To test whether the removal of self-splicing mitochondrial introns affects host physiology and fitness, we compared two S. cerevisiae strains that are isogenic with regard to their nuclear genomes (except for a single marker gene, ura3, see Materials and methods) but differ with respect to mitochondrial intron content. The control strain a161 (WT) contains the full complement of seven cox1 and five cob introns, whereas strain a161-U7 (I₀) carries an intronless mitochondrial genome, which was originally constructed by Seraphin and co-workers via serial recombination of natural yeast isolates that lack individual mitochondrial introns (Séraphin et al., 1987).

Contrary to a model where self-splicing introns are dispensable parasitic passengers, we find that I₀ exhibits stark phenotypic differences to the control strain. When the two strains are cultured in isolation, exponential growth on glucose-supplemented YPD medium is ~30% slower for I₀ compared to WT (Figure 1a). I₀ also fares poorly when pitted directly against WT in competitive fitness assay (Figure 1—figure supplement 1). Chronological life span (CLS, see Materials and methods), on the other hand, is almost two-fold longer for I₀ (Figure 1b). At the cellular level, I₀ displays increased mitochondrial mass and volume and a mitochondrial morphology characterized by a large network of branched tubules of homogeneous diameter (Figure 1c–e). Transcript levels of mitofusin (fzo1) and the GTPase mgm1, key nuclearly encoded regulators of mitochondrial fusion, are strongly upregulated (qPCR, t-test, fzo1: 4.02-fold, p=0.002; mgm1: 5.24-fold, p=3.19×10⁻⁵, Figure 2a) while levels of dnm1 and fis1, which orchestrate mitochondrial fission, are only moderately induced (qPCR, t-test, dnm1: 1.67-fold, p=0.026; dnm1: 1.81-fold, p=0.085), suggesting that the changes in mitochondrial morphology result from increased fusion rather than impaired fission. The I₀ strain also exhibits a 2.7-fold increase in mitochondrial DNA copy number (qPCR, t-test p=1.09×10⁻⁷). At the same time, there are no significant differences in mitochondrial inner membrane potential, as measured by 3,3'-Dihexyloxacarbocyanine iodide [DiOC6(3)] fluorescence (Figure 1f), suggesting that mitochondria are functional despite grossly altered morphology. This notion is further supported by the observation that I₀ retains the capacity to grow on glycerol, a non-fermentable carbon source (Figure 1—figure supplement 2), as previously reported for a different nuclear background (Minczuk et al., 2002). In fact, biomarkers of mitochondrial metabolism point to increased mitochondrial activity, with higher oxygen consumption (Figure 1g) and cellular ATP levels (Figure 1h) during exponential growth. Despite increased activity, levels of mitochondrial superoxide are reduced (Figure 1i), likely reflecting a > 7.5-fold upregulation of the mitochondrial ROS-scavenger sod2 (qPCR, t-test p=0.003, Figure 2a). Thus, intron removal challenges but does not terminally compromise mitochondrial function.

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The phenotypic changes we observe suggest an involvement of the retrograde response, as do upregulation of cit2, upregulation of the two rate-limiting members of the TCA cycle, cit1 and idh1, and upregulation of both mitochondrially (cox1, cox2, atp6) and nuclearly (cox4, atp1, sdh1, sdh2) encoded parts of the respiratory chain (Figure 2a). Indeed, deletion of rtg2, the transcriptional master regulator of the retrograde response and a sensor of mitochondrial dysfunction, suppresses the I₀ phenotype (Figure 3). Tubular structure is lost and large spherical shapes become prominent (Figure 3a), suggesting distinct defects in the maintenance of mitochondrial ultrastructure (Paumard et al., 2002; Velours et al., 2009). Deletion of rtg2 in WT strains, where the retrograde response is not activated, has no significant effect on mitochondrial volume and morphology, oxygen consumption and ATP levels (Figure 3a–d). The extension of CLS is abrogated in the absence of rtg2, becoming shorter even than the wildtype (Figure 3e). Upon deletion of hap4, the transcriptional activator of nuclearly encoded components of the respiratory chain (as well as the TCA cycle enzymes under normal conditions), mitochondrial morphology reverts back to the wild-type state (Figure 4a–e). We conclude that an intact retrograde response, including upregulation of nuclear components of the respiratory chain, is necessary to generate the mitochondrial phenotype observed in the I₀ strain.

