Background
Parkinson’s disease (PD) is the second most common age-related neurodegenerative disorder, affecting more than 10 million people worldwide [
1]. Most patients develop the disease in a sporadic manner through a complex interaction between genetic and environmental risk factors during ageing. Roughly 5%–10% of PD patients are caused by highly penetrant variants in genes such as
PINK1 (encoding PTEN-induced putative kinase 1 [PINK1]) and
PARK2 (encoding the E3 ubiquitin ligase Parkin) [
2,
3]. This type of PD is referred to as familial PD, and missense variants of
VPS35 have been linked to the autosomal dominant form of familial PD [
4,
5]. However, the c.1858G > A, p.D620N variant in
VPS35 is the only proven pathogenic variant [
6].
VPS35 encodes the vacuolar protein sorting-associated protein 35 (VPS35) that, together with VPS26 and VPS29, forms the cargo-selective subcomplex of the retromer complex [
7]. The retromer recycles membrane proteins from endosomes to either the Golgi apparatus or the plasma membrane [
8]. The p.D620N variant is located in a domain of VPS35 that is essential for protein–protein interactions [
7]. Although the variant does not affect the formation of the retromer complex, it has impaired interactions with other factors such as the actin-nucleating WASH (Wiskott-Aldrich syndrome and SCAR homolog) complex [
9,
10]. This leads to the altered retromer functioning and deficits in the sorting of cargoes [
9‐
12].
Retromer also participates in the transport of mitochondrial cargoes to lysosomes or peroxisomes
via mitochondrial-derived vesicles (MDVs) [
13‐
15]. Previous reports have shown that VPS35 is involved in mitochondrial dynamics, as it recycles the fission protein DLP1 and regulates the level of the fusion protein MFN2 through the transport of mitochondrial E3 ubiquitin ligase 1 (MUL1) [
14,
15]. Overexpression of the VPS35 D620N mutant augments mitochondrial fragmentation due to the increased DLP1 activity, whereas VPS35 depletion leads to mitochondrial fragmentation as a result of decreased level of MFN2, which correlates with a reduced mitochondrial respiratory capacity and a decrease in mitochondrial membrane potential [
14‐
16].
Mitochondrial dysfunction plays an integral role in the pathogenesis of both sporadic and familial PD [
17‐
19]. For example, loss-of-function variants of mitochondrial quality control genes such as
PINK1 and
PARK2 lead to early-onset autosomal recessive PD [
2,
3,
20‐
22]. To maintain the mitochondrial quality, PINK1 is imported through a membrane potential–dependent process, from the outer mitochondrial membrane (OMM) into the inner mitochondrial membrane, where it is constitutively degraded by mitochondrial proteases [
23,
24]. However, PINK1 import and cleavage is blocked upon mitochondrial depolarization caused by damage, resulting in the accumulation of PINK1 on the OMM. At the OMM, PINK1 phosphorylates ubiquitin and Parkin, leading to stable recruitment and activation of Parkin onto the mitochondrial surface [
21,
24,
25]. Parkin then ubiquitinates different OMM substrates, inducing proteasomal degradation and removal of damaged cargoes
via the MDVs-to-lysosome transport and/or mitophagy [
26‐
28].
Mitophagy is a selective type of autophagy in which mitochondria targeted for degradation are sequestered into double-membrane autophagosomes and delivered into lysosomes [
29,
30]. This process occurs in different physiological contexts [
30]. For instance, most cells continuously undergo basal mitophagy during routine mitochondrial maintenance [
31]. However, mitophagy can also be induced as a response to mitochondrial stressors such as mitochondrial depolarization. Notably, the PD-associated proteins PINK1 and Parkin are directly involved in stress-induced mitophagy [
21,
24] but not in basal mitophagy [
32,
33]. As dopaminergic neurons undergo substantial mitochondrial stress, presumably due to their pacemaker activity [
34,
35], the stress-induced mitophagy
via PINK1/Parkin has been heavily implicated in the pathogenesis of PD [
30].
Given the mitochondrial impairments associated with the p.D620N variant of VPS35 and the role of PINK1 and Parkin in maintaining mitochondrial quality control under stress conditions, we questioned whether the actions of these genes converge into a similar pathway to cause PD. Therefore, we set out to determine whether stress-induced mitophagy via PINK1/Parkin is affected by the VPS35 p.D620N mutant, using VPS35 mutant SH-SY5Y cells carrying the p.D620N variant on one allele, which recapitulates the patient situation.
