Resistance of mitochondrial DNA to cadmium and Aflatoxin B1 damage-induced germline mutation accumulation in C. elegans.

Mitochondrial DNA (mtDNA) is prone to mutation in aging and over evolutionary time, yet the processes that regulate the accumulation of de novo mtDNA mutations and modulate mtDNA heteroplasmy are not fully elucidated. Mitochondria lack certain DNA repair processes, which could contribute to polymerase error-induced mutations and increase susceptibility to chemical-induced mtDNA mutagenesis. We conducted error-corrected, ultra-sensitive Duplex Sequencing to investigate the effects of two known nuclear genome mutagens, cadmium and Aflatoxin B1, on germline mtDNA mutagenesis in Caenorhabditis elegans. Detection of thousands of mtDNA mutations revealed pervasive heteroplasmy in C. elegans and that mtDNA mutagenesis is dominated by C:G → A:T mutations generally attributed to oxidative damage. However, there was no effect of either exposure on mtDNA mutation frequency, spectrum, or trinucleotide context signature despite a significant increase in nuclear mutation rate after aflatoxin B1 exposure. Mitophagy-deficient mutants pink-1 and dct-1 accumulated significantly higher levels of mtDNA damage compared to wild-type C. elegans after exposures. However, there were only small differences in mtDNA mutation frequency, spectrum, or trinucleotide context signature compared to wild-type after 3050 generations, across all treatments. These findings suggest mitochondria harbor additional previously uncharacterized mechanisms that regulate mtDNA mutational processes across generations.

INTRODUCTION

Mitochondria are vital organelles that provide energy and important signaling molecules for almost every form of eukaryotic life on earth. Phylogenetic evidence strongly suggests that mitochondria share ancestry with alpha-proteobacteria and arose by endosymbiosis. Over the course of 1.45 billion years of coevolution, most genes encoding mitochondrial proteins have translocated into the nuclear genome. Nevertheless, mitochondria still maintain a unique genome that encodes various electron transport chain subunit proteins that are essential for energy production (1).

mtDNA mutations are implicated in diseases affecting at least one person in 5000 (2), in addition to potential roles in many neurological and metabolic disorders, various cancers, and aging (3–5). MtDNA mutation rates are often 100 times higher than nuclear DNA mutation rates in aging and over evolutionary time (6,7,8). Currently, replication errors by the sole mitochondrial DNA polymerase, Pol γ, rather than oxidative stress, are thought to be the main source of mtDNA mutations (9–11). Pol γ contains 3’-5’-exonuclease and 5’-deoxyribose lyase activities which allow for proofreading during mtDNA replication and base excision repair (12). Pol γ has limited translesion synthesis capabilities, potentially rendering mtDNA sensitive to damage-induced mutations or copy number changes due to polymerase stalling at bulky adducts (13). Mitochondria have efficient base excision repair mechanisms, but lack nucleotide excision repair and mismatch repair machinery, which may also contribute to higher mutation rates (14).

Chemical exposure can contribute to nuclear DNA mutagenesis, but few studies have investigated the role of genotoxicants in mtDNA mutagenesis. Evidence is limited regarding the effects of exogenous stress on mtDNA damage and somatic mutagenesis (15), and to our knowledge, the effects of chemical exposure on germline mtDNA mutagenesis have yet to be investigated. This is critical to understand because the basic biology of mitochondria renders mtDNA particularly susceptible to the harmful effects of environmental toxicants and stress. The negative charge of the mitochondrial matrix and Ca2+ transporters results in accumulation of cationic heavy metal pollutants, and the phospholipid bilayer attracts lipophilic organic pollutants which can be activated to toxic (particularly genotoxic) metabolites by mitochondrial cytochrome P450s (16–18).

Though mitochondria are limited in DNA repair capacity, organelle dynamics play a significant role in eliminating irreparable damage and maintaining mitochondrial homeostasis. The importance of mitochondrial fission, fusion, and mitophagy in regulating mitochondrial quality are highlighted through manifestation of disease phenotypes from mutations in nuclear encoded genes (19). For example, mutations in nuclear mitophagy genes PARK2 and PINK1 are associated with Parkinson's disease. Mitochondrial dynamics are also important for mediating response to mitochondrial stress, whether metabolic or chemical, as we have reviewed in the context of pollutant stress (20). A damaged organelle can fuse with a healthy organelle, and subsequently undergo fission to create healthy daughter mitochondria (‘complementation’). Alternatively, damaged mitochondria may be selectively targeted for degradation via mitophagy. We have previously demonstrated that genetic disruption of these pathways in Caenorhabditis elegans results in increased sensitivity to various stressors and inability to remove ultraviolet C radiation-induced mtDNA damage (21–24). Until this study, variation in accumulation of mtDNA damage associated with mitophagy function after chemical exposure had not been investigated.

Mitophagy is also thought to regulate the selective inheritance of mitochondrial genomes in the germline, where mtDNA is, in most metazoans, uniparentally inherited. Despite the high mtDNA copy number in oocytes, only a few mtDNA genome copies are distributed into primordial germ cells, an event that is referred to as the germline mitochondrial ‘bottleneck’ (25). Evolutionary models predict that the bottleneck increases the probability of removal of deleterious mtDNA variants (26). Whether transmission of mutant mitochondrial genomes is stochastic, permitting genetic drift, or targeted via purifying selection, is controversial (27–30).

Previous studies in C. elegans have demonstrated that mitophagy can play an important role in regulating the germline transmission of mtDNA variants. For example, progeny of C. elegans that lack functional Parkin (pdr-1) accumulate higher frequencies of the 3.1 kb uaDf5 mtDNA deletion compared to wild-type (31). However, the role of mitophagy in regulating the transmission of de novo single nucleotide mutations through the germline remains an exciting area of investigation, particularly in the context of exposure to chemicals that cause mtDNA damage. A recent study by Haroon et al. suggests that mitophagy may play a role in mtDNA single nucleotide mutagenesis, as C. elegans deficient in pdr-1 accumulate higher frequencies of mtDNA point mutations than wild-type, though only in a Pol γ exonuclease mutant background (32). Therefore, we hypothesized that exposure to mtDNA-damaging chemicals would result in higher rates of mtDNA mutation accumulation, and that this would be exacerbated in mitophagy-deficient C. elegans.

C. elegans is a well-established organism in which fundamental knowledge of spontaneous mtDNA mutational processes and other evolutionary insights (including genotoxin-induced nuclear mutation studies) has been achieved through classical mutation accumulation line experiments (33). Two independent mutation accumulation line (MA) experiments have been conducted in wild-type C. elegans, though only Konrad et al. conducted next-generation Illumina sequencing (8,34). A third MA experiment was conducted in the C. elegans mutant gas-1, which renders complex I of the ETC dysfunctional, thereby resulting in increased reactive oxygen species production and reduced ATP production (35). However, no other MA approach has investigated the effect of either chemical stressors or mitophagy on mtDNA mutagenesis in C. elegans.

A limitation of previous MA studies in C. elegans is that only a few mtDNA variants that arose to high frequencies or fixation were detected, with one very recent exception (36). However, it is now known that most mtDNA variants are not only heteroplasmic, but exist at very low frequencies that are below the error rate of standard next-generation Illumina sequencing assays. Therefore, we used a powerful sequencing approach, Duplex Sequencing, that permits accurate detection of mtDNA variants at frequencies as low as one mutation in ∼108 bp (37–39). Using this ultra-sensitive, mtDNA-targeted sequencing approach, we were able to detect more heteroplasmic mtDNA mutations at lower frequencies than ever before reported in C. elegans, and also investigated the mutational rates, spectrum, context, and genomic sites of mutagenesis after a long-term mutation accumulation experiment.

