Lifespan regulation by targeting heme signaling in yeast

The yeast strains used in this study and their genotypes are listed in Table. One-step polymerase chain reaction (PCR)-mediated gene disruption was performed using standard techniques to delete genes of interest. The genotypes of the resulting strains were verified using colony PCR. S1

To generate a yeast heme auxotroph strain expressing ceHRG-4, the sequence containing ceHRG-4 was first PCR amplified from pYES-DEST52-ceHRG-4 plasmid (pPP96) [17] and integrated into p416-GPD [18] vector using BamHI and XhoI restriction sites to generate pPP97 plasmid. The sequence encoding ceHRG-4 under the control of the glyceraldehyde-3-phosphate dehydrogenase promoter (GPD1pr-ceHRG-4) along with the URA3 marker was then amplified from pPP97 and integrated into the hem1Δ strain using homologous recombination at URA3 locus. The resulting transformants were selected on SC-URA plates and verified by colony PCR and Sanger sequencing.

To generate the HMX1-OE strain expressing HMX1 under the control of the ADH1 promoter (ADH1pr-HMX1), a 705 bp sequence of the ADH1 promoter was amplified by PCR from yeast genomic DNA and integrated into the pRS306 [19] plasmid using NotI and XhoI restriction sites. Subsequently, the fragment encompassing the URA3 marker and the ADH1 promoter was amplified via PCR using the oPP399 and oPP400 oligonucleotides. These oligonucleotides have homology to the 3′ end of the HMX1 promoter and the 5′ end of the HMX1 ORF, respectively. The resulting PCR product was then utilized for genomic integration. To create the HAP4-OE strain, the fragment containing the URA3 marker and the ADH1 promoter was amplified using the oPP274 and oPP275 oligonucleotides and was integrated into the genome upstream of the HAP4 ORF. To create the yeast strain encoding an HA-tagged Hap4 protein, HA-tag sequence along with KanMX4 marker was amplified from pPP111 plasmid using the primers flanked on the 5′ side by sequence homologous to HAP4 ORF and the 3′ side by sequence homologous to HAP4 3′-UTR for genomic integration. The resulting transformants were selected on YPD plates containing 200 µg/mL G418 and verified by colony PCR and Sanger sequencing. The plasmids used in this study and sequences of primers used for generating yeast strains are listed in Table S2 and Table S3.

Yeast strains were cultured at 30 °C in standard YPD medium (1.0% yeast extract, 2.0% peptone, and 2.0% glucose) unless otherwise stated. hem1Δ cells were maintained in YPD supplemented with 250 µM 5-aminolevulinic acid (ALA). To test the effect of exogenous heme on replicative and chronological lifespan, the media was supplemented with hemin chloride. The stock hemin, chloride solution was prepared by dissolving hemin in 0.3 M NH₄OH. The pH was adjusted to 8 with 6N HCl, and the mixture was filter-sterilized through a 0.2-μm filter.

The spot assays were used to assess the growth of strains under various growth conditions. Each strain to be tested was inoculated into a liquid culture medium and allowed to grow to the exponential phase. Once the cultures reached OD₆₀₀ = 0.6, 10 × serial dilutions for each strain were spotted on YPD agar plates (containing 2% glucose) or YPG plates (contaminating 3% glycerol) supplemented with indicated concentrations of ALA or heme. The plates were incubated at 30 °C for 48 h before they were imaged.

Replicative lifespan assays were carried out as described previously [20]. Cells were cultured on freshly prepared YPD plates at 30 °C. Cells were monitored for cell divisions, and the subsequent daughter cells were removed using a micromanipulator. The replicative lifespan was determined by counting the number of divisions each mother cell underwent before it ceased dividing. The lifespan assays were conducted at least two times, and the data from separate biological replicates were combined. The number of cells assayed, and statistical analysis of the lifespan data are shown in Table S4.

Yeast CLS assays were performed as described [21]. Briefly, single colonies for each strain were inoculated into 10 mL of synthetic complete dextrose media (SCD) and cultured for 4 days at 30 °C. Aliquots of each culture were serially diluted in SCD media until achieving a cell density of roughly 200 cells per 100 μL. Following this, 100 μL of the dilution was plated onto YPD agar plates, which were then kept at 30 °C for the indicated time, and colonies were manually counted. Colony forming units (CFU) were normalized relative to the CFUs counted on day 0. The averages and standard errors of the mean were calculated based on at least six biological replicates.

Total RNA was extracted using hot acid phenol method followed by purification using Direct-zol RNA Miniprep Kit (Zymo Research). The RNA was treated with DNaseI, and 2 µg of RNA was used for cDNA synthesis using SuperScript III reverse transcriptase (Thermo Fisher Scientific) with random hexamer primers according to the manufacturer's instructions. To analyze mRNA expression, real-time PCR was performed using SYBR Fast qPCR Master Mix (Kapa Biosystems) and the CFX-96 Touch Real-Time PCR Detection System (Bio-Rad Laboratories). To normalize the gene expression, ACT1 was used as the reference gene. The primers used for RT-qPCR are listed in Table S3. Results are represented as means ± SEM from at least three independent experiments.

