Parkinson's disease mutations in PINK1 result in decreased Complex I activity and deficient synaptic function

Abstract

Mutations of the mitochondrial PTEN (phosphatase and tensin homologue)-induced kinase1 (PINK1) are important causes of recessive Parkinson disease (PD). Studies on loss of function and overexpression implicate PINK1 in apoptosis, abnormal mitochondrial morphology, impaired dopamine release and motor deficits. However, the fundamental mechanism underlying these various phenotypes remains to be clarified. Using fruit fly and mouse models we show that PINK1 deficiency or clinical mutations impact on the function of Complex I of the mitochondrial respiratory chain, resulting in mitochondrial depolarization and increased sensitivity to apoptotic stress in mammalian cells and tissues. In Drosophila neurons, PINK1 deficiency affects synaptic function, as the reserve pool of synaptic vesicles is not mobilized during rapid stimulation. The fundamental importance of PINK1 for energy maintenance under increased demand is further corroborated as this deficit can be rescued by adding ATP to the synapse. The clinical relevance of our observations is demonstrated by the fact that human wild type PINK1, but not PINK1 containing clinical mutations, can rescue Complex 1 deficiency. Our work suggests that Complex I deficiency underlies, at least partially, the pathogenesis of this hereditary form of PD. As Complex I dysfunction is also implicated in sporadic PD, a convergence of genetic and environmental causes of PD on a similar mitochondrial molecular mechanism appears to emerge.

Figure 1. Drosophila pink1 mutants fail to maintain normal synaptic transmission during intense activity

A–D. Neurotransmitter release in controls (w pink1^REV) and Drosophila pink1 mutants (w pink1^B9) in 2 mM (C) and 0.5 mM Ca²⁺ (A, B, D). Excitatory Junctional Potential traces (EJPs) (A, B) and quantification of EJP amplitudes (C, D) do not show a significant difference, indicating that basal neurotransmitter release is not affected in pink1 mutants. E–H. Spontaneous vesicle fusion (mEJPs) recorded in 0.5 mM Ca²⁺ in the presence of TTX in controls (E) and pink1 mutants (F). Quantification of mEJP amplitude (G) and mEJP frequency (H) did not show a significant difference between control and pink1. We did not observe a statistical difference between the two genotypes. I. Relative EJP amplitudes measured in 2 mM Ca²⁺ during 10 min of 10 Hz stimulation in controls and pink1 mutants. EJP amplitudes were binned per 30 s and normalized to the average amplitude of the first 10 EJPs. EJPs in pink1 mutants gradually declined to a level lower than that in controls. n = 4; ns = non-significant. The number of animals tested is indicated in the bar graphs and error bars indicate SEM.

Figure 2. Drosophila pink1 mutants show defects in reserve pool function that can be rescued by ATP

