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Mutant C. elegans mitofusin leads to selective removal of mtDNA heteroplasmic deletions across generations to maintain fitness

BMC Biology volume 20, Article number: 40 (2022) Cite this article

Abstract

Background

Mitochondrial DNA (mtDNA) is present at high copy numbers in animal cells, and though characterized by a single haplotype in each individual due to maternal germline inheritance, deleterious mutations and intact mtDNA molecules frequently co-exist (heteroplasmy). A number of factors, such as replicative segregation, mitochondrial bottlenecks, and selection, may modulate the exitance of heteroplasmic mutations. Since such mutations may have pathological consequences, they likely survive and are inherited due to functional complementation via the intracellular mitochondrial network. Here, we hypothesized that compromised mitochondrial fusion would hamper such complementation, thereby affecting heteroplasmy inheritance.

Results

We assessed heteroplasmy levels in three Caenorhabditis elegans strains carrying different heteroplasmic mtDNA deletions (ΔmtDNA) in the background of mutant mitofusin (fzo-1). Animals displayed severe embryonic lethality and developmental delay. Strikingly, observed phenotypes were relieved during subsequent generations in association with complete loss of ΔmtDNA molecules. Moreover, deletion loss rates were negatively correlated with the size of mtDNA deletions, suggesting that mitochondrial fusion is essential and sensitive to the nature of the heteroplasmic mtDNA mutations. Introducing the ΔmtDNA into a fzo-1;pdr-1;+/ΔmtDNA (PARKIN ortholog) double mutant resulted in a skewed Mendelian progeny distribution, in contrast to the normal distribution in the fzo-1;+/ΔmtDNA mutant, and severely reduced brood size. Notably, the ΔmtDNA was lost across generations in association with improved phenotypes.

Conclusions

Taken together, our findings show that when mitochondrial fusion is compromised, deleterious heteroplasmic mutations cannot evade natural selection while inherited through generations. Moreover, our findings underline the importance of cross-talk between mitochondrial fusion and mitophagy in modulating the inheritance of mtDNA heteroplasmy.

Background

Unlike the nuclear genome, mitochondrial DNA (mtDNA) is present in multiple copies per animal cell. For instance, each human somatic cell contains an average of ~ 1000 mitochondria, with each mitochondrion harboring 1–10 mtDNA copies [1]. Although this intracellular mtDNA population is inherited from the maternal germline and hence carries a single major haplotype, mtDNA molecules can differ in sequence (heteroplasmy) either due to inheritance of mutations from the ovum or due to the accumulation of changes over an individual lifetime [2,3,4,5]. Some of these changes may have pathological consequences [6, 7], as reflected in a variety of mitochondrial disorders, yet only upon crossing a threshold of prevalence in the cell [8]. Accordingly, the penetrance of disease-causing mutations ranges between 60 and 80%, depending on the symptoms and tissues that display the specific phenotype [9].

The repertoire of heteroplasmic mutations varies among cells and tissues, mainly due to replicative segregation (drift) of the mitochondria during cell division and mitochondrial bottlenecks that appear during embryo development [10]. However, it has been suggested that heteroplasmy can be modulated by non-random factors, including selection [2, 4, 5, 11, 12]. Indeed, it has been shown that mitophagy, a mechanism of mitochondrial quality control, partially provides selection against defective mitochondria and maintains disease-causing mtDNAs below the threshold levels both in human cells [13] and in a Caenorhabditis elegans model system [14,15,16,17]. Mitophagy requires proper fission-fusion cycles of the mitochondrial network to allow the removal of dysfunctional mitochondria [18, 19]. In agreement with this notion, elevated heteroplasmy levels of pathological mtDNA molecules were observed when the fission machinery was disrupted in cell culture [20]. Furthermore, reduction in heteroplasmy levels of potentially deleterious mtDNA mutations was observed when components of the fusion machinery were compromised in Drosophila model systems, especially in germ cells [12, 21, 22]. In consistence with this notion, cell culture experiments revealed that a mixture of mtDNA molecules differing in sequence in the same cell can complement each other by the diffusion of products via the mitochondrial network, which in turn leads to restoration of mitochondrial function [1, 15, 16, 23]. Hence, mitochondrial fusion likely allows the survival of mtDNA disease-causing mutations in cells and, in turn, their transmission to the next generation [1, 8, 24]. Although these experiments suggest a molecular mechanism for the control of heteroplasmy, it remains unclear whether such a mechanism also affects the transmission of heteroplasmy through generations. Investigating this problem will allow explaining the relatively high abundance of low-level disease-causing heteroplasmic mutations in the general population [25, 26]. We, therefore, hypothesized that interfering with the intracellular mitochondrial network by compromising the fusion machinery would hamper mitochondrial functional complementation and thus impede the inheritance of heteroplasmic mutants.

Here, we took the first steps towards testing this hypothesis by crossing C. elegans harboring mitofusin mutant (fzo-1) to animals carrying either of three heteroplasmic mtDNA deletions, which differed in size and mtDNA positions. These experiments resulted in embryonic lethality and developmental delay, which were alleviated in subsequent generations concomitant with a complete loss of the truncated mtDNA molecules. Since the rate of truncated mtDNA loss diverged between the heteroplasmic strains, the sensitivity of the fusion machinery to different mtDNA mutations, in addition to its interaction with mitophagy and relevance to human diseases are discussed.

Results

A heteroplasmic deletion reduces the fitness of C. elegans mitofusin (fzo-1) mutant

The stable heteroplasmic C. elegans strain uaDf5/+ harbors a mixture of intact (+mtDNA) and ~ 60% of a 3.1 kb mtDNA deletion (ΔmtDNA) [27]. Although lacking four essential genes (i.e., mt-ND1, mt-ATP6, mt-ND2, and mt-Cytb) and seven tRNAs (i.e., K, L, S, R, I, Q and F), this strain is viable and displays some mitochondrial dysfunction [16, 27, 28]. High heteroplasmy levels are likely not maintained due to mtDNA duplication but by stably maintaining +mtDNA copy number [16]. We showed that dysfunctional PDR-1, the worm orthologue of the key mitophagy factor Parkin (PARK2), led to elevated levels of the truncated mtDNA, suggesting that mitochondrial quality control can modulate the levels of dysfunctional mitochondria [14]. In conjunction with this finding, RNAi knockdown of fzo-1, the C. elegans orthologue of MFN1/2, led to a slight reduction in the levels of the heteroplasmic ΔmtDNA, although without any phenotypic consequences [15]. We, therefore, asked what would be the impact of the fzo-1(tm1133) deletion (hereafter designated as fzo-1(mut)) on the inheritance of the ΔmtDNA.

To this end, we crossed uaDf5/+ heteroplasmic hermaphrodites (+/ΔmtDNA) with fzo-1(mut) heterozygote males (Fig. 1A). After self-cross of the F1 progeny, the distribution of the genotypes in the F2 heteroplasmic progeny did not deviate from the expected Mendelian ratio, namely 26% homozygous fzo-1(mut), 48.7% fzo-1 heterozygotes (ht), and 25.3% fzo-1 wild type (wt) (chi-square test, P = 0.960; Additional file 1: Table S1). However, we noticed that only 13 ± 5% of the progeny of the self-crossed fzo-1(mut);+/ΔmtDNA worms hatched, as compared to fzo-1(mut) animals (67 ± 5%, ANOVA followed by a Tukey’s post hoc test, P < 0.001; Fig. 1B). Although mitochondrial organization and TMRE uptake of self-crossed fzo-1(mut);+/ΔmtDNA adults were similar to fzo-1(mut) (Additional file 1: Fig. S1A-B), fzo-1(mut);+/ΔmtDNA animals were developmentally delayed, and none of them reached adulthood after six days as compared to fzo-1(mut) (ANOVA followed by a Tukey’s post hoc test, P < 0.001). This was in contrast to self-crossed fzo-1(wt);+/ΔmtDNA animals, all of which reached adulthood after six days (Fig. 1C). These findings demonstrate that the interaction between the heteroplasmic ΔmtDNA and the nuclear DNA-encoded fzo-1 mutant led to a severe reduction in fitness.