To test the effects of chemical stress and the role of mitophagy in the origin and transmission of mtDNA mutagenesis in C. elegans, we performed mutation accumulation experiments in wild-type and two mitophagy mutant strains, pink-1 and dct-1, under various environmental conditions. All independent lineages (replicates) for each of the three strains were bottlenecked for 50 generations in control conditions or with chronic exposure for 50 generations to the heavy metal pollutant, cadmium (Cd, in the form of cadmium chloride, CdCl2) or the mycotoxin aflatoxin B1 (AfB1), both known nuclear mutagens and human carcinogens that have two different mechanisms of DNA damage. Cd inhibits many DNA repair enzymes and interferes with antioxidant enzymes, resulting in increased levels of ROS that can damage mtDNA (40,41). AfB1 metabolites form bulky DNA adducts, which can inhibit replication and result in somatic mutations in the nuclear genome (42). The accumulation of AfB1 in mitochondria, in addition to lack of nucleotide excision repair (NER), results in higher levels and greater persistence of mtDNA lesions compared to nuclear DNA damage, as we and others have previously measured (43–45). Therefore, we used these two toxicants as models to investigate how two different mechanisms of mtDNA damage may affect mtDNA mutational processes.

Duplex Sequencing of wild-type C. elegans mtDNA revealed a strong signature of oxidative damage. Exposure to a level of CdCl2 that is comparable to human blood levels and resulted in high levels of mtDNA damage, and a level of AfB1 that caused mtDNA damage and a significant increase in nuclear DNA mutation rate, did not increase the frequency of mtDNA single nucleotide mutations (SNMs) in wild-type C. elegans. Surprisingly, inhibiting mitophagy did not result in an increase in mtDNA mutations in control conditions or after exposure to CdCl2 or AfB1, despite declines in fitness compared to wild-type C. elegans. These results suggest that the mitochondrial genome harbors robust mechanisms of avoiding chemical-induced mtDNA mutagenesis that may be independent of mitophagy.

MATERIALS AND METHODS

C. elegans strains and maintenance

This work used the C. elegans wild-type Bristol N2 strain and two mitophagy deficient strains, dct-1 and pink-1. The dct-1(tm376) mutant harbors a 912 bp deletion in the promoter region of dct-1 (DAF-16/FOXOControlled, germline Tumour affecting-1, putative orthologue to mammalian mitophagy receptor, BNIP3) and has been characterized previously (46). The pink-1(1779) mutant harbors a 350 bp deletion in pink-1 (PTEN-induced kinase 1) and exhibits altered mitochondrial morphology (22). These deletion strains were acquired from the National Bioresource Project (Tokyo, Japan), genotyped, and backcrossed into N2 six times (dct-1) and eight times (pink-1) prior to any experiments. All C. elegans were maintained following standard procedures (47). We replaced the potassium phosphate in traditional nematode growth medium (NGM plates) with KCl (‘K-agar plates’) in order to prevent buffering which reduces the bioavailability of CdCl2 (48,49). All strains were maintained at 20°C on OP50 Escherichia coli.

CdCl2 and AfB1 treatment

Stocks of CdCl2 and AfB1 (Sigma-Aldrich, St. Louis, MO) were made in ddH2O and DMSO vehicles, respectively, and stored at 4°C. Treatment plates were always made fresh: OP50 was spiked with either CdCl2 and AfB1 immediately prior to seeding, and plates were prepared two days prior to experiments. Reported concentrations are the concentrations in the spiked OP50. 100 μl of OP50 was seeded on 6cm plates (containing 8 ml of K-Agar) and 300 μl of OP50 on 10cm plates (containing 20 ml K-agar).

Mitochondrial morphology

To assess effects of exposure on mitochondrial morphology, wild-type C. elegans harboring an extrachromosomal array Pmyo-3::GFP; which expresses GFP in the mitochondrial matrix of body wall muscle cells, were exposed to 10 μM and 50 μM of CdCl2 and 2 μM and 10 μM AfB1. Images were taken on a Keyence BZ-X700, and analyzed in ImageJ using the Blinder software (50). Blinded images were analyzed qualitatively by an established classification system as previously described (51), with Class 1 indicating highly networked, fused mitochondrial morphology, and Class 5 indicating extremely fragmented mitochondrial morphology. Two experimental replicates were conducted, and the total number of individuals analyzed is displayed above each stacked bar plot in Supplementary Figure S5. Statistical differences in the distribution of classes of mitochondrial morphology were determined by Fisher's exact test followed by Bonferroni correction for multiple comparisons.

Mitochondrial respiration

Age-synchronized L1 C. elegans were transferred to plates seeded with OP50 containing 50 μM and 100 μM CdCl2, or 10 μM and 25 μM AfB1. Approximately 48 h later, L4 C. elegans were washed off the plates and respiration parameters were quantified using the Seahorse Extracellular Flux Bioanalyzer, as previously described (52). The number of individual nematodes per well was counted, such that the Oxygen Consumption Rate (OCR) measurements were normalized per worm. The mean of technical replicates per plate was then determined. Two to six independent experiments were conducted per strain per treatment. Two-way ANOVAs were run for each chemical in order to compare Cd to control and AfB1to control across three strains.

Life history trait analyses

Growth was measured in three independent experiments in the ancestral (G0) wild-type, dct-1 and pink-1 strains on control and treatment plates. About 300 age-synchronized L1s were plated on each 10cm plate with the following conditions: control, 50, 200, 1000 μM CdCl2, and 10, 50 and 200 μM AfB1. After 48 h, L4 C. elegans were washed off the treatment plates and transferred to unseeded plates. Images of each plate were then captured with a Keyence BZ-X700 using brightfield, and analyzed in ImageJ with the Fiji plug-in WormSizer (53). Length data were normalized to the mean control length of each strain for each biological replicate. Box and whisker plots of length distributions were plotted in R and the means of biological replicates were used for statistical analysis (2-way ANOVA, Tukey's HSD post-hoc analysis). Total brood size was also determined in all three strains on control, 50 μM CdCl2 and 10μM AfB1 plates (N = 9–10 individuals per strain per treatment). On day 1 of adulthood, each individual worm was transferred to a new 6cm plate with the respective treatment until reproduction ceased. Plates containing offspring were stored at 20°C for 2 days, and then counted.

Population growth rate was measured on all MA lines at G0 and G50 to measure the effects of mutation accumulation on fitness. Population growth rate was determined by an eating race experiment as originally described by Hodgkin and Barnes (54). Each plate was monitored hourly up to the time when all bacteria had been consumed and the population dispersed. As an additional fitness measurement, total reproduction and lifespan were also determined on a subset of 20 MA lines per strain per treatment at G50. The 20 MA lines were determined with a random number generator. Of these 20 sublines, three L4 individuals (G50 sisters) were picked onto individual control plates. Founders were transferred to a new plate daily until cessation of egg-laying, and total brood was counted as described above. Biological replicates were then averaged per MA line. Individuals that were sterile were included in our analysis. Lifespan was determined by scoring individuals every day until they stopped responding to touch. Reproduction and lifespan results were censored if the animal was lost, dehydrated, or bagged. This only occurred in four of the 180 MA lines.