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To determine the ultimate molecular trigger(s) of the retrograde response, we examined how intron removal affects host gene expression. As previous work had found the omega intron to be ‘optional’ – present in some yeast strains but absent in others without obvious phenotypic effects (Wolters et al., 2015; Dujon, 1980) – we focused on cox1 and cob. For both these genes, mRNA levels are strongly elevated in I₀ (10.9-fold and 5.8-fold for cox1 and cob, respectively; Figure 2a) but also in ∆rtg2 (Figure 3f) and ∆hap4 (Figure 4f), suggesting that this is a direct effect of intron removal rather than a downstream consequence of activating the retrograde response. To investigate whether elevated cox1 and/or cob transcript levels might underpin the wider transcriptional, metabolic and phenotypic changes, we introduced attenuating point mutations into the promoters of cox1 and cob (see Materials and methods). We find that simultaneous attenuation of both promoters in I₀ fully reverses the I₀-characteristic suit of morphological and molecular phenotypes (see I₀pp throughout Figure 5). Attenuation of either cob (I₀cobp) or cox1 (I₀cox1p) in isolation only partially reverses the phenotype, although I₀cobp has a larger relative effect than I₀cox1p (Figure 5).

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Next, we sought to establish why the wholesale deletion of self-splicing introns leads to increased abundance of the host transcripts. We considered two main possibilities. First, in deleting introns, we might have inadvertently removed DNA-/RNA-level regulatory elements that affect expression of the host genes. Alternatively, the act of short-circuiting the splicing process itself might interfere with normal expression. Specifically, we hypothesized that normal levels of transcription might be tuned to accommodate a certain proportion of transcripts that fail to splice correctly. Group II introns in particular are known for low splicing efficiency, even in the presence of auxiliary proteins (Karunatilaka et al., 2010). As a corollary, a large fraction of pre-mRNAs might be targeted and degraded by mitochondrial quality control, either because splicing is erroneous (mis-splicing) or does not occur in a timely manner so that the transcript is shunted into degradation (kinetic coupling). In I₀, splicing does not occur so that erroneous splicing products do not arise. As a result, production of functional cox1/cob mRNAs might overshoot its target and trigger a system-wide response, for example because altered COX1/COB levels upset dosage balance amongst respiratory complexes. To test this abnormally-efficient-maturation hypothesis and simultaneously rule out that mitochondrial stress is caused by the removal of DNA-/RNA-level regulatory elements, we sought to alter splicing efficiency by orthogonal means. To this end, we overexpressed the nuclearly encoded DEAD box RNA helicase Mss116, which promotes splicing of all S. cerevisiae mitochondrial introns by remodeling or stabilizing splice-relevant RNA structures in an ATP-dependent manner (Karunatilaka et al., 2010; Huang et al., 2005). We confirmed overexpression and mitochondrial localization of Mss116 by flow cytometry and immunofluorescence, respectively, using an N-terminal His-tagged version of the protein (Figure 1—figure supplement 3), and then characterized the effects of Mss116 overexpression in the Y258 strain using an untagged version of the protein. Remarkably, the Mss116 overexpression strain (Mss116_OE) phenocopies I₀.