Materials and methods
Cell culture, transient transfections and treatments
Human SH-SY5Y neuroblastoma cells were maintained in Dulbecco’s Modified Eagle’s Medium (Invitrogen, Waltham, MA) supplemented with 15% fetal bovine serum (Invitrogen) and 1% Penicillin-Streptomycin (Gibco, Waltham, MA) in a 37 °C incubator with 5% CO2. Transient plasmid transfections were performed with plasmid DNAs using Lipofectamine (Thermo Fischer Scientific, Waltham, MA), according to the manufacturer’s instructions. To induce mitochondrial depolarization, the SH-SY5Y cells were treated with 10 μM or 20 μM carbonyl cyanide m-chlorophenylhydrazone (CCCP) (Sigma-Aldrich, Saint Louis, MO), 1 μM oligomycin (Sigma-Aldrich), 1 μM antimycin A (Sigma-Aldrich), or 1 μM antimycin A and 1 μM oligomycin (AO), for the indicated times, prior to cell harvesting or fixation. DMSO treatment was used as a control.
Expression plasmids and antibodies
The plasmids used were pEGFP-Parkin [
36] (a gift from Prof. Edward Fon (McGill University, Montreal, Quebec, Canada), Addgene plasmid #45875) and pEGFP-LC3 (a gift from Prof. Toren Finkel (University of Pittsburgh, Pittsburgh, PA), Addgene plasmid #24920) constructs [
37]. The primary antibodies used for immunoblotting were mouse anti-ATPIF1 (1:1000, Abcam, Cambridge, UK, ab110277), mouse anti-β-actin (1:5000, MP Biomedicals, Irvine, CA, 8691001), mouse anti-β-tubulin (1:5000, Sigma-Aldrich T4026), mouse anti-Parkin (1:500, Santa-Cruz Biotechnology, Dallas, TX; sc-32,282), rabbit anti-PINK1 (1:1000, Cell signaling, Danvers, MA, #6946), mouse anti-TOM20 (1:500, BD Biosciences, San Jose, CA; 612278) and goat anti-VPS35 (1:1000, Abcam, ab10099). The primary antibodies used for immunofluorescence (IF) were mouse anti-TOM20 (1:200, Santa-Cruz Biotechnology sc-17764) and rabbit anti-PINK1 (1:200, Abcam ab216144). Secondary antibodies for immunoblotting were HRP-conjugated goat anti-rabbit IgG (H + L) (1:10000, Bio-Rad, Hercules, CA), HRP-conjugated goat anti-mouse IgG (H + L) (1:10000, Bio-Rad) and HRP-conjugated donkey anti-goat IgG (H + L) (1:10000, Abcam). Secondary antibodies for IF were Cy3-conjugated donkey anti-mouse IgG (H + L) (1:250, Jackson ImmunoResearch, West Grove, PA) and Alexa Fluor 488-conjugated donkey anti-rabbit IgG (H + L) (1:250, Jackson ImmunoResearch).
Generation of VPS35 D620N/wild-type (WT) SH-SY5Y cells
The D620N mutation in the
VPS35 gene was obtained by Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)-Cas9–mediated genome editing in the SH-SY5Y neuroblastoma cell line, as previously described [
38]. Briefly, a 20-nt single guide RNA (sgRNA) sequence that targets exon 15 of the
VPS35 gene and is predicted to cut approximately 9 base-pairs (bp) upstream of the GAT triplet encoding the aspartic acid residue on location 620 was cloned into the pSpCas9(BB)-2A-GFP (PX458) plasmid (a gift from Prof. Feng Zhang (Broad Institute, Cambridge, MA); Addgene plasmid #48138) using the
BbsI restriction enzyme to form the targeting plasmid expressing Cas9-GFP. In addition, a single-stranded oligodeoxynucleotide sequence was designed to facilitate homology-directed repair of the endogenous locus and included the substitution of five nucleotides: a nucleotide substitution G > A that leads to the D620N mutation of
VPS35 and four synonymous substitutions that create a novel
EcoRI restriction site that also destroys the protospacer-adjacent motif sequence to avoid repetitive cutting of Cas9 by the repair template. Following validation, the PX458-sgRNA plasmid and the single-stranded oligonucleotides were transfected into the SH-SY5Y cells following the manufacturer’s protocol (Lonza, Basel, Switzerland). GFP-positive cells were single-cell sorted 48 h post-transfection using a SH800S cell sorter (Sony Biotechnology, San Jose, CA) and grown in separate cultures that were subsequently screened for the D620N mutation using the restriction enzyme
EcoRI. In parallel, we mock-electroporated and sorted the same batch of cells, which were used as WT control in the following experiments. Finally, we sequenced the top three predicted off-target genomic regions within coding regions (obtained from
http://crispr.mit.edu) of genes
POU6F1, ZNF318 and
KY, but found no off-target edits (not shown). Detailed primer and template sequences are provided in Table S1.