mtDNA copy number and damage

OP50 was spiked with either CdCl2 or AfB1, such that the final concentrations of chemicals in the bacterial lawn were 50 μM of CdCl2 and 10 μM of AfB1. Plates were seeded with 100 μl of each treatment, and control, and allowed to dry for 2 days. 20–30 gravid adults of each strain, wild-type, pink-1 and dct-1, were then picked onto each treatment plate. Adults were left on each plate for a 2-h egg-lay, after which all of the adults were removed from the plate. After 56 h, synchronized L4 C. elegans were then analyzed for mtDNA CN and damage, as described (55). Two to three biological replicates of six individual L4 C. elegans per strain per treatment were picked into 90 μl of lysis buffer, placed at −80°C for at least overnight, then lysed at 65°C for 1 h, followed by 95°C for 15 min to inactivate Proteinase K. Long-amplicon PCR was then performed (55), with one modification that the final reaction volume was halved (25 μl instead of 50 μl). mtDNA CN was measured using a plasmid-based standard curve and real-time PCR as described (56). mtDNA CN was normalized to the number of L4 individuals per sample (six) because nematodes have an invariant number of somatic cells (∼1000 cells), and thus invariant nuclear genome copies at this life stage. Long-amplicon PCR products were normalized to mtDNA copy number and mtDNA lesions were calculated relative to the control within each strain. Each biological sample was amplified in duplicate or triplicate. This experiment was performed twice. A two-way ANOVA followed by Tukey's HSD post-hoc analysis was used to determine statistical differences in mtDNA lesions between strains and treatment. Results of mtDNA lesions following exposure to additional concentrations of CdCl2 and AfB1 that were not used in the mutation accumulation line experiment are included in Supplementary Materials (Supplementary Figure S1). The effects of exposure on mtDNA CN are described in Supplementary Figure S2.

Mutation accumulation lines

Our criteria to determine a single concentration of CdCl2 and AfB1 to use for mutagenesis experiments were: (i) the concentration caused significant mtDNA damage, but (ii) did not have significant effects on organismal fitness (growth, fecundity and mitochondrial toxicity). Once we determined a concentration for each exposure, we performed a mutation accumulation line experiment in wild-type, pink-1 and dct-1 C. elegans, in control conditions, 50 μM CdCl2 or 10 μM AfB1 (Figure 2). One random young adult individual of each strain was isolated (G0 founder) and allowed to reproduce. Once the offspring reached gravid adulthood, about 20 individuals were picked onto a plate. After a 2-h egg lay, the adults were picked off the plate. When these offspring reached L4, one individual (G1 ‘subline’ with shared lineage from the founder) was randomly picked to an individual plate to begin the mutation accumulation experiment. The MA experiment began with 50 individual G1 sublines per strain per treatment. All lines were maintained at 20°C. Every four days, one individual L4 was randomly picked and transferred to a new plate to propagate the next generation. This population bottlenecking was conducted for 50 generations per strain per treatment per subline, resulting in a total of 2500 generations of mutation accumulation per strain per treatment. The previous generation was always maintained at 15°C, such that if an individual was sterile or dead, a back-up ‘sister’ individual could be transferred to a new plate to continue the MA line. As in previous C. elegans MA experiments, if three attempts were not viable, the line was then considered extinct (57). At the end of the experiment, the average percentage of extinctions per strain per treatment was 4.5% (Supplementary Figure S3). All 450 MA lines were cryopreserved every 5 generations for 50 generations.

Figure 2.

Schematic of mutation accumulation line experimental design. Offspring of a single founding ancestor (G0) were isolated onto individual plates. Every t = 4 days, a single L4 nematode was randomly selected and transferred to a new plate. This was repeated every generation (G) for 50 generations. We conducted MA lines on control plates, and plates that contained OP50 that was spiked with a final concentration of 50 μM CdCl2 and 10 μM AfB1. We conducted MA line experiments in wild-type C. elegans and two mitophagy mutant strains, dct-1 and pink-1. 50 replicate ‘sublines’ were passaged for each strain*treatment. MA lines were randomly selected after 50 generations per subline (9–14 MA lines/strain/treatment) for life history analysis and targeted mtDNA Duplex-Sequencing. Image created with BioRender.com.

DNA isolation, duplex sequencing library preparation and sequencing of mtDNA

After 50 generations of mutation accumulation per MA line, an individual L4 from each MA line was transferred onto a control 6-cm plate. As has been previously described in other MA experiments, as soon as all of the OP50 had been consumed and the population was composed largely of synchronized L1s, the C. elegans were immediately washed off of the plate with ddH2O, transferred to a 1.7 ml Eppendorf tube, pelleted, and flash frozen in liquid nitrogen (34–36). Total DNA was isolated with the DNeasy Blood & Tissue kit (Qiagen, Germany) following the manufacturer's instructions. DNA quantity and purity was analyzed using a Thermo Scientific™ NanoDrop™ One Microvolume UV-Vis Spectrophotometer (Thermo Fisher Scientific) and Qubit 2.0 (Perkin Elmer Victor x2).

Illumina Sequencing libraries were prepared following the original Duplex Sequencing (DS) protocol with slight modifications (58,59). An input of 50 ng of total DNA was used for library preparation. Total DNA was sheared (Covaris E210), followed by preparation and repair of DNA fragments using the NEBNext Ultra II Library prep kit, in which 2 μl of 15μM of the unique DS adapters were added directly to the Ultra II Ligation Master Mix, according to manufacturer's instructions (New England Biolabs). Relative mtDNA copy number of the post-adapter ligated sample was then determined via RT-qPCR (Applied Biosystems StepOnePlus). This critical step determined the amount of post-ligated DNA to use as input for the pre-capture indexing PCR to optimize the number of Duplex Consensus Sequences formed for each Unique Molecular Identifier (59). The optimal family size was determined by earlier sequencing of a C. elegans wild-type sample compared to a known reference standard in order to determine volume of post-adapter-ligated for pre-capture indexing PCR. On average, 0.338 μl of template was required (range of 0.164–0.632). After the pre-capture indexing-PCR, the entire product was lyophilized with the addition of uniquely designed blocking oligos. An enrichment of the C. elegans mitochondrial genome was then performed following the IDT xGen Hybridization Capture of DNA libraries for NGS target enrichment protocol with a custom designed Discovery Pool probe panel that covered 13 091 bp of the 13 991 bp C. elegans mitochondrial genome, capturing 93.5% of the genome. The AT-rich, highly repetitive non-coding region of the genome was excluded in order to minimize sequencing fragments that would not accurately map to the reference genome. Post-capture PCR and clean-up were then performed using according to manufacturer's instructions. Libraries were sequenced on an Illumina NovaSeq 6000 platform to obtain 150 bp paired-end reads (20 million per library). The DS adapter sequences, qPCR and Illumina primer sequences, blocking oligo sequences, and the C. elegans mitochondrial genome custom probe panel oligonucleotide sequences are provided in the Supplementary Data File.

Bioinformatics processing and analysis of mitochondrial genome sequences

Sequencing data were processed on the Duke Computer Cluster using the custom Duplex-Seq Pipeline (v1.1.4) workflow developed by the Kennedy Lab (github.com/Kennedy-Lab-UW/Duplex-Seq-Pipeline) and described in detail in Supplementary Figure 3 of Kennedy et al. (58) and again in Sanchez-Contreras et al. (11). Only reads mapping to the C. elegans mitochondrial reference genome were analyzed, and only single nucleotide mutations were analyzed, while insertions and deletions were ignored. Possible artifacts were removed by 12-bp end clipping at the beginning of each read. A minimum of three reads were required to call a variant, with a minimum heteroplasmy cutoff of 70% per Duplex Consensus Sequence (DCS) compared to the reference genome. Other input parameters are included in the Configuration File (Supplementary Data File). A summary output file of mutations, sequencing depth, and genome coverage per library is included (Supplementary Data File). Sequencing data is uploaded and can be accessed at SRA SRP350474 (PRJNA787252). We detected three polymorphisms that were completely fixed in our C. elegans strains compared to the reference genome (NC_001328.1), two of which have been previously detected (36). These polymorphisms were not included in our analysis and are listed in Supplementary Table S1.