Mss116_OE exhibits increased generation times, extended chronological life span, lower competitive fitness, increased mitochondrial fusion, 2.9-fold increased mtDNA copy number (qPCR, t-test p=5.5×10⁻⁷), elevated oxygen consumption, and altered ATP and ROS production (Figure 1, Figure 1—figure supplement 1). In addition, we observe longer replicative life span (RLS) in Mss116_OE (Figure 1—figure supplement 4), a more direct proxy of ageing that we were not able to measure accurately in I₀, where separating mother and daughter cells in a timely fashion proved challenging (see Materials and methods). Overexpression of a DEAD box mutant of Mss116 (Mss116^E268K), which lacks ATPase and therefore helicase activity, does not phenocopy I₀ (Figure 1). This suggests that the role of Mss116 in splicing – which relies on helicase activity – is critical rather than a recently suggested ATP-independent role in transcription elongation (Markov et al., 2014). More generally, the fact that Mss116_OE – which encodes a full complement of introns – phenocopies I₀ indicates that the stress phenotype in these strains is not caused by missing DNA-level functionality and instead points toward a critical role for splicing. Overexpressing Mss116 in I₀ did not reveal additional phenotypes and the strain behaved like I₀ (Figure 2—figure supplement 1) further supporting the notion that intron deletion and Mss116 overexpression act through the same pathway.

We suspected that phenotypic effects of Mss116 overexpression (and intron deletion) are linked to altered transcript maturation of the host genes, cob and cox1. However, we first wanted to rule out an alternative hypothesis. In both Mss116_OE and I₀, Mss116 is in excess relative to need (in Mss116_OE because Mss116 is overexpressed, in I₀ because its usual targets – the introns – are absent). Phenotypic effects could therefore be caused by excess Mss116 interacting with RNAs that it would not normally target or not target to the same extent. This is conceptually related to the idea that splicing dynamics can change simply as a result of altered competition between mRNAs for access to the spliceosome (Munding et al., 2013). To test this hypothesis, we deleted Mss116 in the I₀ strain. We observe the same suite of phenotypes we see in I₀ (Figure 2—figure supplement 2), demonstrating that neither absolute nor relative excess of Mss116 are responsible for the stress phenotype described above.

Having ruled out this hypothesis, we then focused on characterizing the effects of Mss116 overexpression on cob/cox1 splicing dynamics in greater detail. Using qPCR, we first measured the levels of total cox1 and cob transcripts as well as individual introns at steady state (Figure 2a, 240 mins after Mss116 induction, see Figure 2—figure supplement 3). For cox1, we additionally monitored exon and intron (aI2, aI5β) levels using RNA fluorescent in situ hybridization (RNA FISH, Figure 2b, see Materials and methods). We find that most mitochondrial introns are strongly depleted in Mss116_OE compared to the empty vector control and relative to exons. The relative depletion of individual introns is somewhat variable and one intron – aI5α - is equally abundant in Mss116_OE and the Y258 WT (Figure 2a). We then examined shifts in the abundance of pre-mRNA and mature mRNA over the course of Mss116 induction, using different primer combinations to monitor unspliced RNAs and spliced exon-exon junctions. For both cox1 and cob, Mss116 overexpression shifts the balance between unspliced (or partially spliced) pre-mRNA transcripts, which dominate the uninduced steady state, towards mature mRNA transcripts (Figure 2c), while total transcript abundance (mRNA plus pre-mRNA) remains largely unchanged. These observations are consistent with a model where nascent transcriptional output is unchanged in Mss116_OE compared to the empty vector control and differential steady state levels are the result of post-transcriptional events.

Based on these findings, we suggest that Mss116_OE phenocopies I₀ because eliminating introns at the DNA level (I₀) and facilitating accurate and efficient excision at the RNA level (Mss116_OE) both result in abnormally efficient transcript maturation. That is, fewer transcripts are eliminated by mitochondrial quality control because splicing is erroneous or does not proceed in a timely manner, resulting in a greater number of mature cox1/cob mRNAs. For reasons that remain to be elucidated, increased transcript levels are then perceived as stressful and trigger the retrograde response, culminating in a multifaceted stress phenotype. We speculate in this regard that elevated protein levels of COB (I₀: 9.3-fold; Mss116_OE: 5.0-fold upregulation, as determined by quantitative label-free mass spectrometry) and COX1 (I₀: 11.6-fold; Mss116_OE: 12.1-fold upregulation) might interfere with proper assembly and function of complex III and complex IV, respectively, and therefore constitute a deleterious dosage imbalance phenotype.

