes get shorter (unless the cell has an active telomerase which most cells doesn't) and it reaches the Hayflick limit where every further division will lead to a shortage of coding chromosmal regions and thus cause problems. If cells do not acquire any other chromosomal damage and have enough ressources, this would be the limit for their lifetime.

Of course there are DNA damages occuring over time. Most of them are either repaired by the highly efficient DNA damage repair system of eukaryotic cells, which consists of different checkpoints during the cell cycle. These checkpoints are located at transition between the different phases of the cell cycle. If a DNA damage is identified, the cell cycle is stopped to enable a DNA damage repair by special proteins and to prevent that damaged cells undergo mitosis (here the G2-M checkpoint is important). This works schematically like in this figure (from the Wikipedia article on DNA repair):

If the DNA damage cannot be resolved, cells will either go into senescence (basically go into a permanent dormant state) or undergo apoptosis. If there are damages which are (for whatever reasons) are either not spotted or occur during the repair of DNA double strand breaks by non-homologous end joining, these are then replicated and inherited by further generations. Unless these errors occur during the replication process mother and daughter cells are genetically identical.

So eukaryotic cells acquire DNA damage over time (one of the reasons why late pregnancies pose a greater risk than early), but we are different from yeast in one point: Yeast has an active telomerase, which is not active in most of our cells (mainly except in stem cells). If the cells collect too much chromosmal damages than they will die.

The references listed below go deeper into the topic and are interesting to read.


  1. Negative regulation of yeast telomerase activity through an interaction with an upstream region of the DNA primer
  2. Cell cycle control in the kidney.
  3. The evolution of diverse biological responses to DNA damage: insights from yeast and p53
  4. DNA damage and decisions: CtIP coordinates DNA repair and cell cycle checkpoints
  5. Cellular Responses to DNA Damage: One Signal, Multiple Choices

Background to the different theories of ageing.

This video, from a senior lecturer at the University of Liverpool who specialises in ageing, discusses the theories of ageing. He touches on the DNA damage theory.

DNA damage theory of ageing.

Note that when talking about DNA damage theory, we are specifically talking about damage to the process of cell renewal by DNA damage repair mechanisms in stem cells.

Mutation ageing is not the same as DNA damage causing ageing and cannot really cause ageing in yeast. The idea that cells acquire damage over time due to mutated DNA is fundamentally different to the DNA damage theory of ageing. The mechanisms that lead to DNA damage are covered very well in @Chris' answer.

The suggested systemic effect that DNA damage has in the human body causes an inability to renew cells due to a lack of viable stem cells. This contributes to the process of ageing in the organism.

In yeast, this stem cell depletion process does not happen (they are single cellular mostly). In the comments it appears that you are actually talking about mutation based ageing in your original question. Mutations are passed to daughter cells. But these daughter cells don't appear to age or to die quicker (If the parental cell is 5 days old, the daughter cell does not appear to be 5 days old). This is the fundamental reason why this form of ageing remains less studied - it doesn't really make sense that general mutations cause ageing.


To directly answer the question: "How does DNA damage cause ageing in yeast?"

The question doesn't quite capture the whole picture and confuses DNA damage theory of animal ageing with general DNA damage at a cellular level.

There are many theories of ageing. Nobody knows which ones have more or less influence on ageing. Yeast is a good model for some kinds of age theory modelling, but perhaps not in the case of DNA damage based ageing. Although they experience DNA damage in the form of mutation, this doesn't cause ageing in the same way that DNA damage might cause ageing in humans because they are single celled organisms, and don't experience the same inter-tissue relationships that more complex organisms have.


This answer has been changed extensively since its first draft based on additional questions and queries from the OP in the comments. I decided to keep some important points that were raised and dismissed.

Why there is no DNA damage transfer at all, to daughter cell?

DNA damage can occur and is transferred to the daughter cell. Damage is very rare though.

So how yeast survive? Human stem cells aging and die, or get cancer. But yeast no. Human start from one cell. And die. Yeast start from one cell and live forever (dividing again and again)

Edit After Comments:

Single cell

In yeast there are no inter-tissue systems. Mutated damage is simply passed on if the cell is viable. Also the assumption that they live forever isn't entirely accurate. But the DNA is passed on in its mutated or damaged form.

Multi cell

DNA damage based ageing is a big problem for animals. (Dolle et al 2013) Showed that DNA damage to a single repair pathway causes accelerated ageing, (but note that there was not an increase in mutation during this ageing).

but you talking about cancer, it's not so good example, my mistake. But in humans aging NOT like cancer - it occur in ALL the body and all the cells. So in all cells there are some DNA damage.

Another edit: You were right to talk about cancer in this context. The DNA damage theory of ageing is that small changes (damage) in our DNA accumulate over time and damage cell types by damaging various DNA repair pathways.

Damage accumulates and over time less stem cells for renewal are viable and cellular repair of other damaged cells slows down.

Also this same fundamental principle of mutation damage happening over time can lead to mutations that switch off cell control mechanisms and lead to cancer. Although these these mutations do not increase in rate with age (Hill et al., 2005).

For example the naked mole rat lives 10 times that of comparable species (less ageing) and have not been observed to naturally acquire cancer (less cancer). The theory implies that the two things are quite linked because of the way DNA changes, but not that specifically mutation is a cause or result of ageing.

I talking more about aging. Because there is aging in yeast. And one of aging theories for human - accumulating DNA damage. So what Chris writing is that it's really rare.

**Stem cells are damaged. ** I'm no longer entirely sure of what you mean by ageing in yeast. Yeast aren't reliant on a limited number of stem cells for cellular renewal, they don't age in the way you might be thinking. They simply proliferate the mutated cell type. There is no impact to a larger organism like in human ageing.

DNA damage. Yeast is a very helpful organism for understanding the underlying biochemistry and genetics of ageing, not the overall ageing systems. These are two different fields of study. It is unclear which you are talking about at this point.

Also there is aging in places where no stem cells - like brain - no renew of cells, just existing cells age, damaged, die, and not replaced.

