Loss of α-Synuclein Does Not Affect Mitochondrial Bioenergetics in Rodent Neurons

Abstract Increased α-synuclein (αsyn) and mitochondrial dysfunction play central roles in the pathogenesis of Parkinson’s disease (PD), and lowering αsyn is under intensive investigation as a therapeutic strategy for PD. Increased αsyn levels disrupt mitochondria and impair respiration, while reduced αsyn protects against mitochondrial toxins, suggesting that interactions between αsyn and mitochondria influences the pathologic and physiologic functions of αsyn. However, we do not know if αsyn affects normal mitochondrial function or if lowering αsyn levels impacts bioenergetic function, especially at the nerve terminal where αsyn is enriched. To determine if αsyn is required for normal mitochondrial function in neurons, we comprehensively evaluated how lowering αsyn affects mitochondrial function. We found that αsyn knockout (KO) does not affect the respiration of cultured hippocampal neurons or cortical and dopaminergic synaptosomes, and that neither loss of αsyn nor all three (α, β and γ) syn isoforms decreased mitochondria-derived ATP levels at the synapse. Similarly, neither αsyn KO nor knockdown altered the capacity of synaptic mitochondria to meet the energy requirements of synaptic vesicle cycling or influenced the localization of mitochondria to dopamine (DA) synapses in vivo. Finally, αsyn KO did not affect overall energy metabolism in mice assessed with a Comprehensive Lab Animal Monitoring System. These studies suggest either that αsyn has little or no significant physiological effect on mitochondrial bioenergetic function, or that any such functions are fully compensated for when lost. These results implicate that αsyn levels can be reduced in neurons without impairing (or improving) mitochondrial bioenergetics or distribution.

␣Syn and mitochondria also directly affect each other. In mice and humans, a fraction of ␣syn associates with mitochondria in DA neurons (Martin et al. 2006;Li et al. 2007;Devi et al. 2008). Additionally, supra-physiologic levels of ␣syn disrupt mitochondrial morphology in vitro and in vivo (Kamp et al. 2010;Butler et al. 2012), promote excessive mitophagy (Choubey et al. 2011;Sampaio-Marques et al. 2012), disrupt mitochondrial protein import (Di Maio et al. 2016) and influence mitochondrial Ca 2ϩ homeostasis and apposition between the endoplasmic reticulum and mitochondria (Cali et al. 2012;Guardia-Laguarta et al. 2014). Increased ␣syn also inhibits mitochondrial complex I in cell lines and mouse brains (Devi et al. 2008;Chinta et al. 2010;Loeb et al. 2010), and it correlates with decreased complex I activity in mitochondrial fractions from the SN of PD patients (Devi et al. 2008).
The effects of increased ␣syn on mitochondria may result from a toxic gain-of-function, such as the accumulation of oligomeric ␣syn species that interact preferentially with mitochondria (van Rooijen et al. 2009;Nakamura, 2013;Luth et al. 2014). Moreover, even basal levels of ␣syn may predispose mitochondria to dysfunction. In ␣syn knockout (KO) mice, SN DA neurons resist toxicity from mitochondrial toxins, including 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP) (Dauer et al. 2002;Klivenyi et al. 2006), and reducing endogenous ␣syn protects against toxicity from the complex I inhibitor rotenone (Zharikov et al. 2015) and may improve mitochondrial protein import (Di Maio et al. 2016). However, excessively lowering ␣syn could also adversely affect mitochondria. While increased ␣syn causes mitochondrial fragmentation, decreased ␣syn can produce excessive mitochondrial tubulation (Kamp et al. 2010;Norris et al. 2015). Additionally, ␣syn KO mice have decreased levels of the mitochondrial lipid cardiolipin (Ellis et al. 2005) and lowering ␣syn impairs complex I/III activity, perhaps by directly interacting with complex I (Ellis et al. 2005;Devi et al. 2008). In addition, syn TKO mitochondria have decreased mitochondrial membrane potential but increased oxygen consumption, and lower activity of ATP synthase (Ludtmann et al. 2016).
Both ␣syn and mitochondria are major targets for PD therapy. Lowering ␣syn is under study in clinical trials (Dehay et al. 2015), but whether this approach is safe remains unknown. Interestingly, although single-syn KO (␣, ␤, or ␥) and double-syn KO mice (␣ and ␤) have normal lifespans (Chandra et al. 2004), triple-syn KO mice (␣, ␤, and ␥) die early (Greten-Harrison et al. 2010). These results suggest that ␣syn mediates essential functions, but other syn isoforms can compensate for its loss, at least over the lifespan of a mouse. In addition, the capacity to compensate for ␣syn loss may decrease once neuronal maturation is complete. Indeed, some studies found that lowering ␣syn in adult rodents (Benskey et al. 2016b) or non-human primates (Collier et al. 2016) using shRNA is toxic to nigral DA neurons, while other studies have not (McCormack et al. 2010;Zharikov et al. 2015), with the discrepancies likely due to varying degrees of ␣syn knockdown. Therefore, we must better understand the consequences of lowering ␣syn, particularly on mitochondrial function in axons where ␣syn concentrates and energy failure can selectively occur .
In this study, we aimed to determine if ␣syn is required for normal mitochondrial bioenergetics, and if ␣syn levels can be safely lowered without affecting mitochondrial bioenergetics. We comprehensively evaluated how the loss of ␣syn impacts mitochondrial function, including respiration in neurons and isolated nerve terminals, mitochondrial-derived ATP levels specifically at synapses of intact neurons, localization of mitochondria to DA synapses in vivo, and total bioenergetic function.