A, B. Exo–endo cycling vesicle pool (ECP) labelling in controls and pink1 mutants. Third instar larval fillets incubated for 5 min in HL-3 with 90 mM KCl, 2 mM Ca²⁺ and 4 µM FM1-43 to label the exo–endo cycling pool were washed in Ca²⁺ free HL-3 and imaged (A, ‘Load’) (Nikon FN-1 with DS-2MBWc digital camera, 40× water objective, NA 0.8). ECP vesicles were subsequently re-mobilized by incubating preparations for 5 min in 90 mM KCl, 2 mM Ca²⁺ and after washing in Ca²⁺ free HL-3, the same synapses imaged after loading were imaged again (A, ‘unload’). Fluorescence intensity of loading and unloading, normalized to loading intensity of controls, was quantified (B), but did not show a statistically significant difference. Scale bar = 0.2 µm. C, D. Reserve pool (RP) labelling in controls, pink1 mutants (pink1^B9) and pink1 mutants neuronally expressing human PINK1 (hpink1). Both the ECP and RP were labelled by electrically stimulating motor neurons of third instar fillets in HL-3 with 2 mM Ca²⁺ for 10 min and then leaving the dye with the preparation to rest for 5 min following stimulation (C, ‘load’). ECP vesicles, but not RP vesicles, were subsequently re-mobilized by incubating preparations for 5 min in 90 mM KCl, 2 mM Ca²⁺ and after washing in Ca²⁺ free HL-3, the same synapses imaged after loading were imaged again (C, ‘unload’). Fluorescence intensity of loading and unloading, normalized to loading intensity of controls, was quantified (D). Note the much weaker RP labelling in pink1 mutants compared to controls. Scale bar = 0.2 µm. E, F. Reserve pool (RP) labelling with forward-filling ATP in controls and pink1 mutants. Loading of FM1-43 after forward-filling control and pink1^B9 neurons with 1 mM ATP was performed as described above. Both ECP and RP vesicles are labelled after loading (E, ‘load’) and subsequently the ECP vesicles, but not RP vesicles, were re-mobilized by incubating preparations for 5 min in 90 mM KCl, 2 mM Ca²⁺ (E, ‘unload’). Fluorescence intensity of loading and unloading, normalized to loading intensity of controls, was quantified (F). Statistical analysis was performed using the unpaired Student's t-test, where *p < 0.05; **p < 0.01 when compared to control. ns = non-significant. The number of animals tested is indicated in the bar graphs and error bars indicate SEM. White bars: load; black bars: unload. Scale bar = 0.2 µm.

Figure 3. Morphology of mitochondria is not altered in the absence of PINK1

A, B. Confocal images of mitochondrial green fluorescent protein (mitoGFP) labelling (top panel) in control (w pink1^REV) and pink1 mutant (w pink1^B9). Drosophila third instar larval NMJs were also labelled with discs large (DLG/PSD95) (bottom panel) marking pre- and post-synaptic areas of synaptic boutons (A). Mitochondria at the synapse were quantified (B) by measuring mitochondrial surface to synaptic surface in maximum intensity projections (adapted from Verstreken et al (2005)). We did not observe a statistical difference between the two genotypes; ns = non-significant. The number of animals tested is indicated in the bar graphs (eight synapses from w pink1^REV and 10 synapses from w pink1^B9) and error bars indicate SEM. Scale bar = 0.2 µm. C, D. Mitochondrial morphology of wild-type (Pink1^+/+) and Pink1 knock-out (Pink1^−/−) fibroblast cells transfected with a mitochondrially targeted YFP (mtYFP). Twenty-four hours post-transfection, cells were imaged on an inverted Olympus IMT-2 microscope equipped with a Cell®R imaging system (C). The corresponding morphometric analysis (D) was performed as described previously in Cipolat et al (2004). We did not observe a statistical difference between the two genotypes. Data represent the average ±SEM of three independent experiments (100 images for Pink1^+/+, 120 images for Pink1^−/−); ns = non-significant. E. High voltage transmission electron microscopy of Pink1^+/+ and Pink1^−/− mouse brain and muscle (heart) tissue does not show morphological changes in the mitochondria. Ultrathin tissue sections (70 nm) were cut with an ultramicrotome and negatively stained with uranyl acetate and lead citrate. Samples were examined in a JEOL JEM-2100. Scale bar = 0.2 µm.

Figure 4. Absence of PINK1 causes deficits in mitochondrial membrane potential (Δψ_m) and increases susceptibility to apoptotic stimuli

