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Received: 30 November 2024
Revised: 12 January 2025
Accepted: 15 January 2025
Published: 17 January 2025
Citation: Pohl, K.A.; Zhang, X.; Ji, J.J.;
Stiles, L.; Sadun, A.A.; Yang, X.-J.
Derivation and Characterization of
Isogenic OPA1 Mutant and Control
Human Pluripotent Stem Cell Lines.
Cells 2025,14, 137. https://doi.org/
10.3390/cells14020137
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Article
Derivation and Characterization of Isogenic OPA1 Mutant and
Control Human Pluripotent Stem Cell Lines
Katherine A. Pohl 1,2, Xiangmei Zhang 1, Johnny Jeonghyun Ji 1,2, Linsey Stiles 3, Alfredo A. Sadun 4
and Xian-Jie Yang 1,2,*
1Jules Stein Eye Institute, Department of Ophthalmology, David Geffen School of Medicine, University of
California, Los Angeles, CA 90095, USA; kdpohl@g.ucla.edu (K.A.P.); xmzhang@jsei.ucla.edu (X.Z.)
2Molecular Biology Institute, University of California, Los Angeles, CA 90095, USA
3Department of Molecular and Medical Pharmacology, Davide Geffen School of Medicine, University of
California, Los Angeles, CA 90095, USA
4Doheny Eye Center, Department of Ophthalmology, University of California, Los Angeles, CA 91103, USA;
alfredo.sadun@gmail.com
*Correspondence: yang@jsei.ucla.edu; Tel.: +1-310-825-7020
Abstract: Dominant optic atrophy (DOA) is the most commonly inherited optic neuropa-
thy. The majority of DOA is caused by mutations in the OPA1 gene, which encodes a
dynamin-related GTPase located to the mitochondrion. OPA1 has been shown to regulate
mitochondrial dynamics and promote fusion. Within the mitochondrion, proteolytically
processed OPA1 proteins form complexes to maintain membrane integrity and the respi-
ratory chain complexity. Although OPA1 is broadly expressed, human OPA1 mutations
predominantly affect retinal ganglion cells (RGCs) that are responsible for transmitting
visual information from the retina to the brain. Due to the scarcity of human RGCs, DOA
has not been studied in depth using the disease affected neurons. To enable studies of DOA
using stem-cell-derived human RGCs, we performed CRISPR-Cas9 gene editing to gener-
ate OPA1 mutant pluripotent stem cell (PSC) lines with corresponding isogenic controls.
CRISPR-Cas9 gene editing yielded both OPA1 homozygous and heterozygous mutant ESC
lines from a parental control ESC line. In addition, CRISPR-mediated homology-directed
repair (HDR) successfully corrected the OPA1 mutation in a DOA patient’s iPSCs. In com-
parison to the isogenic controls, the heterozygous mutant PSCs expressed the same OPA1
protein isoforms but at reduced levels; whereas the homozygous mutant PSCs showed a
loss of OPA1 protein and altered mitochondrial morphology. Furthermore, OPA1 mutant
PSCs exhibited reduced rates of oxygen consumption and ATP production associated with
mitochondria. These isogenic PSC lines will be valuable tools for establishing OPA1-DOA
disease models
in vitro
and developing treatments for mitochondrial deficiency associated
neurodegeneration.
Keywords: OPA1 gene; dominant optic atrophy; CRISPR-Cas9 editing; isogenic human
pluripotent stem cell lines; mitochondria
1. Introduction
Dominant optic atrophy (DOA) is the most common inherited optic neuropathy
worldwide [
1
,
2
]. The disease prevalence is 1:25,000–1:35,000 in most populations, but it
can be as high as 1:10,000 in areas with an established founder effect [
3
,
4
]. The visual
impairment of DOA usually begins in the first two decades of life due to the loss of
retinal ganglion cells (RGCs) [
3
,
5
]. RGCs are the essential projection neurons that extend
Cells 2025,14, 137 https://doi.org/10.3390/cells14020137
Cells 2025,14, 137 2 of 20
axons through the optic nerve to transmit visual signals from the retina to the brain. The
majority of DOA is caused by mutations in the gene optic atrophy 1 (OPA1; OMIM:*605290),
which encodes a dynamin-related GTPase located to the mitochondrion [
6
–
8
]. Although
OPA1 is broadly expressed by somatic tissues, most cases of DOA are non-syndromic
and patients only exhibit symptoms related to RGC degeneration—namely progressive,
bilateral vision loss, including reduced visual acuity, color vision defects, and central visual
field defects [
3
,
9
–
11
]. A hallmark of DOA aiding in its diagnosis is temporal optic nerve
head pallor, which is attributed to the preferential loss of RGCs in the papillomacular
bundle [
12
,
13
]. Although inherited in an autosomal dominant manner, OPA1 mutations
are only ~43-88% penetrant, leading to a high degree of heterogeneity in symptoms [
11
,
14
].
Patients vary widely in their disease presentations from asymptomatic to legally blind,
even among family members harboring the same mutation [15,16].
The human OPA1 gene encodes 31 exons and can potentially express eight mRNA
isoforms resulting from alternate splicing of exons 4, 4b, and 5b [
17
,
18
]. All OPA1 pre-
cursor proteins contain an N-terminal mitochondrial targeting sequence (MTS) that al-
lows for the entry into mitochondria where they are further processed into OPA1 protein
isoforms [19–22]
. The cleavage of the MTS generates long isoforms (L-OPA1) that are
anchored to the inner mitochondrial membrane (IMM) [
23
–
25
]. L-OPA1 can be further pro-
cessed by mitochondrial peptidases at several downstream cleavage sites to generate short
isoforms (S-OPA1) that are attached to the IMM or distributed in the intermembrane space.
The ratio of OPA1 long and short isoforms are dynamically regulated by the mitochondrial
inner membrane peptidases OMA1 and YME1L [
22
], which can respond to stress signals,
such as decreased mitochondrial membrane potential and nutrient deprivation [22,26,27].
OPA1 plays critical roles in regulating mitochondrial dynamics, structure, and cellular
bioenergetics. Deletion of the Opa1 gene in mice causes early lethality at embryonic
day 9.5 [
28
]. In contrast, genetically engineered mice with splice site or missense Opa1
mutations have been shown to mimic human DOA symptoms [
29
–
34
]. Using Opa1 null
mouse embryonic fibroblasts, it was demonstrated that neither long nor short isoforms
of OPA1 can function alone [
20
]. OPA1 promotes mitochondrial fusion along with the
mitofusin proteins MFN1 and MFN2 [
21
,
35
], and cells carrying Opa1 mutations or with
reduced levels of OPA1 protein show a fragmented mitochondrial network [
36
]. In addition,
OPA1 is required to maintain cristae complexity and structural integrity, thus stabilizing
the respiratory chain complexes and controlling cytochrome C release [
23
,
37
–
40
]. Defects
in Opa1 have been associated with decreases in mitochondrial ATP synthesis and the
bioenergetic efficiency of the respiratory complexes [41–43].
To date, over 500 pathogenic mutations distributed throughout the OPA1 gene have
been reported (http://www.LOVD.nl/OPA1, accessed on 1 August 2023). Depending
on the type and location of the mutation, DOA may occur via dominant negative or
haploinsufficiency mechanisms [
17
,
40
,
44
]. Despite the ubiquitous expression of OPA1,
human cell types other than RGCs are not affected in ~80% of individuals with OPA1
mutations [
45
]. Due to the scarcity of human retinal tissues and the rarity of RGCs, which
only comprise ~2% of the total human retinal cells [
46
–
48
], OPA1-DOA has not been studied
in depth using human RGCs. The high susceptibility of human RGCs to degenerate when
OPA1 function is compromised remains not fully understood. Furthermore, recent single-
cell transcriptome profiling data have revealed that RGC subtype distribution in primates
differs significantly from rodents [
49
,
50
]. It is thus necessary to examine human RGCs in
order to elucidate the pathological mechanisms responsible for OPA1 mutation-induced
optic nerve degeneration.
Here, we describe the generation of human OPA1-mutant pluripotent stem cell (PSC)
lines with corresponding isogenic controls as tools for establishing DOA disease models
Cells 2025,14, 137 3 of 20
in vitro
. Furthermore, we characterized these PSC lines for OPA1 protein expression,
mitochondrial morphology, and cellular respiration and energy output. These isogenic PSC
lines will be useful tools to investigate how OPA1 mutations impact PSC-derived human
RGCs and facilitate studies of DOA disease mechanisms in vitro.
2. Materials and Methods
2.1. Human Pluripotent Stem Cell Cultures
Human ESCs and iPSCs were maintained in mTeSR plus medium (Stemcell Technolo-
gies, Vancouver, BC, Canada) supplemented with 1% Antibiotic Antimycotic (ThermoFisher
Scientific, Canoga Park, CA, USA) on Matrigel (Corning, Corning, NY, USA)-coated plates
at 37
◦
C with 5% CO
2
as the standard culture condition. PSCs were passaged by dissoci-
ating monolayer cells into a single-cell suspension with Accutase (Stemcell Technologies,
Vancouver, BC, Canada) and plated in the standard medium containing 10
µ
M Y-27632
(Stemcell Technologies, Vancouver, BC, Canada) for 24 h. Afterwards, PSCs were returned
to the standard medium, which was changed every other day.
2.2. CRISPR/Cas9 Gene Editing to Generate OPA1-Mutant ESC Lines
Human UCLA1 (NIH-0058) ESCs with wild-type OPA1 gene were grown till 80%
confluence under the standard condition and dissociated to a single-cell suspension using
Accutase (Stemcell Technologies, Vancouver, BC, Canada). RNPs composed of 300 pmol of
the synthetic guide RNA, “sgRNA_exon1” (Synthego, Redwood City, CA, USA) (Table S1)
and 40 pmol of Cas9 protein (Synthego, Redwood City, CA, USA) were mixed with 5
×
10
5
UCLA1 ESCs in P3 Primary Cell Nucleofector Solution (Lonza Bioscience, Walkersville, MD,
USA) and nucleofected in a Lonza Nucleofector S cuvette using a Lonza 4D-nucleofector X-
unit with the CA-137 electroporation program. Nucleofected cells were cultured in mTeSR
plus, 1% Antibiotic-Antimycotic, and 10% Clone R (Stemcell Technologies, Vancouver, BC,
Canada) for 24 h before returning to the standard medium. To assess the efficiency of
CRISPR-Cas9 editing, genomic DNA was extracted using the Purelink genomic DNA mini
kit (Invitrogen, Carlsbad, CA, USA) and amplified using Hot Star Taq DNA Polymerase
(Qiagen, Hilden, Germany) and primers flanking the double-stranded DNA break site
(XJY1349 and XJY1350, Table S2). The resulting PCR products were sequenced using primer
XJY1350 (Table S2). The rate of editing in the nucleofected population was assessed using
the Inference of CRISPR Edits (ICE) tool (https://ice.synthego.com; V2) [
51
]. PCR products
amplified from UCLA1 ESCs electroporated without any CRISPR reagents were sequenced
to generate a control file.
