Mitochondrial Function in Hereditary Spastic Paraplegia: Deficits in SPG7 but Not SPAST Patient-Derived Stem Cells

Wali, Gautam; Kumar, Kishore Raj; Liyanage, Erandhi; Davis, Ryan L.; Mackay-Sim, Alan; Sue, Carolyn M.

doi:10.3389/fnins.2020.00820

BRIEF RESEARCH REPORT article

Front. Neurosci., 20 August 2020

Sec. Neurodegeneration

Volume 14 - 2020 | https://doi.org/10.3389/fnins.2020.00820

Mitochondrial Function in Hereditary Spastic Paraplegia: Deficits in SPG7 but Not SPAST Patient-Derived Stem Cells

$\r\nGautam Wali,*&#x;$ Gautam Wali^1,2*†

Kishore Raj Kumar^1,2,3,4,5

Erandhi Liyanage¹

Ryan L. Davis^1,2,4

Alan Mackay-Sim^2,6†

Carolyn M. Sue^1,2,5*†

¹Department of Neurogenetics, Royal North Shore Hospital, Kolling Institute of Medical Research, The University of Sydney, Sydney, NSW, Australia
²Faculty of Medicine and Health, Sydney Medical School, The University of Sydney, Sydney, NSW, Australia
³Molecular Medicine Laboratory, Department of Neurology, Concord Hospital, Sydney, NSW, Australia
⁴Kinghorn Centre for Clinical Genomics, Garvan Institute of Medical Research, Sydney, NSW, Australia
⁵Department of Neurology, Royal North Shore Hospital, Northern Sydney Local Health District, Sydney, NSW, Australia
⁶Griffith Institute for Drug Discovery, Griffith University, Brisbane, QLD, Australia

Mutations in SPG7 and SPAST are common causes of hereditary spastic paraplegia (HSP). While some SPG7 mutations cause paraplegin deficiency, other SPG7 mutations cause increased paraplegin expression. Mitochondrial function has been studied in models that are paraplegin-deficient (human, mouse, and Drosophila models with large exonic deletions, null mutations, or knockout models) but not in models of mutations that express paraplegin. Here, we evaluated mitochondrial function in olfactory neurosphere-derived cells, derived from patients with a variety of SPG7 mutations that express paraplegin and compared them to cells derived from healthy controls and HSP patients with SPAST mutations, as a disease control. We quantified paraplegin expression and an extensive range of mitochondrial morphology measures (fragmentation, interconnectivity, and mass), mitochondrial function measures (membrane potential, oxidative phosphorylation, and oxidative stress), and cell proliferation. Compared to control cells, SPG7 patient cells had increased paraplegin expression, fragmented mitochondria with low interconnectivity, reduced mitochondrial mass, decreased mitochondrial membrane potential, reduced oxidative phosphorylation, reduced ATP content, increased mitochondrial oxidative stress, and reduced cellular proliferation. Mitochondrial dysfunction was specific to SPG7 patient cells and not present in SPAST patient cells, which displayed mitochondrial functions similar to control cells. The mitochondrial dysfunction observed here in SPG7 patient cells that express paraplegin was similar to the dysfunction reported in cell models without paraplegin expression. The p.A510V mutation was common to all patients and was the likely species associated with increased expression, albeit seemingly non-functional. The lack of a mitochondrial phenotype in SPAST patient cells indicates genotype-specific mechanisms of disease in these HSP patients.

Introduction

Hereditary spastic paraplegia (HSP) is an inherited, progressive neurodegenerative condition causing spasticity and weakness in the lower limbs as a consequence of corticospinal tract degeneration (Hedera et al., 2005; Salinas et al., 2008; Kumar et al., 2015). Mutations in SPG7 and SPAST are the most common causes of HSP. SPG7 encodes for paraplegin, a mitochondrial matrix protease embedded in the inner mitochondrial membrane (Casari et al., 1998). Paraplegin is involved in multiple mitochondrial processes including mitochondrial protein quality surveillance, ribosome assembly, and mitochondrial biogenesis (Quirós et al., 2015). SPG7 mutations can cause loss of paraplegin (Atorino et al., 2003) or increased paraplegin expression (Thal et al., 2015; Pfeffer et al., 2014). The effects of SPG7 mutations leading to a loss of paraplegin expression on mitochondrial function have been characterized. For example, fibroblasts derived from a patient carrying a homozygous 9.5-kb deletion within the SPG7 gene that caused paraplegin deficiency exhibited impaired respiratory chain complex I activity. When the cells were exposed to hydrogen peroxide, to stress mitochondrial function, they were found to have lower ATP synthesis rate and reduced cell viability (Atorino et al., 2003). Consistent with this, a SPG7-null mutant Drosophila model displayed reduced activity of respiratory chain complexes I and II, severely swollen and aberrantly shaped mitochondria in the synaptic terminals of photoreceptors, and neuronal and muscular degeneration (Pareek et al., 2018).