A, B. Imaging of Δψ_m at third instar Drosophila larval NMJs in controls (w pink1^REV) and pink1 mutants (w pink1^B9) using the ratiometric dye JC-1 as described previously (Verstreken et al, 2005). Pre-treatment of the preparations with 2 µM FCCP (an uncoupler) resulted in a lack of dye accumulation in mitochondria (A, third panel). Quantification of red JC-1 fluorescence emission to green emission (in the same area) showed a significant decrease in Δψ_m in pink1 mutants compared to controls (B). C. Quantification of the third instar NMJ morphology by length (µm⁻¹) and bouton number (z/µm²) both normalized to muscle surface area was similar in controls and pink1 mutants. D, E. Imaging of Δψ_m in Pink1^+/+ (black squares) and Pink1^−/− (grey squares) fibroblast cells grown on cover slips and loaded for 30 min at 37°C with 10 nM TMRM in the presence of (D) 2 µg/ml cyclosporine H (CsH), a P-glycoprotein inhibitor, or (E) 1 µg/ml cyclosporine A (CsA), a P-glycoprotein and PTP inhibitor. Cells were then placed on the stage of an Olympus IMT-2 inverted microscope equipped with a Cell®R imaging system. Sequential images of TMRM fluorescence were acquired every 60 s over a 40 min time course. TMRM fluorescence over mitochondrial regions of interest was measured. When indicated (arrows), 2 µg/ml oligomycin (an ATP synthase inhibitor) and 2 µM FCCP (an uncoupler) were added. Data represent the average ± SEM of eight independent experiments. F, G. Pink1^+/+ and Pink1^−/− fibroblasts were treated with increasing concentrations of H₂O₂ (F) and arachidonic acid (G) and after 2 h apoptosis was determined by the percentage of Annexin-V positive cells by flow cytometry. Data represent the average ± SEM of three independent experiments. H. Cytochrome c release assays were performed on mitochondria isolated by differential centrifugation from Pink1^+/+ and Pink1^−/− mouse liver. Mitochondria were treated for 0, 10, 20, 40 and 60 min with p7/p15 recombinant BID (cBID). The supernatant was analysed on a cytochrome c ELISA and plotted as the percentage of total cytochrome c released. Data represent the average ± SEM of three independent experiments. Statistical analysis was performed using the unpaired Student's t-test, where *p < 0.02; ***p < 0.0005 when compared to control (Graph Pad Prism5 software). ns = non-significant, a.u., arbitrary units; CsH, cyclosporine H; CsA, cyclosporine A; FCCP, carbonylcyanide-p-trifluoromethoxyphenylhydrazone; TMRM, tetramethylrhodamine methyl ester.

Figure 5. PINK1 deficit results in defects in Complex I of the electron transport chain

A. To measure the respiration rates between stimulated and basal ADP (ratio State3/State4), mitochondria were isolated from Pink1^+/+ and Pink1^−/− mice by differential centrifugation and were incubated in the experimental buffer. Mitochondrial oxygen consumption assays were performed in the presence of 5 mM glutamate/2.5 mM malate for the analysis of Complex I-driven respiration; 5 mM succinate in the presence of 2 µM rotenone for Complex II-driven respiration; or 3 mM ascorbate plus 150 µM TMPD in the presence of 0.5 µg/ml antimycin A for Complex IV-driven respiration. Mitochondrial oxygen consumption was measured by using a Clarke-type oxygen electrode (Hansatech Instruments). Data represent the average ± SEM of three independent experiments. B–G. Respiratory chain measurements performed on mitochondrial homogenates from Pink1^+/+, Pink1^−/−, and also the rescue cell line with human PINK1 (hPINK1_res) fibroblast cells were analysed by spectrophotometric assays for the activity measurement of (B) Complex I (NADH:ubiquinone oxidoreductase, rotenone sensitive), (C) Complex II (succinate:ubiquinone oxidoreductase, malonate sensitive), (D) Complex II+III, (E) Complex III (ubiquinone:cytochrome c oxidoreductase, antimycine sensitive), (F) Complex IV (cytochrome c oxidase). Measurements of (G) Complex V (ATPsynthase, oligomycine sensitive) and citrate synthase enzyme activities were also performed. The protein concentration was 2–4 mg/ml. Values were plotted according to the ratio between the specific complex's activity and citrate synthase activity. H. Respiratory chain Complex I measurements were performed on mitochondrial homogenates from Pink1^+/+ and Pink1^−/− mouse brain tissue by spectrophotometric assays (NADH:ubiquinone oxidoreductase, rotenone sensitive). Citrate synthase enzyme activities were also performed. Values were plotted according to the ratio between the specific Complex I activity and citrate synthase activity. I. Respiratory chain Complex I measurements were performed on fly mitochondrial homogenates from control (w pink1^REV) and pink1 mutant (w pink1^B9) brain enriched (heads) and muscle-enriched (thorax) tissues by spectrophotometric assays (NADH:ubiquinone oxidoreductase, rotenone sensitive). Citrate synthase enzyme activities were also performed. Values were plotted as a ratio of the specific Complex I activity and citrate synthase activity. Statistical analysis was performed using the unpaired Student's t-test, where *p = 0.03, **p < 0.005 and ^***p = 0.00006 when compared to wild-type (Graph Pad Prism5 software). Data represent average ± SEM of at least three independent experiments. ns = non-significant.

