Editing the Powerhouse: Mitochondrial Genome Editing for Human Gene Therapy

 

Why Mitochondrial DNA Matters

Mitochondria are the energy-producing organelles of human cells. Unlike most cellular structures, mitochondria possess their own small genome known as mitochondrial DNA (mtDNA). The human mitochondrial genome is only about 16.5 kilobases long, yet it encodes 13 essential proteins of the respiratory chain, 22 transfer RNAs, and two ribosomal RNAs required for mitochondrial protein synthesis. Because this genome is extremely compact, even a single mutation can disrupt mitochondrial function and impair cellular energy production. Mutations in mtDNA have been associated with a wide range of disorders, including primary mitochondrial diseases, Parkinson’s disease, cancer, ageing, diabetes, and other conditions linked to mitochondrial dysfunction.

One important example is Leber’s hereditary optic neuropathy (LHON), the most common inherited mitochondrial optic neuropathy. LHON causes central vision loss, usually beginning at a young age. The most common mutation, 11778G>A, occurs in the mitochondrial ND4 gene, which encodes a subunit of respiratory chain complex I. This mutation changes arginine at position 340 to histidine (R340H), disrupting normal mitochondrial function. Despite decades of research, effective treatments for mitochondrial diseases remain limited. One notable exception is idebenone, a coenzyme Q analogue approved in Europe that has shown benefit in approximately half of patients with early-stage LHON. These limitations have driven the search for technologies capable of directly correcting disease-causing mutations in mitochondrial DNA.

Early Mitochondrial Gene Editing Technologies

Genome editing has revolutionized nuclear genetics, but editing mitochondrial DNA has proven much more challenging. Recent advances have led to the development of mitochondrial base-editing systems based on DNA deaminases. The DdCBE (DddA-derived cytosine base editor) enables conversion of C:G base pairs into T:A base pairs, while TALED (TALE-linked deaminase) systems enable A:T-to-G:C base-pair conversions in mammalian mitochondria. These technologies represented major breakthroughs because they made direct editing of mammalian mtDNA possible.

Despite these important advances, deaminase-based editors have significant limitations. They can introduce only transition mutations, in which one purine is replaced by another purine or one pyrimidine by another pyrimidine. They cannot efficiently generate transversion mutations, insertions, deletions, inversions, or other complex DNA modifications. In addition, these editing systems may produce off-target and bystander edits. Consequently, many pathogenic mitochondrial mutations remain beyond the reach of current mitochondrial genome-editing technologies.

The Challenge of Editing Mitochondrial DNA

Editing mitochondrial DNA presents unique biological challenges because mammalian mitochondria lack the classical non-homologous end joining (NHEJ) pathway used by the nucleus to repair DNA double-strand breaks. Instead, linearized mitochondrial DNA is typically degraded by components of the mitochondrial replication machinery. However, previous studies have shown that mitochondria possess alternative DNA repair activities, including microhomology-mediated end joining (MMEJ) and homologous recombination, raising the possibility that these pathways could be exploited for mitochondrial genome engineering.

The primary objective of this study was to determine whether the endogenous mitochondrial MMEJ machinery could be harnessed for site-directed mutagenesis of human mtDNA. Unlike deaminase-based editing systems, an MMEJ-driven strategy has the theoretical potential to introduce virtually any type of genetic modification, including multiple nucleotide substitutions, insertions, deletions, and other sequence changes. Such an approach could substantially expand the range of mutations that can be corrected in mammalian mitochondrial DNA.

Demonstration of MMEJ Activity in Human Mitochondria

To determine whether human mitochondria possess functional microhomology-mediated end joining (MMEJ) activity, the researchers synthesized DNA duplexes containing complementary microhomology sequences and incubated them with mitochondrial extracts prepared from HEK293T cells. Successful joining of these DNA fragments through the homologous regions would indicate the presence of active MMEJ repair. The experiments demonstrated detectable MMEJ activity in mitochondrial extracts, whereas classical NHEJ activity was absent. In contrast, nuclear extracts exhibited both MMEJ and NHEJ repair activities, confirming that the observed mitochondrial MMEJ activity was not due to nuclear contamination.

Importantly, the 22-nucleotide microhomology arm was designed to correspond to the wild-type ND4 region containing nucleotide G11778, the site of the most common LHON-associated mutation. In a second experiment, engineered DNA fragments recombined through MMEJ to generate a recombinant DNA molecule containing a newly introduced BamHI restriction site, demonstrating that mitochondrial MMEJ machinery could recombine donor DNA fragments in vitro and introduce new DNA sequences into the recombinant product. Together, these findings established that human mitochondria possess endogenous DNA repair machinery that could potentially be exploited for mitochondrial genome editing.