Generation of stable COX8-EGFP-mCherry reporter SH-SY5Y cells
The COX8-EGFP-mCherry sequence was obtained from the pCLBW COX8-EGFP-mCherry construct [
39] (a gift from Prof. David Chan (Caltech, Pasadena, CA), Addgene plasmid #78520) through restriction enzyme digestion with
ApaI and
EcoRI, and was ligated into the mammalian expression vector pcDNA 3.1(+). Subsequently, the vector was transfected into WT and VPS35
D620N SH-SY5Y cells using Lipofectamine 3000 (Thermo Fischer Scientific), following the manufacturer’s protocol. Forty-eight hours after transfection, the growth medium was replaced with selection medium containing 800 ng/μl G-418 (Sigma-Aldrich). The selection medium was refreshed every other day for 10 days until only cells with the plasmid remained. Stable cell lines were cultured for three passages before performing the experiments.
Protein extraction and immunoblotting
SH-SY5Y cells were harvested in 2% sodium dodecyl sulfate (SDS)/phosphate-buffered saline (PBS) buffer containing a proteinase inhibitor cocktail (Roche, Basel, Switzerland) and sonicated. Crude mitochondrial fractions were isolated as previously described [
40]. Briefly, SH-SY5Y cells were collected and homogenized using a Dounce homogenizer in ice-cold isolation buffer containing 320 mM sucrose and a proteinase inhibitor cocktail. The homogenized samples were differentially centrifuged at 1500 g for 15 min and 17,000 g for 30 min to obtain nuclei and crude mitochondria, respectively. The cytosolic fraction was obtained from the final supernatant. Protein concentrations were quantified using the Pierce™ BCA protein assay kit (Thermo Fischer Scientific), and samples were mixed with loading buffer containing 10% β-mercaptoethanol before being boiled at 95 °C for 5 min. Subsequently, equal amounts of total protein extracts were subjected to SDS-PAGE, transferred to nitrocellulose membranes, blocked for 1 h in skimmed milk, incubated overnight with primary antibody at 4 °C and then with the corresponding secondary antibody for 1 h at room temperature (RT). The blots were imaged on a Chemidoc™ MP Imaging System (Bio-Rad). Protein levels were quantified by densitometry using the ImageJ software (NIH, Bethesda, MD).
Immunofluorescence
WT and VPS35D620N SH-SY5Y cells that were seeded on glass coverslips in 24-well plates were fixed in 4% paraformaldehyde in PBS for 10 min at RT. Cells were then permeabilized in 0.1% Triton X-100 in PBS for 10 min and blocked with 5% donkey serum (Abcam) in PBS for 1 h. The coverslips were then incubated overnight at 4 °C with the primary antibodies diluted in blocking buffer and for 1 h at RT for secondary antibody incubation. Coverslips were finally mounted onto glass slides in 4′,6-diamidino-2-phenylindole (DAPI)-containing mounting medium (Vector Laboratories, Burlingame, CA). The slides were analyzed using either structured illumination microscopy (SIM) or confocal microscopy. SIM images were acquired with an AxioObserver Z1 compound microscope (Carl Zeiss, Oberkochen, Germany) equipped with an Apotome, 63x oil objective and an AxioCam MRm3 CCD camera (Carl Zeiss). Confocal images were acquired with a TCS SP8 high-resolution confocal laser scan microscope (Leica Microsystems, Wetzlar, Germany) and an HC PL APO CS2 63x/1.4 oil objective. For quantitative analysis, maximum intensity projections were generated from all Z-stacks, which were captured for each condition with identical exposure times or laser settings.
Image analysis
All image analyses were performed using the ImageJ software (NIH). Colocalization analyses of PINK1 and TOM20 were performed using ImageJ plugin Coloc 2 (
https://imagej.net/Coloc_2). Regions of interest (ROIs) were created per cell in the TOM20 channel (
n = ~ 80–100 cells per condition in each experiment). Pearson’s correlation coefficients were subsequently determined per ROI using the Costes method for threshold regression [
41].