Figure 6. PINK1 clinical mutations also show reduced Complex I function and concomitant defects in synaptic function

A. PD-related PINK1 mutants were transfected in Pink1^−/− fibroblasts and expression levels were analysed by reverse transcriptase-PCR for mRNA detection. B. Respiratory chain measurements performed on mitochondrial homogenates from Pink1 deficient fibroblast cell lines stably expressing human PINK1 (hPINK1_res), PD-related PINK1 mutant hPINK1_G309D and hPINK1_W437X, and the artificial mutant hPINK1_K219A. Spectrophotometric assays for Complex I activity (NADH:ubiquinone oxidoreductase, rotenone sensitive) were performed. Statistical analysis was done using the unpaired Student's t-test, where ^***p = 0.00006, where **p < 0.004 and *p < 0.01 (Graph Pad Prism5 software). Data represent the average ± SEM of five independent experiments. ns = non-significant. Values presented for Pink1^+/+, Pink1^−/− and hPINK1_res were previously shown in Fig 4A. C, D. Reserve pool (RP) labelling in pink1 mutants that neuronally express wild-type human hPINK1 (w pink1^B9/Y; elav-GAL4/UAS-hPink1) or the PD-related hPINK^G309D mutant (w pink1^B9/Y; elav-GAL4/UAS-hPink1^G309D). Both the ECP and RP in these animals were labelled by electrically stimulating motor neurons of third instar fillets in HL-3 with 2 mM Ca²⁺ for 10 min and then leaving the dye with the preparation to rest for 5 min following stimulation (C, ‘load’). ECP vesicles, but not RP vesicles, were subsequently re-mobilized by incubating preparations for 5 min in 90 mM KCl, 2 mM Ca²⁺ and after washing in Ca²⁺ free HL-3, the same synapses imaged after loading were imaged again (C, ‘unload’). Fluorescence intensity of loading and unloading, normalized to loading intensity of pink1 mutants that express hPink1 in their neurons, was quantified (D). Note that wild type hPink1 can rescue RP labelling while hPink1^G309D cannot. Comparison of RP labelling in w pink1^B9 and w pink1^B9/Y; elav-GAL4/UAS-hPink1^G309D is not statistically significantly different. Statistical analysis was performed using the unpaired Student's t-test, where *p < 0.05; **p < 0.01 when compared to control. ns = non-significant. The number of animals tested is indicated in the bar graphs and error bars indicate SEM. White bars: load; black bars: unload.

Figure 7. Schematic representation of upstream and downstream defects caused by PINK1 deficiency

As discussed in the text and illustrated in this figure, Complex I deficiency can explain almost any alteration previously found to be associated with PINK1 loss of function. The decreased electron transfer at the level of Complex I can lead to reduced Δψ_m (present study), decreased ATP generation (Clark et al, 2006) and increased ROS production (Gautier et al, 2008). Deficient ATP levels affect dopamine release (Kitada et al, 2007) and RP mobilization (present study) causing reduced RP mobilization. Additionally, reduced Δψ_m can destabilize cytochrome c release (present study), and dephosphorylation of Drp1 that causes alterations in mitochondrial fusion/fission rates and morphology (Cereghetti et al, 2008). Recruitment of Parkin to such mitochondria (Narendra et al, 2008) will partially act protectively, as it will remove the most severely affected mitochondria, also explaining the genetic interaction between Parkin and Pink (Poole et al, 2008).