There is limited information on mitochondrial function in patient cells and tissues with SPG7 mutations that do not cause a loss of paraplegin. Post-mortem analysis of a patient with homozygous p.Ala510Val mutations showed an increased expression of paraplegin in neurites of the cerebellar and cerebral cortex (Thal et al., 2015), but mitochondria function was not evaluated. In SPG7 patients carrying compound heterozygous mutations with increased paraplegin expression, the skeletal muscle biopsies showed increased mitochondrial biogenesis, multiple mitochondrial DNA deletions, and cytochrome c oxidase-deficient fibers (Pfeffer et al., 2014). Mitochondrial morphology evaluation of fibroblasts from these patients showed an increased mitochondrial mass and an increased number and length of mitochondrial networks; however, mitochondrial respiratory chain function was not evaluated. Inhibition of the respiratory chain function can lead to excess generation of reactive oxygen species, subsequently leading to cell death mediated by oxidative stress (Keane et al., 2011). This link has been shown for many neurodegenerative diseases (Moreira et al., 2010; Keane et al., 2011). Here, we characterize mitochondria morphology, respiratory chain function, and oxidative stress in paraplegin expressing SPG7 patient cells.

SPAST encodes for spastin, a microtubule-severing protein (Errico et al., 2002; Orlacchio et al., 2004). SPAST patient cells have impaired organelle transport, which includes mitochondria (Abrahamsen et al., 2013; Wali et al., 2020). However, it remains to be determined whether mitochondrial respiratory chain function is affected in SPAST patient cells. A study investigating muscle biopsies from SPAST patients did not show any aberrant activity of complexes I, II, and IV or citrate synthase (McDermott et al., 2003). Histochemical reactions, cytochrome c oxidase, and succinate dehydrogenase were also normal (McDermott et al., 2003). However, a recent study in a single SPAST patient fibroblast cell line showed that the patient cells had less tubular and more fragmented mitochondria, reduced mitochondrial membrane potential, and altered mitochondrial distribution (Dong et al., 2018).

In the present study, we quantified paraplegin expression and a range of mitochondrial morphology measures (fragmentation, interconnectivity, and mass), mitochondrial functions (membrane potential, oxidative phosphorylation, and oxidative stress), and cell proliferation. These measures were performed on olfactory neurosphere-derived stem cells (ONS cells) derived from HSP patients with SPG7 mutations, from healthy controls, and from HSP patients with SPAST mutations.

Materials and Methods

Participants

SPG7 and SPAST HSP patients involved in this study were examined by neurologists CMS and KRK. Details of the patient clinical characteristics and causative gene mutations are listed in Table 1. Control and patient samples used were matched for age and gender. Mean age range of SPG7 patients was 60 years, and those of SPAST patients and healthy controls were 51 and 60 years, respectively. Olfactory neurosphere-derived cells (ONS cells) were prepared from olfactory mucosa biopsies of participants (Féron et al., 1998, 2013) with their informed and written consent.

TABLE 1

Table 1. Clinical features and mutation details of HSP patients with SPG7 and SPAST mutations.

Targeted Next-Generation Sequencing

All patients except patient 3 have been previously reported (refer to Table 1). Patient 3 was identified using targeted next-generation sequencing, as described elsewhere (Kumar et al., 2013). In brief, patients with an apparent autosomal recessive or sporadic mode of inheritance were screened for nine autosomal recessive HSP-associated genes using a PCR-based library preparation and Roche Junior 454 next-generation sequencing, after exclusion of SPAST mutations. Amplicons that did not achieve 30 reads were resequenced with a conventional Sanger sequencing method. All pathogenic or potentially pathogenic variants were confirmed by Sanger sequencing on an ABI Prism 3130xl Genetic Analyzer platform (Applied Biosystems).