Mitochondrial morphology was quantified as previously described [
42]. Briefly, images of single cells were pre-processed and binarized, followed by particle analysis and computation of several metrics. The number of mitochondria was determined as the number of individual particles. The aspect ratio was determined by dividing the major axis by the minor axis of each particle. A total of 80–100 cells were quantified per condition in each experiment.
For the EGFP-Parkin translocation experiment, a blinded observer scored each cell for either diffuse EGFP-Parkin or mitochondria-localized EGFP-Parkin (
n = ~ 50 cells per condition in each experiment), as previously described [
36].
Mitophagy in COX8-EGFP-mCherry stable cell lines was quantified by determining the ratio of the number of particles obtained from the mCherry channel (mitophagolysosomes) to the number of particles obtained from the EGFP channel (mitochondria) per cell (n = ~ 50–70 cells per condition in each experiment). Particles were analyzed in a similar fashion to the mitochondrial morphology quantification.
EGFP-LC3 puncta on mitochondria were quantified as follows: a mask was created from the TOM20 mitochondrial staining and used as overlay over the EGFP-LC3 image. The puncta were subsequently counted for each cell (n = ~ 40–50 cells per experiment).
Mitochondrial membrane potential quantification
Mitochondrial membrane potential was measured using fluorescence-activated cell sorting (FACS). WT and VPS35D620N SH-SY5Y cells were incubated for 30 min with 100 nM tetramethylrhodamine methyl ester (TMRM) dye (Thermo Fischer Scientific) and 100 nM MitoTracker Green FM dye (Thermo Fischer Scientific) diluted in culture medium. Cells were rinsed, dissociated with 0.05% Trypsin-EDTA (Thermo Fischer Scientific) and aliquoted in multiple FACS tubes. FACS measurements were performed with a FACSCalibur flow cytometer (BD Biosciences) or a Novocyte Quanteon flow cytometer (Agilent, Santa Clara, CA ). For the timeline measurements, baseline measurements were taken, after which CCCP was added to a final concentration of 10 μM, upon which measurements were taken at each time point. For the dose-response measurements, CCCP was added to the indicated final concentrations, and 1 min later measurements were made. Slopes were determined using a simple linear regression method. For 24-h treatments, cells were treated with 10 μM CCCP or 1 μM antimycin A and 1 μM oligomycin for 24 h, then the cells were dissociated and TMRM was measured as indicated above. Three independent sorts measuring at least 10,000 cells were performed per clone per data point for all experiments. Data analysis was performed using the Kaluza Analysis software (Beckman Coulter, Brea, CA). Mitotracker Green median fluorescence intensity was used to correct for mitochondrial mass fluctuations.
Ultrastructural analyses
For conventional transmission electron microscopy (TEM), WT and VPS35
D620N SH-SY5Y cells were treated with DMSO or 10 μM CCCP for 6 h. Then an equal volume as the culture media of double-strength fixative (4% paraformaldehyde, 5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4) was then added to the cells and incubated for 20 min at RT, followed by further fixing the cells with the same volume of single-strength fixative (2% paraformaldehyde, 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4) for 2 h at RT. After five washes with 0.1 M sodium cacodylate buffer (pH 7.4), the cells were scraped and embedded as previously described [
43]. Subsequently, 70-nm ultrathin sections were cut using a Leica EM UC7 ultra microtome (Leica Microsystems) and stained with uranyl acetate and lead citrate as previously described [
43]. The cell sections were analyzed using an 80 kV transmission electron microscope CM100bio TEM (FEI, Eindhoven, The Netherlands).
The analysis of the different mitochondrial profiles per cell type was performed by random screening of sections derived from at least three different grids per sample. The mitochondrial profiles were categorized as follows: classical mitochondria with well-defined cristae (category I), dark mitochondria with well-defined cristae often swelling (category II), mitochondria with undefined cristae (category III), dark mitochondria with undefined contours and cristae (category IV), and large mitochondria with very light content and few remnant cristae (category V). The number of each mitochondrial type per condition was determined by counting 665, 579 and 521 mitochondria profiles from the DMSO-treated WT and VPS35D620N cell (clones 1 and 2) sections, respectively, as well as 727, 1028 and 914 mitochondria profiles from the CCCP-treated WT and VPS35D620N cell (clones 1 and 2) sections, respectively.
Cell viability assay
Cell viability upon treatment with CCCP was determined using a 3-(4,5-demethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) reduction assay (Abcam). The SH-SY5Y cells were plated in 96-well plates 12 h prior to incubation with CCCP for 24 h. MTT assay compounds were added following the manufacturer’s protocol and absorbance was measured using a Synergy HT optical plate reader (Biotek, Winooski, VT).