Fig. 1
figure 1

ΔmtDNA reduces the fitness of fzo-1(mut) animals. A Establishing fzo-1 mutant and wild type heteroplasmic lines. Heteroplasmic hermaphrodites carrying intact and truncated mtDNA [uaDf5/+] were crossed with fzo-1(tm1133) heterozygotes males, fzo-1(ht). Cross progeny F1 were allowed to self-propagate. Single F2 animals were isolated, allowed to lay eggs, and their genotypes were determined using a single worm PCR. Heteroplasmic (+/ΔmtDNA) mutant (mut) or wild type (wt) fzo-1 progeny were then monitored. B The percent of hatched embryos of parental strains: N2 (WT; N = 13, n = 869), +/ΔmtDNA (N = 7, n = 265) and fzo-1(mut) (N = 13, n = 386), of fzo-1(ht);+/ΔmtDNA animals (N = 5, n = 152) and of mutant, fzo-1(mut);+/ΔmtDNA (N = 7, n = 236) or wild type fzo-1(wt);+/ΔmtDNA (N = 4, n = 104) cross progeny. Data are means ± 1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test. (*) denotes P < 0.05 and (**) denotes P < 0.001 compared with WT animals. C The percent of gravid adults six days after egg laying of parental strains: N2 (WT) (N = 5, n = 114), +/ΔmtDNA (N = 9, n = 232) and fzo-1(mut) (N = 4, n = 278), of fzo-1(ht);+/ΔmtDNA animals (N = 5, n = 69), and of mutant, fzo-1(mut);+/ΔmtDNA (N = 4, n = 152) or wild type fzo-1(wt);+/ΔmtDNA (N = 5, n = 245) cross progeny. Data are means ±1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test. (**) denotes P < 0.001 compared with WT animals. Individual data values are presented in Additional file 2

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The adverse effects of the interaction between ΔmtDNA and fzo-1(mut) are reversed across generations

To better characterize the phenotypic impact of the interactions between ΔmtDNA and fzo-1(mut), we monitored the development of progeny of the self-crossed fzo-1(ht);+/ΔmtDNA worms, followed by genotyping the resultant adult animals (generation 1, G1). We continued to follow the hatching and development of their progeny, i.e., fzo-1(mut);+/ΔmtDNA (G2m) and fzo-1(wt);+/ΔmtDNA (G2wt), across four generations (Fig. 2A). Specifically, we measured the duration of the larva-to-adulthood period during development in the G1m-G4m generations (Fig. 2B). While ~ 75% of the G1m animals reached adulthood after 6 days, the development of G2m animals was 1.9-fold delayed (Cox proportional-hazards regression, P < 0.001), with 75% of the animals reaching adulthood only after 9 days. Surprisingly, the G3m animals showed significant improvement (Cox proportional-hazards regression, P < 0.001), with ~ 60% of this population reaching adulthood after six days. Moreover, G4m animals showed a full reversal of ΔmtDNA-associated adverse effects (Cox proportional-hazards regression, P = 0.750; Fig. 2B and Additional file 1: Table S2). We noted a similar pattern across generations when hatching was considered: In contrast to the 13.5% hatching observed among G2m embryos, 60 ± 8% hatching of the G3m embryos was observed (ANOVA followed by a Tukey’s post hoc test, P < 0.001). The hatching percentage of the G4m generation was similar to that of G1 animals (71 ± 9% and 67 ± 5%, respectively, ANOVA followed by a Tukey’s post hoc test P = 0.993) and remained stable over subsequent generations (Fig. 2C). Finally, no phenotypic changes were observed for G1wt-G4wt animals while tracing their developmental pace (Cox proportional-hazards regression, P > 0.140; Additional file 1: Table S2) and hatching percentage (ANOVA followed by a Tukey’s post hoc test, P > 0.266; Additional file 1: Fig. S2A-B). Taken together, our findings demonstrate a full reversal of the adverse effects of the interaction between the ΔmtDNA and the nuclear DNA-encoded mutant fzo-1 gene.

Fig. 2
figure 2

ΔmtDNA levels are selectively eliminated, and their adverse effects reversed in fzo-1(mut);+/ΔmtDNA animals across generations. A Schematic representation of the experimental setup. The fzo-1 heterozygotes progeny of heteroplasmic hermaphrodites (fzo-1(ht);+/ΔmtDNA) was identified and maintained using self-propagation and single worm genotyping to establish heteroplasmic lines carrying fzo-1(ht);+/ΔmtDNA. Progeny animals (generation 1; G1) were isolated, allowed to lay eggs, and their genotypes were determined. Heteroplasmic mutant fzo-1(mut);+/ΔmtDNA or wild type fzo-1(wt);+/ΔmtDNA progeny were then monitored over several generations (G2m-G4m and G2wt-G4wt, respectively). B The percent of gravid adults of fzo-1(mut);+/ΔmtDNA mutant progeny across generations (G1m-G4m) at the indicated times after egg laying (G1m N = 3, n = 40, G2m N = 4, n = 152, G3m N = 3, n = 155 and G4m N = 3, n = 82). Data were analyzed using Cox proportional-hazards regression (Additional file 1: Table S2). G2m and G3m were slower to reach adulthood than G1m (P < 0.001) but not G4m (P = 0.750). C The percent of hatched embryos of fzo-1(mut);+/ΔmtDNA progeny across generations (G2m N = 7, n = 236, G3m N = 6, n = 155 and G4m N = 4, n = 107) and the stable line (> 20 generations) fzo-1(mut);+/ΔmtDNA (N = 4, n = 93). Data are means ±1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test, (**) denotes P < 0.002 compared with G2m animals. D, E Box plot showing the percent of ΔmtDNA (N > 3 biological repeats) determined in individual animals (D) of the parental heteroplasmic strain +/ΔmtDNA (n = 23), the heteroplasmic fzo-1(mut) mutant cross-progeny strains (G1ht n = 13, G1m-G4m n = 20, 31, 21 and 21, respectively) and the progeny of G4m animals crossed with fzo-1(wt), (Gm→Gwt n = 21); (E) of the heteroplasmic fzo-1(wt) cross progeny strains (G1wt-G4wt n = 17, 20, 27 and 21, respectively). In the boxplot representation, center line, median; box limits, upper and lower quartiles; whiskers, minimum and maximum; points, data. Data were analyzed using Fractional regression (Additional file 1: Table S3). ΔmtDNA levels of G2m-G4m and Gm->Gwt were significantly lower than those of the parental heteroplasmic strain +/ΔmtDNA, (**) denotes P < 0.001. F The percent of ΔmtDNA determined for a population of animals from the stable cross lines (> 20 generations), fzo-1(wt);+/ΔmtDNA (N = 7), fzo-1(mut);+/ΔmtDNA (N = 3) and +/ΔmtDNA;fzo-1(Gm→Gwt) (N = 4). Data are means ±1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test, (**) denotes P < 0.001 compared with fzo-1(wt);+/ΔmtDNA animals. Individual data values are presented in Additional file 2

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fzo-1(mut) leads to selection against ΔmtDNA heteroplasmy in C. elegans

We next asked how the deleterious interactions between fzo-1(mut) and ΔmtDNA were abrogated. We hypothesized that if the ΔmtDNA is not tolerated in the background of fzo-1(mut), then selection against the ΔmtDNA should occur. To test this prediction, we assessed the levels of ΔmtDNA by quantitative PCR (qPCR, see Methods) across the G1m-G4m generations (using gravid adults) in both the fzo-1(mut);+/ΔmtDNA and the fzo-1(wt);+/ΔmtDNA strains. We found that ΔmtDNA levels declined by 10-fold in the G2m fzo-1(mut) animals, as compared to +/ΔmtDNA parental strain (Fractional regression, odds ratio = 0.08; P < 0.001). Values of ΔmtDNA reached below detection levels in most G3m (N = 18/21) and G4m (N = 20/21) animals (fractional regression, odds ratio < 0.007, P < 0.001; Fig. 2D and Additional file 1: Table S3), while the relative levels of intact +mtDNA molecules increased (Additional file 1: Fig. S2C and Table S3). In contrast, ΔmtDNA and +mtDNA levels did not significantly change across the G1wt-G4wt generations of fzo-1(wt);+/ΔmtDNA animals (fractional regression, P = 0.852; Fig. 2E and Additional file 1: S2D and Table S3).

These results suggest that the ΔmtDNA was completely lost during the G1m-G4m generations. To test this hypothesis, we crossed G4m hermaphrodites with wild type males to isolate fzo-1(wt) progeny (Gm→Gwt). Since traces of ΔmtDNA were neither detected in Gm->Gwt animals (fractional regression, P < 0.001; Fig. 2D) nor in subsequent generations (ANOVA followed by a Tukey’s post hoc test, P < 0.001; Fig. 2F), we concluded that disrupting fzo-1 function indeed resulted in a complete and specific loss of the deleterious heteroplasmic ΔmtDNA. These results provide a proof of concept that mitochondrial fusion is critical for regulating the transmission of the uaDf5 ΔmtDNA heteroplasmy across generations.

Selection against heteroplasmic truncations depends on deletion size or mtDNA position

We next asked whether the deleterious interactions between fzo-1(mut) and ΔmtDNA depend on the size or genomic location of mtDNA deletions. To achieve this goal, we characterized two additional mtDNA deletions obtained from the Million Mutation Project strain collection [29]. Specifically, these deletions comprise two new stable heteroplasmic C. elegans strains: bguDf1 (derived from strain VC40128), harboring a mixture of intact +mtDNA along with mtDNA molecules lacking ~ 1 kb (1kbΔmtDNA) encompassing two essential mtDNA genes (i.e., mt-ATP6 and mt-ND2) and three tRNAs (i.e., K, L, and S); the second strain, bguDf2 (derived from VC20469), harbors in addition to the +mtDNA, a ~4.2 kb mtDNA deletion (4kbΔmtDNA) encompassing four different essential genes (i.e., mt-CO1, mt-CO2, mt-ND3, and mt-ND5) and five tRNAs (i.e., C, M, D, G, and H). Notably, the levels of the 1kbΔmtDNA and 4kbΔmtDNA were stable over > 100 generations (80% and 55%, respectively) in the presence of functional (wild type) fzo-1. Reanalysis of whole-genome sequencing data for the two mutant strains identified +mtDNA and deletion sequences, as previously described [29, 30]. These analyses did not reveal any evidence for duplicated regions, confirming the mtDNA deletion heteroplasmy in these strains (Additional file 1: Fig. S3A-B). As previously observed for uaDf5/+ [16], truncated mtDNA levels highly varied between animals, while intact +mtDNA levels were more constant (Additional file 1: Fig. S3C-D). The animals displayed neither embryonic nor developmental phenotypes, and no impact on mitochondria fusion was observed (Additional file 1: Fig. S3E-G and Table S2).