RNA Mitochondrial Import Signals (RMIS): Delivering DNA into Mitochondria

One of the greatest challenges in mitochondrial genome engineering is delivering donor DNA into mitochondria within living cells. To overcome this obstacle, the researchers used RNA mitochondrial import signals (RMIS), specialized RNA hairpin structures that had previously been shown to facilitate RNA import into human mitochondria. The RMIS used in this study was derived from the modified D-arm of yeast tRNA-Lys(CUU) and was fused to donor DNA molecules designed to target the mitochondrial ND4 gene. The investigators asked whether this strategy could deliver larger donor DNA molecules than had previously been reported.

The experiments demonstrated that DNA duplexes composed of approximately 49-base-pair donor sequences could be imported into human mitochondria. To verify true mitochondrial localization, purified mitochondria and mitoplasts were treated with a mixture of nucleases to degrade any nucleic acids remaining outside the organelle. Donor DNA remained protected from nuclease digestion in intact mitochondria and mitoplasts but was completely degraded after mitochondrial membrane disruption, supporting its localization within the mitochondrial matrix. Remarkably, when only one strand of the DNA duplex carried the RMIS sequence, both strands were still detected inside mitochondria, suggesting that the duplex crossed the mitochondrial membranes without complete strand separation. These findings significantly expand our understanding of mitochondrial nucleic acid transport and provide a promising strategy for delivering donor DNA for mitochondrial genome engineering.

Mitochondrial Genome Editing by Donor DNA Delivery

The researchers next investigated whether imported donor DNA could directly modify mitochondrial genomic sequences. HEK293T cells were transfected with single-stranded donor DNA or preformed donor DNA duplexes designed to target the mitochondrial ND4 gene. Six days after transfection, mitochondria were isolated, the target region of mtDNA was amplified by PCR, and the products were analyzed by deep sequencing. To ensure that only genuine mitochondrial sequences were analyzed, reads originating from nuclear mitochondrial DNA sequences (NUMTs) were carefully excluded.

Deep sequencing detected a low but statistically significant number of mitochondrial genomes containing the designed four-nucleotide modification. Editing efficiencies ranged from approximately 210 to 424 counts per million, corresponding to 0.021% to 0.042% edited mitochondrial genomes. Although the efficiency was low, this study provided proof-of-concept that donor DNA delivery can introduce predefined multi-nucleotide changes into human mtDNA. Notably, RMIS-bearing double-stranded donor DNA produced significantly higher editing efficiencies than the corresponding single-stranded donor oligonucleotides. This finding is particularly important because current deaminase-based mitochondrial editing technologies cannot generate such four-nucleotide modifications, demonstrating the broader editing potential of this donor DNA-based approach.

Combining Donor DNA with CRISPR-Based Mitochondrial Cleavage

To improve mitochondrial genome editing efficiency, the researchers combined donor DNA delivery with site-specific mitochondrial DNA cleavage using mito-AsCas12a, a mitochondria-targeted CRISPR nuclease. Their rationale was that targeted cleavage of mitochondrial DNA might generate DNA ends that could facilitate donor DNA-mediated recombination through the endogenous microhomology-mediated end joining (MMEJ) pathway. To test this strategy, the researchers designed a new set of donor DNA molecules targeting a different region of the ND4 gene. These donor molecules contained five engineered nucleotide changes located within the CRISPR recognition sequence. As a result, successful editing would prevent the modified mitochondrial DNA from being recognized and cleaved again by the mito-AsCas12a/crRNA complex. Importantly, all five nucleotide substitutions were introduced at the third position of codons, producing synonymous codons that preserved the amino acid sequence of the ND4 protein.

To further improve the likelihood of successful recombination, the researchers reduced the expression of MGME1, a mitochondrial exonuclease involved in degrading linearized mitochondrial DNA. They hypothesized that decreasing MGME1 activity would stabilize the ends of cleaved mitochondrial DNA molecules and provide more time for donor DNA-mediated repair. T-REx-293-Su9-AsCas12a cells were transfected with donor single-stranded or double-stranded DNA together with the CRISPR RNA, while expression of mito-AsCas12a was induced with tetracycline. After seven days, mitochondrial DNA was isolated, amplified using unique molecular identifiers (UMIs), and analyzed by deep sequencing.