Cell Culture

ONS cells were derived from olfactory mucosa biopsies using our published techniques (Matigian et al., 2010; Féron et al., 2013). Cells were cultured in Dulbecco’s Modified Eagle Medium (DMEM)/F12 (Gibco) with 10% fetal bovine serum at 37°C and 5% CO₂. Please refer to our previous publication (Féron et al., 2013) for a detailed protocol of generating olfactory neurosphere-derived cells from olfactory mucosal biopsies.

Paraplegin Expression

Protein Isolation

Cells cultured in T-25 flasks were washed three times with Dulbecco’s phosphate-buffered saline (Gibco 14190250) to completely remove culture media. Cells were lysed with cell lysis buffer (Sigma, C3228) supplemented with a protease inhibitor (Thermo Fisher Scientific, 78429). Cells were harvested by scraping and maintained on ice over a rocker for 30 min. Then, the samples were centrifuged at 10,000 g for 10 min at 4°C. The protein samples were then collected and quantified using the BCA pierce TM BCA Protein Assay kit (Thermo Fisher Scientific, 23225).

Western Blot

Total cellular protein (20 μg) was resolved on a 12% Bis-Tris polyacrylamide gel and transferred onto a nitrocellulose membrane. The membrane blot was incubated overnight at 4°C with the primary mouse anti-human SPG7 monoclonal antibody (dilution 1/10,000; OriGene). Following this, the membrane was incubated with goat anti-mouse secondary antibody coupled to horseradish peroxidase (dilution 1/4,000, Sigma-Aldrich). Bands were visualized using Pierce SuperSignal West Femto Chemiluminescent Substrate on a Bio-Rad ChemiDoc XL. Paraplegin expression was normalized to GAPDH (dilution 1/10,000; Abcam) expression levels using ImageJ Version 1.52u software (Schneider et al., 2012).

Western Blot Positive Control

Human HEL293 cells overexpressing paraplegin were used to validate the antibody (OriGene, LC401085). The paraplegin band size was 88 kDa for both the control samples tested and the positive control, showing antibody specificity (Supplementary Figure S1).

Flow Cytometry

Cells were fixed and permeabilized with the BD Cytofix/Cytoperm reagent for 20 min at room temperature. Cells were washed with BD Perm/Wash buffer and then incubated with mouse anti-human SPG7 monoclonal antibody (dilution 1/10,000; OriGene) or the IgG1 isotype control for 1 h. Cells were washed and incubated with Alexa Fluor secondary antibody for 30 min. Cells were washed, and the relative fluorescence intensity was measured for 10,000 events per sample on a BD LSRFortessa, and the data were analyzed using BD FACS Diva software. Mean fluorescence intensity was calculated as the fluorescence intensity of the paraplegin antibody to the intensity of the isotype control antibody.

Mitochondrial Morphology

Cells were treated with 100 nM MitoTracker green for 30 min at 37°C to label active mitochondria (Park et al., 2014). Images of mitochondria-labeled cells were captured using a 63× objective on a confocal microscope. Using ImageJ image analysis software, raw images were binarized, and mitochondrial morphological characteristics of the aspect ratio (AR), indicating mitochondrial length, form factor (FF), and mitochondrial branching, were calculated as described before (Mortiboys et al., 2008). AR is the ratio of the major to minor axis of the mitochondria, and FF is calculated as (perimeter of mitochondria²)/(4π ^∗ area of mitochondria) (Mortiboys et al., 2008). Thirty cells were imaged and analyzed per cell line per technical replicate experiment.

Mitochondrial Mass and Mitochondrial Membrane Potential

Mitochondrial mass and membrane potential were assessed by labeling mitochondria with 100 nM MitoTracker green for 30 min (as above) and 50 nM non-quenching TMRM for 30 min at 37°C, respectively. The cells were washed twice with PBS before measuring fluorescence using a BD LSRFortessa^™. Fluorescence was measured for 10,000 events per sample, and the data were analyzed using the BD FACS Diva^™ software (Gautier et al., 2012). Mitochondrial membrane potential was normalized to MitoTracker green fluorescence, a relative estimate of the mitochondrial mass per cell.

MitoSOX and CM-H₂DCFDA

ONS cells (10,000) were plated in each well of a black-walled 96-well plate and cultured for 24 h before being assessed. The cells were stained with 5 μM MitoSOX Red for 15 min (as an indication of mitochondrial oxidative stress) (Teves et al., 2018) or 5 μM CM-H₂DCFDA for 15 min (a measure of general cellular oxidative stress) (Wali et al., 2016) at 37°C. The cells were washed twice with PBS before fluorescence was measured using a VICTOR Nivo Multimode Microplate Reader. MitoSOX and CM-H₂DCFDA fluorescence levels were normalized to mitochondrial mass, using dual MitoTracker green staining.