To enable clonal selection, a portion of the nucleofected cells was plated at a density
of 1 cell/well in 96-well plates and cultured in mTeSR plus, 1% Antibiotic-Antimycotic,
and 10% Clone R. Once the colonies reached ~20 cells, Clone R was removed. Individual
colonies were expanded, and their genomic DNA was isolated and PCR-amplified using
primers XJY1361 and XJY1362 (Table S2) as described above. The PCR products were
sequenced using primer XJY1361 (Table S2) to identify the specific insertion or deletion
mutations. Both strands of the genomic DNA around the sgRNA_exon1 of OPA1 from E10
and D9 mutant ESC lines were sequenced. In addition, all OPA1-coding exons of E10 and
D9 ESC lines were sequenced to rule out any unintended OPA1 mutations (Table S2).
2.3. CRISPR-HDR Correction of the OPA1 Mutation in 1iDOA iPSC
Prior to nucleofection, 1iDOA iPSCs [52] were dissociated to a single-cell suspension
using Accutase (Stemcell Technologies, Vancouver, BC, Canada), and resuspended at a
concentration of 25,000 cells/
µ
L in P3 Primary Cell Nucleofector solution (Lonza Bioscience,
Walkersville, MD, USA). The solution of RNP composed of 200 pmol of the synthetic
Cells 2025,14, 137 4 of 20
sgRNA, “sgRNA_exon19” (Synthego, Redwood City, CA, USA) (Table S1), and 120 pmol
of Cas9 protein (ThermoFisher Scientific, Canoga Park, CA, USA) was constituted and
incubated at room temperature for 15 min. The single-stranded Alt-R modified donor
template (Integrated DNA Technologies, Carolville, IA, USA) (Table S1) was added to the
RNP mix to a final concentration of 3
µ
M along with 5
×
10
5
1iDOA iPSCs in P3 Primary
Cell Nucleofector solution (Lonza Bioscience, Walkersville, MD, USA). The single-cell
suspension was then nucleofected in a Lonza Nucleofector cuvette S using a Lonza 4D-
nucleofector X-unit with electroporation program CA-137. Afterwards, cells were cultured
as a population in mTeSR plus, 1% Antibiotic-Antimycotic, and 10% Clone R for 24 h before
returning to the standard medium.
To facilitate clonal selection, the population of electroporated cells was subsequently
plated in Matrigel-coated 96 well plates at a density of 1 cell/well in standard medium
containing 10% Clone R. Clone R was removed once colonies reached ~20 cells. Genomic
DNA was isolated from expanded colonies using the Purelink genomic DNA mini kit
(ThermoFisher Scientific, Canoga Park, CA, USA). The region surrounding the G insertion
mutation in exon 19 of OPA1 was amplified via PCR using primers XJY1366 and XJY1367
(Table S2). PCR products were incubated with or without BstBI (New England Biolabs,
Ipswich, MA, USA) and resolved by agarose gel electrophoresis. One iPSC clone (1iDOA-
CR) was identified to carry one allele with BstBI site and was further verified by DNA
sequencing with primers XJY1424-2 and XJY1367 (Table S2). Standard G-band karyotyping
was performed by the iPSC core at Cedars Sinai Medical Center (Los Angeles, CA, USA)
as previously described [
53
] to verify that 1iDOA-CR iPSCs displayed a 46, XY normal
male karyotype.
2.4. Immunofluorescent Labeling, Confocal and Super-Resolution Imaging
PSCs grown on Matrigel-coated plastic coverslips (ThermoFisher Scientific, Canoga
Park, CA, USA) were fixed in 4% paraformaldehyde in PBS for 2 min and then incubated
in blocking solution (0.1% TritonX-100, 2% donkey serum, 10% FBS in DMEM). Cover-
slips were sequentially incubated with primary antibodies, followed by incubation with
secondary antibodies and 10
µ
g/mL 4
′
, 6-diamidino-2-phenylindole (DAPI) diluted in
blocking solution (Table S3). All incubations were for one hour at room temperature, and
followed by three, 5-min washes in PBS with 0.1% Tween 20. Coverslips were mounted on
glass slides and imaged using the Olympus BX61 scanning laser confocal microscope with
Plan-APO objectives.
For imaging performed using structured illumination microscopy (SIM), PSCs were
grown on #1.5 coverslips (Warner Instruments, Hamden, CT, USA) coated with Matrigel
(Corning, Corning, NY, USA). Fixation and immunofluorescent labeling were as described
above (Table S3). Coverslips were mounted on glass slides using Vectashield (Vectorlabs,
Burlingame, CA, USA) and sealed with CoverGrip (Biotium, Fremont, CA, USA). SIM im-
ages were captured using General Electric DeltaVision OMX microscope with a PlanApoN
60
×
/1.42 NA oil objective (Olympus, Tokyo, Japan). Immersion oil with a refractive index
of 1.516 was used. Images were acquired in 3D-SIM mode using a Z-spacing of 0.125
µ
m
and reconstructed using SoftWoRx software 7.2.1 (GE Healthcare Technology, Chicago,
IL, USA).
2.5. Western Blot
PSCs were washed twice in cold PBS and then incubated with lysis buffer (10
µ
M
HEPES, 10
µ
M KCL, 0.1% NP40, 1.3 mM MgCl
2
) supplemented with 1
×
protease and
phosphatase inhibitor (Cell Signaling Technology, Danvers, MA, USA) for 2 min at room
temperature. Cells were manually dissociated and agitated at 4
◦
C in lysis buffer for
Cells 2025,14, 137 5 of 20
15 min. Cell extracts were centrifuged at 13,000 rpm at 4
◦
C for 10 min, after which
supernatants were collected. Protein concentration was quantified using the micro-BCA
protein assay kit (ThermoFisher Scientific, Canoga Park, CA, USA). 20
µ
g of protein lysate
per sample was loaded on a 4–12% NuPAGE gel (Invitrogen, Carlsbad, CA, USA). Following
electrophoresis, the gel was transferred to a PVDF membrane (MilliporeSigma, Temecula,
CA, USA) under reducing conditions. The membrane was incubated sequentially with
primary and secondary antibodies (Table S3) according to the Near Infrared Western Blot
Detection technical guide (LI-COR Biosciences, Lincoln, NE, USA). The Western blots were
imaged and quantified using the Odyssey
®
CLx Imaging System (LI-COR Biosciences,
Lincoln, NE, USA).
2.6. Cell Respiration Assays
Human PSCs were dissociated using Accutase and seeded at a density of 10,000 cells/well
in a Matrigel-coated Seahorse XF 96-well plate (Agilent, Santa Clara, CA, USA) in 60
µ
L
of standard PSC medium and 10
µ
M Y-27632. The following day when cells were ap-
proximately 80% confluent, the oxygen consumption rate (OCR) and the extracellular
acidification rate (ECAR) were measured in parallel in a Seahorse XF96 Extracellular Flux
Analyzer (Agilent, Santa Clara, CA, USA). Approximately 1 h prior to analysis, PSC
medium was changed to XF assay media (unbuffered DMEM supplemented with 10 mM
glucose, 2 mM glutamine, 1 mM pyruvate, and 5 mM HEPES) and the plate was incubated
at 37
◦
C, without CO
2
. Compounds were injected sequentially throughout the assay via
injection ports A-D. Final concentrations of injected compounds included: 2
µ
M oligomycin
(Port A), 0.5
µ
M (Port B) and 0.9
µ
M (Port C) FCCP, and 2
µ
M antimycin A and 2
µ
M
rotenone (Port D). Upon assay completion, the plate was washed with PBS and fixed with
4% PFA. Nuclei were stained with 10 ng/mL of Hoechst 33,342 (ThermoFisher, Canoga
Park, CA, USA) and counted with an Operetta High-Content Imaging System (PerkinElmer,
Tempe, AZ, USA). Rate measurements were normalized to the number of Hoechst-positive
nuclei stained before data analysis. Data were analyzed and plotted using the Seahorse
Wave Desktop XF software (Agilent, Santa Clara, CA, USA) and exported to Microsoft
Excel and GraphPad Prism #9.4.1. ATP production rates were calculated as previously
described [54,55].
2.7. Statistical Analysis
Seahorse cell respiration data were analyzed using GraphPad Prism #9.4.1 software.
Ordinary one-way ANOVA and Tukey’s multiple comparisons tests were used. All error
bars are presented as mean value ±SEM. p< 0.05 was considered statistically different.
3. Results
3.1. Generation of Isogenic OPA1-Mutant ESC Lines Using CRISPR-Cas9 Gene Editing
To provide tools for the study of OPA1-DOA
in vitro
, we first generated OPA1 het-
erozygous and homozygous mutant ESC lines using CRISPR-Cas9 gene editing. The
wild-type (WT) UCLA1 human ESC line was electroporated with ribonucleotide-protein
(RNP) complexes consisting of the Cas9 protein and a small guide RNA (sgRNA) targeting
exon 1 of the OPA1 gene (sgRNA_exon1), which was designed to utilize a PAM site near
the OPA1 translation initiation codon to maximize the disruption of protein production
from one or both alleles (Figure 1a; Table S1). Following electroporation, the genomic DNA
sequence tracing from the edited population were compared to the parental UCLA1 ESCs
using the Inference of CRISPR Edits (ICE) tool, which calculates the percentage of insertion
or deletion (INDEL) mutations generated by the non-homologous end joining (NHEJ)
pathway after Cas9 creates a double-stranded break [
51
]. The ICE analysis indicated that
Cells 2025,14, 137 6 of 20
the RNP-mediated editing was 97% efficient, and 82% of the edits generated were predicted
to disrupt OPA1 protein function (not shown).