A single colony was inoculated in 3 mL of YPD medium and incubated overnight at 30 °C. The next day, the cells were diluted to an OD₆₀₀ of 1 in 10 mL of fresh medium and grown until they reached the log phase. The cells were collected by centrifugation and washed with sterile water, and the pellet was frozen in liquid nitrogen and stored at − 80 °C for later use. For protein extraction, the cell pellet was re-suspended in 300 µL of lysis buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA, and 1% DMSO) containing 1 mM phenylmethylsulfonyl fluoride (PMSF). The samples were homogenized with glass beads by vortexing at maximum speed for seven 30-s cycles and centrifuged for 5 min at 12,000 rpm at 4 °C to obtain the supernatant. Equal amounts of proteins were resolved in 10% SDS-PAGE gels and transferred to PVDF membrane. The membrane was then probed with horseradish peroxidase (HRP)-conjugated HA tag monoclonal antibody (2–2.2.14) (ThermoFisher) using 1:1000 dilution, and signal quantification was performed using ImageJ software. Mouse anti-Pgk1 monoclonal antibody (1:5000, Life Technologies) and HRP-conjugated secondary anti-mouse antibody (1:100,000, Santa Cruz Biotechnology) were used to detect Pgk1 protein levels as a loading control.

Intracellular heme levels were assessed using the oxalic acid method as previously described with minor changes [22]. Briefly, overnight yeast cultures were diluted to OD₆₀₀ = 0.2 units/mL, and grown until cells reached OD₆₀₀ = 0.8 units/mL. Subsequently, 8 OD₆₀₀ units of cells were harvested by centrifugation at 2500 × g and washed with distilled water, and the pellet was resuspended in 500 µL of 20 mM oxalic acid. An additional 500 µL of 2 M oxalic acid was added, and the suspension was divided equally into two tubes. One set of sample tubes was placed in a heating block at 100 °C for 45 min, while the other set of samples was kept at room temperature for the same duration as a baseline. Following incubation, samples were cooled to room temperature, and 100 µL of suspension was transferred per well into a black-well 96-well plate in duplicates. The fluorescence of porphyrin was measured using a BioTek plate reader with 400 nm excitation and 662 nm emission. Baseline values (from parallel unheated samples in oxalic acid) were subtracted, and the relative fluorescence intensity in arbitrary units (A.F.U.) was plotted.

Statistical analysis was performed using Prism 9.3.1 (GraphPad Software, Inc.). The statistical significance of the heme quantification and RT-qPCR data was determined by calculating p values using one-way ANOVA. Error bars represent standard errors of the mean (SEM). The statistical significance of the lifespan data was evaluated using the Wilcoxon Rank-Sum test [23].

Given the beneficial effect of heme supplementation on lifespan, we sought to investigate the effects of decreasing intracellular heme levels on lifespan. To this end, we overexpressed the HMX1 gene, which encodes heme oxygenase involved in intracellular heme degradation. HMX1 overexpression (HMX1-OE) decreased lifespan by 16.4% compared to wild-type cells (Fig. 2E). Furthermore, supplementing the HMX1-OE strain with heme was able to rescue the shortened lifespan caused by the HMX1 overexpression, and this increase in lifespan correlated with intracellular heme concentration (Fig. 2F). Together, these findings suggest that heme plays a critical role in the regulation of replicative lifespan in yeast.

It has been previously shown that Hap4 is susceptible to proteasome-mediated degradation [28]. To test whether decreased Hap4 protein levels can be attributed to its increased proteasomal degradation, we treated cells with heme in the presence or absence of MG132 proteasome inhibitor [29]. Because S. cerevisiae is resistant to MG132 [30], we employed the pdr5Δ strain with reduced drug efflux for experiments using proteasome inhibitors [31]. When pdr5Δ cells were supplemented with 75 µM MG132, degradation of Hap4 by proteasome was prevented (Supplementary Figure S2A). However, MG132 treatment did not counteract the negative impact of heme supplementation on Hap4 protein levels (Fig. 3B), suggesting that the decrease in Hap4 levels is not due to proteasomal degradation. Furthermore, the addition of the proteasome inhibitor was not able to rescue the growth of ceHRG-4-expressing cells treated with heme on glycerol-containing medium (YPG) (Supplementary Figure S2B). Additionally, we analyzed the abundance of HAP4 mRNA and expression of the Hap4 transcription factor targets, including KGD1, ACO1, and SDH1 (Fig. 3C) using RT-qPCR. We observed a significant decrease in HAP4 mRNA levels (p < 0.001) and diminished expression of the Hap4 targets when yeast cells were exposed to high heme concentrations.

To directly test if the increased lifespan in response to heme supplementation requires Hap4, we analyzed replicative lifespan in wild-type cells and the HAP4 deletion mutant (hap4Δ) in the absence and presence of heme supplementation (Fig. 3D). Although the deletion of HAP4 resulted in a shortened replicative lifespan (p < 0.001) compared to the wild-type strain, the supplementation of hap4Δ with heme was sufficient to significantly extend the lifespan. Taken together, our findings suggest that heme supplementation causes a reduction in HAP4 expression, and that the lifespan extension effect of heme cannot be attributed to the increased activity of the Hap4 transcription factor.

Together, our results suggest the existence of a previously unanticipated role of heme in the regulation of lifespan in yeast and raise an exciting possibility that increased heme levels may extend lifespan in other organisms. Previous studies have shown that inhibition of mitochondrial function as well as components of the ETC and TCA cycle can extend the lifespan in several species [39], including yeast [40], worms [41], flies [42], and mice [43]. For example, the downregulation of aconitase and isocitrate dehydrogenase, two enzymes in the TCA cycle, leads to lifespan extension in worms [44]. Additionally, a genetic screen in yeast measuring the replicative lifespan of 4698 deletion mutants identified mitochondrial translation and components of TCA among the most enriched functional groups [45]. However, specific genes and stress response pathways that connect these changes to longevity remain unknown. Further studies analyzing age-dependent changes in transcriptional network of cells in response to heme treatment may provide additional insights into pathways associated with lifespan extension. It is also important to note that yeast do not depend on mitochondrial respiration in the same way that many metazoan cells do under physiological conditions. Therefore, further research is needed to fully understand the mechanisms underlying the effect of heme on lifespan in mammalian cells and its potential implications for human aging.

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