Strain a161 (also known as ID41-6/161, or sometimes simply 161), described by Wenzlau et al. (1989), and the intronless a161-U7 (I₀) were gifts from Alan Lambowitz. These strains were used previously to show that splicing of group I and II introns is Mss116-dependent (Huang et al., 2005). a161 and a161-U7 are isogenic except for the mitochondrial genome and a single marker gene (a161: MATa ade1 lys1; a161-U7: MATa ade1 lys1 ura3).

Strain Y258 and the pBG1805 plasmid bearing Mss116 for overexpression were purchased from Thermo Scientific (Dharmacon, Lafayette, CO). Mss116^E268K was purchased from DNA 2.0 and cloned into pBG1805 using standard cloning techniques (Sambrook and Maniatis, 1989). Mss116 and Mss116^E268K were overexpressed in the Y258 nuclear background. The expression of Mss116 and Mss116^E268K was induced from the plasmids using 2% galactose (final), added at OD 0.2.

All strains were grown on YPD medium with 2% (w/v) glucose at 30°C with shaking. All experiments were performed on exponentially growing cells: cells were grown to OD 0.6–0.7 for WT, I₀, ΔHap4, and ΔRtg2 and to OD 0.9–1.0 for Mss116_OE and Mss116^E268K, harvested by centrifugation at 4000 × g for 5 min, washed and further treated as required.

In order to test growth on glycerol, strains were grown in YPD medium until saturation, at 30°C with shaking. Stationary cells were serially diluted and 5 μL drops plated onto YPEG agar plates (containing 3% ethanol and 3% glycerol). Growth was observed after 4 days.

Deletion of hap4 and rtg2 was performed as previously described (Rothstein, 1983), using a hygromycin cassette for selection in the WT background and a nourseothricine cassette in the I₀ background. Primers used for the deletions are listed in Supplementary file 1.

Overnight cultures of Mss116_OE were diluted to OD 0.1. Galactose (2% final) was added at OD 0.2 to induce the expression of Mss116, and this is designated as time 0. Aliquots of the culture were harvested at 30, 60, 90, 120, 135, 150, 180, 210 and 240 min post-induction. The cells were pelleted and used for RNA isolation and cDNA preparation for qPCR. Multiple primer pairs were used to monitor unspliced intron-exon fragments and spliced exon-exon junctions (Figure 2—figure supplement 4, Supplementary file 1). Additional aliquots were harvested at 60, 150, 300, 360, 420, and 540 min post-induction and were used to measure Mss116 expression levels by flow cytometry (see below).

In order to introduce promoter-attenuating mutations into mitochondrial DNA, we followed the protocol described in Bonnefoy et al., 2007) for the integration of altered mtDNA sequences by homologous double crossovers. Briefly, a mutant fragment of mtDNA (in this case promoter sequences) flanked by WT mtDNA sequence is first transformed into a rho⁰ strain, which is then mated with a recipient rho⁺ strain. Upon mating, mitochondria from the two strains fuse and recombination between the two mtDNAs produces recombinant rho⁺ strains in which the new mtDNA sequence is integrated by double crossover. For transformation, tungsten powder was used as a carrier of DNA (Tungsten M-10 Microcarriers #1652266, BioRad, Hercules, CA). Bombardment was performed using the Biolistic PDS-1000/He particle delivery system (BioRad). Cells were transformed with linear DNA fragments obtained by ligation of each mutated promoter region with 500 bp of up- and downstream flanking DNA (Supplementary file 1). SacI and SalI restriction sites were added by PCR for ligation between the 3’-end of the upstream flanking region and the 5’-end of the promoter sequence, and 3’-end of the promoter sequence and 5’-end of the downstream flanking region, respectively. The mutations introduced here (highlighted in Figure 5—figure supplement 1) have been previously shown to reduce the strength of cox1 and cob promoters (Turk et al., 2013).