Dolle 2006 found that spontaneous DNA damage in relation to age happened in the liver but not the brain.

Oxidative stress and aging: Learning from yeast lessons

The yeast Saccharomyces cerevisiae has played a vital role in the understanding of the molecular basis of aging and the relationship of aging process with oxidative stress (non-homeostatic accumulation of Reactive Oxygen Species, ROS). The mammalian and yeast antioxidant responses are similar and over 25 % of human-degenerative disease related genes have close homologues in yeast. The reduced genetic redundancy of yeast facilitates visualization of the effect of a deleted or mutated gene. By manipulating growth conditions, yeast cells can survive only fermenting (low ROS levels) or respiring (increased ROS levels), which facilitates the elucidation of the mechanisms involved with acquisition of tolerance to oxidative stress. Furthermore, the yeast databases are the most complete of all eukaryotic models. In this work, we highlight the value of S. cerevisiae as a model to investigate the oxidative stress response and its potential impact on aging and age-related diseases.


Stability of chromosome ends in yeast cells is maintained by a telomere-specific DNA–protein complex (Lydall, 2003). A key component of this telomere cap is a single-stranded DNA-binding protein, Cdc13p (Nugent et al., 1996). In a cdc13-1 mutant incubated at the nonpermissive temperature, the uncapped telomeres are recognized as double strand breaks, leading to accumulation of single-stranded DNA at the ends of chromosomes (Garvik et al., 1995 Maringele and Lydall, 2002), inducing a DNA-damage checkpoint signal (Garvik et al., 1995) and promoting a terminal cell cycle arrest at G2/M (Weinert and Hartwell, 1993). When these cells are returned to permissive temperature after prolonged arrest, only a small proportion recover. This cell death has been attributed to loss of essential genes located in the subtelomeric regions (Garvik et al., 1995 Booth et al., 2001), conversion of single-stranded DNA at telomeres to unrepaired lethal lesions (Jia et al., 2004), and/or the formation of chromosome end-to-end fusions (DuBois et al., 2002).

Recently, Qi et al. (2003) have demonstrated that dying cdc13-1 cells display phenotypic markers of apoptosis such as exposure of phosphatidylserine on the outer leaflet of the plasma membrane, accumulation of reactive oxygen species (ROS), and induction of caspase activity based on retention of the caspase inhibitor FITC-VAD-FMK. Binding of the valyl-alanyl-aspartyl (VAD) sequence to an activated mammalian caspase allows the fluoromethyl ketone (FMK) moiety to react with the active site cysteine, resulting in inactivation and fluorescent labeling of the protein (Grabarek and Darzynkiewicz, 2002). Yeast express a single caspase-like protease, Yca1p. Like mammalian caspases, Yca1p undergoes cleavage, which activates its proteolytic activity toward several synthetic caspase substrates in vitro (Madeo et al., 2002a). Yca1p activity can be inhibited by Z-VAD-FMK (Madeo et al., 2002a). Uncleaved FITC-VAD-FMK can be washed out readily from viable cells, but dead cells retain FITC and can be detected by green fluorescence in flow cytometry. Thus, in vivo labeling of yeast with FITC-VAD-FMK has been attributed to bona fide caspase activation (Madeo et al., 2002a Qi et al., 2003 Herker et al., 2004 Wadskog et al., 2004). Deletion of MEC1, a yeast ATM/ATR kinase homologue, prevented this cdc13-1 apoptosis, suggesting that unprotected telomeres generate a MEC1-dependent apoptotic death signal (Qi et al., 2003).

In this report, we show that cell death induced by the inactivation of Cdc13p is not dependent on the caspase-like protease Yca1 or increased ROS production. We further demonstrate that flow cytometric measurements of caspase activity and ROS production, using fluorochrome-conjugated caspase inhibitors and dihydrorhodamine 123 (DHR123), respectively, is confounded by binding to already-dead yeast cells. This work suggests that flow cytometric analysis of dying yeast cells is subject to significant artifacts and indicates a need to reinterpret prior results on mechanisms of yeast cell death.


Yeast Strains and Media

A wild-type yeast strain (4741) and an isogenic DNA replication mutant (4741dna2-1) ( Kuo and Campbell, 1983 ) were used for all experiments. Liquid YPD media (2% glucose, 1% yeast extract, 2% peptone) was used for growing yeast cells at 24ଌ, the permissive growth temperature for the dna2-1 strain.

Isolation of Young and Old Cells by Elutriation

For the isolation of young and old cells, the WT4741 strain and the isogenic strain containing the dna2-1 mutation, 4741dna2-1, were diluted appropriately and grown at 24ଌ in two liters YPD containing 35 μg/ml chloramphenicol. Wild-type cells, WT4741, were first grown for � generations (A600 = 10). Then 3 × 10 8 10-generation-old cells were isolated by elutriation and then inoculated into two liters fresh YPD medium and grown again for 𢏈 generations (A600 = 1.5). Both young (1 to 3 generations) and old cells (16-18 generations) were isolated by a second elutriation, yielding 2 × 10 9 young cells and 4.8 × 10 8 old cells. Cells were quick frozen in liquid nitrogen and stored at -80ଌ. Note that RNA extractions were not performed on the 1- to 3-generation-old and 10-generation-old wild-type cells harvested at A600 = 10. The latter culture of wild-type cells was grown to such a high density to avoid doing three elutriations for each culture, thus minimizing the preparation time. The young wild-type cells and the 18-generation-old wild-type cells used for RNA extraction were harvested well below the diauxie limit based on the density of the culture at the time of collection (σ.2) ( DeRisi et al., 1997 ).

For isolation of young and old dna2-1 mutant cells, 5 × 10 8 4741dna2-1 cells growing exponentially were inoculated into two liters of YPD containing 35 μg/ml chloramphenicol and grown for 𢏈 generations at 24ଌ (A600 = 1.5-2). Young and old 4741dna2-1 cells were separated by elutriation. We collected 1.2 × 10 9 1- to 3-generation-old cells, and the yield of 8-generation-old cells was 3.6 × 10 8 . Again, care was taken to harvest well before the diauxic shift. Samples of elutriated young and old cells were quick frozen in liquid nitrogen and stored at -80ଌ.