Mutation frequency and spectrum analysis

Each mutation was only called once at each genomic position. Overall mutation frequency was calculated for each library with the following equation: [total number of unique mutations]/[total number of error-corrected nucleotides sequenced]. For mutational signature calculations, the total number of each type of the six possible transition and transversion mutations was divided by the coverage of the reference nucleotides sequenced (i.e.: [total # C:G → A:T mutations]/[total number of cytosines + guanines sequenced]). Mean mutation frequencies were calculated for each strain and treatment, followed by parametric statistical analysis and appropriate multiple comparisons corrections, as described in Results.

Trinucleotide context mutational signature

Trinucleotide context mutations were calculated using the Bioconductor Package MutationalPatterns (v1.1.0) (60) after forging a C. elegans mitochondrial reference genome (BSgenome v1.58.0). The number of mutations in each of the 96 possible trinucleotide contexts were determined in reference to pyrimidines C and T, hence six possible mutation types instead of 12. The number of each trinucleotide context mutation was normalized to the average number of wild-type control mutations in each context. A Welch two-sample t-test was performed on a specific mutation only after a significant ANOVA was determined.

Synonymous and nonsynonymous mutation analysis

The dNdS ratio was calculated in order to determine if mtDNA evolution departed from neutrality in our MA experimental design. We used the R package dNdScv, a statistical modeling approach that allows for normalization to depth of coverage as a covariate, and incorporation of the invertebrate mitochondrial genetic code (translation Table 5) (61).

Whole genome sequencing, processing, and de novo variant calling for nuclear DNA mutagenesis

We conducted whole genome sequencing of four control, four CdCl2, and four AfB1 wild-type MA samples to estimate nuclear DNA SNM rates. Sequencing libraries were created by the Duke University Genomics and Computational Biology Core from the same DNA templates that were used for Duplex Sequencing of the mitochondrial genome. Libraries were sequenced on an Illumina NovaSeq 6000 platform to obtain 150 bp PE reads.

Data quality control for the NGS raw fastq files was performed using the FastQC v0.11.8 to ensure no fundamental errors occurred during sequencing steps or library preparation of the samples. Reads were processed by Trimmomatic v0.39 to remove the 3’ end adapter sequences and trim low-quality sequences. Subsequently, sequence alignment of the processed reads was performed by mapping to the WBcel235 C. elegans Bristol N2 strain reference genome utilizing the BWA-MEM v0.7.17 aligner. SAMtools v1.14 flagstat was used to compute the number and percent of reads that mapped to the genome. Depth and breadth of coverage was computed using BEDTools v2.29.2. Post-alignment processing for the removal of read duplicates, indel realignment, and base quality score recalibration was completed using Picard v2.18.16 MarkDuplicates and the GATK v4.1.9.0 tools InDelRealigner and BaseRecalibrator. These post-alignment steps ensured high-quality aligned reads remained, which are necessary for the accurate detection of genomic variants including SNVs and CNVs. Germline variants were called using strelka v2.9.2 and population level SNPs and ancestral variants shared across all MA lines were filtered out. All variants were annotated with gene names, predicted function, population frequencies, and other variant annotations using Ensembl Variant Effect Predictor. Remaining de novo variants were filtered for depth, quality and 100% variant allele frequency.

Statistical analysis

All statistical analysis and data visualization were conducted in RStudio (v1.1.463; R v4.0.3). Values are conveyed as means and standard error, unless otherwise indicated. We performed Welch Two Sample T-test to determine significance between two groups, and ANOVA (one way if more than two groups and two way if multiple factors, as indicated in further detail in the figure legends) followed by Tukey's HSD to correct for multiple comparisons. Fisher's exact test was performed to determine variation in the proportion of each type of mutation (Supplementary Figure S4).

RESULTS

CdCl2 and AfB1 exposure causes mtDNA damage in wild-type C. elegans

We have previously shown that exposure to CdCl2 and AfB1 results in significantly higher levels of mtDNA damage than nuclear DNA damage in wild-type C. elegans (43). In order to determine a single dose of CdCl2 and AfB1 to use for mutation accumulation lines and sequencing where there was detectable mtDNA damage but no evident effect on fitness, we first conducted a dose response and quantified mtDNA damage (Supplementary Figure S1), followed by measuring growth, reproduction, mitochondrial morphology, and mitochondrial respiration. Exposure to 50 μM CdCl2 induced 0.92 lesions/10 kb, and 10 μM AfB1 induced 0.25 lesions/10 kb relative to control in wild-type C. elegans (P < 0.01, Figure 1A). These concentrations of CdCl2 and AfB1 did not result in statistically significant growth delay (8.3% and 6.3% growth inhibition, respectively, Figure 1B) or fecundity (Figure 1C). We also observed little effect on mitochondrial morphology (Supplementary Figure S5) and no decrease in mitochondrial function (Supplementary Figure S6). Therefore, these concentrations were used for the mutation accumulation experiments to avoid any effects on fitness that may cause selective pressure within MA lines and therefore compromise mutation rate calculations.

Figure 1.

50μM CdCl2 and 10 μM AfB1 exposure induces mtDNA lesions in wild-type C. elegans, but has no effect on growth or reproduction. (A) mtDNA damage from pools of six individual age-synchronized L4 C. elegans was quantified after chronic exposure to control (N = 8), 50 μM CdCl2 (N = 8) or 10 μM AfB1 (N = 4) (** P< 0.01, one-way ANOVA). (B) A dose–response was conducted to measure effects on growth after 48 h of exposure (L1–L4). Each individual worm length was first normalized to the mean control length for each experiment. Dots represent technical replicates (individual nematodes, total N displayed) and the boxplots display the median and upper and lower quartiles. The mean normalized length was then calculated within each experimental replicate. There was no effect of growth after exposure to 50 μM or 200 μM CdCl2, but we observed 22% growth inhibition at 1000 μM CdCl2 compared to control (P = 0.15, P= 0.2, P= 0.009, respectively; one-way ANOVA, Tukey HSD). We did not observe growth inhibition after 10 μM or 50 μM AfB1 compared to control, but did observe 11% growth inhibition at 200 μM AfB1, though trending (P= 0.27, P= 0.59, P= 0.09, respectively; one-way ANOVA, Tukey HSD). (C) There was no effect of either 50 μM CdCl2 or 10 μM AfB1 on total brood size compared to control (N = 9–10, P= 0.9, P= 0.63, respectively; one-way ANOVA, Tukey HSD). Error bars indicate standard error (* P< 0.05; ** P< 0.01).

Wild-type C. elegans exhibit resistance to mtDNA damage-induced single nucleotide mutations

The strong increase in mtDNA damage after CdCl2 and AfB1 suggested that mutations should be correspondingly increased. To test this hypothesis, we conducted Duplex Sequencing after 50 generations of mutation accumulation to determine the frequency and spectrum of mtDNA single nucleotide mutations on wild-type C. elegans in control and genotoxicant-treated conditions (Figure 2). We sequenced 11 wild-type MA lines and identified a total of 760 SNMs (an average of 70 mutations/MA line) under control conditions. The overall mtDNA mutation frequency of wild-type C. elegans was 10.1 × 10–7 SNMs/bp. Despite significant mtDNA lesions (Figure 1A), there was no effect of CdCl2 (14 MA lines, 874 total SNMs) or AfB1 (10 MA lines, 636 total SNMs) on the overall mtDNA SNM frequency in wild-type C. elegans (9.75 × 10–7 and 9.47 × 10–7, respectively) (Figure 3A, Supplementary Table S2). WGS was conducted in a random subset of four of the wild-type MA lines per condition in order to investigate the potential effects of CdCl2 and AfB1 on nuclear genome mutagenesis. We detected 189, 177 and 301 single nucleotide mutations in control, CdCl2 and AfB1-treated MA lines respectively after 50 total generations of mutation accumulation per treatment. We estimated a wild-type nuclear DNA mutation rate of 9.45 × 10–9 bp/site, which is very similar to previous estimates of nuclear DNA mutation rates in C. elegans mutation accumulation line experiments (62,63). We observed a 1.6-fold increase in the SNM rate in the AfB1 MA lines (15 × 10–9 bp/site, P = 0.016). There was no detectable effect of CdCl2 on overall nuclear DNA mutation rate (8.85 × 10–9 bp/site, P = 0.93) (Figure 3B).