All strains were grown to saturation as described above and pelleted at 4000 × g for 5 min. Cells were then washed twice and resuspended in sterile deionized water (10⁶ cells in 10 mL in order to avoid cell growth on the debris of dead cells) and incubated at 30°C with shaking. Every 2–3 days, cells were serially diluted and plated onto YPD plates in order to evaluate cell growth.

Replicative lifespan (RLS) was determined by micromanipulation for Mss116_OE and Mss116^E268K in the Y258 background, as well as for Mss116_OE in the Δhap4 and Δrtg2 nuclear backgrounds, counting the number of daughters produced by individual mother cells. We were unable to reliably determine RLS for the I₀ strain as daughter and mother cells could not be separated in a timely manner, which is critical for RLS measurements. For unknown reasons, and perhaps specific to the nuclear background, cells were unusually sticky so that the first daughter often could not be separated from the mother until the mother had already produced other buds, making it difficult to track mother/daughter identity over time, an essential prerequisite for reliably determining replicative lifespan. Cells were incubated at 30°C on YPD (WT) or -URA (mutants) plates for the duration of the experiment. Using a microscope equipped with a microdissection apparatus suitable for S. cerevisiae (Singer Instruments, Watchet, UK), cells were transferred to defined places on agar plates and virgin daughter cells collected. Each cell was monitored continuously over several days every 60–90 min until all mother cells stopped budding. The total number of daughter cells was noted for each mother cell. The total number of monitored mother cells is as follows: 90 cells for the empty vector control, 86 cells for Mss116_OE, and 91 cells for the Mss116^E268K. The measurements were pooled from three independent experiments.

Oxygen uptake was monitored polarographically using an oxygraph equipped with a Clark-type electrode (Oxygraph, Hansatech, Norfolk, UK). Cells were harvested during exponential growth phase, spun and resuspended in growth medium (as above) at the density of 30 × 10⁶ cells/mL. 500 μL of culture were transferred to an airtight 1.5 mL oxygraph chamber. Cells were assayed in conditions closely similar to the ones in a flask culture (30°C and stirring). Oxygen content was monitored for at least 4 min. To ensure that the observed oxygen consumption was due to the mitochondrial activity, complex III inhibitor antimycin (final concentration 10 μg/mL) was routinely added to the cultures and compared to the rate observed without antimycin.

Competition experiments were carried out between I₀ (a161-U7) and its control strain (a161) as well as between Mss116_OE and its corresponding empty vector control strain. Prior to the competition, we plated 5 × 10⁹ cells of each mutant (I₀ or Mss116_OE) strain on YPD agar plates with 200 μg/mL geneticin, followed by a 4-day incubation at 30°C, to select for spontaneous geneticin resistance. In doing so, we can subsequently determine relative fitness in a relatively simple fashion using a plating method (rather than, for example, sequencing barcodes). The two strains to be competed were then grown independently on YPD medium in 2% glucose until saturation. The next day, an equal number of cells from each WT culture (geneticin-sensitive) and each mutant (geneticin-resistant) were mixed in fresh YPD medium, 2% glucose so that each was diluted 200x. In mid-exponential phase, OD 0.4–0.6 (approximately 5–6 hr after dilution), aliquots of cells were harvested, and serial dilutions were plated on YPD-agar plates without geneticin. Next, dilutions with between 50 and 200 colonies were replica-plated on YPD-agar plates with geneticin (200 μg/mL). Colonies were counted on both types of plates and the ratio of geneticin-resistant (mutant) to geneticin-sensitive (WT) colonies was calculated as a measure of relative fitness.

Flow cytometry was carried out on a Becton-Dickinson (Franklin Lakes, NJ) FACSCalibur machine equipped with a 488 nm Argon laser and a 635 nm red diode laser.