Deep sequencing detected the intended five-nucleotide modification, but the overall editing efficiency was unexpectedly extremely low, ranging from approximately 0.7 to 2.3 counts per million, corresponding to 7 × 10⁻⁵% to 2.3 × 10⁻⁴% edited mitochondrial genomes. No corresponding three- or four-nucleotide modifications were detected, demonstrating that the observed five-nucleotide changes were not random sequencing artifacts. Furthermore, no editing events were detected in control cells that did not receive donor DNA, indicating that donor DNA delivery was required for editing. As observed in the earlier experiments, double-stranded donor DNA consistently produced higher editing levels than single-stranded donor DNA.

The authors proposed several possible explanations for the unexpectedly low editing efficiency. First, the T-REx-293-Su9-AsCas12a cell line used in these experiments has reduced respiratory capacity, which may adversely affect mitochondrial metabolism, transfection efficiency, and DNA repair activity. Second, previous work suggested that mitochondrial DNA cleavage by mito-AsCas12a itself may occur with relatively low efficiency. Finally, the engineered mutations were distributed across a larger 16-nucleotide region than in the donor DNA-only experiments, which may have reduced the probability of successful recombination. Although substantial optimization is still required, these experiments provide important proof-of-concept evidence that CRISPR-assisted mitochondrial genome editing may eventually become feasible.

Proposed Mechanism of MMEJ-Based Mitochondrial Editing

The researchers proposed a mechanistic model to explain how donor DNA-mediated editing of mitochondrial DNA may occur. According to this model, naturally occurring mitochondrial DNA breaks generated by reactive oxygen species (ROS) or replication fork stalling may occasionally provide entry points for donor DNA-mediated repair through the MMEJ pathway. Because these events occur naturally within mitochondria, donor DNA may sometimes become incorporated even without deliberate CRISPR-mediated cleavage. The authors emphasized that this represents a proposed mechanism rather than one that has been experimentally demonstrated in this study.

In this model, the imported donor DNA molecules are first processed by the mitochondrial nucleases EndoG and ExoG, which may remove the RNA mitochondrial import signal (RMIS) or process the RNA-DNA junctions. The repair process may then proceed similarly to classical MMEJ. PARP1 is proposed to recruit MRE11, BRCA1, and CtIP, which generate 3′ single-stranded DNA overhangs through end resection. RAD51 and POLQ may subsequently promote strand invasion, alignment of homologous sequences, and DNA synthesis. Finally, FEN1 removes non-homologous DNA flaps, while DNA ligase III completes DNA synthesis and ligates the repaired DNA ends, thereby introducing the designed nucleotide changes into the mitochondrial genome.

The proposed model is supported by the observation that double-stranded donor DNA consistently produced higher editing efficiencies than single-stranded donor DNA. The authors suggest that double-stranded donor DNA may more closely resemble naturally occurring double-strand breaks, making it more effective at recruiting the mitochondrial DNA repair machinery. However, they also note that single-stranded donor DNA may undergo more rapid degradation within mitochondria, further reducing its editing efficiency. Although additional studies will be required to validate each step of this mechanism, the proposed model provides a plausible explanation for how programmable donor DNA-mediated mitochondrial genome editing may occur in living human cells.

Future Perspectives and Clinical Potential

Although the editing efficiencies achieved in this study remain extremely low, the work represents a significant conceptual advance in mitochondrial genome engineering. Existing mitochondria-targeted nucleases, including restriction enzymes, zinc-finger nucleases, TALENs, and ARCUS meganucleases, have primarily been developed to achieve heteroplasmy shifting by selectively eliminating mutant mitochondrial genomes. This strategy can increase the proportion of healthy mitochondrial DNA but is unlikely to be effective for nearly homoplasmic diseases, such as many cases of Leber hereditary optic neuropathy (LHON), in which almost all mitochondrial genomes already contain the pathogenic mutation.

The strategy described in this study offers a fundamentally different approach because it seeks to rewrite mitochondrial DNA rather than simply eliminate mutant genomes. If editing efficiency can be substantially improved, donor DNA-mediated genome editing could potentially correct disease-causing mutations directly within the mitochondrial genome. Moreover, unlike current deaminase-based editing systems that are limited primarily to transition mutations, this approach has the theoretical potential to introduce transversion mutations, multiple nucleotide substitutions, insertions, deletions, and other complex genomic modifications. The authors further suggest that combining donor DNA delivery with more efficient mitochondria-targeted nucleases could significantly improve editing efficiency in the future.