A complete description of the elutriation technique can be found in ( Diamond, 1991 ). Briefly, pellets of cells harvested from the exponentially growing cultures were washed in phosphate-buffered saline (PBS 136.89 mM NaCl, 2.68 mM KCl, 5.37 mM Na2HPO4, 1.76 mM KH2PO4, pH 7.4), and the cells resuspended in 20 ml of PBS and bovine serum albumin (2 mg/ml). Cell clumps were separated by sonic irradiation (30 s), and the cell suspension was filtered with mesh (3-64/32 NITEX Tetko, Briarcliff Manor, N.Y.). The elutriation chamber was then loaded with 20 ml of cell suspension corresponding to 𢏃 × 10 11 cells (2 liters of culture, A600 = 10, first elutriation) for WT4741 and 𢏇 × 10 10 cells for 4741 dna2-1 (2 liters of culture, A600 = 2). The young wild-type and dna2-1 cells were isolated by fixing the flow rate of the water injected into the centrifuge at 45 ml/min and the speed of the rotor of the centrifuge at 1400 rpm. The old wild-type and dna2-1 cells were elutriated by fixing the flow rate at 85 ml/min and the speed of the rotor, respectively, at 900 rpm and 1550 rpm (J-6M centrifuge with JE-10 × rotor Beckman Coulter, Fullerton, CA). The preparation of the cells before loading into the elutriator took 𢏁 h. Once loaded, the young cells were isolated within 30 min at 24ଌ and then placed at 4ଌ. It took 𢏃 h to isolate the fractions of old cells. As for the young cells, we performed the elutriation of the old cells at 24ଌ, and the cells were then placed at 4ଌ. Flow cytometric analysis revealed that the cell cycle distribution was the same for elutriated and nonelutriated cells.

Calcofluor Staining

We stained the elutriated young and old cells with fluorescent brightener (Calcofluor White M2R Tinopal UNPA-GX #F3543) to count the bud scars and establish the age of the cells isolated by elutriation. About 10 6 cells were washed once in 150 μl of 1 M sorbitol and resuspended in 150 μl of fluorescent brightener (10 mg/ml) for 5 min at 4ଌ. After being washed four times with fresh 1 M sorbitol, the cells were resuspended in 20 μl of 1 M sorbitol and then observed under a fluorescence axioscope (Carl Zeiss, Jena, Germany).

Indirect Immunofluorescence

Nucleoli were monitored by indirect immunofluorescence by using monoclonal antibodies directed against the abundant nucleolar protein Nop1 ( Aris and Blobel, 1988 Mays Hoopes et al., 2002 ). After 1-h incubation with potassium phosphate buffer (KPO4), magnesium chloride (MgCl2), and 37% formaldehyde, cells were placed in 2 M sorbitol buffer and treated with Zymolyase (0.3 mg/ml Zymolyase 20T, 0.1% β-mercaptoethanol) at 30ଌ for 45 min for the WT4741 strain and 15 min for the 4741dna2-1 strain. The cells were then fixed on eight-well, Teflon-coated slides pretreated with 0.1% polylysine 6040805 ICN Biomedicals, Costa Mesa, CA). After 15 min in blocking solution (PBS, 0.5% bovine serum albumin, 0.5% chicken albumin, 0.5% Tween 20), the cells were incubated with the primary antibodies diluted at 1:2000 in the block solution for 2 h at room temperature. The cells were then washed four times in blocking buffer and incubated for 2 h with the secondary antibody, rabbit anti-mouse diluted at 1:5000 in blocking solution. After incubation, the cells were washed four times in blocking solution and immediately costained with 4′,6′-diamidino-2-phenylindole (DAPI). DAPI was used at 50 ng/ml as described previously ( Mays Hoopes et al., 2002 ). Cells were then observed under a fluorescence axioscope (Carl Zeiss) coupled with a Orca 2 camera (Hamamatsu, Bridgewater, NJ).

Flow Cytometry

For flow cytometry analysis, samples were fixed in 70% ethanol. After an overnight digestion with RNase I (8 mg/ml in PBS) at 37ଌ and a 4-h digestion with proteinase K (1 mg/ml) at 50ଌ, cells were resuspended in propidium iodide in PBS (50 μg/ml), sonicated (output 3-20% duty cycle, 15 s) and filtered with mesh (3-64/32 NITEX Tetko). The control strain (WT4741) was grown asynchronously at 24ଌ (with �% of control strain cells having 1C and 48% having 2C content) and until saturation to identify the 1C peak (100% having 1C content).

DNA Microarray Experiments and Analysis

Old and young wild-type cells were isolated as described under “Isolation of Young and Old Cells by Elutriation.” The elutriated cells from six such experiments (a total of 12 liters), yielding 4.8 × 10 8 cells each, were pooled. Samples of these cells were used for multiple RNA extractions (see below). Old and young dna2-1 cells were isolated as described under “Isolation of Young and Old Cells by Elutriation.” The elutriated cells from eight such experiments (a total of 16 liters), yielding 3.6 × 10 8 cells each, were pooled. Samples of these cells were used for multiple RNA extractions (see below).

RNA extractions were performed with hot/phenol chloroform followed by an Rneasy column (74104 QIAGEN, Valencia, CA). cDNA was synthesized by using oligo(dT) primer (18418-012 Invitrogen, Carlsbad, CA) and Powerscript reverse transcriptase (8460-1 BD Biosciences Clontech, Palo Alto, CA) by using 60 μg of total RNA for each reaction. The cDNA was labeled during reverse transcription with either Cy5-labeled dCTP (red, PA55021 Amersham Biosciences, Piscataway, NJ), or Cy3-labeled dCTP (green, PA53021 Amersham, Biosciences), according to the directions of the supplier (BD Biosciences Clontech). Independent RNA preparations were used for each cDNA preparation.