The expression level of Mss116 in the Mss116_OE strain was measured by using a rabbit polyclonal anti-His tag antibody (Abcam, ab137839, 1:10000) and secondary IgG goat anti-rabbit labeled with Alexa 488 (Thermo Fisher Scientific, Waltham, MA, A11034, 1:2000). The signal obtained by flow cytometry (mean fluorescence over 10000 cells) was compared to the Mss116 tagged with GFP (Thermo Fisher Scientific) endogenous expression level estimated by using flow cytometry measurement based on the GFP signal. The mean fluorescence intensity in Mss116_OE was normalized to the mean fluorescence intensity detected in wild-type cells with endogenous expression of Mss116.

Isolated mitochondrial fractions containing 100 µg of protein were loaded onto Microcon 30kD centrifugal filters (Merck Millipore, Burlington, MA, MRCF0R030). Samples were then digested using a Filter Aided Sample Preparation (FASP) protocol (Wiśniewski et al., 2009). Briefly, samples were concentrated on the filter unit by centrifugation and buffer exchanged using sequential washing and centrifugation with 8M urea, 100 mM TRIS/HCL buffer (pH8.5). Proteins were reduced and alkylated sequentially with 10 mM Dithiothreitol and 50 mM Iodoacetamide (in 8M urea buffer), respectively. Samples were buffer exchanged to remove salts using sequential washing with 50 mM ammonium bicarbonate (AmBic). Trypsin Gold (Promega, Fitchburg, WI, V5280) was added to the samples in 50 mM ammonium bicarbonate to an approximate 1:50, protease:protein ratio. Digestions were incubated at 37°C overnight (17 hr). Digest extracts were recovered from FASP filters via centrifugation and acidified with 1% trifluoroacetic acid (TFA). Acidified protein digests were desalted using Glygen C18 spin tips (Glygen Corp, Columbia, MD, TT2C18.96) according to the manufacturer’s recommendation and peptides eluted with 60% acetonitrile, 0.1% formic acid (FA). Eluents were then dried using a vacuum centrifuge.

Protein digests were redissolved in 0.1% TFA by shaking (1200 rpm) for 30 min and sonication on an ultrasonic water bath for 10 min, followed by centrifugation (14,000 rpm, 5°C) for 10 min. LC-MS/MS analysis was carried out in technical duplicates (1 µg on column) and separation was performed using an Ultimate 3000 RSLC nano liquid chromatography system (Thermo Fisher Scientific) coupled to a Orbitrap Velos mass spectrometer (Thermo Fisher Scientific) via an Easy-Spray source. For LC-MS/MS analysis protein digests were injected and loaded onto a trap column (Acclaim PepMap 100 C18, 100 μm × 2 cm) for desalting and concentration at 8 μL/min in 2% acetonitrile, 0.1% TFA. Peptides were then eluted on-line to an analytical column (Easy-Spray Pepmap RSLC C18, 75 μm × 50 cm) at a flow rate of 250 nL/min. Peptides were separated using a 120 min gradient, 4–25% of buffer B for 90 min followed by 25–45% buffer B for another 30 min (composition of buffer B – 80% acetonitrile, 0.1% FA) and subsequent column conditioning and equilibration. Eluted peptides were analysed by the mass spectrometer operating in positive polarity using a data-dependent acquisition mode. Ions for fragmentation were determined from an initial MS1 survey scan at 30,000 resolution, followed by CID (Collision-Induced Dissociation) of the top 10 most abundant ions. MS1 and MS2 scan AGC targets were set to 1e6 and 3e4 for maximum injection times of 500 ms and 100 ms respectively. A survey scan m/z range of 350–1500 was used, normalised collision energy set to 35%, charge state screening enabled with +1 charge states rejected and minimal fragmentation trigger signal threshold of 500 counts.