ATPlite Assay Measuring ATP Content

ONS cells (10,000 per well) were plated in a white-walled 96-well plate for 24 h before being assessed. The assay was performed as per the manufacturer’s instructions and as described before (Wali et al., 2016). Luminescence was measured using the VICTOR Nivo Multimode Microplate Reader. The ATPlite luminescence values were normalized to mitochondrial mass, using dual MitoTracker green staining.

Oxygen Consumption Rate

A Seahorse XF24-3 Bioanalyzer (Agilent Technologies) was used to measure mitochondrial oxygen consumption as described previously (Koentjoro et al., 2017). On day 1, 70,000 ONS cells per well were plated in a 24-well XF24 V7 cell culture microplate and cultured in DMEM/F12 (Gibco) with 10% fetal bovine serum at 37°C and 5% CO₂ for 24 h. On day 2, the DMEM/F12 media was replaced with Seahorse XF Base media supplemented with 2 mM L-glutamine, and the pH was adjusted to 7.35 ± 0.05 (basal media). Cells were incubated in the basal media for 1 h at 37°C in a non-CO₂ incubator. The plate was then assembled with a hydrated sensor cartridge and placed in the Seahorse flux analyzer. The oxygen consumption rate (OCR) was measured at baseline and after subsequent stepwise injection of the mitochondrial respiratory chain inhibitors, oligomycin (1 μM), carbonyl cyanide-4-(trifluoromethoxy)phenylhydrazone (FCCP) (1 μM) and rotenone/antimycin A (0.5 μM). The OCR readings were normalized to cell number measured by CyQUANT assay in a sister plate. Multiple parameters of mitochondrial respiration were calculated based on the response of the cells to the respiratory chain inhibitors: (a) basal respiration, basal OCR - non-mitochondrial respiration rate; (b) ATP production, difference in OCR before and after oligomycin treatment; (c) maximal respiration, OCR measurement after FCCP treatment - OCR measurement after oligomycin treatment; and (d) spare respiratory capacity, maximal respiration - basal respiration.

Cell Proliferation

ONS cells (5,000) were seeded into each well of a 96-well plate. On day 0 (4 h post seeding), day 1 (24 h post seeding), and day 2 (48 h post seeding), the relative cell numbers were assessed using the CyQUANT^® NF Cell Proliferation Assay Kit, following the manufacturer’s instructions.

Statistical Analysis

Three technical replicates were performed for all experiments: three control cell lines, three SPG7 patient lines, and three SPAST patient cell lines. One-way analysis of variance with Tukey’s multiple comparisons test was performed to compare control, SPG7, and SPAST sample results. Statistical analysis and graphing were performed with GraphPad (Prism 8).

Results

Clinical Phenotype

All patients with mutations in SPG7 had complex HSP phenotypes (clinical features detailed in Table 1). Complicating features in SPG7 patients included ophthalmoplegia (patient 1), ptosis (patient 1), ataxia (patients 2 and 3), and mitochondrial cytopathy on muscle biopsy (patient 1). In contrast, no SPAST patients (patients 4, 5, and 6) had typical mitochondrial disorder-related clinical signs (such as ocular myopathy, present in a range of mitochondrial myopathy syndromes; Pfeffer and Chinnery, 2013), and only one of three (patient 4) had additional features of dysarthria and upper limb weakness indicative of complicated HSP.

Genetic Studies

Six HSP patients, three with SPG7 mutations and three with SPAST mutations, were recruited to the study. These patients carried a variety of mutations (Table 1). SPG7 patients were patient 1 [c.1454_1462del (het); c.1529C > T (het)], patient 2 [c.1529C > T (het); c.1727C > G (het)], and patient 3 [c.1449 + 1G > A (novel het); c.1529C > T (het)]. SPAST patients were patient 4 (c.1392A > T), patient 5 (c.583C > G), and patient 6 (c.1096G > A). All patients except patient 3 have been reported previously (Abrahamsen et al., 2013; Kumar et al., 2013). SPG7 patient 3 was a 59-year-old man with a severe HSP phenotype complicated by upper limb cerebellar ataxia and cerebellar atrophy on MRI brain (Figures 1A,B). He was found to have the known common mutation NM_003119.3:c.1529C > T (p.Ala510Val) (Brugman et al., 2008; Bonn et al., 2010) and a novel canonical splice variant NM_003119.3:c.1449 + 1G > A, classified as pathogenic by the ACMG criteria (Richards et al., 2015) (PVS1, PM2, and PP3). Both variants were also present in a similarly affected sister (Figure 1C).