RNA levels in young and old cells were determined in duplicate for each strain by a ratiometric method by using microarray hybridization analysis. We performed two control experiments (one for wild-type and one for dna2-1), which consisted of a determination of RNA levels in young cells of the same strain, alternatively labeled with Cy3 and Cy5. We also performed four hybridization experiments that consisted of a comparison between RNA levels in young cells and old cells of the same strain. For each strain, wild-type and dna2-1, two hybridization experiments were performed. In the first experiment, Cy3 was used to prepare the cDNA from young cells and Cy5 for old cells. In the second experiment, the dyes were switched to remove any bias that may have been introduced by gene-specific differences in incorporation of the two dyes. For instance, equal amounts of the Cy3 and Cy5-labeled dyes, attached to their respective cDNAs, were used for each array (at least 20 pmol). In addition, as just mentioned, independent RNA isolations were used for each cDNA preparation to control for possible differences due to extraction procedures. Hybridizations were performed on DNA microarrays from Corning Microarrays Technology (Fountain Valley, CA). The CMT Yeast-S228c Gene Array version 1.31 contains � polymerase chain reaction (PCR) products (𢏁 kb) representing open reading frames (ORFs) from the fully sequenced S. cerevisiae genome, plus 108 control genes from Bacillus subtilis. For hybridization, the microarrays were incubated in 50 μl of hybridization solution (1% sonicated salmon sperm, 24.75% formamide, 4.95× SSC, 0.099% SDS, and 0.99 mM dithiothreitol) for 10-18 h at 42ଌ. After washing (1 min in 2× SSC, 0.1% SDS at 42ଌ 5 min in a new solution of 2× SSC, 0.1% SDS at 42ଌ 10 min in 0.1× SSC, 0.1% SDS 15 s, 2 min, 2 min, and 1 min in 0.1× SSC 15 s in 0.01× SSC), separate images were acquired for each dye, and fluorescence intensity ratios were obtained for all genes (Genepix-Pro3.0 Axon scanner 4000A).

We performed the analysis using MArray, a software that allows the user to analyze single or paired microarray data sets ( Wang et al., 2002 ). MArray defines the quality of microarray experiments and assesses the reproducibility of replicate experiments. The analysis consisted of three steps: filtering, normalization, and interpretation. During the filtering, all empty or negatively flagged spots (the flag coefficient is given by the Genepix file) were removed (see supplementary data

lesur/old_and_young_cells/MArray_output.pdf). We eliminated the genes that did not show a consistent expression profile in the duplicates (for example, genes that were more highly expressed in the young wild-type cells in the first experiment but were then more highly expressed in the old wild-type cells in the repeated experiment). After all filtering, 5733 wild-type genes and 6141 dna2-1 genes were left. The minimum signal intensity of each spot was set to zero. We used two control experiments to quantify the noise due to the technique by itself, especially dye bias, and to set the thresholds for identifying the genes significantly differentially expressed in old or young cells in each strain. This sets the intensity limits for interpreting the experimental arrays. We then normalized the intensity young/old cells by using intensity-dependent normalization of the ratios as described by Yang et al. ( 2002 ). The data are presented as the background subtracted, normalized young/old ratios for each array. For each strain, reported expression ratios are the average of the two expression ratios young cells/old cells after normalization of the two hybridizations in which the Cy3 and the Cy5 fluorescent dyes were reversed (see tables in supplementary data). To demonstrate the reproducibility of the hybridizations, we calculated the correlation coefficients for each set of reversed experiments.

The entire experiment was also performed again comparing young cells isolated without elutriation with old cells isolated by elutriation. The genes activated fell in the same pathways identified in the data reported. The results are not included because the young cells were not isolated by elutriation as they were in the data set presented here, making averaging of the data impossible. However, it is noteworthy that the results overlap significantly and lead to identical interpretations.

The Yeast Proteome Database (, the Saccharomyces Genome Database (, and the annotations of the MIPS Comprehensive Yeast Genome Database ( were used for interpretation of the expression profiles. In addition, comparison of our results with databases generated recently describing transcriptional responses in yeast to various environmental stresses, conditions that extend life span, and to various mutations that lead to chromosome damage or repair was valuable ( Jelinsky and Samson, 1999 Gasch et al., 2000 , 2001 Jelinsky et al., 2000 Rep et al., 2000 Kaeberlein et al., 2002 Lin et al., 2001 Lin et al., 2002 ).

Real-Time Quantitative Reverse Transcription RT-PCR

Confirmation of differentially expressed transcripts was performed using the iCycler IQ real time PCR detection system (Bio-Rad, Hercules, CA) on cDNA obtained from old and young cells isolated with the elutriation system. Total RNA was obtained from each sample and treated with deoxyribonuclease (catalog no. 1906 Ambion, Austin, TX) to remove DNA contamination. cDNA was synthesized using 5 μg of RNA and the iScript cDNA synthesis kit from Bio-Rad (catalog no. 170-8891).

Four microliters of a serial 10-fold dilution (10, 100, 1,000, and 10,000) of cDNA obtained by reverse transcription was amplified in a 25-μl reaction mix containing 1× SYBR Green Supermix (catalog no. 170-8880 Bio-Rad) and 25 pM of each primer. Each sample was run in triplicate. After a 3-min Taq activation step at 95ଌ, reactions were subjected to 50 cycles of 10-s denaturation at 95ଌ and 10-s extension at 54ଌ. Primers were purchased from IDT Integrated Technologies (Coraville, IA). Primer pairs were chosen to minimize primer dimerization and to generate an amplicon between 100 and 150 base pairs. Optical data were collected during the annealing step of each cycle. After PCR, reaction products were melted for 1 min at 95ଌ and then the temperature was set to 54ଌ and increased to 94ଌ in 0.5ଌ increments. Optical data were collected over the duration of the temperature increase. This was done to ensure that only one PCR product was amplified per reaction.