Cells2025,14,xFORPEERREVIEW6of21
Figure1.GenerationofOPA1heterozygousandhomozygousmutantsisogenictotheWTESCline
UCLA1.(a)SchematicdrawingofthehumanOPA1gene,whichcontains31exons(exon1‐29,4b,and
5b).Theexonsarerepresentedasboxeswithprotein‐codingregionsshadedinblack.Partialsequence
ofexon1isenlargedtoshowtheATGtranslationinitiationcodon(green),theguideRNA(magenta
box),thePAMsite(redunderline),andthepotentialCAS9cleavagesite(blackarrowhead).(b)Align‐
mentsofthegenomicDNAandpredictedproteinsequencesofthecontrolUCLA1andthetwo
CRISPR‐Cas9editedOPA1mutantESClines.ThecontrolUCLA1(OPA1+/+)ESClineshowsidentical
DNAsequencesforbothalleles.TheUCLA1‐D9ESC(OPA1−/−)containsasingle‐baseCinsertion
(boxed)inbothalleles,resultinginaframeshiftandearlystopafter11aminoacids.TheUCLA1‐E10
ESC(OPA1R5H/‐)hasaG>Amissensemutation(boxed),resultinginArg‐to‐Hischange(grey
shadedbox)inallele1,whereastheallele2hasa16‐basedeletion,whichisreplacedbya3‐base‐pair
insertion(3Csbetweenthetwoasterisks),disruptingtheATGstartcodon.(c)Brightfieldimagesshow
Figure 1. Generation of OPA1 heterozygous and homozygous mutants isogenic to the WT ESC line
UCLA1. (a) Schematic drawing of the human OPA1 gene, which contains 31 exons (exon 1-29, 4b,
and 5b). The exons are represented as boxes with protein-coding regions shaded in black. Partial
sequence of exon 1 is enlarged to show the ATG translation initiation codon (green), the guide RNA
(magenta box), the PAM site (red underline), and the potential CAS9 cleavage site (black arrowhead).
(b) Alignments of the genomic DNA and predicted protein sequences of the control UCLA1 and the
two CRISPR-Cas9 edited OPA1 mutant ESC lines. The control UCLA1 (OPA1+/+) ESC line shows
identical DNA sequences for both alleles. The UCLA1-D9 ESC (OPA1
−
/
−
) contains a single-base C
insertion (boxed) in both alleles, resulting in a frame shift and early stop after 11 amino acids. The
UCLA1-E10 ESC (OPA1 R5H/-) has a G > A missense mutation (boxed), resulting in Arg-to-His
change (grey shaded box) in allele 1, whereas the allele 2 has a 16-base deletion, which is replaced by a
3-base-pair insertion (3 Cs between the two asterisks), disrupting the ATG start codon. (c) Brightfield
Cells 2025,14, 137 7 of 20
images show that E10 and D9 display normal pluripotent stem cell morphology comparable to the
control UCLA1 ESC line from which they were derived. Scale bar, 500
µ
m. (d) Immunofluorescent
labeling of UCLA1, E10, and D9 ESC lines for pluripotent stem cell markers SOX2, OCT3/4, NANOG,
and nuclear dye DAPI. Scale bar, 50 µm.
To identify edited ESC clones that carry either heterozygous or homozygous OPA1
mutations, the edited UCLA1 cell population was plated as single cells at clonal density.
The genomic DNA of expanded ESC clones was analyzed by DNA sequencing of both
strands using primers flanking the sgRNA_exon1 cut site (Table S2). The analysis showed
that the majority of the ESC clones contained same INDEL mutations on both OPA1 alleles
in the vicinity of the guide RNA targeting site resulting in homozygous OPA1 mutants. As
an example, the ESC clone UCLA1-D9, hereby referred to as D9, contained a single-base
C insertion on both alleles (Figure 1b). This mutation caused a frame shift resulting in a
premature termination codon after eleven amino acid residuals from the translation start
codon (Figure 1b). Since this short peptide abolishes the normal function of OPA1, the ESC
clone D9 is predicted as an OPA1 null mutant (Table 1).
Table 1. OPA1 genotypes of pluripotent stem cell lines.
Line OPA1 Genotype OPA1 Mutation(s) Effect
UCLA1 +/+ c.473G>A(;)2274T>C
p.(Ser158Asn)(;)(Ala758=)
UCLA1 carries two polymorphic mutations
that do not affect OPA1 protein function.
UCLA1-E10 #R5H/−Allele 1: c.14G>A
p.(Arg5His)
Allele 2: c.3_18delinsCCC
p. No translation initiation
Arg5His is a conserved amino acid change.
The deletion and insertion mutations result in
disrupted ATG start codon.
UCLA1-D9 #−/−c.[13dup];[13dup]
p.[ Arg5ProfsTer8];[Arg5ProfsTer8]
Both alleles have the same single base insertion
resulting in premature termination
(11 vs. 1015 amino acids)
1iDOA +/−c.[1948dup];[1948=]
p.[Glu650GlyfsTer4];[Glu650=]
One allele has a single base insertion leading to
a truncated protein
(652 vs. 1015 amino acids)
1iDOA-CR +/+ c.[1947T>C];[1947=]
p.(Phe649=)
One allele has a silent mutation that results in a
novel BstBI restriction site, but normal
proteins.
2iDOA +/−c.[1417_1418del];[1417_1418=]
p.[ Ile473PhefsTer12];[Ile473=]
One allele has a single base insertion leading to
a truncated protein
(483 vs. 1015 amino acids)
H9 +/+ None reported N/A
The base positions and amino acid positions refer to reference transcript NM_130837.3 and protein sequence
NP_570850.2, respectively. +/+: OPA1 WT; +/
−
:OPA1 heterozygous mutant;
−
/
−
:OPA1 homozygous mutant.
# ESC lines derived from UCLA1 also contain its polymorphic OPA1 gene changes at positions 473 and 2274 in
exons 4 and 21, respectively.
Cells 2025,14, 137 8 of 20
Due to the high efficiency of Cas9-sgRNA_exon1 editing, the resulting OPA1 heterozy-
gous loss of function clones were rare. We identified one ESC clone UCLA1-E10, here
by referred to as E10, harboring compound heterozygous OPA1 mutations (Figure 1b).
Genomic DNA sequencing analysis showed that one allele of E10 carries a G > A missense
mutation, which changes the fifth amino acid from arginine to histidine (R5H). The other
allele contains an ATG start codon-disrupting deletion resulting in the loss of the normal
translation initiation (Table 1). Because protein function prediction software scored the
R5H mutation as having a very low likelihood of being pathogenic [
56
], the ESC clone E10
is thus considered to be similar to an OPA1 heterozygous mutant and can be used to model
DOA disease caused by true haploinsufficiency.
The OPA1 mutant D9 and E10 ESC lines can be grown and passaged under standard
pluripotent stem cell (PSC) culture conditions and show typical ESC morphology compared
to the WT parental UCLA1 (Figure 1c). Further, the D9 and E10 ESCs continue to express
pluripotent stem cell markers SOX2, OCT3/4, and NANOG as their isogenic parental line
UCLA1 (Figure 1d), suggesting that they have retained pluripotent features.
3.2. Generation of Isogenic iPSC Lines Using CRISPR-Mediated Homology-Directed Repair
To establish patients’ iPSC-based DOA disease models, we have previously generated
DOA patients’ iPSC lines carrying OPA1 heterozygous mutations [
52
]. To reduce impacts of
genetic backgrounds on DOA disease phenotype analysis, we used CRISPR-Cas9-mediated
homology-directed repair (HDR) to correct the OPA1 gene mutation carried in the iPSC
line 1iDOA, generated from a patient with the classic DOA symptoms.
The iPSC line 1iDOA carried a heterozygous single G insertion in exon19 of the OPA1
gene (Figure 2a). To carry out CRISPR HDR correction, 1iDOA iPSCs were nucleofected
with RNPs consisting of sgRNA_exon19 and Cas9 protein, along with a single-stranded
oligodeoxynucleotide (ssODN) repair template. The sgRNA_exon19 was designed to take
advantage of the 1iDOA iPSCs’ OPA1 mutation, which creates a PAM site unique to the
mutant allele (Figure 2a), thus allowing for specific targeting by the RNP complex. The
ssODN/HDR donor template contained 60 base pair homology arms (Figure 2a) and Alt-R
HDR modifications to increase oligo stability and rate of repair [
57
] (Table S1). In addition
to eliminating the G insertion, the HDR donor template also introduced a T > C silent
mutation to create a novel BstBI restriction site on the corrected allele (Figure 2a,b), which
distinguishes the correctly edited clones from other OPA1 WT lines. The removal of the G
insertion mutation from the 1iDOA also eliminated the PAM site, thus preventing further
editing of the corrected allele after successful recombination.
Through CRISPR-HDR, we identified a correctly edited clone, 1iDOA-CR (for 1iDOA
CRISPR Corrected). Genomic DNA sequencing confirmed that 1iDOA-CR carried the
corrected OPA1 allele eliminating the G insertion and the premature translation stop codon
(Figure 2b). In addition, 1iDOA-CR contained the newly created BstBI restriction site, which
does not exist in 1iDOA mutant iPSC and control wild-type ESC lines (Figure 2c). The
HDR corrected 1iDOA-CR iPSCs showed a normal karyotype (Figure 2d) and expressed
the pluripotency markers OCT3/4, SOX2, and NANOG (Figure 2e).
Cells 2025,14, 137 9 of 20
Cells2025,14,xFORPEERREVIEW9of21
Figure2.CRISPR‐Cas9mediatedcorrectionoftheOPA1mutationinaDOAiPSCline1iDOA.(a)Sche‐
maticdrawingdepictingtheregionofthe1iDOAgenomecarryingaGinsertion(yellowhighlight)in
OPA1exon19,thesgRNA_exon19(underlinedinmagenta),andthePAMsite(underlinedinred)absent
inthewild‐typeallele.The124‐nucleotidesingle‐strandedHDRdonortemplate(green)removestheG
insertionandintroducesasilentT>Cmutation(bluehighlight),whichcreatesaBstBIrestrictionsite
(blueoverline)ontheeditedallele.Uppercaselettersinthe1iDOAgenome(blacktext)indicatethese‐
quenceofOPA1exon19whereaslowercaselettersrepresentintronicsequences.(b)Alignmentsofpartial
OPA1exon19genomicDNAandpredictedproteinsequencesofthewild‐typecontrol,themutant
1iDOA,andtheCRISPR‐HDRcorrected1iDOA‐CR.DNAsequencesforbothOPA1allelesareshown
aboveofSangersequencingprofiles.Theallele2of1iDOAcontainsaGinsertion(boxed),whichleadsto
aprematurestopcodon.DNAsequencingconfirmedtheT>Creplacement(boxed)andtheBstBIsite
(blueunderline)in1iDOA‐CR.Bothallelesof1iDOA‐CRencodethewild‐typeOPA1proteinsequence.
AminoacidsthatdifferfromtheWTproteinareshadedingrey.(c)Gelimageshowsthepresenceofthe
Figure 2. CRISPR-Cas9 mediated correction of the OPA1 mutation in a DOA iPSC line 1iDOA.
(a) Schematic drawing depicting the region of the 1iDOA genome carrying a G insertion (yellow
highlight) in OPA1 exon19, the sgRNA_exon19 (underlined in magenta), and the PAM site (underlined
in red) absent in the wild-type allele. The 124-nucleotide single-stranded HDR donor template (green)
removes the G insertion and introduces a silent T > C mutation (blue highlight), which creates a BstBI
restriction site (blue overline) on the edited allele. Uppercase letters in the 1iDOA genome (black
text) indicate the sequence of OPA1 exon19 whereas lowercase letters represent intronic sequences.