Figure 3.

The mtDNA single nucleotide mutation signature in wild-type C. elegans is consistent with oxidative damage and demonstrates resistance to point mutations caused by CdCl2 and AfB1 mtDNA lesions. (A) Overall mtDNA SNM frequency was determined by Duplex Sequencing after 50 generations of mutation accumulation per subline in wild-type C. elegans in control conditions (gray dots; N = 11), as well as after 50 generations per subline of exposure to 50 μM CdCl2 (gold dots; N = 14) and 10 μM AfB1 (green dots, N = 10). There was no effect of either 50 μM CdCl2 or 10 μM AfB1 on overall mtDNA mutation frequency compared to control (P= 0.96, P =0.90, respectively); one-way ANOVA, Tukey HSD). Horizontal lines indicate mean values. (B) Overall nuclear SNM rates of a subset of MA lines in each treatment (N = 4 per treatment). There was no effect of 50 μM CdCl2 on overall SNM rate, but there was a 1.6-fold increase in overall SNM rate after exposure to 10 μM AfB1 (P = 0.01; ANOVA). Horizontal lines indicate mean rates. (C) mtDNA mutation spectrum of control, CdCl2 and AfB1-treated MA lines. Each dot represents a single MA line. The wild-type C. elegans mtDNA mutational signature was dominated by C:G → A:T and C:G → G:C transversion mutations, and there was no effect of Cd or AfB1 exposure on mtDNA mutational signature (Welch two-sample t-test). Error bars represent standard error of the mean. (D) mtDNA lesions may accumulate disproportionately on mtDNA strands, resulting in mtDNA mutation strand bias. We observed a trend towards an increase in C → T over G → A mutations after exposure to AfB1 compared to control (P = 0.08; two-way ANOVA, Tukey HSD). Open circles indicate one strand (i.e. A → C) and filled circles indicate the other strand (i.e. T → G).

The mtDNA mutation spectrum of C. elegans in all conditions was dominated by C:G → A:T and C:G → G:C transition mutations, and C:G → T:A mutations. There was no effect of CdCl2 or AfB1-induced mtDNA damage on mutation spectrum (Figure 3C; Welch two-sample t-test). Wild-type C. elegans exhibited no strand asymmetry in mtDNA mutation accumulation in control conditions (Figure 3D). There was no mutational strand asymmetry after exposure to CdCl2 (Figure 3D). We did observe a trend of a slightly higher frequency of C → T over G → A transversions after exposure to AfB1 compared to control conditions (P = 0.08; two-way ANOVA) (Figure 3D).

Mitophagy deficiency impairs mtDNA damage removal but does not increase mtDNA mutation accumulation

Given the surprising finding that exposure to well-known mutagens did not result in increased mtDNA mutations over control, despite the increase in mtDNA damage and increased nuclear mutation rates after AfB1 exposure, we hypothesized that mitophagy may be working to remove the damaged mitochondria before having a chance to form mutations. To test this hypothesis, we conducted a similar MA exposure experiment in mitophagy deficient strains. We chose dct-1 and pink-1 deficient strains because dct-1 and pink-1 (BNIP-3 and PINK1 human homologs) are involved in two independent mitophagy pathways (64). BNIP3 is a receptor-mediated pathway that, when activated, directly interacts with LC3 to facilitate autophagosome formation and mitochondrial degradation (65). This pathway is activated in response to hypoxia and nutrient depravation. PINK1 mediated mitophagy is activated by loss of mitochondrial membrane potential, resulting in the stabilization and accumulation of PINK1 on the outer mitochondrial membrane. PINK1 phosphorylates ubiquitin, followed by recruitment of Parkin E3 ubiquitin ligase which then recruits autophagy adapters to deliver mitochondrial to lysosomes for degradation (66). In our experiments, mitophagy-deficient dct-1 mutants did not accumulate significant mtDNA lesions compared to control after exposure to CdCl2 (0.21 lesions/10 kb), but did after exposure to AfB1 (1.33 lesions/10 kb, P < 0.01; two-way ANOVA, Tukey HSD) (Figure 4A). pink-1 mutants accumulated significantly higher levels of mtDNA damage after exposure to CdCl2 and AfB1 respectively compared to control (1.03 and 0.72 lesions/10 kb, P < 0.01; two-way ANOVA, Tukey HSD) (Figure 4A). Neither dct-1 or pink-1 mutants accumulated higher levels of mtDNA damage after exposure to CdCl2 compared to wild-type (two-way ANOVA). dct-1 mutants did accumulate significantly higher levels of mtDNA damage compared to wild-type after exposure to AfB1 (P < 0.05; two-way ANOVA, Tukey HSD) (Figure 4A). pink-1 mutants may accumulate higher levels of mtDNA damage compared to wild-type after AfB1 exposure; this trended towards, but did not reach, significance (P =0.08; two-way ANOVA, Tukey HSD) (Figure 4A).

Figure 4.

Mitophagy deficient mutants accumulate higher levels of mtDNA damage and exhibit greater fitness declines compared to wild-type, but exhibit no differences in mtDNA mutation frequencies after exposures. (A) mtDNA damage from pools of six individual age-synchronized L4 C. elegans after exposure to control (N = 8), 50 μM CdCl2 (N = 8) or 10 μM AfB1 (N = 8). pink-1 mutants accumulated mtDNA damage after exposure to CdCl2, but not more than wild-type, while dct-1 mutants did not accumulate mtDNA damage. Both mutants accumulated high levels of mtDNA damage after exposure to AfB1, and dct-1 accumulated significantly higher levels compared to wild-type exposed C. elegans. Error bars indicate standard error of the mean (* P < 0.05; ** P< 0.01; two-way ANOVA, Tukey HSD). (B) Population growth rate and total brood size as indicators of fitness in ancestors (G0) and after 50 generations of MA in all lines (G50). Each dot represents the mean value of days to starvation (until all of the food was eliminated and the population dispersed on the plate), with error bars as standard error of the mean (N = 3 for G0, N = 39- 48 for G50 MA lines). Lines indicate the rate of change in fitness from G0 to G50 (gray solid = control, gold small dash = 50 μM CdCl2, and green large dash = 10 μM AfB1) for wild-type, dct-1, and pink-1 strains. Y-axis lower limit begins at Day 6. There was no effect of 50 μM CdCl2 or 10 μM AfB1 on population growth rate in any strain at G0. All MA lines had significantly slower population growth rate at G50 compared to G0 (two-way ANOVA, P <2.2e–16). There was no effect of either CdCl2 or AfB1 on wild-type MA lines at G50 (P= 1, P= 0.95, respectively; two-way ANOVA, Tukey HSD). There was no effect of either CdCl2 or 10 μM AfB1 on dct-1 MA lines at G50 (P= 0.98, P= 0.6, respectively; two-way ANOVA, Tukey HSD). There was no effect of either CdCl2 or 10 μM AfB1 on pink-1 MA lines at G50 (P= 1, P= 0.4, respectively; two-way ANOVA, Tukey HSD). Both control and 10 μM AfB1pink-1 MA lines had a significantly greater decline in fitness compared to control and 10 μM AfB1 wild-type MA lines (P= 0.03, P = 0.0008, respectively; two-way ANOVA, Tukey HSD). At G50, three individual L4s per MA line were transferred onto an individual control plate for brood size experiments. Each individual was transferred every day until cessation of egg-laying. Reproduction was counted on the previous plate after 48 h. Each dot represents the mean value total brood size per MA line, with error bars indicating standard error of the mean (N = 9–10 for G0, N = 18–20 for G50 MA lines). Lines indicate the rate of change in fitness from G0 to G50 (gray solid = control, gold small dash = 50 μM CdCl2, and green large dash = 10 μM AfB1) for wild-type, dct-1, and pink-1 strains. The y-axis lower limit begins at 190. All MA lines had significantly lower fecundity at G50 compared to G0 (P =2.596e–11; two-way ANOVA). However, there was no effect of either CdCl2 or 10 μM AfB1 on total brood size in wild-type MA lines (P= 1, P =0.7, respectively; two-way ANOVA, Tukey HSD). pink-1 control MA lines had significantly smaller broods than dct-1 control MA lines at G50 (P< 0.01; two-way ANOVA, Tukey HSD) and pink-1 CdCl2 MA lines had significantly smaller broods than dct-1 CdCl2 MA lines at G50 (P= 0.03), suggesting a decrease in fitness in pink-1 MA lines compared to dct-1 MA lines. (C) Mutation spectrum of control (gray), CdCl2 (gold), and AfB1 (green) treated MA lines in wild-type, dct-1, and pink-1 strains. The mitophagy mutant C. elegans mtDNA mutational signatures were also dominated by C:G → A:T and C:G → G:C transversion mutations, and there was no effect of CdCl2 or AfB1 exposure on dct-1 or pink-1 mtDNA mutational signature (two-way ANOVA). Error bars indicate standard error of the mean.