Relative expression of the RT-PCR products was determined using the mathematical model from M.W. Pfaffl ( Pfaffl, 2001 ). This model calculates relative expression by using the equation ratio = [(Etarget) 㥌t target(control - sample) ]/[(Eref) 㥌t ref(control - sample) ]. The ratio of a target gene is expressed in a sample versus a control in comparison to a reference gene. Etarget is the real-time PCR efficiency of a target gene transcript, Eref is the real-time PCR efficiency of a reference gene transcript, 㥌ttarget is the Ct deviation of control - sample of the target gene transcript, and 㥌tref = Ct deviation of control - sample of reference gene transcript. The reference gene we used is TUB1 because it is a stable unregulated transcript in each of our microarray data sets. Nine target genes belonging to the recombinational pathway were studied in the wild-type and the dna2-1 strains. For the calculation of the ratio old cells/young cells, the individual real-time PCR efficiencies (E) and the deviation 㥌t must be known. We calculated the efficiencies according to E = 10 [-1/slope] . Because each sample was run in triplicate, the mean Ct value was used in the equation and the 㥌t values were the differences in averaged Ct values between old and young cells for the same gene. The control sample containing the same concentration of cDNA was chosen to be compared with the target gene.


Death Induction by Hydrogen Peroxide, Acetic Acid, and Hyperosmotic Shock

We chose hydrogen peroxide and acetic acid for inducing PCD and TUNEL-positive phenotype in S. cerevisiae, to determine the nature of the DNA lesions detected by the TUNEL assay. Apoptosis-like cell death triggered by these two compounds has been extensively characterized by several laboratories (Madeo et al., 1999, 2002 Ludovico et al., 2001a, 2002 Fabrizio et al., 2004 Wissing et al., 2004 Giannattasio et al., 2005). Hyperosmotic shock by 60% (wt/wt) glucose has been recently described also to induce TUNEL-positive phenotype as well as other apoptotic markers (Silva et al., 2005). Cells were grown in preculture overnight and then for two generations in fresh medium before treatment with hydrogen peroxide, acetic acid, or 60% (wt/wt) glucose, as described in Materials and Methods. Samples were taken at time zero and at various time points, as indicated in Figure 1. Cells were spread on solid YPD medium and colony-forming units (c.f.u.) were counted after 3 d of incubation. The survival percentage was calculated as the number of c.f.u. obtained at each time point divided by the number of c.f.u. obtained at time 0. Treatment with 10 mM hydrogen peroxide initiated cell death (Figure 1A) without lag phase. Cell death by 175 mM acetic acid at pH 3.0 showed an initial lag phase of ∼30 min, before cell death was evident (Figure 1B). Treatment for 200 min with 175 mM acetic acid at pH 3.0 or 10 mM hydrogen peroxide or 60% (wt/wt) glucose for 10 h (Figure 1C) resulted in ∼10% survival, which in our and others' experience (Madeo et al., 1999) gives the highest yield of TUNEL-positive cells. Treatment for 20–25 min with 150 mM hydrogen peroxide (Figure 1D) gave about the same remaining survival as 10 mM for 200 min, thus representing a cell death induction approximately 10 times faster.

Figure 1. S. cerevisiae BY4741 was incubated with 10 mM hydrogen peroxide (A) or 175 mM acetic acid, pH 3.0 (B), 60% (wt/wt) glucose (C) or 150 mM hydrogen peroxide (D). Samples were taken at the time points indicated and plated on solid YPD media. c.f.u. were counted after 2 d of incubation at 30°C. Relative survival is plotted on the x-axis (100% corresponds to the number of c.f.u. at time 0). Values are mean ± SEM of three independent experiments.

TUNEL Stains Both SSBs and DSBs in S. cerevisiae

Cells grown as described in Materials and Methods were fixed with formaldehyde and subsequently digested with Zymolyase to remove the cell wall. Cells were treated with DNaseI to generate chromosomal DSBs. DNaseI cleaves both DNA strands at approximately the same site, producing DNA fragments with blunt ends or to a lesser extent protruding termini of one or two nucleotides. The cells were then stained with TUNEL protocol according to previously published protocols (Madeo et al., 1997). The nuclei was intensely stained by TUNEL in cells treated with DNaseI (Figure 2, DNaseI) This staining does not occur in nontreated cells (Figure 2, negative control). Positive TUNEL staining was also obtained by treatment with N.BbvCIA nicking endonuclease (Figure 2, N.BbvCIA). N.BbvCIA is a restriction enzyme engineered to cut one DNA strand only. This result shows that the free 3′-OH exposed in SSBs in S. cerevisiae is a good substrate for the terminal transferase in the TUNEL reaction. Cells treated with hydrogen peroxide or acetic acid (as described in Materials and Methods), but no added restriction enzymes, did also show TUNEL staining (Figure 2, hydrogen peroxide and acetic acid). TUNEL staining is more diffuse and has slightly higher background for the hydrogen peroxide-treated cells. Nevertheless, nuclei are clearly visible against the background.

Figure 2. Bright field and epifluorescence images of TUNEL and in situ ligation-stained cells. Left column, TUNEL staining of exponentially growing cells, fixed and treated with DNaseI or N.BbvCIA nicking endonuclease. Right column, in situ ligation staining of exponentially growing cells, fixed and treated with BsuRI (blunt endonuclease) or DNaseI and N.BbvCIA (nicking endonuclease). Row marked negative control shows exponentially growing cells stained with TUNEL or in situ ligation staining. Rows marked hydrogen peroxide and acetic acid show cells incubated for 200 min with 10 mM hydrogen peroxide or 175 mM acetic acid, pH 3.0. Left, phase-contrast microscopy right, fluorescence microscopy of the same cells. Bar, 5 μm.