(b) Alignments of partial OPA1 exon19 genomic DNA and predicted protein sequences of the wild-
type control, the mutant 1iDOA, and the CRISPR-HDR corrected 1iDOA-CR. DNA sequences for
both OPA1 alleles are shown above of Sanger sequencing profiles. The allele 2 of 1iDOA contains a
G insertion (boxed), which leads to a premature stop codon. DNA sequencing confirmed the
T>C
replacement (boxed) and the BstBI site (blue underline) in 1iDOA-CR. Both alleles of 1iDOA-CR
encode the wild-type OPA1 protein sequence. Amino acids that differ from the WT protein are shaded
in grey. (c) Gel image shows the presence of the BstBI site in the 1iDOA-CR iPSC line. A 704 bp PCR
fragments spanning the area of CRISPR HDR targeting were incubated with or without BstBI and
resolved by electrophoresis. Only 1iDOA-CR iPSCs show both the expected 704 bp and two additional
Cells 2025,14, 137 10 of 20
bands at 436 and 268 bp, indicating the presence of the novel BstBI site. (d) The 1iDOA-CR iPSCs dis-
plays a normal male karyotype after undergoing CRISPR-Cas9 gene editing. (e) Immunofluorescent
labeling of 1iDOA-CR iPSCs with pluripotent stem cell markers SOX2, OCT3/4, NANOG, and nuclei
dye DAPI. Scale bar, 30 µm.
3.3. OPA1 Protein Expression and Localization in Isogenic PSC Lines
We next investigated the protein expression levels and cellular localization of OPA1 in
the established PSC lines. Western blot analysis detected five OPA1 isoforms expressed
by HEK 293T cells and by the wild-type H9 ESCs (Figure 3a). Compared to the isogenic
parental ESC line UCLA1, the mutant E10 expressed the same isoforms but with reduced
OPA1 protein levels, whereas the homozygous mutant D9 showed a loss of OPA1 protein
expression (Figure 3a; also see Figure S1). Similarly, Western blot analysis also revealed
decreased levels of OPA1 protein expression from DOA patients’ iPSC lines 1iDOA and
2iDOA, which carry distinct heterozygous OPA1 gene mutations resulting in premature
terminations
53
(Table 1), compared to the control PSC lines H9 and UCLA1 (Figure 3a; also
see Figure S1). Noticeably, compared to the 1iDOA mutant iPSCs, the isogenic 1iDOA-CR
iPSC line showed increased OPA1 protein at levels comparable to WT controls, indicating
the restoration of OPA1 protein expression after correction of the mutation (Figure 3a).
Cells2025,14,xFORPEERREVIEW10of21
BstBIsiteinthe1iDOA‐CRiPSCline.A704bpPCRfragmentsspanningtheareaofCRISPRHDRtarget‐
ingwereincubatedwithorwithoutBstBIandresolvedbyelectrophoresis.Only1iDOA‐CRiPSCsshow
boththeexpected704bpandtwoadditionalbandsat436and268bp,indicatingthepresenceofthenovel
BstBIsite.(d)The1iDOA‐CRiPSCsdisplaysanormalmalekaryotypeafterundergoingCRISPR‐Cas9
geneediting.(e)Immunofluorescentlabelingof1iDOA‐CRiPSCswithpluripotentstemcellmarkers
SOX2,OCT3/4,NANOG,andnucleidyeDAPI.Scalebar,30μm.
3.3.OPA1ProteinExpressionandLocalizationinIsogenicPSCLines
WenextinvestigatedtheproteinexpressionlevelsandcellularlocalizationofOPA1in
theestablishedPSClines.WesternblotanalysisdetectedfiveOPA1isoformsexpressedby
HEK293Tcellsandbythewild‐typeH9ESCs(Figure3a).Comparedtotheisogenicparen‐
talESClineUCLA1,themutantE10expressedthesameisoformsbutwithreducedOPA1
proteinlevels,whereasthehomozygousmutantD9showedalossofOPA1proteinexpres‐
sion(Figure3a;alsoseeFigureS1).Similarly,Westernblotanalysisalsorevealeddecreased
levelsofOPA1proteinexpressionfromDOApatients’iPSClines1iDOAand2iDOA,which
carrydistinctheterozygousOPA1genemutationsresultinginprematureterminations53
(Table1),comparedtothecontrolPSClinesH9andUCLA1(Figure3a;alsoseeFigureS1).
Noticeably,comparedtothe1iDOAmutantiPSCs,theisogenic1iDOA‐CRiPSCline
showedincreasedOPA1proteinatlevelscomparabletoWTcontrols,indicatingtheresto‐
rationofOPA1proteinexpressionaftercorrectionofthemutation(Figure3a).
Figure3.CharacterizationofOPA1proteinexpressionincontrolandmutantPSClines.(a)Western
blotsshowingOPA1proteinexpression.TheleftpanelshowstheWTcontrolESClinesH9and
UCLA1,theOPA1heterozygousmutantESClineE10,andtheOPA1homozygousmutantESCline
D9.TherightpanelshowstheWTcontrolESClinesH9andUCLA1,theDOApatients’iPSClines
1iDOAand2iDOA,andtheCRISPRcorrectediPSCline1iDOA‐CR.AllPSClinesexceptOPA1homo‐
zygousmutantD9expressOPA1proteinisoforms(~80‐100kDa).GAPDHwasusedasaloadingcon‐
trol.NumbersindicatemolecularweightmarkerinkDa.(b)Immunofluorescentconfocalimagesshow
co‐labelingformitochondrialmarkerTOM20andOPA1inparentalESCUCLA1,andisogenicOPA1
Figure 3. Characterization of OPA1 protein expression in control and mutant PSC lines. (a) Western
blots showing OPA1 protein expression. The left panel shows the WT control ESC lines H9 and
UCLA1, the OPA1 heterozygous mutant ESC line E10, and the OPA1 homozygous mutant ESC line D9.
The right panel shows the WT control ESC lines H9 and UCLA1, the DOA patients’ iPSC lines 1iDOA
and 2iDOA, and the CRISPR corrected iPSC line 1iDOA-CR. All PSC lines except OPA1 homozygous
mutant D9 express OPA1 protein isoforms (~80-100kDa). GAPDH was used as a loading control.
Numbers indicate molecular weight marker in kDa. (b) Immunofluorescent confocal images show
co-labeling for mitochondrial marker TOM20 and OPA1 in parental ESC UCLA1, and isogenic OPA1
mutant ESCs E10 and D9. Scale bar, 20
µ
m. (c) Confocal images show co-labeling for TOM20 and
OPA1 in control H9 ESCs, DOA patient-derived 1iDOA, and isogenic 1iDOA-CR iPSCs. Scale bar,
10 µm.
Cells 2025,14, 137 11 of 20
We next examined the influence of OPA1 mutations on mitochondria in various PSC
lines. Confocal imaging of immunofluorescent labeled mitochondrial outer membrane
protein TOM20 showed similar mitochondria presence regardless of OPA1 genotypes
(Figure 3b,c). However, co-labeling for TOM20 and OPA1 revealed that mitochondria of
mutant E10 cells contained lower levels of OPA1 signals, whereas the homozygous D9
mutant showed minimal OPA1 labeling compared to the isogenic parental control line
UCLA1 (Figure 3b). Consistent with the result of Western blot analysis, the corrected iPSC
1iDOA-CR restored OPA1 labeling in mitochondria compared to the isogenic heterozygous
mutant 1iDOA (Figure 3c).
To assess the impact of OPA1 mutations on mitochondrial morphology in PSCs, we
performed high-resolution imaging using structured illumination microscopy (SIM). In the
isogenic control ESC line UCLA1, most OPA1 signals were colocalized to mitochondria
labeled by TOM20 (Figure 4a). In comparison, SIM imaging revealed reduced TOM20
and OPA1 co-localization in the mutant E10 cells. Furthermore, fragmental mitochondria
were detected in the homozygous mutant D9 cells (Figure 4a). SIM imaging also showed a
reduced colocalization of TOM20 and OPA1 signals in the heterozygous iPSC line 1iDOA
compared with its isogenic control iPSC 1iDOA-CR (Figure 4b).
Cells2025,14,xFORPEERREVIEW11of21
mutantESCsE10andD9.Scalebar,20μm.(c)Confocalimagesshowco‐labelingforTOM20andOPA1
incontrolH9ESCs,DOApatient‐derived1iDOA,andisogenic1iDOA‐CRiPSCs.Scalebar,10μm.
WenextexaminedtheinfluenceofOPA1mutationsonmitochondriainvariousPSC
lines.Confocalimagingofimmunofluorescentlabeledmitochondrialoutermembranepro‐
teinTOM20showedsimilarmitochondriapresenceregardlessofOPA1genotypes(Figure
3b,3c).However,co‐labelingforTOM20andOPA1revealedthatmitochondriaofmutant
E10cellscontainedlowerlevelsofOPA1signals,whereasthehomozygousD9mutant
showedminimalOPA1labelingcomparedtotheisogenicparentalcontrollineUCLA1(Fig‐
ure3b).ConsistentwiththeresultofWesternblotanalysis,thecorrectediPSC1iDOA‐CR
restoredOPA1labelinginmitochondriacomparedtotheisogenicheterozygousmutant
1iDOA(Figure3c).
ToassesstheimpactofOPA1mutationsonmitochondrialmorphologyinPSCs,we
performedhigh‐resolutionimagingusingstructuredilluminationmicroscopy(SIM).Inthe
isogeniccontrolESClineUCLA1,mostOPA1signalswerecolocalizedtomitochondriala‐
beledbyTOM20(Figure4a).Incomparison,SIMimagingrevealedreducedTOM20and
OPA1co‐localizationinthemutantE10cells.Furthermore,fragmentalmitochondriawere
detectedinthehomozygousmutantD9cells(Figure4a).SIMimagingalsoshowedare‐
ducedcolocalizationofTOM20andOPA1signalsintheheterozygousiPSCline1iDOA
comparedwithitsisogeniccontroliPSC1iDOA‐CR(Figure4b).
Figure4.Super‐resolutionimagingofmitochondriainWTandOPA1mutantPSCs.(a)MergedSIM
imagesofESClinesUCLA1,E10,andD9co‐labeledforthemitochondrialmarkerTOM20,OPA1,and
Figure 4. Super-resolution imaging of mitochondria in WT and OPA1 mutant PSCs. (a) Merged SIM
images of ESC lines UCLA1, E10, and D9 co-labeled for the mitochondrial marker TOM20, OPA1,
and nuclear dye DAPI. The insets are 3x in scale. Scale bars, 5
µ
m. (b) Merged SIM images of iPSC
lines 1iDOA-CR, 1iDOA, and 2iDOA co-labeled for the mitochondrial marker TOM20, OPA1, and
nuclear dye DAPI. Scale bars, 5 µm.