Heteroplasmic hermaphrodites of both strains were separately crossed with fzo-1(mut) heterozygote males, followed by self-cross of F1 progeny (cross as in Fig. 1A). Like the approach taken with the uaDf5/+ strain, we examined the distribution of the genotypes in the F2 heteroplasmic progeny. We found that the genotypes distribution for the fzo-1(mut);+/1kbΔmtDNA did not deviate from the expected Mendelian ratio (23.5% homozygous fzo-1(mut), 43.2% fzo-1(ht) and 33.3% fzo-1(wt) (P = 0.14, chi-square test; Additional file 1: Table S1). In contrast, this ratio strongly deviated from the expected Mendelian ratio for fzo-1(mut) animals harboring +/4kbΔmtDNA (6.4% homozygous fzo-1(mut), 59.3% fzo-1(ht) and 34.3% fzo-1(wt) (P < 0.001, chi-square test; Additional file 1: Table S1). Hence, these results indicate that fzo-1(mut) differentially tolerates mtDNA deletions based on size and/or mtDNA position.

We next quantified the levels of ΔmtDNA in mutant versus wild type fzo-1 progeny across four subsequent generations (as in Fig. 2A; Fig. 3 and Additional file 1: Table S3). We found that both truncated mtDNA molecules were undetectable after four generations (Fig. 3A, B), yet the decline rates significantly diverged (Fig. 3C). Specifically, mean ΔmtDNA levels were significantly lower in fzo-1(mut) animals harboring the 4kbΔmtDNA than in animals harboring either 1kbΔmtDNA or 3kbΔmtDNA in both G1m and G2m animals (fractional regression followed by within generation pairwise comparisons, P < 0.001; Fig. 3C and Additional file 1: Table S3). By the G3m generation, mean ΔmtDNA levels of 3kbΔmtDNA were also significantly lower than those observed in animals harboring 1kbΔmtDNA. Indeed, the 1kbΔmtDNA was still detected in most of G3m animals (N = 13/18). It is worth noting that the levels of both types of ΔmtDNA did not significantly change across the G1wt-G4wt generations of the fzo-1(wt);+/ΔmtDNA animals (fractional regression, P > 0.118 in both cases; Additional file 1: Fig. S3H-I and Table S3).

Fig. 3
figure 3

ΔmtDNA levels are differentially eliminated in fzo-1(mut);ΔmtDNA animals based on size or mtDNA position. A, B Box plot showing the percentage of 1kbΔmtDNA (A) or 4kbΔmtDNA (B) ΔmtDNA (N > 3 biological repeats), determined in individual animals of the parental heteroplasmic strain +/1kbΔmtDNA (n = 14) and +/4kbΔmtDNA (n = 15), the fzo-1(mut) mutant cross-progeny strains (1kbΔmtDNA F1(ht) n = 18, G1m-G4m n = 15, 20, 17 and 19 and 4kbΔmtDNA F1(ht) n = 18, G1m-G4m n = 14, 14, 21 and 20) and the progeny of G4m animals crossed with fzo-1(wt), (Gm → Gwt n = 15 and n = 20 respectively). In the boxplot representation, center line, median; box limits, upper and lower quartiles; whiskers, minimum and maximum; points, data. Data were analyzed using fractional regression (Additional file 1: Table S3). ΔmtDNA levels of G1m-G4m and Gm->Gwt were significantly lower than those of the parental strains in both +/1kbΔmtDNA and +/4kbΔmtDNA, (**) denotes P < 0.001. C Box plot comparing the percentage of ΔmtDNA in animals carrying +/3kbΔmtDNA (blue), +/1kbΔmtDNA (yellow) or +/4kbΔmtDNA (red) in each generation (G1m-G4m; data from Fig. 2C, Fig. 3A, and Fig. 3B, respectively). In the boxplot representation, center line, median; box limits, upper and lower quartiles; whiskers, minimum and maximum; points, data. Data were analyzed using fractional regression followed by within generation pairwise comparisons (Additional file 1: Table S3), different letters indicate a significant difference in mean levels of ΔmtDNA levels: G1-G2, 4 kb levels (a) lower than 3 kb and 1 kb (b, P < 0.001); G3, 4 kb levels (a) lower than 3 kb (b, P < 0.01) and 4 kb (a) and 3 kb (b) levels lower than 1 kb (c, P < 0.001); G4, 4 kb and 3 kb levels (a) lower than 1 kb (b, P < 0.001). Individual data values are presented in Additional file 2

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To assess whether the truncated mtDNAs were completely lost, we crossed G4m hermaphrodites with wild type males and isolated fzo-1(wt) progeny (Gm→wt). qPCR analyses revealed no traces of the ΔmtDNA copies in subsequent generations (fractional regression, P < 0.001; Fig. 3A-B and Additional file 1: Table S3). Thus, disrupting fzo-1 function resulted in a complete and specific loss of a variety of heteroplasmic ΔmtDNAs. We interpret these results to mean that fzo-1 function is sensitive to either the size or location of deleterious mtDNA heteroplasmy.

Selection against ΔmtDNA molecules occurs during C. elegans development

In C. elegans, mtDNA copy numbers increase significantly during the fourth larval stage (L4) in association with oocyte production [27, 31]. We, therefore, asked at which point during the C. elegans life cycle selection against ΔmtDNA occurred. Given that the relative levels of ΔmtDNA are maintained during normal development [27], we compared ΔmtDNA levels between embryos and adults in G2m animals. Our results indicate that ΔmtDNA levels were dramatically reduced (~ 5-fold) during the development of G2m animals (ANOVA followed by a Tukey’s post hoc test, P < 0.001; Fig. 4A) but not in G2wt animals (Fig. 4B). This observation suggests that ΔmtDNA is most likely selected against during the fzo-1(mut) worm development, in agreement with the observed adverse effect of heteroplasmy on the hatching and development of G2m animals.

Fig. 4
figure 4

ΔmtDNA molecules are selectively eliminated during C. elegans development and defer between gonad and soma. A The percent of ΔmtDNA determined for a population of animals in generation 2 mutant, G2m, eggs (N = 5), unhatched eggs (N = 5), larvae (N = 3) and adults (N = 4) and generation 3 mutant, G3m eggs (N = 4). Data are means ±1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test, (*) denotes P < 0.05 and (**) denotes P < 0.001 compared with G2m unhatched eggs. B The percent of ΔmtDNA determined for a population of animals (n = 5) in generation 2 wild type, G2wt eggs (N = 3) and adults (N = 3), and generation 3 wild type, G3wt eggs (N = 4). Data are means ± 1 standard error of the mean (1SE). Data were analyzed using one-way ANOVA followed by a Tukey’s post hoc test compared with G2wt eggs. C The percent of ΔmtDNA determined in individual G2m adults that reached adulthood (N = 3 biological repeats) after 7–8 days (mild delay; n = 12) or 9–10 days (severe delay; n = 22). Data were analyzed using the Wilcoxon Mann-Whitney rank sum test (P < 0.005). D–G The relative levels of ΔmtDNA (D, F) or +mtDNA (E, G) were determined for the gonad and soma (normalized to soma) of a population of (D, E) G2m adults (N = 8) or (F, G) G2wt adults (N = 4). Data were analyzed using the Wilcoxon Mann-Whitney rank sum test, (*) denotes P < 0.05. Individual data values are presented in Additional file 2

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To examine the possible association of ΔmtDNA levels with embryo lethality, we compared ΔmtDNA levels of unhatched embryos (unhatched > 48 h after being laid) to newly hatched larvae (L1). As expected, given that ~ 85% of G2 embryos did not hatch, ΔmtDNA levels of unhatched embryos were similar to the relative ΔmtDNA levels of newly laid embryos. In contrast, ΔmtDNA levels in L1 animals were reduced by 2-fold (ANOVA followed by a Tukey’s post hoc test, P < 0.05; Fig. 4A). This suggests that hatching is enabled only in embryos with reduced ΔmtDNA levels.

We next examined whether ΔmtDNA levels associate with developmental delay. To this end, we compared the levels of ΔmtDNA in mildly delayed G2 animals that reached adulthood on days 7–8 to severely delayed animals that reached adulthood on days 9–10. Our results show that ΔmtDNA levels were 2-fold higher in the severely delayed group (Wilcoxon Mann-Whitney rank sum test, P < 0.005 test; Fig. 4C). These data demonstrate increased embryo lethality and aggravation in developmental delay in animals harboring high levels of ΔmtDNA. This supports our interpretation that disruption of mitochondrial fusion in animals carrying ΔmtDNA molecules leads to reduced fitness and suggests that there is selection against such molecules at the level of the organism.