Spontaneous mutagenesis due to mutation accumulation results in negative fitness consequences (67,68). Therefore, we assayed population growth rate and reproduction in all of the mutation accumulation lines after 50 generations of bottlenecks per strain per treatment per subline (Figure 4B). We quantified the time to starvation as the number of days that a population derived from a single L4 consumed all of the food source on the plate (54), as well as brood size (number of progeny per individual). There was no effect of 50 μM CdCl2 or 10μM AfB1 on population growth rate in any strain at G0 (Figure 4B). As expected, all MA lines had significantly slower population growth rate at G50 compared to G0 (two-way ANOVA, P < 2.2e-16). There was no effect of either CdCl2 or AfB1 on wild-type MA lines at G50 (P = 1, P = 0.95, respectively; two-way ANOVA, Tukey HSD), on dct-1 MA lines at G50 (P = 0.98, P = 0.6, respectively; two-way ANOVA, Tukey HSD) or on pink-1 MA lines at G50 (P = 1, P = 0.4, respectively; two-way ANOVA, Tukey HSD). However, both control and 10μM AfB1pink-1 MA lines had a significantly greater decline in fitness compared to control and 10μM AfB1 wild-type MA lines, with about a 10% decline in population growth rate (P = 0.03, P = 0.0008, respectively; two-way ANOVA, Tukey HSD). We quantified total brood size per individual at G50 in a subset of MA lines (N = 20 per strain per treatment). All MA lines had significantly lower fecundity at G50 compared to G0 (P =2.596e–11; two-way ANOVA). However, there was no effect of either CdCl2 or AfB1 on total brood size in wild-type MA lines (P = 1, P =0.7, respectively; two-way ANOVA, Tukey HSD). pink-1 control MA lines had significantly smaller broods than dct-1 control MA lines at G50 (P < 0.01; two-way ANOVA, Tukey HSD) and pink-1 CdCl2 MA lines had significantly smaller broods than dct-1 CdCl2 MA lines at G50 (P = 0.03). The lower fecundity and slower population growth rate suggests a greater decrease in fitness in pink-1 MA lines compared to wild-type and dct-1 MA lines (Figure 4B).

In order to determine if the decline in fitness could be due to accumulation of mtDNA damage-induced mutagenesis in the mitophagy-deficient strains, we sequenced dct-1 and pink-1 MA lines after 50 generations of mutation accumulation in control, CdCl2, and AfB1-treated conditions. A total of 491, 727 and 574 total SNMs were detected after sequencing 11 control, 11 CdCl2 and 9 AfB1dct-1 MA lines, respectively. A total of 530, 696 and 752 total SNMs were detected after sequencing 9 control, 11 CdCl2, and 10 AfB1pink-1 MA lines, respectively. There was no effect of mitophagy deficiency or exposure on overall mtDNA mutation frequency (Supplementary Table S2, Supplementary Figure S7; two-way ANOVA, P =0.3). As in wild-type, the mutational spectra of dct-1 and pink-1 MA lines were also dominated by C:G → A:T and C:G → G:C transitions, and C:G → T:A transversions (Figure 4C). There was no effect of CdCl2 or AfB1 on the mutational spectrum in either dct-1 or pink-1 mutants (two-way ANOVAs).

Trinucleotide context mutagenesis

Mammalian mtDNA has been reported to have a distinctive mutational signature (69). Therefore, we analyzed the identity of the 5’ and 3’ neighboring nucleotides at each SNM to investigate possible enrichment of SNMs in specific sequence contexts to reveal mechanisms of mtDNA mutagenesis in C. elegans. Overall, the C. elegans mitochondrial genome does have a distinct trinucleotide mutational signature (Supplementary Figure S8). Specifically, C:G → T:A mutations occur in a G[C→T]C context, while C:G → A:T mutations occur mainly at A[C/G]A, X[C/G]T, T[C/G]A and T[C/G]T sites. In order to determine the effect of chemical-induced mtDNA damage on trinucleotide context mutagenesis, we normalized the relative contribution of each mutation to the mean wild-type control contribution (Figure 5). In wild-type exposed to CdCl2, there were more C → G mutations in a C[C/G]A context compared to control (P < 0.05; Welch two-sample t-test). There was also a trend of more T → C mutations in a C[T/A]T context compared to control (P = 0.06; one-way ANOVA, Welch two-sample t-test). In AfB1 MA lines, we observed fewer C → G and C → T mutations in a C[C/G]C context, and more T → C mutations in a C[T/A]A context compared to control (P < 0.05; one-way ANOVA, Welch two-sample t-test). In dct-1 mutants, we observed more T → C mutations in CdCl2 MA lines in a C[T/A]A context compared to control (P < 0.05; one-way ANOVA, Welch two-sample t-test), and more C → T in a A[C/G]T context and T → C in A[T/A]T and G[T/A]A contexts after exposure to AfB1 compared to control (P < 0.05; one-way ANOVA, Welch two-sample t-test). In the pink-1 mutants, we observed lower relative contributions of mutations at specific trinucleotide contexts compared to wild-type control, with the exception of more C → A mutations in a A[C/G]C context in AfB1 compared to pink-1 control (P < 0.05; one-way ANOVA, Welch two-sample t-test).

Figure 5.

Trinucleotide context mutational signature of wild-type, dct-1 and pink-1 C. elegans after exposure to CdCl2 or AfB1. Contributions of each of the 96 possible trinucleotide mutations were determined for each MA line, and then normalized to the mean wild-type control contribution (show as a dashed line) to determine effects relative to wild-type control. Bar graphs show the relative mean and standard error of each trinucleotide context mutation, with each mutation represented by a different color and control, CdCl2 and AfB1 treatment as increasing color hues. Potential effects of each exposure compared to control were determined within each mutation type (Welch two-sample t-test) only after a significant ANOVA was determined. * P< 0.05.