3.4. Impact of OPA1 Mutations on Cellular Respiration and ATP Production
To determine if OPA1 mutations affect mitochondrial function in various PSC lines, we
performed cellular respiration analysis by measuring the oxygen consumption rates (OCR)
Cells 2025,14, 137 12 of 20
and the extracellular acidification rates (ECAR). We first examined bioenergetics of the
isogenic ESC lines UCLA1, E10, and D9 with glucose and pyruvate as fuels (Figure 5a,c).
Compared with the isogenic control UCLA1, the E10 and homozygous D9 mutant ESCs
showed reduced basal respiration rates (Figure 5b). After treatments with the ATP synthase
inhibitor oligomycin followed by the uncoupler FCCP, E10 and D9 showed deficits in
maximal respiration rates, as well as corresponding reduction in mitochondrial reserve
capacity and ATP linked respiration (Figure 5b). These significant OCR deficiencies were
proportional to copies of OPA1 mutant alleles. In accordance with the effects of OPA1
mutations on oxygen consumption, the mitochondrial ATP production rates also showed
significant and corresponding decreases in E10 and D9 in comparison to the isogenic control
UCLA1 ESCs (Figure 5e). In contrast, the indicator of cellular glycolytic activities ECAR
was not significantly affected by OPA1 mutations (Figure 5d), and the ATP production
rates associated with glycolysis did not change among the isogenic ESCs (Figure 5e). As
a consequence, the rates of total ATP production in UCLA1, E10, and D9 ESCs did not
show statistically significant differences (Figure 5e). The non-isogenic WT ESC line H9
showed a lower mitochondrial ATP production rate but a higher ECAR and glycolytic ATP
production (Figure 5e), resulting in a similar total ATP production as UCLA1 (Figure 5e).
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Figure5.CellularbioenergeticsofnormalcontrolandOPA1mutantESCs.ThecontrolESClineH9,
theparentalESCUCLA1,andUCLA1‐derivedOPA1mutantESClinesE10andD9weresubjectedto
Seahorsecellularrespirationanalysis.(a)TracingsofOCRsundernormalcellularrespirationandres‐
piratorychainperturbationconditionsareshown.Verticallinesindicatethetimesofinhibitorappli‐
cations.(b)BargraphsshowquantificationsofbasalandmaximalOCRs,mitochondrialreserveca‐
pacities,aswellasOCRslinkedtoATPproduction.(c)TracingsofECARundernormalcellularrespi‐
ration,inhibitingATPsynthase(oligomycin),anduncouplingconditions(FCCP)areshown.(d)Bar
graphspresentquantificationsofbasalECARandECARunderATPsynthaseinhibition.Theratiosof
OCR/ECARreflectrelativeparticipationofmitochondrialrespirationversuscellularglycolysis.(e)
ATPproductionratesduetomitochondrialrespiration,glycolysis,andtotalcellularATPproduction
arepresented.n=5replicatesperESCline.Bargraphsshoweachnasaseparatedatapoint,whichare
presentedasmeanvalues+/‐ SEM.Adjustedp‐valueswereobtainedfromone‐wayANOVAand
Tukeyall‐pairstest.*p<0.05,**p<0.005,***p<0.0005,****p<0.0001.
Figure 5. Cellular bioenergetics of normal control and OPA1 mutant ESCs. The control ESC line H9,
the parental ESC UCLA1, and UCLA1-derived OPA1 mutant ESC lines E10 and D9 were subjected to
Cells 2025,14, 137 13 of 20
Seahorse cellular respiration analysis. (a) Tracings of OCRs under normal cellular respiration and
respiratory chain perturbation conditions are shown. Vertical lines indicate the times of inhibitor
applications. (b) Bar graphs show quantifications of basal and maximal OCRs, mitochondrial reserve
capacities, as well as OCRs linked to ATP production. (c) Tracings of ECAR under normal cellular
respiration, inhibiting ATP synthase (oligomycin), and uncoupling conditions (FCCP) are shown.
(d) Bar graphs present quantifications of basal ECAR and ECAR under ATP synthase inhibition.
The ratios of OCR/ECAR reflect relative participation of mitochondrial respiration versus cellular
glycolysis. (e) ATP production rates due to mitochondrial respiration, glycolysis, and total cellular
ATP production are presented. n= 5 replicates per ESC line. Bar graphs show each nas a separate
data point, which are presented as mean values +/
−
SEM. Adjusted p-values were obtained from
one-way ANOVA and Tukey all-pairs test. * p< 0.05, ** p< 0.005, *** p< 0.0005, **** p< 0.0001.
We also performed cellular respiration analysis using OPA1 mutant iPSC lines 1iDOA
and 2iDOA, the CRISPR-corrected iPSC 1iDOA-CR, and the WT ESC line H9. Both OPA1
heterozygous mutant 1iDOA and 2iDOA iPSCs showed reduced levels of basal, maximal,
and ATP-linked respiration, as well as a decreased reserve capacity, compared to the WT
control 1iDOA-CR and H9 (Figure 6a,b). In addition, 1iDOA and 2iDOA mutant iPSCs also
showed significantly lower levels of ECAR compared to WT control 1iDOA-CR and H9
(Figure 6c,d). In concordance with the reduced OCAR and ECAR, rates of ATP production
by mitochondria and glycolysis, as well as the total ATP production rates were significantly
reduced in OPA1 mutant iPSCs compared to WT H9 and 1iDOA-CR (Figure 6e).
Cells2025,14,xFORPEERREVIEW14of21
Figure6.BioenergeticcharacterizationofcontrolESC,DOApatients’iPSClines,andCRISPR‐HDR
correctediPSCline.ThecontrolESClineH9,DOAmutantiPSClines1iDOAand2iDOA,andthe
CRISPRHDR‐correctediPSCline1iDOA‐CRweresubjectedtoSeahorsecellularrespirationanalysis.
(a)TracingsofOCRsundernormalcellularrespirationandperturbationconditionsareshown.Verti‐
callinesindicatetimesofinhibitorapplications.(b)Bargraphspresentquantificationsofbasaland
maximalOCRs,mitochondrialreservecapacities,aswellasOCRslinkedtoATPproduction.(c)Trac‐
ingsofECARundernormalcellularrespiration,inhibitingATPsynthase(oligomycin),oruncoupling
(FCCP)conditionsareshown.(d)BargraphspresentquantificationsofbasalECARandECARunder
ATPsynthase‐inhibitedconditions.TheratiosofOCR/ECARreflectrelativeparticipationofmitochon‐
driarespirationversuscellularglycolysis.(e)ATPproductionratesduetomitochondrialrespiration,
glycolysis,andtotalcellularATPproductionarepresented.n=5replicatespercellline.Bargraphs
Figure 6. Cont.
Cells 2025,14, 137 14 of 20
Cells2025,14,xFORPEERREVIEW14of21
Figure6.BioenergeticcharacterizationofcontrolESC,DOApatients’iPSClines,andCRISPR‐HDR
correctediPSCline.ThecontrolESClineH9,DOAmutantiPSClines1iDOAand2iDOA,andthe
CRISPRHDR‐correctediPSCline1iDOA‐CRweresubjectedtoSeahorsecellularrespirationanalysis.
(a)TracingsofOCRsundernormalcellularrespirationandperturbationconditionsareshown.Verti‐
callinesindicatetimesofinhibitorapplications.(b)Bargraphspresentquantificationsofbasaland
maximalOCRs,mitochondrialreservecapacities,aswellasOCRslinkedtoATPproduction.(c)Trac‐
ingsofECARundernormalcellularrespiration,inhibitingATPsynthase(oligomycin),oruncoupling
(FCCP)conditionsareshown.(d)BargraphspresentquantificationsofbasalECARandECARunder
ATPsynthase‐inhibitedconditions.TheratiosofOCR/ECARreflectrelativeparticipationofmitochon‐
driarespirationversuscellularglycolysis.(e)ATPproductionratesduetomitochondrialrespiration,
glycolysis,andtotalcellularATPproductionarepresented.n=5replicatespercellline.Bargraphs
Figure 6. Bioenergetic characterization of control ESC, DOA patients’ iPSC lines, and CRISPR-HDR
corrected iPSC line. The control ESC line H9, DOA mutant iPSC lines 1iDOA and 2iDOA, and
the CRISPR HDR-corrected iPSC line 1iDOA-CR were subjected to Seahorse cellular respiration
analysis. (a) Tracings of OCRs under normal cellular respiration and perturbation conditions are
shown. Vertical lines indicate times of inhibitor applications. (b) Bar graphs present quantifications of
basal and maximal OCRs, mitochondrial reserve capacities, as well as OCRs linked to ATP production.
(c) Tracings of ECAR under normal cellular respiration, inhibiting ATP synthase (oligomycin), or
uncoupling (FCCP) conditions are shown. (d) Bar graphs present quantifications of basal ECAR
and ECAR under ATP synthase-inhibited conditions. The ratios of OCR/ECAR reflect relative
participation of mitochondria respiration versus cellular glycolysis. (e) ATP production rates due
to mitochondrial respiration, glycolysis, and total cellular ATP production are presented. n= 5
replicates per cell line. Bar graphs show each nas a separate data point, which are presented as mean
values +/
−
SEM. Adjusted p-values were obtained from one-way ANOVA and Tukey all-pairs test.
*p< 0.05, ** p< 0.005, *** p< 0.0005, **** p< 0.0001.
4. Discussion
Although OPA1 has been extensively studied in readily accessible cell types, these
studies do not solve the conundrum in the field of DOA research: why are RGCs particu-
larly sensitive to OPA1 mutations? To better address this question and to model OPA1-DOA
disease using pluripotent stem-cell-derived human RGCs, we generated OPA1 heterozy-
gous and homozygous mutant ESC lines from a WT control ESC line and corrected the
OPA1 mutation in a DOA patient iPSC line [
52
] using CRISPR-Cas9 gene editing tech-
nology. Because OPA1 mutation-driven DOA shows a high degree of heterogeneity and
incomplete penetrance, the OPA1 mutant PSC lines and their isogenic controls can serve as
useful research tools for reducing variabilities in DOA disease models
in vitro
and provide
opportunities to investigate OPA1 deficiency-driven DOA pathogenesis.