Selection against ΔmtDNA molecules defers between gonad and soma

Previously Lieber et al. demonstrated germline selection acting against high levels of mutant mtDNA in Drosophila oogenesis [22]. In C. elegans, the germline tends to accumulate higher levels of deleterious mitochondrial molecules than somatic tissues, although unfertilized oocytes contain lower levels of ΔmtDNA compared to that of germline tissue [32]. Consistently, we found that ΔmtDNA molecules became undetectable in the resultant embryos of G2m animals (i.e., in G3m animals; Fig. 4A). Hence, it is possible that selection against ΔmtDNA molecules occurred during C. elegans gametogenesis. In support of this claim, a comparison of ΔmtDNA levels between gonads and somatic tissues in G2m animals revealed a two-fold decrease of ΔmtDNA levels in the gonads (Wilcoxon Mann-Whitney rank sum test, P < 0.05; Fig. 4D), whereas the levels of +mtDNA intact molecules were ~1.4-fold higher in the gonad (Fig. 4E). In contrast, both ΔmtDNA and +mtDNA molecules were ~1.5-fold higher in the gonad when comparing gonads and somatic tissues of fzo-1(wt);+/ΔmtDNA animals (P < 0.05 Wilcoxon Mann-Whitney rank sum test; Fig. 4F, G) [32]. Since both the Drosophila experiments [12, 21, 22] and our observations are consistent, we argue that selection against ΔmtDNA molecules during gametogenesis is evolutionarily conserved [12].

PARKIN mutant aggravates the adverse effects of the ΔmtDNA-fzo-1 interactions

Since Parkin mediates the turnover of mitofusins and hence impacts their activity [33, 34], we asked what would be the impact of the pdr-1;fzo1 double mutant on the inheritance of the ΔmtDNA. To address this question, we first crossed +/ΔmtDNA heteroplasmic hermaphrodites with pdr-1(gk448) (here named pdr-1(mut)) males and established a stable pdr-1(mut);+/ΔmtDNA strain. Consistent with previous findings, the ΔmtDNA levels were elevated in this strain (75%) [14,15,16,17]. To establish a strain which is mutant in pdr-1 and fzo-1 in the context of +/ΔmtDNA heteroplasmy, we then crossed pdr-1(mut);+/ΔmtDNA heteroplasmic hermaphrodites with pdr-1;fzo-1 heterozygous males, let the F1 progeny self-cross, and isolated fzo-1(ht);pdr-1(mut) hermaphrodites that are harboring +/ΔmtDNA (Fig. 5A). This strain was allowed to propagate, and the genotypic distribution of the heteroplasmic progeny was assessed. While the genotype distribution of fzo-1(mut);+/ΔmtDNA did not deviate from the expected Mendelian ratio, the genotype distribution of fzo-1(ht);pdr-1(mut);+/ΔmtDNA was strongly affected, as follows: 9% homozygous fzo-1(mut);pdr-1(mut), 55.8% fzo-1(ht);pdr-1(mut), and 35.2% fzo-1(wt);pdr-1(mut) (P < 0.001, chi-square test; Additional file 1: Table S1). This suggests that the heteroplasmic ΔmtDNA cannot be tolerated in the background of fzo-1(mut);pdr-1(mut) double mutant, as reflected in further reduction in fitness.

Fig. 5
figure 5

Mutant Parkin aggravates the adverse effects of the ΔmtDNA;fzo-1. A Schematic representation of experimental setup. The fzo-1 heterozygotes progeny of heteroplasmic pdr-1(mut) hermaphrodites (fzo-1(ht);pdr-1(mut);+/ΔmtDNA) was identified and maintained using self-propagation and single worm genotyping to establish heteroplasmic lines carrying fzo-1(ht);pdr-1(mut);+/ΔmtDNA. Progeny animals (generation 1; G1) were isolated, allowed to lay eggs and their genotypes were determined. The progeny of heteroplasmic pdr-1(mut) hermaphrodites that were fzo-1 mutant (fzo-1(mut);pdr-1(mut);+/ΔmtDNA) or wild type (fzo-1(wt);pdr-1(mut);+/ΔmtDNA) were then monitored over several generations (G2m-G4m and G2wt-G4wt, respectively). B The percent of gravid adults of fzo-1(mut);pdr-1(mut);+/ΔmtDNA mutant progeny across generations (G2m-G4m) and of the stable cross line (> 30 generations, Gm > 30) was determined at the indicated times after egg laying (G2m N = 6, n = 140, G3m N = 5, n = 83, G4m N = 6 n = 89, and Gm > 30 N = 3, n = 116). Data were analyzed using Cox proportional-hazards regression (Additional file 1: Table S2). G2m-G3m were slower to reach adulthood than Gm > 30 (P < 0.001) but not G4m (P = 0.084). C Box plot showing the percent of ΔmtDNA (N > 3 biological repeats) determined in individual animals of the heteroplasmic mutant cross-progeny strains, fzo-1(mut);pdr-1(mut);+/ΔmtDNA (G1(ht) n = 41, G1m-G4m n = 14, 20, 17 and 16, respectively). In the boxplot representation, center line, median; box limits, upper and lower quartiles; whiskers, minimum and maximum; points, data. Data were analyzed using Fractional regression (Additional file 1: Table S3). ΔmtDNA levels of G1m-G4m were significantly lower than those observed in G1(ht), (**) denotes P < 0.001. D Heteroplasmy levels of individual fzo-1(ht);pdr-1(mut);+/ΔmtDNA animals (N > 3 biological repeats) sampled at generations 3–7 (Ght3-Ght7; n = 22) and generations 20–24 (Ght20-Ght24; n = 14). Data were analyzed using the Wilcoxon Mann-Whitney rank sum test (P < 0.001). E–H The relative levels of ΔmtDNA (E, G) or +mtDNA (F, H) determined for the gonad and soma (normalized to soma) of a population of E–F G2m adults (N = 4) or G–H G2wt adults (N = 5). Data were analyzed using the Wilcoxon Mann-Whitney rank sum test, (*) denotes P < 0.05 and (**) denotes P < 0.001. Individual data values are presented in Additional file 2

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We next monitored the development and fecundity of these animals. Phenotypic characterization revealed a developmental delay in the fzo-1(mut);pdr-1(mut);+/ΔmtDNA G1m, similar to the parental strain (11/11 were adults by day 7); and most of the G1m progeny (G2m eggs) hatched (85 ± 8%; Additional file 1: Fig. S4A). However, 66 ± 1% of the G2m were developmentally arrested (Additional file 1: Fig. S4B). Only 33 ± 12% of the remaining animals reached adulthood by day 7 (Cox proportional-hazards regression, P < 0.001; Fig. 5B and Additional file 1: Fig. S4C and Table S2), and their progeny production was severely reduced (laying seven eggs or less over 20 h). Moreover, 22 ± 7.5% of the G3m animals were still developmentally arrested (Additional file 1: Fig. S4B), and G3m development was similarly delayed (Cox proportional-hazards regression, P < 0.001; Fig. 5B and Additional file 1: Fig. S4C and Table S2). However, we noticed a significant recovery of animals’ development during subsequent generations (Fig. 5B and S4C). In contrast, heteroplasmic fzo-1(wt);pdr-1(mut);+/ΔmtDNA hatching and development was unaffected (~ 98% hatched and 100% were adults by day 7; Additional file 1: Fig. S4A and S4C). Thus, the adverse effects of the interaction between fzo-1(mut) and ΔmtDNA on fecundity and developmental timing were aggravated by pdr-1(mut), supporting fzo-1-pdr-1 epistasis.

We next asked whether the selection against ΔmtDNA would strengthen in the background of fzo-1(mut);pdr-1(mut);+/ΔmtDNA. To directly examine this, we compared the levels of ΔmtDNA molecules in fzo-1(mut);pdr-1(mut);+/ΔmtDNA animals (Fig. 5C) to fzo-1(mut);+/ΔmtDNA (Fig. 2D) across G1m-G4m generations. We found that ΔmtDNA levels declined more sharply in fzo-1(mut);pdr-1(mut);+/ΔmtDNA as compared to the single mutant strain fzo-1(mut);+/ΔmtDNA. Specifically, we found lower ΔmtDNA levels in G1m and G2m fzo-1(mut);pdr-1(mut);+/ΔmtDNA animals (fractional regression followed by within generation pairwise comparisons, P < 0.001; Additional file 1: Table S3). In contrast, ΔmtDNA levels did not significantly change across generations of fzo-1(wt);pdr-1(mut);+/ΔmtDNA animals (fractional regression, P > 0.167 in all cases; Additional file 1: Fig. S4D and Table S3). Taken together, the concomitant disruption of fusion and mitophagy strongly selected against ΔmtDNA molecules. In support of this interpretation, we noticed that even in the fzo-1(ht);pdr-1(mut);+/ΔmtDNA animals, where only one genomic copy of fzo-1 remained functional, ΔmtDNA levels declined and in some individuals were lost across ~ 25 generations (Wilcoxon Mann-Whitney rank sum test, P < 0.001; Fig. 5D).