Data was processed using the MaxQuant software platform (v1.5.6.0), with database searches carried out by the in-built Andromeda search engine against the Uniprot S. cerevisiae database (version 20160815, number of entries: 6,729). A reverse decoy database approach was used at a 1% false discovery rate (FDR) for peptide spectrum matches. Search parameters included: maximum missed cleavages set to 2, fixed modification of cysteine carbamidomethylation and variable modifications of methionine oxidation, asparagine deamidation and N-terminal glutamine to pyroglutamate conversion. Label-free quantification was enabled with an LFQ minimum ratio count of 2. ‘Match between runs’ function was used with match and alignment time limits of 1 and 20 min, respectively. Data have been deposited in the PRIDE repository [project accession number PXD008785].

Variations of the mitochondrial transmembrane potential (ΔΨm) were studied using 3,3-dihexyloxacarbocyanine iodide [DiOC6(3)]. This cyanine cationic dye accumulates in the mitochondrial matrix as a function of ΔΨm (Perry et al., 2011). Cells (1 × 10⁶/mL) were incubated in 1 mL culture medium containing 40 nM DiOC6(3) for 30 min in the dark at 30°C with constant shaking. DiOC6(3) membrane potential-related fluorescence was recorded using FL1 height. A total of 10,000 cells were analyzed for each curve. The collected data was analyzed using FlowJo software version 7.2.5 to determine the mean green fluorescence intensity after each treatment. The results are expressed as a percentage of mean fluorescence of the control strain. As a negative control, in each experiment, we preincubated aliquots of cells with carbonyl-cyanide 4-(trifluoromethoxy)- phenylhydrazone (FCCP, Sigma) and antimycin (Sigma, St. Louis, MO) at 100 μM and 5 μg/mL, respectively, 10 min before fluorescent dye staining, which leads to a collapse of mitochondrial membrane potential.

To measure mitochondrial mass, we used 10-N-Nonyl acridine orange (NAO), a dye that binds to cardiolipin, a phospholipid specifically present on the mitochondrial membrane (Perry et al., 2011). Cells (1 × 10⁶/mL) were incubated in 1 mL culture medium containing 100 nM NAO for 30 min in the dark at 30°C with constant shaking, followed by analysis on FACSCalibur flow cytometer with the same photomultiplier settings as used for DiOC6(3).

To image mitochondrial morphology, strains were transformed with a MitoLoc plasmid (Vowinckel et al., 2015) (a gift from Markus Ralser) according to a previously described protocol (Gietz and Schiestl, 2007), with the only difference that the cells were incubated with the plasmid overnight at room temperature. Microscope slides were prepared as follows: 150 μL of YPD media containing 2% agarose was placed on a preheated microscope slide and cooled, before applying yeast cells to obtain a monolayer. The cells were centrifuged at 4000 × g for 3 min, and resuspended in 50 μL YPD. Once dry, the cover slip was placed, sealed, and mounted on a temperature-controlled Nikon Ti-E Eclipse inverted/UltraVIEW VoX (Perkin Elmer, Waltham, MA) spinning disc confocal setup, driven by Volocity software (version 6.3; Perkin Elmer). Images were recorded through a 60xCFI PlanApo VC oil objective (NA 1.4) using coherent solid state 488 nm and 543 nm diode lasers with a DPSS module, and a 1000 × 1000 pixel 14-bit Hamamatsu (C9100-50) electron-multiplied, charge-coupled device (EMCCD). The exposure time was 100 ms for GFP and 300 ms for mCherry, at 5–10% laser intensity. The number of cells with cytosolic mCherry accumulation was counted manually. More than 1000 cells were examined for each strain. Images were analyzed using ImageJ software with the MitoLoc plugin.

Cells were incubated in the dark with 5 μM MitoSOX red mitochondrial superoxide indicator (Molecular Probes, Eugene, OR) for 10 min at 30°C and subsequently analyzed by flow cytometry. Fluorescence (excitation/emission maxima of 510/580 nm) of 10,000 cells resulting from the intracellular red fluorescence was measured in the FL2 channel. The collected data was analyzed using FlowJo software version 7.2.5 for Microsoft (TreeStar, San Carlos, CA) to determine the mean green fluorescence intensity after each treatment. The results are expressed as the mean fluorescence across 10,000 cells.