Using both OPA1 ESC and iPSC lines for disease modeling is beneficial, as each confers
their own specific advantages. Since the advent of gene editing technology, we are no longer
restricted to studying OPA1 mutations that occur naturally in human patients. Editing a
WT ESC line is advantageous in that all derivative lines will be isogenic with the same
genetic background. Comparing isogenic ESC lines with or without OPA1 mutations can
increase the reliability in attributing phenotypic differences observed
in vitro
to a given
OPA1 mutation and avoid issues of incomplete penetrance observed in patient pedigrees.
The OPA1 mutant ESC lines we obtained using CRISPR-Cas9 editing represent the loss
of either one or both functional alleles. The E10 ESC line carries a non-expressing null
allele and an allele with an R5H missense mutation in the expected mitochondrial targeting
sequence. Our results suggest that this mutant line has retained mitochondrial targeting
ability and function, thus may resemble a heterozygous scenario. Together, these ESC lines
can serve to better understand the requirement for OPA1 gene dosage during development
and pathogenesis.
Conversely, iPSC lines derived from DOA patients with pathogenic OPA1 variants
can be corrected
in vitro
using gene editing to generate isogenic control iPSC lines. The
Cells 2025,14, 137 15 of 20
in vitro
phenotypes of DOA patients’ iPSC-derived RGCs can be correlated to patients’
ophthalmological data. Observations from
in vitro
cell cultures can provide insights into
whether the PSC-based disease models recapitulate the DOA disease symptoms and inform
what mechanisms underlie these symptoms. Given the heterogeneity of DOA patient
population, examining a range of PSC lines with different OPA1 mutations will be beneficial
to fully understand OPA1-driven DOA.
Since OPA1 proteins form complexes within mitochondria [
25
], OPA1 mutation-
triggered DOA disease can occur via dominant negative or haploinsufficiency mecha-
nisms [
17
,
45
]. Currently, it remains challenging to correlate the location and type of OPA1
mutations to the pathogenicity of disease and to predict the mechanism of action for OPA1
mutant variants [
3
,
8
]. We have determined the expression levels of OPA1 protein in the
CRISPR-Cas9 editing-generated OPA1 mutant ESCs. The E10 and the homozygous D9
mutant ESC lines showed reduced and none-detectable OPA1 proteins compared to the WT
parental ESC line UCLA1, respectively. The E10 ESC line provides a true loss-of-function
scenario, which can be used to represent haploinsufficiency. The two heterozygous DOA
patient iPSC lines also showed reduction of OPA1 protein compared to WT control PSCs,
while the CRISPR-HDR corrected iPSC restored OPA1 protein expression levels. In OPA1
mutant PSC lines, we observed the same pattern of OPA1 isoforms [
20
,
58
] as their WT
counterparts, at relatively equal ratios. Interestingly, mouse embryonic fibroblasts (MEFs)
and human embryonic kidney (HEK) 293T cells also express the same OPA1 protein iso-
forms [
58
]. These data indicate that these OPA1 mutations do not alter OPA1 splicing
or post-translational processing in PSCs. Therefore, functional differences detected be-
tween WT and OPA1 mutant PSCs likely result from differences in total OPA1 protein
expression levels.
Previous studies have demonstrated that cells lacking OPA1 expression have highly
fragmental mitochondrial networks and reduced cristae complexity [
36
,
40
]. Using super-
resolution SIM imaging, we also detected differences of mitochondrial morphology between
the WT control and the OPA1 homozygous mutant ESC line UCLA1-D9. However, dif-
ferences of mitochondrial morphology between WT and OPA1 heterozygous mutant PSC
lines appears subtle in fixed cells by SIM imaging. Unexpectedly, we also observed some
cytoplasmic distribution of punctate OPA1 labeling signals with SIM imaging, even in the
OPA1 null ESC lines UCLA1-D9. One possibility is that the OPA1 antibody recognizing
the C-terminal portion of OPA1 also binds to other cellular proteins. Alternatively, this
could potentially be attributed to the cryptic translation initiation within exon 3 or exon 4
as reported in the NCBI protein database (NP_001341592.1 and NP_001341593.1). These
predicted OPA1 proteins lack the mitochondrial targeting sequence encoded by exon 1
and exon 2, therefore may still exist in CRISPR-Cas9 edited ESCs as the guide RNA we
used targeted exon 1. However, we have not detected any putative cryptic OPA1 protein
isoforms by Western blot analysis. Currently, it remains unclear whether the low levels of
OPA1 proteins located in the cytoplasm have any biological function.
It is known that PSCs have high anabolic activities and predominately use glycolysis
to provide metabolites for cell proliferation and the maintenance of pluripotency [
59
]. The
mTeSR medium used to culture PSCs contains high glucose (13 mM), which supports
glycolysis and anabolism. Our cellular respiration analysis using glucose and pyruvate
as fuel sources shows that compared to the WT parental ESC line UCLA1, the isogenic
OPA1 mutant ESCs had significant reduction in their basal, maximal, and ATP-linked
respiration. Moreover, these deficits are proportional to the loss of one or both functional
OPA1 alleles (E10 and D9, respectively). However, OPA1 mutant ESCs retained similar
ECARs and glycolytic ATP production as the parental UCLA1 line. This may explain why
Cells 2025,14, 137 16 of 20
the OPA1 null mutant D9 ESCs can continue to proliferate, even though they have highly
fragmental mitochondria.
Interestingly, DOA patients’ heterozygous mutant iPSC lines not only showed reduced
basal, maximal, and ATP-linked respiration rates compared to WT PSCs, they also had
reduced basal and oligomycin-induced ECAR, indicating that both oxidative phosphoryla-
tion and glycolysis were impaired in DOA patients’ iPSCs. Consequently, in OPA1 mutant
iPSC lines both mitochondrial and glycolytic ATP productions were significantly reduced,
causing a decline in total ATP production. It is worth noting that although the DOA patient
from whom 2iDOA iPSCs were derived has very mild clinical symptoms [
52
], the 2iDOA
iPSC line displayed similar cellular respiration defects
in vitro
as 1iDOA, which was de-
rived from a patient with classic DOA symptoms. This observation suggests that OPA1
mutant phenotypes may more readily manifest under
in vitro
conditions, and patients’
innate glycolytic activities may play a role in DOA pathogenesis. It would be interesting to
assess whether ECAR is consistently reduced in iPSCs of other OPA1 pathogenic variants,
as increasing research has been reported using PSCs to study DOA and other neural de-
generative diseases [
60
–
65
]. Despite the observed differences in ECARs between ESCs and
iPSCs, and the reduced cellular respiration capacities of all OPA1 mutant PSC lines, we did
not detect any obvious differences in PSC morphology and growth rates, indicating that
under the high glucose culture condition, the cellular metabolic status including the level
of cellular ATP is sufficient for PSC maintenance.
Our results demonstrate that cells’ individual genetic backgrounds, in addition to their
OPA1 mutation status, influence their OCR, ECAR, and ATP production
in vitro
. These
differences in basal bioenergetics may contribute to the varied severity of DOA symptoms
among individuals with the same OPA1 mutation.
5. Conclusions
We have established human OPA1 mutant ESC lines by the CRISPR-Cas9 gene editing
of a parental control ESC line. In addition, we have developed an isogenic control iPSC
line by correcting the pathogenic OPA1 mutation of a DOA patient’s iPSC using CRISPR-
mediated homology-directed repair. Characterization of the isogenic PSCs revealed the
impact of different mutations on OPA1 protein expression levels, mitochondrial morphol-
ogy, and cellular respiration and ATP production. Since OPA1-DOA shows incomplete
penetrance and varied severity in clinical presentations, these isogenic OPA1 mutant and
control PSCs can serve as useful tools to establish DOA disease models
in vitro
and evalu-
ate the effects of OPA1 mutations on PSC-derived human RGCs, thus facilitating disease
mechanisms studies and therapeutic treatment development.
Supplementary Materials: The following supporting information can be downloaded at: https:
//www.mdpi.com/article/10.3390/cells14020137/s1, Figure S1: Original Western Blots and Quantifi-
cation; Table S1: CRISPR-related reagent sequences; Table S2: PCR and Sequencing Primers; Table S3:
Antibodies and Dyes.
Author Contributions: Conception and design, data collection and analysis, and manuscript writing:
K.A.P.; collection and assembly of data: J.J.J., X.Z. and L.S.; provision of patients: A.A.S.; conception
and design of the research, data analysis and interpretation, manuscript writing, financial support:
X.-J.Y. All authors have read and agreed to the published version of the manuscript.
Funding: This work was in part supported by National Institute of Health (NIH) grant 2R01EY026319
and California Institute of Regenerative Medicine (CIRM) grant DISC2-13475 awarded to X.-J.Y.,
NIH grant F31EY033242 awarded to K.A.P., NIH core grant P30EY000331, and an unrestricted grant
from Research to Prevent Blindness to the Department of Ophthalmology at University of California,
Los Angeles.
Cells 2025,14, 137 17 of 20
Institutional Review Board Statement: The recombinant DNA research described has followed
National Institute of Health guidelines. The research described here was initially approved by the
Institutional Review Board (IRB) of University of California Los Angeles under IRB#19-000879-AM-
00007 on 19 September 2019, and has been reviewed on the annual basis. The stem cell research has
been reviewed and approved by the Human Pluripotent Stem Cell Research Oversight (hPSCRO)
committee of University of California Los Angeles under the permit hPSCRO 2019-003-06.
Informed Consent Statement: Not applicable.
Data Availability Statement: All relevant data are included in the manuscript. No new sequencing
data beyond verification of CRISPR editing were generated in this study. Any additional information
may be obtained by contacting the corresponding author. Requesting of human PSC lines for
research by non-profit institutions will undergo an MTA signing process with University of California
Los Angeles.
Conflicts of Interest: The authors declare no conflicts of interest.
References
1. Kjer, B.; Eiberg, H.; Kjer, P.; Rosenberg, T. Dominant optic atrophy mapped to chromosome 3q region. II. Clinical and epidemio-
logical aspects. Acta Ophthalmol. Scand. 1996,74, 3–7. [CrossRef] [PubMed]
2.
Yu-Wai-Man, P.; Griffiths, P.G.; Chinnery, P.F. Mitochondrial optic neuropathies-disease mechanisms and therapeutic strategies.
Prog. Retin. Eye Res. 2011,30, 81–114. [CrossRef] [PubMed]
3.
Yu-Wai-Man, P.; Griffiths, P.G.; Burke, A.; Sellar, P.W.; Clarke, M.P.; Gnanaraj, L.; Ah-Kine, D.; Hudson, G.; Czermin, B.; Taylor,
R.W.; et al. The prevalence and natural history of dominant optic atrophy due to OPA1 mutations. Ophthalmology 2010,117,
1538–1546.e1. [CrossRef] [PubMed]
4.