FIGURE 1

Figure 1. (A) Sagittal T1-weighted fast field echo and (B) transverse T2-weighted MRI images showing mild cerebellar atrophy of the patient (patient 3) carrying novel SPG7 variant. (C) Pedigree of patient 3 (arrow); both the proband and his affected sister carried two heterozygote variants in SPG7: a common mutation [NM_003119.3:c.1529C > T (p.Ala510Val)] and novel canonical splice variant (NM_003119.3:c.1449 + 1G > A). (D) Linear protein domain diagram of paraplegin. The AAA+ domain is shown as two subdomains (ATPase and Lid), separated by a linker region. Three of the four variants in the SPG7 patients in this study locate to the AAA+ ATPase domain and one to the M41 peptidase domain.

Protein Structure

All three SPG7 patients had a common heterozygous variant (c.1529C > T; p.A510V) in addition to different compounding heterozygous variants. All coding variants were predicted to be pathogenic by several prediction algorithms (e.g., MutationTaster, PolyPhen 2, SIFT, and PROVEAN), likely due to high evolutionary conservation of affected amino acids and the considerable physicochemical or likely structural changes imposed. Assessment of possible splicing changes for the non-coding variant using Human Splicing Finder 3.1 provided possible aberrant splicing outcomes (see below).

The pathogenicity of the c.1529C > T variant (p.A510V) common to all SPG7 patients in this study has been extensively studied (Bonn et al., 2010). It affects a highly conserved and completely buried alanine residue at the beginning of the structural AAA+ Lid domain (Figure 1D). Because of the constrained environment around the alanine residue, the introduction of a bulkier valine side chain (89 vs. 117 Da) would be predicted to destabilize the hydrophobic core of the Lid domain. Inspecting the crystal structure of the paraplegin AAA+ ATPase domain (amino acids 305–565; PDB entry 2qz4; Karlberg et al., 2009), of the three possible conformations of the valine side chain, two are predicted to clash with Arg486, and one could potentially be accommodated. This may afford expression of the mutant p.510V protein, although it would likely be dysfunctional for ATPase activity.

The second variant in patient 1 was a deletion variant (c.1454_1462del; p.485_487delRRE) removing three charged amino acids from the highly conserved alpha helix bundle of the AAA+ Lid domain (Figure 1D). The deletion would remove an entire alpha helix turn and three charged residues necessary for structural bonds to other helices in the domain (Karlberg et al., 2009) and is predicted to cause severe structural destabilization.

The second variant in patient 2 was a missense mutation (c.1727C > G; p.S576W) located within the magnesium metal ion coordination and peptidase active site of the M41 peptidase domain (Figure 1D). The substitution of tryptophan for serine is a very large physicochemical change, a mass change of 105 to 204 Da and a hydrophilic serine residue common in active sites vs. tryptophan, which has a bulky hydrophobic aromatic side chain. This would likely cause structural and functional disruption to the peptidase active site. Based on previous mutagenesis studies of this functional site, if the variant protein were expressed, it would likely form dysfunctional complexes (Shanmughapriya et al., 2015). In this situation, peptidase activity could be compensated for by AFG3L2, as demonstrated in yeast (Bonn et al., 2010).

The second variant in patient 3 was a novel canonical donor splice site disrupting mutation (c.1449 + 1G > A) that would impact the polypeptide chain from the end of the AAA+ ATPase domain at the start of the AAA+ domain linker region (Figure 1D). A number of possibilities for the consequent splicing defect include (1) skipping of exon 10 and a frameshift leading to termination after six amino acids; (2) intronic read-through into intron 10 before an alternate donor splice site is encountered 90 nucleotides downstream, resulting in an in-frame inclusion of 30 amino acids that would alter the length of the AAA+ ATPase domain linker; and (3) intronic read-through into intron 10 before encountering a termination codon 96 nucleotides downstream. Transcript analysis would be required to resolve the resulting splicing mechanism, although the variant protein product would almost certainly not be expressed.