Statistical analyses
Data of Western blot densitometry measurements, mitochondrial membrane potential measurements, TEM and mitochondrial morphology were analyzed using a linear model by one-way or two-way analysis of variance (ANOVA) followed by Tukey’s post-hoc test. Count data, such as the EGFP-LC3 and mitochondrial particle quantification data were modelled using a generalized linear model followed by one-way or two-way ANOVA and Tukey’s post-hoc test. PINK1-TOM20 colocalization data were analyzed using the Kruskal-Wallis test followed by pairwise Mann-Whitney U-test with Benjamini-Hochberg multiple testing correction. Proportional data of COX8-EGFP-mCherry mitophagy and EGFP-Parkin localization were analyzed using beta regression analysis. Data are means ± SEM from at least three independent experiments, unless otherwise specified. P < 0.05 was considered as statistically significant. Statistical analyses were performed in the statistical computing environment R (version 1.3.959; The R Foundation for Statistical Computing; Vienna, Austria).
Discussion
In the present study, we show for the first time that the actions of VPS35 converge on the PINK1/Parkin pathway and that the VPS35D620N cells show deficits in CCCP-induced PINK1/Parkin-mediated mitophagy. Importantly, these data were acquired using a model that closely mimics the situation in PD patients. The mitochondria of VPS35D620N cells seemed desensitized to a CCCP-induced Δψm collapse, as they appeared already damaged/fragmented and had a reduced mitochondrial membrane potential at steady state. Consequently, the mitochondria of CCCP-treated VPS35D620N cells showed almost no accumulation of PINK1 and Parkin, and therefore failed to initiate mitophagy. However, PINK1/Parkin-dependent mitophagy in VPS35D620N cells was still operational, as the VPS35D620N cells displayed PINK1/Parkin-mediated mitophagy upon AO treatment. The results suggest that the mitochondria of VPS35D620N already exhibit a specific type of damage at steady state. This renders them insensitive to CCCP and likely also to other stressors that may initiate PINK1/Parkin-mediated mitophagy in humans. We speculate that individuals carrying the p.D620N variant of VPS35 tend to accumulate damaged mitochondria because of this impairment, and, over time, this could cause neurodegeneration.
The observed failure of VPS35
D620N cells to maintain Δψ
m under steady state is likely linked to the presence of damaged mitochondria and will have deleterious effects on cell viability and functions, as Δψ
m provides the driving force for ATP synthesis [
53]. Maintenance of Δψ
m is important for the inward transport of cations such as Ca
2+ [
54] and is necessary for the import of numerous mitochondrial proteins [
55,
56]. Mitochondrial quality control mechanisms that maintain Δψ
m, such as mitochondrial fragmentation [
57] and the removal of depolarized mitochondria through mitophagy [
29,
58], are thus essential and are likely affected in VPS35
D620N cells, which leads to the observed accumulation of damaged and fragmented mitochondria under steady state conditions. Depletion of VPS35 in neuroblastoma cells also causes reduced basal Δψ
m and an increase in mitochondrial fission at steady state [
15]. Notably, defects in the maintenance of Δψ
m and mitochondrial dynamics have been observed in other models of PD, including those genetically modified for PINK1 and Parkin [
59,
60]. Importantly, while this manuscript was in preparation, a study reported that patient-derived p.D620N-mutant VPS35 dopaminergic neurons exhibit a reduction in Δψ
m at steady state, and show a lysosomal-associated defect in CCCP-induced mitochondrial clearance [
61]. While we were unable to test for mitochondrial clearance in CCCP-treated VPS35
D620N cells, we did not observe an evident impairment in this pathway in VPS35
D620N cells upon AO treatment. However, we cannot exclude that mitochondrial clearance is not affected. A mitophagy impairment downstream of mitophagy induction may therefore contribute to the accumulation of damaged mitochondria at steady state.