Yu-Wai-Man, P.; Chinnery, P.F. Reply: Sensorineural hearing loss in OPA1-linked disorders. Brain 2013,136, e237. [CrossRef]
[PubMed]
5.
Kjer, P. Infantile optic atrophy with dominant mode of inheritance: A clinical and genetic study of 19 Danish families. Acta
Ophthalmol. Suppl. 1959,164, 1–147.
6.
Alexander, C.; Votruba, M.; Pesch, U.E.; Thiselton, D.L.; Mayer, S.; Moore, A.; Rodriguez, M.; Kellner, U.; Leo-Kottler, B.; Auburger,
G.; et al. OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome
3q28. Nat. Genet. 2000,26, 211–215. [CrossRef]
7.
Delettre, C.; Lenaers, G.; Griffoin, J.M.; Gigarel, N.; Lorenzo, C.; Belenguer, P.; Pelloquin, L.; Grosgeorge, J.; Turc-Carel, C.; Perret,
E.; et al. Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat.
Genet. 2000,26, 207–210. [CrossRef]
8.
Thiselton, D.L.; Alexander, C.; Taanman, J.W.; Brooks, S.; Rosenberg, T.; Eiberg, H.; Andreasson, S.; Van Regemorter, N.; Munier,
F.L.; Moore, A.T.; et al. A comprehensive survey of mutations in the OPA1 gene in patients with autosomal dominant optic
atrophy. Investig. Ophthalmol. Vis. Sci. 2002,43, 1715–1724.
9.
Eliott, D.; Traboulsi, E.I.; Maumenee, I.H. Visual prognosis in autosomal dominant optic atrophy (Kjer type). Am. J. Ophthalmol.
1993,115, 360–367. [CrossRef]
10.
Han, J.; Li, Y.; You, Y.; Fan, K.; Lei, B. Autosomal dominant optic atrophy caused by six novel pathogenic OPA1 variants and
genotype-phenotype correlation analysis. BMC Ophthalmol. 2022,22, 322. [CrossRef]
11.
Cohn, A.C.; Toomes, C.; Potter, C.; Towns, K.V.; Hewitt, A.W.; Inglehearn, C.F.; Craig, J.E.; Mackey, D.A. Autosomal dominant
optic atrophy: Penetrance and expressivity in patients with OPA1 mutations. Am. J. Ophthalmol. 2007,143, 656–662. [CrossRef]
[PubMed]
12.
Amati-Bonneau, P.; Milea, D.; Bonneau, D.; Chevrollier, A.; Ferre, M.; Guillet, V.; Gueguen, N.; Loiseau, D.; de Crescenzo, M.A.;
Verny, C.; et al. OPA1-associated disorders: Phenotypes and pathophysiology. Int. J. Biochem. Cell Biol. 2009,41, 1855–1865.
[CrossRef] [PubMed]
13.
Barboni, P.; Savini, G.; Parisi, V.; Carbonelli, M.; La Morgia, C.; Maresca, A.; Sadun, F.; De Negri, A.M.; Carta, A.; Sadun, A.A.; et al.
Retinal nerve fiber layer thickness in dominant optic atrophy measurements by optical coherence tomography and correlation
with age. Ophthalmology 2011,118, 2076–2080. [CrossRef]
14.
Toomes, C.; Marchbank, N.J.; Mackey, D.A.; Craig, J.E.; Newbury-Ecob, R.A.; Bennett, C.P.; Vize, C.J.; Desai, S.P.; Black, G.C.;
Patel, N.; et al. Spectrum, frequency and penetrance of OPA1 mutations in dominant optic atrophy. Hum. Mol. Genet. 2001,10,
1369–1378. [CrossRef] [PubMed]
15. Hoyt, C.S. Autosomal dominant optic atrophy. A spectrum of disability. Ophthalmology 1980,87, 245–251. [CrossRef] [PubMed]
Cells 2025,14, 137 18 of 20
16.
Delettre, C.; Lenaers, G.; Pelloquin, L.; Belenguer, P.; Hamel, C.P. OPA1 (Kjer type) dominant optic atrophy: A novel mitochondrial
disease. Mol. Genet. Metab. 2002,75, 97–107. [CrossRef]
17.
Delettre, C.; Griffoin, J.M.; Kaplan, J.; Dollfus, H.; Lorenz, B.; Faivre, L.; Lenaers, G.; Belenguer, P.; Hamel, C.P. Mutation spectrum
and splicing variants in the OPA1 gene. Hum. Genet. 2001,109, 584–591. [CrossRef]
18.
Olichon, A.; Elachouri, G.; Baricault, L.; Delettre, C.; Belenguer, P.; Lenaers, G. OPA1 alternate splicing uncouples an evolutionary
conserved function in mitochondrial fusion from a vertebrate restricted function in apoptosis. Cell Death Differ. 2007,14, 682–692.
[CrossRef]
19.
Ishihara, N.; Fujita, Y.; Oka, T.; Mihara, K. Regulation of mitochondrial morphology through proteolytic cleavage of OPA1. EMBO
J. 2006,25, 2966–2977. [CrossRef]
20.
Song, Z.; Chen, H.; Fiket, M.; Alexander, C.; Chan, D.C. OPA1 processing controls mitochondrial fusion and is regulated by
mRNA splicing, membrane potential, and Yme1L. J. Cell Biol. 2007,178, 749–755. [CrossRef]
21.
Wang, R.; Mishra, P.; Garbis, S.D.; Moradian, A.; Sweredoski, M.J.; Chan, D.C. Identification of new OPA1 cleavage site reveals
that short isoforms regulate mitochondrial fusion. Mol. Biol. Cell 2021,32, 157–168. [CrossRef]
22.
Anand, R.; Wai, T.; Baker, M.J.; Kladt, N.; Schauss, A.C.; Rugarli, E.; Langer, T. The i-AAA protease YME1L and OMA1 cleave
OPA1 to balance mitochondrial fusion and fission. J. Cell Biol. 2014,204, 919–929. [CrossRef] [PubMed]
23.
Del Dotto, V.; Mishra, P.; Vidoni, S.; Fogazza, M.; Maresca, A.; Caporali, L.; McCaffery, J.M.; Cappelletti, M.; Baruffini, E.; Lenaers,
G.; et al. OPA1 Isoforms in the Hierarchical Organization of Mitochondrial Functions. Cell Rep. 2017,19, 2557–2571. [CrossRef]
[PubMed]
24.
Nyenhuis, S.B.; Wu, X.; Strub, M.P.; Yim, Y.I.; Stanton, A.E.; Baena, V.; Syed, Z.A.; Canagarajah, B.; Hammer, J.A.; Hinshaw, J.E.
OPA1 helical structures give perspective to mitochondrial dysfunction. Nature 2023,620, 1109–1116. [CrossRef] [PubMed]
25.
von der Malsburg, A.; Sapp, G.M.; Zuccaro, K.E.; von Appen, A.; Moss, F.R., 3rd; Kalia, R.; Bennett, J.A.; Abriata, L.A.; Dal Peraro,
M.; van der Laan, M.; et al. Structural mechanism of mitochondrial membrane remodelling by human OPA1. Nature 2023,620,
1101–1108. [CrossRef]
26.
MacVicar, T.; Langer, T. OPA1 processing in cell death and disease—The long and short of it. J. Cell Sci. 2016,129, 2297–2306.
[CrossRef]
27.
Baker, M.J.; Lampe, P.A.; Stojanovski, D.; Korwitz, A.; Anand, R.; Tatsuta, T.; Langer, T. Stress-induced OMA1 activation and
autocatalytic turnover regulate OPA1-dependent mitochondrial dynamics. EMBO J. 2014,33, 578–593. [CrossRef]
28.
Rahn, J.J.; Stackley, K.D.; Chan, S.S. Opa1 is required for proper mitochondrial metabolism in early development. PLoS ONE 2013,
8, e59218. [CrossRef]
29.
Alavi, M.V.; Bette, S.; Schimpf, S.; Schuettauf, F.; Schraermeyer, U.; Wehrl, H.F.; Ruttiger, L.; Beck, S.C.; Tonagel, F.; Pichler, B.J.;
et al. A splice site mutation in the murine Opa1 gene features pathology of autosomal dominant optic atrophy. Brain 2007,130,
1029–1042. [CrossRef]
30.
Heiduschka, P.; Schnichels, S.; Fuhrmann, N.; Hofmeister, S.; Schraermeyer, U.; Wissinger, B.; Alavi, M.V. Electrophysiological
and histologic assessment of retinal ganglion cell fate in a mouse model for OPA1-associated autosomal dominant optic atrophy.
Investig. Ophthalmol. Vis. Sci. 2010,51, 1424–1431. [CrossRef]
31.
Piquereau, J.; Caffin, F.; Novotova, M.; Prola, A.; Garnier, A.; Mateo, P.; Fortin, D.; Huynh le, H.; Nicolas, V.; Alavi, M.V.; et al.
Down-regulation of OPA1 alters mouse mitochondrial morphology, PTP function, and cardiac adaptation to pressure overload.
Cardiovasc. Res. 2012,94, 408–417. [CrossRef] [PubMed]
32.
White, K.E.; Davies, V.J.; Hogan, V.E.; Piechota, M.J.; Nichols, P.P.; Turnbull, D.M.; Votruba, M. OPA1 deficiency associated with
increased autophagy in retinal ganglion cells in a murine model of dominant optic atrophy. Investig. Ophthalmol. Vis. Sci. 2009,50,
2567–2571. [CrossRef] [PubMed]
33.
Alavi, M.V.; Fuhrmann, N.; Nguyen, H.P.; Yu-Wai-Man, P.; Heiduschka, P.; Chinnery, P.F.; Wissinger, B. Subtle neurological
and metabolic abnormalities in an Opa1 mouse model of autosomal dominant optic atrophy. Exp. Neurol. 2009,220, 404–409.
[CrossRef]
34.
Davies, V.J.; Hollins, A.J.; Piechota, M.J.; Yip, W.; Davies, J.R.; White, K.E.; Nicols, P.P.; Boulton, M.E.; Votruba, M. Opa1 deficiency
in a mouse model of autosomal dominant optic atrophy impairs mitochondrial morphology, optic nerve structure and visual
function. Hum. Mol. Genet. 2007,16, 1307–1318. [CrossRef] [PubMed]
35.
Cipolat, S.; Martins de Brito, O.; Dal Zilio, B.; Scorrano, L. OPA1 requires mitofusin 1 to promote mitochondrial fusion. Proc. Natl.
Acad. Sci. USA 2004,101, 15927–15932. [CrossRef]
36.
Olichon, A.; Baricault, L.; Gas, N.; Guillou, E.; Valette, A.; Belenguer, P.; Lenaers, G. Loss of OPA1 perturbates the mitochondrial
inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J. Biol. Chem. 2003,278, 7743–7746.