Finally, we asked whether impaired mitophagy affects selection against ΔmtDNA in the worm germline. To this end, we examined the ratio of ΔmtDNA and +mtDNA between the gonad and soma in double mutant fzo-1(mut);pdr-1(mut);+/ΔmtDNA animals (Fig. 5E, F) and pdr-1(mut);+/ΔmtDNA (Fig. 5G, H). While mtDNA levels, including both ΔmtDNA and +mtDNA, were ~1.5-fold higher in the gonad vs. the soma of pdr-1(mut) animals (Wilcoxon Mann-Whitney rank sum test, P < 0.05; Fig. 5G, H), similar to wild type animals (Fig. 4F, G). We observed that fzo-1(mut);pdr-1(mut);+/ΔmtDNA animals displayed similar mtDNA levels in the gonad compared to soma, for both truncated and intact mtDNA molecules (Fig. 5E, F). On top of the selection against ΔmtDNA, fzo-1(mut);pdr-1(mut);+/ΔmtDNA double mutant displayed a reduction in total mtDNA levels, again supporting epistasis. Taken together, these data suggest that disrupting parkin-mediated mitophagy increased the organismal selection in fzo-1(mut);pdr-1(mut);+/ΔmtDNA individuals, which is associated with reduced fitness.

Discussion

Our cross-generational analyses revealed complete loss of heteroplasmic deleterious mtDNA deletions when mitochondrial fusion is compromised. This demonstrated that heteroplasmy of deleterious mtDNA molecules could not be tolerated unless in the presence of a functional compensatory mechanism inherent to the mitochondrial network and the mitochondrial quality control machinery [14,15,16, 22, 23]. What drives this selection? It was previously found that fzo-1 mutation impacted the developmental pace of the worms [35,36,37]. Nevertheless, we show that introducing heteroplasmic mtDNA deletions strongly aggravated these phenotypes, leading to a sharp decline in survival and fecundity. This was manifested by increased embryonic lethality and delayed or even arrested larval development of worms with high levels of deleterious heteroplasmic mtDNAs (Fig. 6A). This strong selective pressure also positively correlated with the decline in heteroplasmic deletion levels across generations. Since the loss of ΔmtDNA molecules is associated with the improved health condition of the worms within 3–4 generations, mitochondrial fusion is likely critical for tolerating ΔmtDNA molecules to maintain fitness.

Fig. 6
figure 6

A model depicting selective removal of ΔmtDNA during development and gametogenesis of C. elegans. A Schematic representation—high ΔmtDNA levels (red) associate with reduced egg hatching and delayed development. B Schematic representation of mitochondria in the gonad, carrying intact +mtDNA (black) and ΔmtDNA (red). Mitochondria in fzo-1(wt) oocytes form an intracellular mitochondrial network suggested to enable mitochondrial functional complementation, hence allowing ΔmtDNA inheritance. In contrast, in fzo-1(mut) animals, the fusion machinery is compromised, mitochondria are fragmented, suggesting that ΔmtDNA molecules are ‘exposed’ and hence selectivity eliminated in the gonad of fzo-1(mut);+/ΔmtDNA. C Two possible non-mutually exclusive mechanisms can explain the selective removal of ΔmtDNA molecules in the gonad of fzo-1(mut);+/ΔmtDNA. Signals from mitochondria are suggested to trigger programmed cell death of germ cells when the transition from a globular to a tubular organization is disrupted, as in fzo-1(mut) (left). Sperm-derived signals are suggested to trigger lysosome acidification in mature oocytes, which can activate mitophagy (right). These processes can selectively impact the removal of mitochondria with high levels of ΔmtDNA, respectively, in fzo-1(mut) animals. The figure was created in part using BioRender.com

Full size image

In parallel, our discovery that the levels of heteroplasmic deletions are specifically reduced in the gonad when the fusion machinery is impaired (Fig. 6B) is consistent with previous findings of selective forces in the human germline [4, 5, 12]. Moreover, in Drosophila, germline selection of heteroplasmic mutations was directly observed in response to compromised fusion and quality control machinery [12, 21, 22]. We found that disrupting mitophagy in addition to mitochondrial fusion (fzo-1(mut);pdr-1(mut)) in the presence of ΔmtDNA sharply increased lethality already in G1 animals (reflected by the deviation from Mendelian ratios). Likewise, we found that disrupting Parkin-mediated mitophagy did not block germline selection, affecting the specific removal of mtDNA. These findings agree with germline selection in Drosophila, where downregulation of mitophagy factors, BNIP3 and Atg1, significantly blocked selection against fragmented mitochondria in germline cysts [22]. We, therefore, propose that selection against deleterious mtDNA molecules across generations affects the fitness of the organism, at two levels, namely during embryogenesis and larvae development as well as during oogenesis (Fig. 6A, B).

Two quality control processes were previously suggested to impact germline health and may contribute to selection against mutated mtDNA in fzo-1(mut) animals. Firstly, increased apoptosis was observed upon disruption of mitochondrial transition from a globular to a tubular organization in C. elegans oogenesis [38]. This may contribute to selective mitochondrial removal since selective export of mitochondria from germ cells undergoing apoptosis was observed [39]. Secondly, sperm-derived signals were found to induce lysosome acidification in mature oocytes prior to fertilization [40] and could promote mitophagy activation of fragmented mitochondria in fzo-1(mut) animals (Fig. 6C). Compromising either of these two processes led to reduced brood size. Therefore, we argue that the interaction between mitochondrial fusion and quality control machinery is not only critical to cope with ΔmtDNA within the cell but is essential for the organism’s fecundity, development, and survival across generations. Other selection mechanisms, such as selective replication, observed in Drosophila [12], or homologous recombination of a deletion and a corresponding duplication (in case of “triplasmy”) [41] could also contribute to selection against mutated mtDNA in C. elegans. Our results suggest that the latter explanations are less likely for the heteroplasmic strains described in this study.

Our analysis of three different mtDNA deletions in C. elegans revealed significant differences in the pace of their loss and levels of phenotypic severity when grown in the presence of a fzo-1 mutant. Indeed, these three deletions differ in size and encompass different sets of mtDNA genes and tRNAs, suggesting that the fusion machinery is sensitive to differential severity of the phenotypic impact of heteroplasmic mutations. This finding is in line with differences in the penetrance of disease-causing mutations, which range between 60 and 80%, depending on the symptoms [9]. Nevertheless, the question about the functional importance of certain mtDNA regions versus others remains open. This calls for a screen of mtDNA mutants that will systematically enable assessing the sensitivity of the mitochondrial quality control and fusion machinery in differentiating the phenotypic impact of a variety of mutations, locations, and sizes.

If mitochondrial fusion is indeed important for modulating the inheritance of mtDNA heteroplasmy, one could anticipate that dysfunctional mitochondrial fusion, such as in the case of Charcot Marie Tooth type 2A (CMT2A) patients, will affect patterns of heteroplasmy. Our deep mtDNA sequencing analysis of three CMT2A pedigrees lends first clues that this might be the case [42]. Whereas two of the pedigrees did not reveal any potentially functional mtDNA mutations or deletions, we found that the levels of a potentially functional mtDNA mutation in a patient were notably lower than her healthy maternal relatives. We note that these results are in line with the observations in worms, supporting our working hypothesis that the fusion machinery modulates the levels of deleterious mtDNA heteroplasmy across generations to allow tolerance and survival. However, there are two mitofusin genes (MFN1 and MFN2) in humans, and MFN2 as well as DRP1 (inner membrane fusion) also function as tethers at mitochondria-associated ER membranes [43, 44] and could impact Parkin-mediated mitochondrial quality control [34, 45, 46]. Future collection of a larger number of CMT2A pedigrees is required to draw clearer conclusions.

The complete loss of ΔmtDNA across C. elegans generations underlines the fusion machinery as an attractive candidate target for future treatment of mitochondrial disorders. For example, the activity of protein quality control systems, including this machinery, declines during the aging of the individual [47, 48], and the levels/repertoire of mtDNA heteroplasmic deletions increase in tissues from aged individuals [49]. It would, therefore, be of great interest to assess the importance of such three-way interaction (i.e., mitochondrial fusion-mitochondrial quality control and patterns of heteroplasmy) to the tendency to develop age-associated diseases.

Conclusions

Here, by manipulating the fusion machinery in C. elegans, we demonstrated that fzo-1 (mitofusin) is a key modulator of mtDNA heteroplasmy while showing its impact on the transmission of heteroplasmic deletions across generations in living animals. Firstly, we discovered that a fzo-1(mut) led to the complete loss of three different mtDNA deletions, separately. These findings provide experimental support for the hypothesis that functional complementation among mitochondria in the intracellular network likely enables the survival and prevalence of deleterious heteroplasmic mtDNA mutations in the population [25, 26]. Secondly, we found that the fzo-1(mut) was differentially sensitive to the size/nucleotide positions of the different mtDNA deletions. Third, fzo-1(mut);pdr-1(mut) double mutants selected against the survival of animals with heteroplasmic ΔmtDNA deletion and accelerated the loss of ΔmtDNA molecules across generations. Taken together, our results demonstrate the importance of cross-talk between mitochondrial fusion and the mitochondrial quality control machinery in protecting living animals from the adverse impact of inheriting deleterious mtDNA molecules.

Methods

Nematodes and growth conditions

A list of strains used in this work and name abbreviations are found in Additional file 1: Table S4. All strains were outcrosses to our N2 stock at least four times. Nematodes were grown on Nematode Growth Medium (NGM) plates seeded with the Escherichia coli OP50-1 strain at 15 °C.