ISL Assay Specifically Stains DSBs in S. cerevisiae

Cells were also subjected to ISL staining with a fluorescent double-stranded DNA hairpin probe according to the methods published by Didenko and coworkers (Didenko and Hornsby, 1996 Didenko et al., 1999). A small double strand fluorescent DNA probe is ligated to sites of DSBs. It has been shown to specifically stain mammalian apoptotic cells (Didenko et al., 2003). The probe cannot ligate to single-stranded DNA breaks and is therefore in theory more specific for DSBs than TUNEL staining. Specific ISL staining, visually similar to staining by TUNEL, was evident in cells treated with DNaseI and the blunt restriction enzyme BsuRI (Figure 2, in situ ligation, DNaseI and BsuRI). There was no detectable staining in the absence of endonucleases (Figure 2, in situ ligation, negative control). ISL does not stain cells treated with nicking endonuclease N.BbvCIA, confirming that SSBs do not facilitate ligation of the ISL probe (Figure 2, N.BbvCIA). These results show that both TUNEL and ISL procedures are equally efficient at detecting DSBs in S. cerevisiae chromosomal DNA, but TUNEL assay also detects nicked DNA. Cells treated with peroxide or acetic acid did not show any ISL staining (Figure 2, hydrogen peroxide and acetic acid), indicating that these cells do not contain blunt DSBs.

Staggered DSBs Can Be Discriminated by ISL in Combination with Klenow DNA Polymerase

ISL has been used to distinguish between different types of staggered DSBs in combination with Klenow DNA polymerase (Didenko et al., 2003). Theoretically, if DSBs with 5′ overhangs are present, the DNA polymerase activity of Klenow will synthesize the missing DNA in the presence of dNTPs and produce blunt ends. In contrast, if DSBs breaks with 3′ overhangs are present, the Klenow 5′-3′ exonuclease activity will degrade the overhang until the DNA is blunt.

To test whether ISL could discriminate between different types of DSBs in S. cerevisiae, we performed ISL on cells treated with endonucleases producing 5′ or 3′ overhang. Cells were grown, fixed, and Zymolyase treated, as described above, and subsequently treated with restriction endonucleases, creating either 1-base pair 3′ overhangs (HhaI) or 1-base pair 5′ overhangs (Bme1390I). None of the enzyme-treated samples showed any positive staining with ISL assay (Figure 3, − Klenow). This shows that ISL does not stain staggered DSBs. ISL staining is restored to cells treated with HhaI and to cells treated with Bme1390I, by treatment with Klenow DNA polymerase and Klenow DNA polymerase plus dTNPs, respectively (Figure 3, + Klenow). This means that different types of staggered double-stranded DNA ends can be discriminated by the ISL assay in yeast. The intensity is lower than for the DNaseI or BsuRI treatment (Figure 2), probably due to incomplete transformation of staggered DSBs.

Figure 3. Bright field and epifluorescence images of in situ ligation-stained cells. Exponentially growing cells were treated with HhaI (1-base pair 3′ overhang) and Bme1390I (1-base pair 5′ overhang) endonucleases. The cells in the right column were treated with Klenow (+ Klenow, HhaI) or Klenow + dNTPs (+ Klenow, Bme1390I) before in situ ligation staining. Left, phase-contrast microscopy right, fluorescence microscopy of the same cells. Bar, 5 μm.

We reasoned that the lack of ISL staining of cells treated with hydrogen peroxide or acetic acid (Figure 2, hydrogen peroxide, acetic acid) may be that they contain staggered DSBs that are not detectable by ISL. We added Klenow or Klenow and dNTPs to cells treated with hydrogen peroxide or acetic acid in a manner similar to the experiment described in Figure 3, but the ISL staining was still negative (Figure 4). This indicates that hydrogen peroxide- or acetic acid-treated cells do not contain staggered DSBs with 1-base pair overhangs.

Figure 4. Bright field and epifluorescence images of in situ ligation-stained cells. In situ ligation staining of cells incubated for 200 min with 10 mM hydrogen peroxide or 175 mM acetic acid, pH 3.0. The cells were treated with Klenow (left column) or Klenow + dNTPs (right column) before in situ ligation staining. Left, phase-contrast microscopy right, fluorescence microscopy of the same cells, Bar, 5 μm.

Chromosomal DNA Is Damaged in Cells Treated with Hydrogen Peroxide, Acetic Acid, or High Concentrations of Glucose

In our hands, ISL assay together with Klenow or Klenow + dNTPs is limited in the overhang length of staggered DSBs that the method can overcome. We treated cells with enzymes creating four-base pair overhangs with subsequent Klenow transformation to blunt DSBs, and we noted a considerable decrease in the ISL signal (our unpublished data) compared with the treatments with HhaI and Bme1390I (Figure 3). This led to the hypothesis that cells treated with hydrogen peroxide or acetic acid may have staggered DSBs with long overhangs, undetectable by ISL.

To overcome this difficulty, we analyzed chromosomal DNA from cells treated with hydrogen peroxide, acetic acid, or glucose-induced hyperosmotic shock using PFGE (Figure 5A). Samples were taken at six time points between zero and 200 min (hydrogen peroxide, acetic acid) or 10 h (hyperosmotic shock). DNA from cells treated with hydrogen peroxide after 200 min (Figure 5A, hydrogen peroxide, lane 200 min) was completely degraded to a smear of fragments slightly shorter than S. cerevisiae chromosome I (225 kb) but still of a considerable size. The DNA from acetic acid-treated cells showed less degradation but still a visible smear and chromosomal bands of lower intensity (Figure 5A, acetic acid, lanes 60–200 min). DNA from cells killed by hyperosmotic shock showed degradation over 10 h comparable to that of hydrogen peroxide-treated cells (Figure 5A, glucose 60%, lanes 6–10 h). This is the first time that clear DNA degradation has been shown in S. cerevisiae associated with PCD.

Figure 5. Genomic DNA analyzed by PFGE from viable cells (A and C–E) or fixed S. cerevisiae cells (B). Cells were exposed to 10 mM hydrogen peroxide, 175 mM acetic acid, pH 3.0, 60% (wt/wt) glucose, or 150 mM hydrogen peroxide as indicated in the figure. Lanes marked as control were loaded with DNA isolated from exponentially growing cells without further treatment. Samples were collected after various periods as indicated (minutes) except for treatment with 60% (wt/wt) glucose, where time points are indicated in hours. (D) Cells were exposed to 10 mM hydrogen peroxide or 175 mM acetic acid, pH 3.0, and isolated chromosomal DNA was subsequently treated with S1 nuclease to degrade nicked DNA. Lane marked control S1 is similar to control lanes, but the DNA was treated with S1 nuclease before PFGE. (E) Lanes 1 and 2 show chromosomal DNA from nontreated cells, digested with N.BbvCIA nicking endonuclease (1) or DNaseI (2).