Total RNA was isolated using the NucleoSpin RNA kit (Macherey and Nagel, Dueren, Germany) according to the manufacturer’s instructions for up to 3 × 10⁸ yeast cells, which includes incubation with 50–100U of zymolyase for 1 hr at 30°C. The quality of the resulting total RNA was tested on 1% agarose gels.

Cells from saturated cultures (approximately 10⁹ cells) of WT Y258, WT a161, I₀, and Mss116_OE were harvested by centrifugation and washed, as described above. Cell wall was digested for 1 hr at 30°C in the presence of 50–100U of zymolyase. Spheroplasts were then resuspended in 500 μL of cell lysis buffer (75 mM NaCl, 50 mM EDTA, 20 mM HEPES pH 7.8, 0.2% SDS). Next, 10 μL of 20 mg/mL proteinase K was added and the mixture incubated for 2 hr at 50°C. DNA was precipitated by the addition of isopropanol at room temperature, followed by centrifugation at 4°C and 11,000 g for 30 min. The DNA pellet was then washed with ice-cold 70% ethanol, air-dried and resuspended in 50 μL of DNase-free water at 55°C.

cDNA was synthesized from 1000 ng of total RNA using the iScript cDNA Synthesis Kit (Biorad). The cDNA was diluted 100-fold, mixed with primer pairs for each gene and SYBRgreen (BioRad). All primer pairs were designed to have a melting temperature of 60°C and are listed in Supplementary file 1. The qPCR reaction was run on a QuantFlexStudio 6 (Life Technologies, Carlsbad, CA) using 40 cycles, after which the melting curves for each well were determined. Final fold change values were estimated relative to the UBC6 gene in the control strain replicates. mtDNA copy number was assessed by qPCR using genomic DNA as template and primers against cox1 and cox3 (mtDNA) and rpl32 (nuclear DNA, for normalization).

Individual cells (approximately 100 for each strain) were placed on agar plates of appropriate growth medium, as described above, using a micromanipulator. Next, an image of each original mother cell was taken every 10 min for 8–9 hr. The images were then analyzed and division time of each cell was extracted.

Yeast cultures were grown as described above, fixed with 37% formaldehyde for 45 min at room temperature, digested with 2.5 μL of zymolyase (Zymo Research, Irvine, CA, 2000 U) at 30°C for 60 min and permeabilized with 70% ethanol overnight at 4°C. Cells were hybridized in the dark at 30°C using Stellaris RNA-FISH probes (Biosearch Technologies, Novato, CA). Forty-five probes targeting intron aI2 and 40 probes targeting intron aI5b cox1 were coupled to Quasar 670 dye (red). Forty-three probes targeting cox1 exons were coupled to Quasar 570 dye (green). Yeast cells were placed on microscope slides with Vectashield Mounting Medium and imaged with an Olympus IX70 wide-field fluorescence microscope. A series of z-stacks was acquired with a step size of 0.3 μm. The images were analysed using Image J. The number of green (exon), red (intron) and yellow (colocalized) foci was manually counted and normalized per 100 cells in each of three biological replicates. At least 300 cells were analyzed per replicate per strain.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

The authors would like to thank Tea Copic, Dirk Schwitters, Iva Pesun Medjimorec, Martin Billman, Sam Marguerat, and Xi Ming Sun for experimental assistance, Holger Kramer and the LMS Proteomics facility for proteomics work, and Nuno Raimundo, Ira Milosevic, Santiago Vernia, and Laurence Hurst for discussions. This work was supported by funding from the NAOS Group and the Mediterranean Institute for Life Sciences (MR, MM, MK, AK), an Imperial College Junior Research Fellowship (TW) and UK Medical Research Council core funding (TW).

1. Received: January 23, 2018
2. Accepted: March 19, 2018
3. Accepted Manuscript published: March 23, 2018 (version 1)
4. Version of Record published: April 13, 2018 (version 2)

© 2018, Rudan et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.