Statistical analyses

To test the null hypothesis that the heteroplasmic deletions reduce the fitness of WT (Additional file 1: Figs. S2B and S3E), fzo-1(mut) (Fig. 1B, C and Fig. 2C) or fzo-1(mut);pdr-1(mut); (Additional file 1: Fig. S4A-C) as compared to wild type or fzo-1(mut) animals, we used one-way analysis of variance (ANOVA) followed by a Tukey’s post hoc test. We used the same test to compare the levels of ΔmtDNA, +mtDNA, and TMRE staining in fzo-1(mut) and fzo-1(wt) strains (Fig. 2F, Fig. 4A, B, and Additional file 1: S1B). Data are presented as bar graphs showing means ±1 standard error of the mean (1SE). To compare the mtDNA (ΔmtDNA and/or +mtDNA) levels between two conditions and assess statistical significance (Fig. 4C–G, Fig. 5D–H, and Additional file 1: S3C-D), we used the Wilcoxon Mann-Whitney rank sum test. Data are presented as scatter dot plots showing points for data or bar graphs showing means ± 1SE. To test whether heteroplasmic progeny deviated from the expected Mendelian ratio, we used χ2 goodness of fit test, data are presented in Additional file 1: Table S1. To examine differences in developmental rate across generations of fzo-1 mutant (Figs. 2B and 5B) or wild type animals (Additional file 1: Figs. S2A and S3F), we used Cox proportional-hazards regressions (Additional file 1: Table S2). To control for the dependency of individuals within biological repeats, a robust jackknife variance estimator grouped by observations per experimental plate was used. Data points showing the percent of total animals that reached adulthood within the experimental time (11 days) are presented as line graphs. To test for changes in heteroplasmic deletions (Fig. 2D, E, Fig. 3A, B, Fig. 5C, and Additional file 1: S3H-I, S4D) or +mtDNA (Additional file 1: Fig. S2C-D) levels across generations of fzo-1 mutant (Fig. 2D, Fig. 3A, B, Fig. 5C, and Additional file 1: S2C) or wild type animals (Fig. 2E and Additional file 1: S2D, S3H-I, and S4D), we used fractional regressions with logit link function (Additional file 1: Table S3). To compare the change in heteroplasmic deletion levels across generations in the different genetic backgrounds (Fig. 3C), we used a fractional regression with logit link function followed by within generation pairwise comparisons (Additional file 1: Table S3). To control for the dependency of individuals within biological repeats, a robust variance estimator grouped by observations per experimental plate was used. Odds ratios were calculated to determine the likelihood of heteroplasmy in a given generation. Data are presented in box plot representation: center line, median; box limits, upper and lower quartiles; whiskers, minimum and maximum; points, data. The numbers of biological repeats (N) and individuals (n) in each condition tested are noted in the figure legends (Figs. 1, 2, 3, and 5 and Additional file 1: S1-S4).

Single worm genotyping

Animal genotype was determined using a single worm PCR Phire Animal Tissue Direct PCR Kit (Thermo Scientific) with primers to detect fzo-1 or pdr-1 deletions. The list of PCR primers is found in Additional file 1: Table S5. The resultant amplification products were visualized by gel electrophoresis to determine the genotype.

Establishing and maintaining fzo-1 heterozygotes heteroplasmic lines

Mutant fzo-1(tm1133) animals (strain CU5991) are very poor in mating and, therefore, were first crossed with males expressing a yellow fluorescent protein marker (unc-54p::YFP). Heteroplasmic hermaphrodites (uaDf5/+, bguDf1/+ or bguDf2/+) were then crossed with fzo-1(tm1133);unc-54p::YFP heterozygote males to ensure maternal inheritance of mtDNA deletions (Fig. 1A). Heteroplasmic (uaDf5/+, bguDf1/+ or bguDf2/+) animals that were heterozygotes for fzo-1(tm1133), fzo-1(ht), were identified and maintained using single worm genotyping, establishing independent heteroplasmic lines carrying fzo-1(ht).

Mutant pdr-1(gk448) animals (strain VC1024) were first crossed with heteroplasmic hermaphrodites (uaDf5/+) and with fzo-1(tm1133) to establish double mutant strains. fzo-1(tm1133);pdr-1(gk448) double mutants were crossed with unc-54p::YFP (marker); then, the F1 heterozygote males, fzo-1(tm1133);pdr-1(gk448);unc-54p::YFP, were crossed with heteroplasmic hermaphrodites, ΔmtDNA;pdr-1(gk448) to ensure maternal inheritance of uaDf5 (Fig. 5A). Heteroplasmic (uaDf5/+) animals that were homozygous to pdr-1(gk448), pdr-1(mut), and heterozygotes for fzo-1(tm1133), were identified using PCR genotyping. fzo-1(ht) were then maintained using single worm genotyping to establish a heteroplasmic line carrying fzo-1(ht);pdr-1(mut).

Monitoring animals across generations

Single animals from the heterozygotes heteroplasmic lines (uaDf5/+, bguDf1/+, or bguDf2/+) were isolated, allowed to lay eggs, and genotyped using a single worm PCR for fzo-1. The progeny of fzo-1(ht) was again isolated, allowed to lay eggs, and screened to identify mutant or wild type fzo-1 animals (G1m and G1wt, respectively). Heterozygous progeny was maintained to generate G1. The progeny of mutant or wild type animals (G2m and G2wt, respectively) was then monitored and/or isolated and allowed to lay eggs. This was repeated over several generations (G2m-G4m and G2wt-G4wt, respectively; Fig. 2A).

To test for residual ΔmtDNA (uaDf5, bguDf1, or bguDf2), G4m hermaphrodites were crossed with wild type males; the progeny was allowed to self-propagate, isolated, allowed to lay eggs, and genotyped. Heteroplasmy levels in fzo-1(wt) animals (Gm→Gwt) were then examined.

Embryo hatching

Gravid animals were moved to a fresh plate for 2–12 h and then removed from the plates. Hatching was examined after 48 h. The numbers of biological repeats (N) and individuals examined (n) in each condition tested are noted in the figure legends (Fig. 1B, Fig. 2C and Additional file 1: S2B, S3E and S4A). Individual data values are included in Additional file 2.

Developmental timing

Single embryos were placed on fresh plates and allowed to grow at 15 °C. The animals’ developmental stage was examined every day, and the number of animals reaching reproductive adulthood on each day was recorded. Developmentally arrested animals that did not reach adulthood in over 11 days were excluded. The numbers of biological repeats (N) and individuals examined (n) in each condition tested are noted in the Figure legends (Fig. 1C, Fig. 2B, Fig. 5B and Additional file 1: S2A, S3F and S4B-C). Individual data values are included in Additional file 2.

Mitochondria staining and membrane potential assay

Age-synchronized adults were placed on NGM plates seeded with the E. coli OP50-1 and containing 100 μM MitoTracker Deep Red FM (Thermofisher) or 100 μM TMRE (tetramethylrhodamine, ethyl ester) (Biotum). The animals were kept on the plates for 24 h in the dark and then recovered on regular plates for 2 h. Animals were then fixed with 4% paraformaldehyde and imaged using a LEICA DM5500 B epifluorescence microscope. MitoTracker was imaged using a × 40 or a × 60 numerical aperture objective with a 633-nm line for excitation. TMRE was imaged using a × 10 numerical aperture objective with a 549-nm line for excitation. TMRE staining was quantified using CellProfiler cell image analysis software. The numbers of individuals examined (n) in each condition tested are noted in the figure legend of Additional file 1: S1B. Individual data values are included in Additional file 2.

DNA purification and extraction

Total DNA was extracted using a QuickExtract kit (Lucigen). Unless otherwise indicated, DNA was extracted from a single worm. When populations were examined, ~ 5 animals were collected. For embryos, DNA was extracted from 15 to 30 embryos. For gonad-soma analysis, gonads were dissected from 5 to 10 animals per biological repeat. DNA was extracted separately from the gonads and soma.

Quantification of mtDNA copy numbers

mtDNA levels were measured by qPCR performed on a C1000 Thermal Cycler (Bio-Rad) with KAPA SYBRFAST qPCR Master Mix (KAPA Biosystems). Analysis of the results was performed using CFX Manager software (Bio-Rad). To quantify the different mtDNA molecules, three sets of primers were used for truncated (ΔmtDNA), intact (+mtDNA), and total mtDNA molecules for each of the three deletions examined (Additional file 1: Table S5). ΔmtDNA levels were determined using primers located in the boundaries of the deletions and thus amplified only from the truncated copies. +mtDNA levels were determined using one primer located within the deletion and a second primer located outside of the deletion and thus amplified only from the intact copies. Total mtDNA levels were determined using primers located outside of the deletion area. For each sample, the average CT (threshold cycle) of triplicate values obtained for these mtDNA molecules was normalized to a nuclear DNA marker using the 2−ΔΔCT method [50]. Truncated/total or intact/total ratio was defined as the ratio of the normalized CT values of truncated to total mtDNA for a given animal or strain. The numbers of biological repeats (N) and individuals examined (n) in each condition tested are noted in the figure legends (Figs. 2D–F, 3A, B, 4A–G and 5C–H and Additional file 1: S2C-D, S3C-D, S3H-I and S4D). Individual data values are included in Additional file 2.