DNA Fragmentation by Hydrogen Peroxide Requires Active Enzymes

Cells were fixed with 3.7% (vol/vol) formaldehyde before treatment with 10 mM hydrogen peroxide for 200 min or 150 mM for 25 min (Figure 5B) to test whether the presence of active enzymes is necessary for DNA fragmentation to proceed. Fixing cells with formaldehyde preserves the structure of cells and enzymes, but it eliminates enzymatic activity. The fixed cells treated with hydrogen peroxide show no DNA damage, indicating that the DNA fragmentation requires some enzymatic activity within the cell and that fragmentation is not due to a direct chemical reaction between hydrogen peroxide and DNA (Figure 5B). In addition, purified yeast chromosomes were not degraded by 10 mM hydrogen peroxide in vitro for 200 min (our unpublished data).

DNA Fragmentation by Hydrogen Peroxide Does Not Occur after Treatment with High Concentrations of Hydrogen Peroxide

A common observation regarding PCD is that specific markers of PCD usually only occur within a rather limited window of treatment intensity. At too high intensity of the treatment, the cell is presumed to die from complete breakdown (necrosis) before any PCD process can be initiated. Chromosomal DNA was fragmented during 200-min treatment with 10 mM hydrogen peroxide (Figure 5A). During this process, the fraction of viable cells decreases from 100 to ∼5–10% (Figure 1A). The DNA in S. cerevisiae cells treated with 150 mM hydrogen peroxide remained intact (Figure 5C) for the 25 min necessary for a decrease from 100 to ∼1% surviving cells (Figure 1C). This result shows that cell death is only associated with DNA fragmentation for treatment of relatively low intensity.

DNA Damage in Hydrogen Peroxide- or Acetic Acid-treated Cells Is Primarily Made Up of SSBs

Lack of ISL staining of cells treated with hydrogen peroxide or acetic acid combined with the finding that TUNEL assay is not specific for DSBs led us to the hypothesis that the main type of DNA damage detected by TUNEL in our experiments consists of SSBs. This hypothesis can be tested by treatment of the DNA with S1 nuclease that will preferably attack at sites of single-stranded DNA breaks, reducing SSBs to DSBs, causing the DNA to migrate faster on the gel. Figure 5D shows a PFGE gel with DNA from cells treated with hydrogen peroxide or acetic acid, with and without treatment with S1 nuclease. Untreated DNA (Figure 5D, lane control) shows little degradation with S1 nuclease treatment (Figure 5D, lane control S1), indicating that no or few single-stranded breaks are present in untreated cells. Comparing the same time points for hydrogen peroxide- and acetic acid-treated cells without (Figure 5A) and with S1 treatment (Figure 5D, lanes marked +), there is an enhanced degradation upon S1 treatment. We conclude that the main form of DNA damage in hydrogen peroxide- and acetic acid-treated cells consists of single-stranded DNA breaks.

N.BbvCIA Nicking Endonuclease and Hydrogen Peroxide Treatment Yield Similar DNA Damage

It is evident from the images showing DNA damage (Figure 5A) that although the chromosomes are broken down to the point of not being visible, the resulting fragments are still several hundreds of kilobases and do not seem to break down into smaller fragments. We treated isolated DNA with DNaseI and N.BbvCIA nicking endonuclease (Figure 5E). Isolated DNA treated with DNaseI broke down into fragments too small to be visible on the gel (Figure 5E, lane 2), whereas DNA treated with N.BbvCIA (Figure 5E, lane 1) revealed DNA fragmentation with the same visual appearance on the gel as cells treated with hydrogen peroxide for 200 min (Figure 5A, lane 200 min). This observation is consistent with the conclusion that DNA damage in hydrogen peroxide-treated cells are primarily a consequence of accumulation of SSBs.

Boy Is Clue to What Causes Aging

One of Niederhofer's colleagues on the study was Jan Hoeijmakers, PhD, of Erasmus Medical Center in Rotterdam, Netherlands.

Hoeijmakers had heard from doctors in Germany about a 15-year-old Afghan boy with an extreme form of premature aging, a condition called progeria.


The boy was extremely sensitive to sunlight and had had an old, wizened appearance since age 10.

He was gaunt, had hearing loss and vision problems, and had had hepatitis A and tuberculosis as a young child.

The boy died when he was 16 after getting severe pneumonia and having organ failure. Genetic tests showed the boy had a severe mutation in his XPF gene, a gene involved in DNA repair.


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Deleting 'Anti-Aging' Gene From Yeast Greatly Lengthens Life Span

A counterintuitive experiment has resulted in one of the longest recorded life-span extensions in any organism and opened a new door for anti-aging research in humans.

Scientists have known for several years that an extra copy of the SIR2 gene can promote longevity in yeast, worms and fruit flies.

That finding was covered widely and incorporated into anti-aging drug development programs at several biotechnology companies.

Now, molecular geneticists at the University of Southern California suggest that SIR2 instead promotes aging.

Their study, "Sir2 Blocks Extreme Life-Span Extension," appears in the Nov. 18 edition of the biology journal Cell. The lead author is Valter Longo, assistant professor in the Leonard Davis School of Gerontology and the USC College of Letters, Arts and Sciences.

Rather than adding copies of SIR2 to yeast, Longo's research group deleted the gene altogether.

The result was a dramatically extended life span - up to six times longer than normal - when the SIR2 deletion was combined with caloric restriction and/or a mutation in one or two genes, RAS2 and SCH9, that control the storage of nutrients and resistance to cell damage.

Human cells with reduced SIR2 activity also appear to confirm that SIR2 has a pro-aging effect, Longo said, although those results are not included in the Cell paper.