We used the software MutationalPatterns to determine the cosine similarities of trinucleotide context mutational signatures between strain and treatment. As there were very few effects in the enrichment of trinucleotide mutations due to treatment as described above, it was unsurprising that the trinucleotide context mutational signatures were highly similar between strains and treatment (Figure 5 and Supplementary Figure S8). Indeed, the cosine similarity values across the three strains and three conditions were all close to 1, indicating high similarity (Supplementary Table S3). We next determined the similarity of these mutational signatures to known mutational processes as described in the Catalogue of Somatic Mutations in Cancer (COSMIC) database (http://cancer.sanger.ac.uk/cosmic/signatures) (Supplementary Table S4) (70). It is critical to note that the COSMIC signatures are derived from somatic nuclear mutagenesis from human cancer genomes, and C. elegans germline mitochondrial mutational processes are likely very different. Taking this into consideration, it was compelling that across all of our samples we detected high cosine similarity to the single base substitution (SBS) mutational signature that is associated with nucleotide excision repair deficiency (SBS24), which is absent in mitochondria. There was also high similarity with signatures associated with tobacco exposures (SBS4, SBS29). Tobacco products contains high levels of cadmium and smoke contains high levels of benzo[a]pyrene, which results in the formation of DNA adducts similar to those caused by AfB1. There also exists high similarity between signatures in which high levels of ROS is the proposed etiology (SBS18), in addition to defective base excision repair due to ROS-induced DNA damage (SBS36) and replication error across abasic sites (SBS13a). The similarities of C. elegans mtDNA signatures with these COSMIC signatures are likely due to the high C:G → A:T mutational bias in the C. elegans mitochondrial genome. Cosine similarity values are in Supplementary Table S4 and S5.

DISCUSSION

Mutation accumulation experiments in C. elegans and other organisms have determined that chemical exposures contribute to mutagenesis in the nuclear genome (71,72). However, the role of chemicals in mtDNA mutagenesis is not well studied (73). mtDNA is resistant to single nucleotide mutations in somatic tissues after exposure to the polycyclic aromatic hydrocarbon benzo[a]pyrene and hydrogen peroxide (9,15), and in germ cells after exposure to hydrogen peroxide (74). Mitophagy is responsible for degrading organelles with damaged mtDNA (23) and purifying selection against highly deleterious deletion mutations in the germline (31,75). Indeed, placing a Pol γ mutator strain in a mitophagy-deficient background resulted in increased mtDNA mutation frequency in C. elegans (76,77). However, to our knowledge, empirical evidence for or against a role of mitophagy in ameliorating chemical-induced mtDNA damage, or mutations caused by damage in the germline, had not been reported. We found that C. elegans mtDNA is resistant to germline mutagenesis from exposure to known nuclear mutagens Cd and AfB1, despite accumulating high levels of mtDNA damage. Unexpectedly, we also found that accumulation of spontaneous single nucleotide germline mutations appears to be independent of mitophagy.

Previous C. elegans mutation accumulation line experiments have been instrumental in our understanding of mtDNA mutation rates (8,34). However, various intricacies of mitochondrial genomics in C. elegans and other species complicate the study of mtDNA mutagenesis (78). The mitochondrial genome exists in multiple copies per organelle and cell, meaning that multiple genomes can pass through the germline bottleneck (∼60 copies in C. elegans) (34), potentially allowing selection to still act on the organelle and cellular level even in a neutral mutation accumulation design (79). Because mtDNA is polyploid, the frequency of each de novo single nucleotide mutation is often lower than the limit of detection of traditional next-generation sequencing (error rate of 1 in 103 bp). Furthermore, mutations that result from DNA damage are likely even more rare than spontaneous mutagenesis (80).

We sequenced 96 MA lines after about 50 generations of mutation accumulation each, resulting in a total of ∼4,800 generations of mutation accumulation. Highly sensitive Duplex Sequencing allowed us to detect a total of 6040 SNMs, which is about 250-fold higher than any previous C. elegans mitochondrial mutation accumulation line experiment to date. Our results were likely not biased by selection, as dN/dS ratios were ∼1 across all MA lines (Supplementary Table S6) (81,82). We observed a mutation spectrum similar to that reported in a recent Duplex Sequencing study of wild-type C. elegans, though roughly a 5-fold higher mutation frequency, which might be attributed to the higher number of generation bottlenecks in our experiment (50 generations of one individual versus three generations of passaging 10 individuals) or other differences in methodological and computational parameters (36). High frequencies of C:G → A:T and C:G → G:C transversion mutations support the conclusion that oxidative damage may drive mtDNA mutagenesis in C. elegans. The mechanism underlying this spectrum may relate to the apparent absence of some BER enzymes (83–85) or highly oxidizing intracellular environment under extreme drift (86). Mitochondrial redox state and oxidative mtDNA damage have not been measured in C. elegans, but could explain the high C:G → A:T mutation frequency in our MA experiment.

We observed a dominant G[C→T]C trinucleotide mutation across all strains and treatments (Supplemental Figure S6). This signature has been detected in various studies including aged human populations and has also been observed in some cancers (87). This suggests that either deoxycytidine deamination readily occurs in the C. elegans mitochondrial genome, or oxidized cytosines contribute to this mutational pattern. Cytosine deamination due to oxidative stress results in an excess of C → T over G → A mutations on the mtDNA heavy strand in many different organisms including Drosophila (9), mouse (88) and human mtDNA (89), and is likely the primary source of Pol γ error-induced mtDNA mutagenesis (11). However, our results did not reveal any strand asymmetry in mutation frequencies in wild-type C. elegans as would be predicted, particularly since oxidative damage is the prevailing model of mtDNA mutagenesis in C. elegans (36). Lack of strand-asymmetric mutagenesis could be because the mechanism of mtDNA replication in C. elegans may deviate from the single-strand displacement theory of other organisms (90), perhaps restricting exposure of damage-prone single-stranded mtDNA to oxidative stress and thus preventing strand bias. The potential compensatory mechanisms that have evolved to evade the consequences of high rates of oxidative damage-induced mutagenesis in C. elegans remains an intriguing area of study (91,92).

Mitochondria are a target of Cd (93), but the genotoxic and mutagenic effects of Cd on mtDNA are unknown. Cd is a ubiquitous heavy metal that is released from a range of sources, including mining, smelting, and combustion. The most significant sources of human exposure to Cd are contaminated food, smoking, e-waste sites, and products such as toys, jewelry, and plastics (94). Cd is a known carcinogen (95). It inhibits nuclear DNA repair mechanisms such as mismatch repair (40,96–98) and base excision repair (99), as well as zinc-dependent enzymes. Cd induces oxidative stress, likely via antioxidant depletion mechanisms (41). We found that exposure to 50 μM CdCl2 resulted in high levels of mtDNA damage across all genotypes, yet to our surprise, we observed no effect of genotoxicity on mtDNA SNM frequency or spectrum (Figures 1A, and 4A). There was almost no effect on the trinucleotide context mutational signature after exposure to CdCl2, except for an enrichment of a C → G transversion mutation in a C[N]A context compared to control in wild-type C. elegans. This may be indicative of oxidative damage, as 2,5-diamino-4H-imidazole-4-one (Iz) lesions contribute to C → G transversions in E coli (100). A recent study determined that human Pol γ is more prone to preferentially misincorporate a G opposite 2,2,4-triamino-5(2 H)-oxazolone, the common hydrolyzed product of Iz (101,102). We did not have the capability to identify the specific type of mtDNA lesions that are a result of Cd exposure, but the preponderance of this C → G transversion mutation could suggest that Cd causes a forms of oxidative damage, beyond 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) lesions which are likely repaired in the mitochondria, and thus contributes to C → A/G → T transversions. The mechanism by which oxidative damage and repair contributes to mtDNA mutational spectrum in C. elegans mtDNA mutagenesis is unclear, but C. elegans is able to repair at least some forms of oxidative mtDNA damage (85,103–106), despite the aforementioned absence of clear homologues of some BER enzymes.