Reanalysis of whole-genome sequencing

The occurrence of deletions and duplications in the mtDNA was assessed by analyzing whole-genome sequencing reads from the NCBI’s Sequence Read Archive (SRA) database, corresponding to two of the strains used in this study: SRR801606 - SRR801609 for strain VC40128, and SRR793379 - SRR793382 for strain VC20469 [30]. Reads were downloaded, trimmed according to quality scores (default parameters), and filtered for excluding read-through adapters sequences. The processed reads were mapped against the entire genome of the N2 wild type strain to exclude contamination of nuclear mitochondrial DNA (NUMTs) in our results. N2 genome information was downloaded from NCBI’s RefSeq database (accession numbers: Chr_1 - NC_003279.8, Chr_2 - NC_003280.10, Chr_3 - NC_003281.10, Chr_4 - NC_003282.8, Chr_5 - NC_003283.11, Chr_X - NC_003284.9, and Chr_M - NC_001328.1). We used a genome browser to visualize reads coverage and detect the previously reported deletions in the examined strains [29]. Deletions and duplications were determined by two parameters; a relative change in sequencing coverage and the mapping of broken read pairs at the edges of the deletions (i.e., new sites formed by the deletion or duplication). The algorithm would identify pairs as broken if the observed distance post mapping between the pairs was significantly larger than the expected distance, according to the size selection procedure during library preparation. Analyses described above were performed using CLC Genomics workbench 20, QIAGEN.

Gonad dissection

G2 wild type or mutant animals were placed in a drop of ultra-pure water on a coverslip slide, and a 25-gauge needle was used to remove the gonads from the body of the animals. Gonads or the remaining carcasses were then transferred to a DNA extraction buffer.

Availability of data and materials

All data generated or analyzed during this study are included in this published article and its supplementary information files. Individual data values are included in Additional file 2. The sequencing datasets analyzed in the current study are available in the NCBI Sequence Read Archive repository accession number SRP018046 [30], http://www.ncbi.nlm.nih.gov/sra.

References

  1. Schon EA, Gilkerson RW. Functional complementation of mitochondrial DNAs: mobilizing mitochondrial genetics against dysfunction. Biochimica et Biophysica Acta (BBA)-General Subjects. 2010;1800(3):245–9.

    CAS  Google Scholar 

  2. Avital G, Buchshtav M, Zhidkov I, Tuval Feder J, Dadon S, Rubin E, et al. Mitochondrial DNA heteroplasmy in diabetes and normal adults: role of acquired and inherited mutational patterns in twins. Hum Mol Genet. 2012;21(19):4214–24.

    CAS  PubMed  PubMed Central  Google Scholar 

  3. Goto H, Dickins B, Afgan E, Paul IM, Taylor J, Makova KD, et al. Dynamics of mitochondrial heteroplasmy in three families investigated via a repeatable re-sequencing study. Genome Biol. 2011;12(6):R59.

    CAS  PubMed  PubMed Central  Google Scholar 

  4. Wei W, Tuna S, Keogh MJ, Smith KR, Aitman TJ, Beales PL, et al. Germline selection shapes human mitochondrial DNA diversity. Science. 2019;364(6442).

  5. Zaidi AA, Wilton PR, Su MS, Paul IM, Arbeithuber B, Anthony K, et al. Bottleneck and selection in the germline and maternal age influence transmission of mitochondrial DNA in human pedigrees. Proc Natl Acad Sci U S A. 2019;116(50):25172–8.

    CAS  PubMed  PubMed Central  Google Scholar 

  6. Sharpley MS, Marciniak C, Eckel-Mahan K, McManus M, Crimi M, Waymire K, et al. Heteroplasmy of mouse mtDNA is genetically unstable and results in altered behavior and cognition. Cell. 2012;151(2):333–43.

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Latorre-Pellicer A, Lechuga-Vieco AV, Johnston IG, Hamalainen RH, Pellico J, Justo-Mendez R, et al. Regulation of mother-to-offspring transmission of mtDNA heteroplasmy. Cell Metab. 2019;30(6):1120–1130 e1125.

    CAS  PubMed  PubMed Central  Google Scholar 

  8. Hahn A, Zuryn S. The cellular mitochondrial genome landscape in disease. Trends Cell Biol. 2019;29(3):227–40.

    CAS  PubMed  Google Scholar 

  9. Craven L, Alston CL, Taylor RW, Turnbull DM. Recent Advances in Mitochondrial Disease. Annu Rev Genomics Hum Genet. 2017;18:257–75.

    CAS  PubMed  Google Scholar 

  10. Floros VI, Pyle A, Dietmann S, Wei W, Tang WCW, Irie N, et al. Segregation of mitochondrial DNA heteroplasmy through a developmental genetic bottleneck in human embryos. Nat Cell Biol. 2018;20(2):144–51.

    CAS  PubMed  PubMed Central  Google Scholar 

  11. Burgstaller JP, Johnston IG, Jones NS, Albrechtova J, Kolbe T, Vogl C, et al. MtDNA segregation in heteroplasmic tissues is common in vivo and modulated by haplotype differences and developmental stage. Cell Rep. 2014;7(6):2031–41.

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Jeedigunta SP, Minenkova AV, Palozzi JM, Hurd TR. Avoiding extinction: recent advances in understanding mechanisms of mitochondrial DNA purifying selection in the germline. Annu Rev Genomics Hum Genet. 2021;22:55–80.

    PubMed  Google Scholar 

  13. Suen DF, Narendra DP, Tanaka A, Manfredi G, Youle RJ. Parkin overexpression selects against a deleterious mtDNA mutation in heteroplasmic cybrid cells. Proc Natl Acad Sci U S A. 2010;107(26):11835–40.

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Valenci I, Yonai L, Bar-Yaacov D, Mishmar D, Ben-Zvi A. Parkin modulates heteroplasmy of truncated mtDNA in Caenorhabditis elegans. Mitochondrion. 2015;20:64–70.

    CAS  PubMed  Google Scholar 

  15. Lin Y-F, Schulz AM, Pellegrino MW, Lu Y, Shaham S, Haynes CM. Maintenance and propagation of a deleterious mitochondrial genome by the mitochondrial unfolded protein response. Nature. 2016;533(7603):416.

    CAS  PubMed  PubMed Central  Google Scholar 

  16. Gitschlag BL, Kirby CS, Samuels DC, Gangula RD, Mallal SA, Patel MR. Homeostatic responses regulate selfish mitochondrial genome dynamics in C. elegans. Cell Metab. 2016;24(1):91–103.

    CAS  PubMed  PubMed Central  Google Scholar 

  17. Ahier A, Dai CY, Kirmes I, Cummins N, Hung GCC, Gotz J, et al. PINK1 and parkin shape the organism-wide distribution of a deleterious mitochondrial genome. Cell Rep. 2021;35(9):109203.

    CAS  PubMed  Google Scholar 

  18. Mao K, Wang K, Liu X, Klionsky DJ. The scaffold protein Atg11 recruits fission machinery to drive selective mitochondria degradation by autophagy. Dev Cell. 2013;26(1):9–18.

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Twig G, Elorza A, Molina AJA, Mohamed H, Wikstrom JD, Walzer G, et al. Fission and selective fusion govern mitochondrial segregation and elimination by autophagy. Embo J. 2008;27(2):433–46.

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Malena A, Loro E, Di Re M, Holt IJ, Vergani L. Inhibition of mitochondrial fission favours mutant over wild-type mitochondrial DNA. Hum Mol Genet. 2009;18(18):3407–16.

    CAS  PubMed  Google Scholar 

  21. Kandul NP, Zhang T, Hay BA, Guo M. Selective removal of deletion-bearing mitochondrial DNA in heteroplasmic Drosophila. Nat Commun. 2016;7:13100.

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Lieber T, Jeedigunta SP, Palozzi JM, Lehmann R, Hurd TR. Mitochondrial fragmentation drives selective removal of deleterious mtDNA in the germline. Nature. 2019;570(7761):380–4.

    CAS  PubMed  PubMed Central  Google Scholar 

  23. Gilkerson RW, Schon EA, Hernandez E, Davidson MM. Mitochondrial nucleoids maintain genetic autonomy but allow for functional complementation. J Cell Biol. 2008;181(7):1117–28.

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Busch KB, Kowald A, Spelbrink JN. Quality matters: how does mitochondrial network dynamics and quality control impact on mtDNA integrity? Philos Trans R Soc Lond B Biol Sci. 2014;369(1646):20130442.

    PubMed  PubMed Central  Google Scholar 

  25. Rebolledo-Jaramillo B, Su MS-W, Stoler N, McElhoe JA, Dickins B, Blankenberg D, et al. Maternal age effect and severe germ-line bottleneck in the inheritance of human mitochondrial DNA. Proc Natl Acad Sci. 2014;111(43):15474–9.

    CAS  PubMed  PubMed Central  Google Scholar 

  26. Ye K, Lu J, Ma F, Keinan A, Gu Z. Extensive pathogenicity of mitochondrial heteroplasmy in healthy human individuals. Proc Natl Acad Sci. 2014;111(29):10654–9.

    CAS  PubMed  PubMed Central  Google Scholar 

  27. Tsang WY, Lemire BD. Stable heteroplasmy but differential inheritance of a large mitochondrial DNA deletion in nematodes. Biochem Cell Biol. 2002;80(5):645–54.

    CAS  PubMed  Google Scholar 

  28. Liau W-S, Gonzalez-Serricchio AS, Deshommes C, Chin K, CW LM. A persistent mitochondrial deletion reduces fitness and sperm performance in heteroplasmic populations of C. elegans. BMC Genet. 2007;8(1):8.