Since all mammals share key aging-related genes, the paper points to a new direction for human anti-aging research.

Longo proposes that SIR2 and possibly its counterpart in mammals, SIRT1, may block the organism from entering an extreme survival mode characterized by the absence of reproduction, improved DNA repair and increased protection against cell damage. Organisms usually enter this mode in response to starvation.

The long-lived organisms in Longo's experiment showed extraordinary resilience under stress.

"We hit them with oxidants, we hit them with heat," Longo said. "They are highly resistant to everything. What they're doing is basically saying, 'I cannot afford to age. I still have to generate offspring, but I don't have enough food to do it now."

Longo predicted that as molecular geneticists master the levers of aging, they will be able to design drugs that coax the body into entering chosen aspects of a starvation-response mode, such as stress resistance, even when food is plentiful.

If enough food is available, an organism might be programmed both to reproduce normally and to maximize its survival systems.

Longo urged caution in extrapolating the result to humans.

"We have been very successful with simple organisms," he said. "Naturally, mammals are complex, and it will be a great challenge to get major life-span extension."

A "really exciting" implication, Longo said, is that cells may be able to speed up their DNA repair efforts. All organisms have the ability to repair harmful mutations in their DNA, whether caused by age, radiation, diet or other environmental factors. Cancer often begins when DNA mutations outstrip a cell's ability to remain differentiated.

Many researchers believe DNA repair systems are already running flat out. The organisms in Longo's experiment say otherwise.

"In our paper, we show that age-dependent mutations increase at a much slower pace in organisms lacking RAS2 or SCH9 and at a remarkably low pace in organisms lacking both SCH9 and SIR2, raising the possibility that the mutations that cause human cancers can be delayed or prevented," Longo said.

"Notably, mutations that increase the activity of human homologs of the yeast SCH9 and RAS2 genes play central roles in many human cancers." Homologs are genes descended from a common ancestral DNA sequence.

Joining with researchers at the USC Norris Comprehensive Cancer Center, Longo is studying the feasibility of reducing or preventing the age-dependent DNA mutations that cause cancer.

Longo and his collaborators began studying SIR2 in 2000, soon after a well-known set of experiments by Leonard Guarente at the Massachusetts Institute of Technology. Guarente was the first to show that over-expression of the SIR2 gene could extend life span beyond its natural limit.

However, Longo said, "We were convinced that SIR2 had the potential to be a more potent pro-aging than an anti-aging gene. And the reason was in part because of the similarity with this other gene, called HST1, which negatively regulated so-called protective genes. So we set out to test whether SIR2 could do the opposite of what everybody said it does."

The researchers do not quarrel with Guarente's finding of a moderate increase in life span when SIR2 is over-expressed. But their work shows that much greater potential gains lie in the opposite direction.

Longo's co-author was USC research scientist Paola Fabrizio. The other USC authors were Cristina Gattazzo, Luisa Battistella, Min Wei, Chao Cheng and Kristen McGrew.

Funding for this research came from the American Federation for Aging Research and from the National Institute of Aging of the National Institutes of Health.

The type of vacuole found in yeast cells is somewhat analogous to the lysosome that we animals possess in that it is involved in breaking down waste products and recycling broken cellular components (via the process of autophagy) that would otherwise harm the cell. It is an agent of cellular housekeeping, in other words. There the similarities end, however, as the vacuole performs many other vital tasks that the more specialized lysosome does not.

So here, researchers show that they can extend life in yeast by reversing a change that occurs in the vacuole. Because the vacuole has many more tasks than the lysosome, it's not immediately clear that this has any application to our biology of aging, however. It is still worth keeping an eye on this research as we know that decline in lysosomal function (and thus of cellular housekeeping) is important in animal aging. You might recall, for example, that researchers managed to reverse the age-related loss of liver function in mice by finding a way to keep lysosomal function running at youthful rates. Similarly, reversing the root causes of lysosomal decline is on the SENS agenda - to be achieved by breaking down the build up of metabolic waste products that accumulate in lysosomes and cause them to malfunction.

Normally, mitochondria [in yeast] are beautiful, long tubes, but as cells get older, the mitochondria become fragmented and chunky. The changes in shape seen in aging yeast cells are also observed in certain human cells, such as neurons and pancreatic cells, and those changes have been associated with a number of age-related diseases in humans.

The vacuole - and its counterpart in humans and other organisms, the lysosome - has two main jobs: degrading proteins and storing molecular building blocks for the cell. To perform those jobs, the interior of the vacuole must be highly acidic. [Researchers] found that the vacuole becomes less acidic relatively early in the yeast cell's lifespan and, critically, that the drop in acidity hinders the vacuole's ability to store certain nutrients. This, in turn, disrupts the mitochondria's energy source, causing them to break down. Conversely, when [researchers] prevented the drop in vacuolar acidity, the mitochondria's function and shape were preserved and the yeast cells lived longer.

Until now, the vacuole's role in breaking down proteins was thought to be of primary importance. We were surprised to learn it was the storage function, not protein degradation, that appears to cause mitochondrial dysfunction in aging yeast cells. . The unexpected discovery prompted [the researchers] to investigate the effects of calorie restriction, which is known to extend the lifespan of yeast, worms, flies and mammals, on vacuolar acidity. They found that calorie restriction - that is, limiting the raw material cells need - delays aging at least in part by boosting the acidity of the vacuole.


  • Use the procedure above to make plates with diluted yeast cultures and expose the yeast to sunlight at various times of day for example, at 10:00 AM, noon, and 2:00 PM. Use the same duration of exposure.
  • Expose the yeast for various durations at the same time of day, for example 0, 0.5, 1, 2, 4, and 8 minutes at noon.
  • Compare wild-type and DNA-repair-deficient yeast strains for UV sensitivity. Obtain wild-type S. cerevisiae from a science supply company for example, Carolina Biological, item # 898900. Culture this strain and plate out dilutions, as described in the procedure. Compare the sensitivity of the wild-type and DNA-repair-deficient strains to ultraviolet light from the sun.


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