Neither of the mitophagy mutant strains accumulated significantly higher levels of mtDNA damage after exposure to 50μM CdCl2 compared to exposed wild-type. Because CdCl2 by itself did not result in detectable mtDNA mutations, the lack of an increase in mutations in mitophagy mutants after CdCl2 cannot be interpreted to rule out mitophagy-mediated removal of CdCl2-induced mutations. dct-1 mutants do have an enrichment of one T → C transition mutation in a C[N]A context in Cd conditions compared to control, but the process underlying this mutation is unknown. It is possible that the quantity of lesions formed at this concentration of Cd was not enough to contribute to mtDNA mutagenesis. However, the internal body burden of Cd in C. elegans after exposure to 50μM CdCl2 (Supplementary Table S7) is equivalent to levels detected in human blood (95), suggesting that this concentration is relevant to human exposures.

As mentioned, C. elegans can repair at least some oxidative mtDNA damage. To investigate the effect of irreparable mtDNA damage on mutagenesis, we used AfB1. AfB1 is a mycotoxin produced by the fungus Aspergillus flavus and is one of the leading causes of hepatocellular carcinoma worldwide. A. flavus grows on staple grains and legumes, and is metabolized when consumed to an epoxide that forms stable AfB1-N7-G adducts that cause G → T mutations in the nuclear genome, primarily in CGC trinucleotide contexts (42). The C. elegans and mammalian nuclear mutational spectra are similar (62,72). However, to our knowledge, no study has investigated effects on mtDNA mutagenesis, even though it has been known for decades that AfB1 preferentially attacks mtDNA over nuclear DNA (45).

Mitophagy is required to remove AfB1-induced lesions, yet we measured only very small effects of 10μM AfB1 exposure on mtDNA mutational spectrum in wild-type and mitophagy-deficient C. elegans. We did observe a trend towards an increase in C:G → A:T transition mutations in AfB1-exposed pink-1 mutants compared to wild-type (1.4-fold greater mutation frequency, P = 0.07, two-way ANOVA, Tukey HSD). This suggests that pink-1 may play a role in ameliorating mtDNA damage or mediating purifying selection against deleterious mutations. Over 50 generations of mutation accumulation, pink-1 mutants had a greater fitness decline than wild-type and dct-1 mutants. This is consistent with increased accumulation of deleterious mutations, but we cannot rule out mutations in the nuclear genome or nongenetic effects (109).

The trinucleotide context mutational signature in AfB1 MA lines did not reveal the clear signature that has been observed in nuclear genome studies. There is a significant but small enrichment of C → A mutations in a A[C/G]C context in AfB1 MA lines compared to control in pink-1 mutants, which varies slightly from the known nuclear C[G]C context, suggesting a subtle but unique mechanism of aflatoxin damage-induced mutagenesis in the C. elegans mitochondrial genome. The unique AfB1 signature may result from the fact that damage removal processes are different for the two genomes; AfB1–induced damage in the nuclear genome is repaired by the enzyme NEIL1 as well as NER (107), but NER does not operate in the mitochondria and C. elegans does not appear to have a NEIL1 gene.

Mitophagy does regulate the transmission and accumulation of mtDNA mutations (31,108), yet our results suggest that this is perhaps limited to high-frequency, highly deleterious large deletion contexts. We found, for the first time, that constitutive mitophagy is responsible for removing mtDNA damage caused by AfB1 exposure (Figure 3A). A limitation of our work is that we did not measure mitophagy per se after exposure, though exposure to Cd and AfB1 induces mitophagy in other species (109–112). However, previous work in our lab has demonstrated that mitophagy is required to remove mtDNA damage caused by UVC exposure in C. elegans, but induction of mitophagy is not required (23,24,113). It is also possible that mtDNA damage exacerbates polymerase error-mediated accumulation of deletion mutations that we are unable to detect with Duplex Sequencing. Future studies should include technologies such as LostArc that can estimate frequencies of de novo deletions (114). Taken together, the current evidence suggests that de novo single nucleotide mutations present at low frequency do not have severe phenotypic consequences on organelle function and therefore evade targeted degradation via mitophagy.

Overall, our study suggests that in C. elegans, mitochondria harbor important quality control processes that are perhaps complementary or redundant to the two mitophagy pathways investigated in this study, resulting in a remarkable resistance to chemical-induced mutagenesis. A recent study in Drosophila found that unhealthy organelles failed to import nuclear-encoded factors essential for mtDNA replication and biogenesis. This resulted in the replication of wild-type mtDNA in healthy organelles, and not mutant mtDNA (115). Perhaps a ‘replication-competition’ model remains to be discovered in C. elegans. Direct degradation of damaged or mutant mtDNA may also play a protective role (116), though to our knowledge, C. elegans lacks a homolog for mitochondrial genome maintenance exonuclease (MGME-1) that degrades linear and damaged mtDNA at the mtDNA replisome. Some mechanisms that target paternal mitochondria and mtDNA for degradation in C. elegans are known, and an intriguing possibility is that similar pathways that target damaged or mutant mtDNA are present in the germline, which could suppress accumulation of deleterious mtDNA variants (117,118). On the organelle level, mitochondrial fusion and fission play a significant role in mediating mitochondrial quality control and are prerequisites for mitophagy (21,24). At 50μM CdCl2 and 10μM AfB1 exposure, we did not observe fragmentation of the C. elegans body wall muscle mitochondrial network in G0 wild-type C. elegans compared to control. We observed increased mitochondrial fusion at lower levels of Cd and AfB1exposure, with recovery at higher doses, which suggests that mitochondrial dynamics responded to Cd and AfB1-induced toxicity (Supplementary Figure S5). Future studies should investigate the role of mitochondrial dynamics in organelle and mtDNA turnover and biogenesis in the context of other chemical exposures. Lastly, another potential mechanism of ameliorating the removal of damaged mtDNA in the germline on the organismal level is germline apoptosis, which has previously been reported to increase after Cd and AfB1 exposure in C. elegans, though not at the concentrations that were used in our study (119,120).

The use of chemicals in society has resulted in over 86,000 chemicals of unknown toxicity in production (121), in addition to the hundreds of thousands of tons of pharmaceutical and industrial waste, increases in air pollution, and other sources of pollution that impact the health of hundreds of millions of people around the globe daily (122). Many chemicals can cause genome instability, which drives cancer development and other diseases; therefore it is necessary to better understand how low-level exposures to environmental agents can promote mutagenesis (123,124). mtDNA mutagenesis contributes to cancer and other diseases (69,125–128), yet is far less understood than nuclear DNA mutagenesis. Mitochondria are vital organelles that are highly susceptible to toxicity and genomic damage (17,129,130). MtDNA mutations accumulate with age, yet there remains conflicting evidence in the literature for the role of genotoxicant exposure in mtDNA mutagenesis (73). Surprisingly, we found that in C. elegans, after hundreds of generations of continuous chemical exposure, with hundreds of SNMs detected, and in the context of loss of two key mitophagy genes, mtDNA is resistant to damage-induced single nucleotide mutations. Future studies to investigate the effect of more chemicals in various genetic backgrounds may elucidate mechanisms in which mitochondria resist chemical-induced point mutations.

DATA AVAILABILITY

The Duplex-Seq-Pipeline is written in Python and R, but has dependencies written in other languages. The Duplex-Seq-Pipeline software has been tested to run on Linux, Windows WSL1, Windows WSL2 and Apple OSX. The software can be obtained at https://github.com/Kennedy-Lab-UW/Duplex-Seq-Pipeline under the BSD license.

The data are available as raw reads under BioProject PRJNA787252.

Click here for additional data file.