    PubMed  PubMed Central  Google Scholar 

  29. Thompson O, Edgley M, Strasbourger P, Flibotte S, Ewing B, Adair R, et al. The million mutation project: a new approach to genetics in Caenorhabditis elegans. Genome Res. 2013;23:1749–62.

    CAS  PubMed  PubMed Central  Google Scholar 

  30. The Million Mutation Project – A C. elegans Mutation Resource. In: NCBI Sequence Read Archive repository; 2012. https://tracencbinlmnihgov/Traces/sra/?study=SRP018046.

    Google Scholar 

  31. Bratic I, Hench J, Henriksson J, Antebi A, Burglin TR, Trifunovic A. Mitochondrial DNA level, but not active replicase, is essential for Caenorhabditis elegans development. Nucleic Acids Res. 2009;37(6):1817–28.

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Ahier A, Dai CY, Tweedie A, Bezawork-Geleta A, Kirmes I, Zuryn S. Affinity purification of cell-specific mitochondria from whole animals resolves patterns of genetic mosaicism. Nat Cell Biol. 2018;20(3):352–60.

    CAS  PubMed  Google Scholar 

  33. Tanaka A, Cleland MM, Xu S, Narendra DP, Suen DF, Karbowski M, et al. Proteasome and p97 mediate mitophagy and degradation of mitofusins induced by Parkin. J Cell Biol. 2010;191(7):1367–80.

    CAS  PubMed  PubMed Central  Google Scholar 

  34. Basso V, Marchesan E, Peggion C, Chakraborty J, von Stockum S, Giacomello M, et al. Regulation of ER-mitochondria contacts by Parkin via Mfn2. Pharmacol Res. 2018;138:43–56.

    CAS  PubMed  Google Scholar 

  35. Yasuda K, Hartman PS, Ishii T, Suda H, Akatsuka A, Shoyama T, et al. Interrelationships between mitochondrial fusion, energy metabolism and oxidative stress during development in Caenorhabditis elegans. Biochem Biophys Res Commun. 2011;404(3):751–5.

    CAS  PubMed  Google Scholar 

  36. Byrne JJ, Soh MS, Chandhok G, Vijayaraghavan T, Teoh JS, Crawford S, et al. Disruption of mitochondrial dynamics affects behaviour and lifespan in Caenorhabditis elegans. Cell Mol Life Sci. 2019;76(10):1967–85.

    CAS  PubMed  PubMed Central  Google Scholar 

  37. Luz AL, Rooney JP, Kubik LL, Gonzalez CP, Song DH, Meyer JN. Mitochondrial morphology and fundamental parameters of the mitochondrial respiratory chain are altered in caenorhabditis elegans strains deficient in mitochondrial dynamics and homeostasis processes. Plos One. 2015;10(6):e0130940.

    PubMed  PubMed Central  Google Scholar 

  38. Charmpilas N, Tavernarakis N. Mitochondrial maturation drives germline stem cell differentiation in Caenorhabditis elegans. Cell Death Differ. 2020;27(2):601–17.

    CAS  PubMed  Google Scholar 

  39. Raiders SA, Eastwood MD, Bacher M, Priess JR. Binucleate germ cells in Caenorhabditis elegans are removed by physiological apoptosis. Plos Genet. 2018;14(7):e1007417.

    PubMed  PubMed Central  Google Scholar 

  40. Bohnert KA, Kenyon C. A lysosomal switch triggers proteostasis renewal in the immortal C. elegans germ lineage. Nature. 2017;551(7682):629–33.

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Tang Y, Manfredi G, Hirano M, Schon EA. Maintenance of human rearranged mitochondrial DNAs in long-term cultured transmitochondrial cell lines. Mol Biol Cell. 2000;11(7):2349–58.

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Meshnik L, Bar-Yaacov D, Kasztan D, Neiger T, Cohen T, Kishner M, et al. Mutant C. elegans mitofusin leads to selective removal of mtDNA heteroplasmic deletions at different rates across generations. bioRxiv. 2020. https://doi.org/10.1101/610758.

  43. de Brito OM, Scorrano L. Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature. 2008;456(7222):605–10.

    PubMed  Google Scholar 

  44. Friedman JR, Lackner LL, West M, DiBenedetto JR, Nunnari J, Voeltz GK. ER tubules mark sites of mitochondrial division. Science. 2011;334(6054):358–62.

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Gautier CA, Erpapazoglou Z, Mouton-Liger F, Muriel MP, Cormier F, Bigou S, et al. The endoplasmic reticulum-mitochondria interface is perturbed in PARK2 knockout mice and patients with PARK2 mutations. Hum Mol Genet. 2016;25(14):2972–84.

    CAS  PubMed  Google Scholar 

  46. Larrea D, Pera M, Gonnelli A, Quintana-Cabrera R, Akman HO, Guardia-Laguarta C, et al. MFN2 mutations in Charcot-Marie-Tooth disease alter mitochondria-associated ER membrane function but do not impair bioenergetics. Hum Mol Genet. 2019;28(11):1782–800.

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Sharma A, Smith HJ, Yao P, Mair WB. Causal roles of mitochondrial dynamics in longevity and healthy aging. Embo Rep. 2019;20(12).

  48. Shai N, Shemesh N, Ben-Zvi A. Remodeling of proteostasis upon transition to adulthood is linked to reproduction onset. Curr Genomics. 2014;15(2):122–9.

    CAS  PubMed  PubMed Central  Google Scholar 

  49. Levin L, Mishmar D. A genetic view of the mitochondrial role in ageing: killing us softly. Adv Exp Med Biol. 2015;847:89–106.

    CAS  PubMed  Google Scholar 

  50. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 2001;25(4):402–8.

    CAS  Google Scholar 

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Acknowledgements

Strains, CU5991, VC1024, and LB138 (after outcrossing ABZ271, ABZ283, and ABZ270, respectively), were provided by the Caenorhabditis Genetics Center, which is funded by the NIH National Center for Research Resources (NCRR).

Strains, VC40128 and VC20469 (after outcrossing ABZ275 and ABZ279, respectively), were provided by the C. elegans Reverse Genetics Core Facility at the University of British Columbia, which is part of the international C. elegans Gene Knockout Consortium.

Funding

This study was funded by the Israel Science Foundation (ISF) grant 278/18 to ABZ and grant 372/17 to DM. L.M. was supported by the Ministry of Science and Technology, Yitzhak Navon Ph.D. fellowship, and Kreitman Biotech scholarship. D.B. and T.C. were supported by the Kreitman Negev scholarships.

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Affiliations

  1. Department of Life Sciences, Ben-Gurion University of the Negev, Beer Sheva, Israel

    Lana Meshnik, Dan Bar-Yaacov, Dana Kasztan, Tali Neiger, Tal Cohen, Mor Kishner, Itay Valenci, Sara Dadon, Ofer Ovadia, Dan Mishmar & Anat Ben-Zvi

  2. Department of Microbiology, Immunology and Genetics, Ben-Gurion University of the Negev, Beer Sheva, Israel

    Dan Bar-Yaacov

  3. Department of Neurology, Department of Laboratory Medicine and Pathology, Mayo Clinic, Rochester, MN, USA

    Christopher J. Klein

  4. Dr. John T. Macdonald Foundation Department of Human Genetics and Hussman Institute for Human Genomics, Miller School of Medicine, University of Miami, Miami, FL, USA

    Jeffery M. Vance & Stephan Züchner

  5. Institute of Neurology, Schneider Children’s Medical Center of Israel, Tel-Aviv University, Petach Tikva, Israel

    Yoram Nevo

Authors
  1. Lana Meshnik
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  2. Dan Bar-Yaacov
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Contributions

Conceptualization, A.B. and D.M.; experimental design, A.B. L.M., and I.V.; data acquisition, L.M., D.K., T.N., M.K., and S.D.; data analysis, L.M., D. B, T. C, C.J.K., J.M.V., Y.N., S. Z, and O.O.; writing and revising the text, A.B. and D.M. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Anat Ben-Zvi.

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Supplementary Information

Additional file 1: Table S1.

Genotypes distribution of F2 progeny in different heteroplasmic strains. Table S2. Cox proportional-hazards regression analyses. Table S3. Fractional regression analyses. Table S4. A list of C. elegans strains used in this study. Table S5. A list of primers used in this study. Figure S1. Characterization of +/ΔmtDNA animals. Figure S2. Characterization of fzo-1(wt);+/ΔmtDNA; animals. Figure S3. Characterization of the 1kbΔmtDNA and 4kbΔmtDNA animals. Figure S4. Characterization of fzo-1(wt);pdr-1(mut);+/ΔmtDNA animals.

Additional file 2.

Individual data values. Spreadsheets of numerical data for Figures 1, 2, 3, 4 and 5 and Supplementary Figures S1-S4.

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Meshnik, L., Bar-Yaacov, D., Kasztan, D. et al. Mutant C. elegans mitofusin leads to selective removal of mtDNA heteroplasmic deletions across generations to maintain fitness. BMC Biol 20, 40 (2022). https://doi.org/10.1186/s12915-022-01241-2

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  • DOI: https://doi.org/10.1186/s12915-022-01241-2

Keywords

  • C. elegans
  • fzo-1
  • Heteroplasmy inheritance
  • Mitofusin
  • mtDNA
  • PARKIN
  • pdr-1

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