[CrossRef]
37.
Varanita, T.; Soriano, M.E.; Romanello, V.; Zaglia, T.; Quintana-Cabrera, R.; Semenzato, M.; Menabo, R.; Costa, V.; Civiletto, G.;
Pesce, P.; et al. The OPA1-dependent mitochondrial cristae remodeling pathway controls atrophic, apoptotic, and ischemic tissue
damage. Cell Metab. 2015,21, 834–844. [CrossRef]
Cells 2025,14, 137 19 of 20
38.
Frezza, C.; Cipolat, S.; Martins de Brito, O.; Micaroni, M.; Beznoussenko, G.V.; Rudka, T.; Bartoli, D.; Polishuck, R.S.; Danial,
N.N.; De Strooper, B.; et al. OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion. Cell 2006,126,
177–189. [CrossRef]
39.
Zaninello, M.; Palikaras, K.; Sotiriou, A.; Tavernarakis, N.; Scorrano, L. Sustained intracellular calcium rise mediates neuronal
mitophagy in models of autosomal dominant optic atrophy. Cell Death Differ. 2022,29, 167–177. [CrossRef]
40.
Cartes-Saavedra, B.; Lagos, D.; Macuada, J.; Arancibia, D.; Burte, F.; Sjoberg-Herrera, M.K.; Andres, M.E.; Horvath, R.; Yu-Wai-
Man, P.; Hajnoczky, G.; et al. OPA1 disease-causing mutants have domain-specific effects on mitochondrial ultrastructure and
fusion. Proc. Natl. Acad. Sci. USA 2023,120, e2207471120. [CrossRef]
41.
Quintana-Cabrera, R.; Manjarres-Raza, I.; Vicente-Gutierrez, C.; Corrado, M.; Bolanos, J.P.; Scorrano, L. Opa1 relies on cristae
preservation and ATP synthase to curtail reactive oxygen species accumulation in mitochondria. Redox Biol. 2021,41, 101944.
[CrossRef] [PubMed]
42.
Quintana-Cabrera, R.; Quirin, C.; Glytsou, C.; Corrado, M.; Urbani, A.; Pellattiero, A.; Calvo, E.; Vazquez, J.; Enriquez, J.A.; Gerle,
C.; et al. The cristae modulator Optic atrophy 1 requires mitochondrial ATP synthase oligomers to safeguard mitochondrial
function. Nat. Commun. 2018,9, 3399. [CrossRef] [PubMed]
43.
Amini, P.; Stojkov, D.; Felser, A.; Jackson, C.B.; Courage, C.; Schaller, A.; Gelman, L.; Soriano, M.E.; Nuoffer, J.M.; Scorrano, L.;
et al. Neutrophil extracellular trap formation requires OPA1-dependent glycolytic ATP production. Nat. Commun. 2018,9, 2958.
[CrossRef] [PubMed]
44.
Le Roux, B.; Lenaers, G.; Zanlonghi, X.; Amati-Bonneau, P.; Chabrun, F.; Foulonneau, T.; Caignard, A.; Leruez, S.; Gohier, P.;
Procaccio, V.; et al. OPA1: 516 unique variants and 831 patients registered in an updated centralized Variome database. Orphanet
J. Rare Dis. 2019,14, 214. [CrossRef]
45.
Yu-Wai-Man, P.; Griffiths, P.G.; Gorman, G.S.; Lourenco, C.M.; Wright, A.F.; Auer-Grumbach, M.; Toscano, A.; Musumeci, O.;
Valentino, M.L.; Caporali, L.; et al. Multi-system neurological disease is common in patients with OPA1 mutations. Brain 2010,
133, 771–786. [CrossRef]
46. Mellough, C.B.; Bauer, R.; Collin, J.; Dorgau, B.; Zerti, D.; Dolan, D.W.P.; Jones, C.M.; Izuogu, O.G.; Yu, M.; Hallam, D.; et al. An
integrated transcriptional analysis of the developing human retina. Development 2019,146, dev.169474. [CrossRef]
47.
Hoshino, A.; Ratnapriya, R.; Brooks, M.J.; Chaitankar, V.; Wilken, M.S.; Zhang, C.; Starostik, M.R.; Gieser, L.; La Torre, A.; Nishio,
M.; et al. Molecular Anatomy of the Developing Human Retina. Dev. Cell 2017,43, 763–779.e4. [CrossRef]
48. Hendrickson, A. Development of Retinal Layers in Prenatal Human Retina. Am. J. Ophthalmol. 2016,161, 29–35.e1. [CrossRef]
49.
Hahn, J.; Monavarfeshani, A.; Qiao, M.; Kao, A.; Kolsch, Y.; Kumar, A.; Kunze, V.P.; Rasys, A.M.; Richardson, R.; Baier, H.; et al.
Evolution of neuronal cell classes and types in the vertebrate retina. bioRxiv 2023. [CrossRef]
50.
Peng, Y.R.; Shekhar, K.; Yan, W.; Herrmann, D.; Sappington, A.; Bryman, G.S.; van Zyl, T.; Do, M.T.H.; Regev, A.; Sanes,
J.R. Molecular Classification and Comparative Taxonomics of Foveal and Peripheral Cells in Primate Retina. Cell 2019,176,
1222–1237.e22. [CrossRef]
51.
Conant, D.; Hsiau, T.; Rossi, N.; Oki, J.; Maures, T.; Waite, K.; Yang, J.; Joshi, S.; Kelso, R.; Holden, K.; et al. Inference of CRISPR
Edits from Sanger Trace Data. CRISPR J. 2022,5, 123–130. [CrossRef] [PubMed]
52.
Pohl, K.A.; Zhang, X.; Pham, A.H.; Chan, J.W.; Sadun, A.A.; Yang, X.J. Establishing induced pluripotent stem cell lines from
two dominant optic atrophy patients with distinct OPA1 mutations and clinical pathologies. Front. Genet. 2023,14, 1251216.
[CrossRef] [PubMed]
53.
Toombs, J.; Panther, L.; Ornelas, L.; Liu, C.; Gomez, E.; Martin-Ibanez, R.; Cox, S.R.; Ritchie, S.J.; Harris, S.E.; Taylor, A.; et al.
Generation of twenty four induced pluripotent stem cell lines from twenty four members of the Lothian Birth Cohort 1936. Stem
Cell Res. 2020,46, 101851. [CrossRef] [PubMed]
54.
Divakaruni, A.S.; Paradyse, A.; Ferrick, D.A.; Murphy, A.N.; Jastroch, M. Analysis and interpretation of microplate-based oxygen
consumption and pH data. Methods Enzymol. 2014,547, 309–354. [CrossRef]
55.
Divakaruni, A.S.; Wallace, M.; Buren, C.; Martyniuk, K.; Andreyev, A.Y.; Li, E.; Fields, J.A.; Cordes, T.; Reynolds, I.J.; Bloodgood,
B.L.; et al. Inhibition of the mitochondrial pyruvate carrier protects from excitotoxic neuronal death. J. Cell Biol. 2017,216,
1091–1105. [CrossRef]
56.
Ioannidis, N.M.; Rothstein, J.H.; Pejaver, V.; Middha, S.; McDonnell, S.K.; Baheti, S.; Musolf, A.; Li, Q.; Holzinger, E.; Karyadi, D.;
et al. REVEL: An Ensemble Method for Predicting the Pathogenicity of Rare Missense Variants. Am. J. Hum. Genet. 2016,99,
877–885. [CrossRef]
57.
Skarnes, W.C.; Pellegrino, E.; McDonough, J.A. Improving homology-directed repair efficiency in human stem cells. Methods 2019,
164–165, 18–28. [CrossRef]
58.
Akepati, V.R.; Muller, E.C.; Otto, A.; Strauss, H.M.; Portwich, M.; Alexander, C. Characterization of OPA1 isoforms isolated from
mouse tissues. J. Neurochem. 2008,106, 372–383. [CrossRef]
59.
Gu, W.; Gaeta, X.; Sahakyan, A.; Chan, A.B.; Hong, C.S.; Kim, R.; Braas, D.; Plath, K.; Lowry, W.E.; Christofk, H.R. Glycolytic
Metabolism Plays a Functional Role in Regulating Human Pluripotent Stem Cell State. Cell Stem Cell 2016,19, 476–490. [CrossRef]
Cells 2025,14, 137 20 of 20
60.
Sladen, P.E.; Perdigao, P.R.L.; Salsbury, G.; Novoselova, T.; van der Spuy, J.; Chapple, J.P.; Yu-Wai-Man, P.; Cheetham, M.E. CRISPR-
Cas9 correction of OPA1 c.1334G>A: P.R445H restores mitochondrial homeostasis in dominant optic atrophy patient-derived
iPSCs. Mol. Ther. Nucleic Acids 2021,26, 432–443. [CrossRef]
61.
Chen, J.; Riazifar, H.; Guan, M.X.; Huang, T. Modeling autosomal dominant optic atrophy using induced pluripotent stem cells
and identifying potential therapeutic targets. Stem Cell Res. Ther. 2016,7, 2. [CrossRef] [PubMed]
62.
Garcia-Lopez, M.; Jimenez-Vicente, L.; Gonzalez-Jabardo, R.; Dorado, H.; Gomez-Manjon, I.; Martin, M.A.; Ayuso, C.; Arenas, J.;
Gallardo, M.E. Creation of an Isogenic Human iPSC-Based RGC Model of Dominant Optic Atrophy Harboring the Pathogenic
Variant c.1861C>T (p.Gln621Ter) in the OPA1 Gene. Int. J. Mol. Sci. 2024,25, 7240. [CrossRef] [PubMed]
63.
Lei, Q.; Xiang, K.; Cheng, L.; Xiang, M. Human retinal organoids with an OPA1 mutation are defective in retinal ganglion cell
differentiation and function. Stem Cell Rep. 2024,19, 68–83. [CrossRef] [PubMed]
64.
Sladen, P.E.; Jovanovic, K.; Guarascio, R.; Ottaviani, D.; Salsbury, G.; Novoselova, T.; Chapple, J.P.; Yu-Wai-Man, P.; Cheetham,
M.E. Modelling autosomal dominant optic atrophy associated with OPA1 variants in iPSC-derived retinal ganglion cells. Hum.
Mol. Genet. 2022,31, 3478–3493. [CrossRef] [PubMed]
65.
Jonikas, M.; Madill, M.; Mathy, A.; Zekoll, T.; Zois, C.E.; Wigfield, S.; Kurzawa-Akanbi, M.; Browne, C.; Sims, D.; Chinnery,
P.F.; et al. Stem cell modeling of mitochondrial parkinsonism reveals key functions of OPA1. Ann. Neurol. 2018,83, 915–925.
[CrossRef]
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