Our data also suggest that the VPS35
D620N cells are less able to respond to a collapse in Δψ
m, as indicated by reduced PINK1/Parkin-mediated mitophagy upon Δψ
m loss due to CCCP treatment. The fact that the AO treatment was able to induce PINK1/Parkin-mediated mitophagy in VPS35
D620N cells, while CCCP could not, can be explained by the difference in how the compounds affect mitochondrial depolarization. CCCP can dissipate Δψ
m by removing the proton gradient over the mitochondrial membrane through increasing the permeability of protons across the inner mitochondrial membrane. As such, the duration and the amount of CCCP dictate the extent of Δψ
m loss, and consequently the amount of PINK1 stabilization and accumulation. Antimycin A and oligomycin both block the function of the mitochondrial electron transport chain that actively maintains the Δψ
m, causing a loss of Δψ
m. Oligomycin also blocks the reverse ATP synthase activity of the F
1F
0 ATPase, which is normally utilized to counteract the loss of Δψ
m by actively pumping protons into the intermembrane space, thereby causing a further decrease in Δψ
m [
51,
62]. In addition, it is known that the inhibition of electron transport chain subcomplex III by antimycin A can cause an increase in oxidative stress through the production of reactive oxygen species [
48]. Importantly and in line with our data, AO has been previously shown to cause less mitochondrial membrane depolarization compared to CCCP [
50]. Therefore, the robust PINK1 accumulation in response to AO treatment is not solely due to the dissipation of Δψ
m, which rather probably works in conjunction with increased oxidative stress to form a more substantial mitophagy stimulus than CCCP. Notably, a recent study has shown that the AO-induced PINK1 accumulation can be inhibited by antioxidants [
63]. This could explain why the VPS35
D620N cells accumulated PINK1 when treated with AO, but not when exposed to CCCP. Furthermore, the increased ATPIF1 levels in VPS35
D620N cells probably contributed to the diminished Δψ
m dissipation in response to CCCP as the F
1F
0 ATPase was more inhibited. Thus, it is likely that in VPS35
D620N cells, which already have a lower Δψ
m at steady state conditions, the Δψ
m collapses induced by CCCP are too small to provoke additional mitochondrial fragmentation and thereby induce PINK1/Parkin-mediated mitophagy, while this is not the case with AO treatment due to the different mode of PINK1 recruitment.
So why was PINK1/Parkin-mediated mitophagy impaired in VPS35D620N cells upon CCCP treatment? Since VPS35D620N cells already display loss of Δψm at steady state, PINK1/Parkin-mediated mitophagy may not be activated upon mild stress, which prevents continuous turnover of mildly damaged mitochondria. As these damaged mitochondria are chronically stressed at steady state, we hypothesize that they may be desensitized by a yet unknown regulatory feedback loop or factor to prevent depletion of the mitochondrial population. Although we did not elucidate how VPS35 modifies PINK1/Parkin-mediated mitophagy, our data advocate that this impairment is associated with the already damaged/fragmented mitochondria with lower Δψm in VPS35D620N cells at steady state.
Furthermore, as our study focused only on PINK1/Parkin-mediated mitophagy, we cannot rule out that other forms of mitophagy are affected in VPS35
D620N cells. Of note, multiple studies have shown that mitophagy can be induced with CCCP
via other ubiquitin E3 ligases, independent of Parkin [
64,
65]. Moreover, receptor-mediated mitophagy can occur without ubiquitin E3 ligases through direct interactions of autophagic receptors present on the OMM with LC3, thereby circumventing the PINK1/Parkin pathway [
66,
67]. Future studies are necessary to investigate whether these alternative routes of mitophagy induction are also affected in VPS35
D620N cells.
One established role of VPS35 and retromer in mitochondrial physiology is to retrieve mitochondrial proteins
via MDVs [
13], and it has been shown that the D620N mutation of
VPS35 affects the sorting of MUL1 and DLP1 [
14,
16]. With the expanding research on the proteome of MDVs [
68], it is likely that the retromer is involved in trafficking of additional mitochondrial proteins. It is unclear to what extent this is regulated by retromer and, importantly, which cargoes are being transported. However, the sorting of other mitochondrial proteins is probably affected and could cause mitochondrial impairments, e.g. damage/fragmentation, in VPS35
D620N cells. For example, Δψ
m loss in the VPS35
D620N cells at steady state might be caused by changes in regulatory proteins involved in maintaining Δψ
m, including ATPIF1, and other proteins such as the components of the mitochondrial permeability transition pore complex and the oxidative phosphorylation machinery [
69,
70]. Notably, ATPIF1 has also been found enriched in MDVs [
68]. Whether VPS35 plays an active role in the regulation of ATPIF1 has to be determined in future studies. In addition, a recent study has shown that VPS35 interacts with Parkin, suggesting that members of the PINK1/Parkin pathway are directly affected by VPS35-mediated MDV trafficking [
71]. Although the role of VPS35 in MDV transport has been established [
13,
14], the field of MDV-mediated transport is still emerging and many questions remain regarding their role in mitochondrial quality control and regulation of different cargoes [
72].