The authors also note that recent advances in mitochondrial CRISPR technologies, including studies using mito-AsCas12a and recent CRISPR-based editing of yeast mitochondrial DNA, support the feasibility of future mitochondrial genome editing. Continued improvements in donor DNA design, mitochondrial DNA delivery, CRISPR-mediated DNA cleavage, RNA targeting, and manipulation of mitochondrial DNA repair pathways may eventually transform this early proof-of-concept into a practical technology for both basic research and therapeutic applications.

Conclusion

This study introduces a novel strategy for editing human mitochondrial DNA by combining RNA mitochondrial import signals (RMIS) with the intrinsic microhomology-mediated end joining (MMEJ) repair machinery present in human mitochondria. The researchers demonstrated that donor DNA duplexes can be imported into mitochondria of living human cells and that delivery of these donor molecules can introduce predefined multi-nucleotide modifications into the mitochondrial ND4 gene. Although the editing efficiency remains low, the study provides important proof-of-concept evidence that programmable donor DNA-mediated mitochondrial genome editing is possible in human cells.

Unlike currently available deaminase-based mitochondrial genome-editing technologies, which are largely restricted to transition mutations, this donor DNA-based strategy has the theoretical potential to introduce virtually any type of genetic modification, including multiple nucleotide substitutions, transversions, insertions, deletions, and other complex sequence changes. Considerable optimization will be required before this approach becomes therapeutically useful, but the work establishes an important foundation for the future development of mitochondrial genome-editing technologies. If these challenges can be overcome, programmable editing of mitochondrial DNA may ultimately provide new treatment options for mitochondrial diseases such as LHON and may also advance research into many other disorders associated with mitochondrial dysfunction.

Reference

Vadim V.M., et al. Mitochondrial genome microhomology-mediated editing by donor DNA delivery into mitochondria in human cells. Molecular Therapy Nucleic Acids, 2026,102959, https://doi.org/10.1016/j.omtn.2026.102959.

Image credit: Portions of the figure in this article were generated using ChatGPT (OpenAI) or Google Gemini.

Disclaimer: This blog post is intended solely for educational and scientific informational purposes. Any mention of therapeutic drug names, including FDA-approved medications, is for the purpose of accurate reporting and discussion of biomedical research and does not constitute medical advice, endorsement, or promotion. Readers should not interpret the content as a recommendation for any specific treatment. Always consult a qualified healthcare professional for medical advice or treatment decisions.

About Marin Biologic Laboratories

Our Recent Publication/Meeting Presentation on Gene Therapy

1. Development of VNX-101, an Adeno-Associated Virus with Less Immunogenicity and Efficient Long-Term Expression of a CD19 T-Cell Engager. Molecular Therapy Methods & Clinical Development, published online July 24, 2025.

3. Cell-Based Potency Assay for Anti-CD3-Anti-CD19 Diabody. Journal of Immunological Methods. 2025. 545-114004.

3. Development of a Pharmacokinetic (PK) Mouse Serum GLP ELISA for an Anti–CD19–AntiCD3 Diabody
bioRxiv 2025.03.19.644217; doi: https://doi.org/10.1101/2025.03.19.644217.

4. American Society of Hematology (ASH) Annual Meeting 2024.
Abstract link: Using Gene Therapy to Solve Challenges with CAR-T Cell Immunotherapy: Lead Selection and Preclinical Development of an Adeno-Associated Virus with Reduced Immunogenicity Exhibiting Efficient and Long-Term Expression of an Anti-CD19 T-Cell Engager.

Comprehensive Assay Solutions for In Vitro and Cell Based Potency Assays and Pharmacokinetics Studies- Our Expertise

With 30 years of expertise in cell culture, cell-based assays, and preclinical/clinical PK/PD analysis, we specialize in offering assay services essential for a wide variety of therapeutic drug development programs, preclinical studies, IND/BLA applications, and commercialization. Our comprehensive services include both preclinical non-GLP and GLP assays, as well as non-GMP and GMP assays, providing critical support throughout the entire development pipeline.

Watch the following video and explore our latest presentation on the development and validation of potency and pharmacokinetic (PK) assays for AAV vectors, highlighting innovative methodologies and industry-leading expertise.

 

 

Download the full presentation: Development of Custom Cell Based and In vitro Potency and Pharmacokinetics (PK) Assays for AAV vectors- Marin biologic Laboratories

 

Development of Cell-Based Potency Assays: Case Studies and Blogs from Marin Biologic Laboratories (MarinBio)

 

Drug Discovery & Development Assays Offered by Marin Biologic Laboratories (MarinBio)