Molecular Biology
All constructs used for transient transfection were subcloned or cloned into the pCAGGS vector downstream of the chicken actin promoter (Voglmaier et al. 2006). The AT1.03 YEMK FRET sensor was a kind gift from Dr Hiroyuki Noji at Osaka University (Imamura et al. 2009a). VGLUT1-pHluorin-mCherry is derived from the VGLUT1-pHluorin fusion (Voglmaier et al. 2006) and was a kind gift from Dr Timothy Ryan (Weil Cornell Medical School), mCherrysynaptophysin has been described (Voglmaier et al. 2006;Hua et al. 2011), and mTagBFP was a kind gift from Dr Vladislav Verkhusha at the Albert Einstein College of Medicine (Subach et al. 2008 and recombinant adeno-associated virus (AAV)1 was made by the Vector Core at the University of North Carolina.
Viral constructs for ␣syn silencing were similar to those used before (Gorbatyuk et al. 2010); the siRNA sequences [␣syn: GAAGGACCAGATGGGCAAG, scrambled (SCR): GTCGACAATTCATATTTGC] were expressed as a shRNA by incorporating the loop structure TTCAAGAGA. The shRNA cassette was inserted behind an H1 promoter, and the viral genome also contained mTagBFP2 under the control of the hybrid chicken ␤-actin/cytomegalovirus enhancer promoter (pCBA) as a transduction marker. The viral vectors were packaged into AAV5 capsids by transfection of 293 cells with the viral genome and the pXYZ5 helper plasmid. Viral particles were purified using an iodixanol gradient followed by column chromatography, and titers were determined by dot-blot (Benskey et al. 2016a).

Knockout and Transgenic Mice
␣Syn KO mice on a C57BL/6N background (strain 016123, The Jackson Laboratory) (Baptista et al. 2013) were used for most experiments. Before these mice were available, ␣syn KO mice on a mixed C57BL/6 and 129 ϫ 1/SvJ background (strain 003692, The Jackson Laboratory, backcrossed one generation with C57BL/6N controls) were used for experiments outlined in Fig. 2B. DATcre (Backman et al. 2006) mice were also obtained from The Jackson Laboratory. C57BL/6N mice served as controls for all studies. Mice were group-housed in a colony maintained with a standard 12-h light/dark cycle and given food on the cage floor and water ad libitum. All experiments were performed on age-and sex-matched mice. Experiments were conducted in accordance with the Guide for the Care and Use of Laboratory Animals, as adopted by the National Institutes of Health and with approval of the Authors' University Institutional Animal Care and Use Committee.

Synaptosome Isolation
Cortical synaptosomes were isolated from cerebral cortices of 6-month-old mice as described (Gerencser et al. 2009). Briefly, cortices were quickly dissected, rinsed, and gently homogenized in ice-cold sucrose buffer (320 mM sucrose, 1 mM EDTA, 0.25 mM dithiothreitol, pH 7.4). Homogenates were centrifuged at 1000 ϫ g for 10 min at 4°C. The supernatant was layered on top of a discontinuous Percoll gradient of 3, 10, and 23% Percoll layers in sucrose medium and centrifuged at 32500 ϫ g for 10 min at 4°C. Synaptosomes accumulated as a band between the 10% and 23% Percoll layers and were gently aspirated and washed in HBS medium containing 20 mM HEPES, 10 mM D-glucose, 1.2 mM Na 2 HPO 4 , 1 mM MgCl 2 , 5 mM NaHCO 3 , 5 mM KCl, and 140 mM NaCl at pH. 7.4. The final synaptosomal pellet was resuspended in HBS medium.
To prepare dopaminergic synaptosomes, mice striata were quickly dissected and homogenized using ice-cold sucrose buffer . Homogenates were then incubated with antibodies against the dopamine transporter (Alpha Diagnostic International; 25 g/sample) for 60 min at 4°C and washed three times in sucrose buffer at 10 000 ϫ g for 2 min. Pellets were then incubated with 150 l of secondary IgG magnetic beads (Miltenyi) for 45 min at 4°C and then poured into a magnetic column (MACS LS; Miltenyi) to separate the magnetic bead-labeled dopaminergic synaptosomes (bound to column, DA) from the nondopaminergic fraction that flows through the column.

Respiration and Glycolysis
The extracellular-acidification rates (ECAR, a surrogate for glycolysis) and oxygen-consumption rates (OCR, assesses mitochondrial respiration) were measured in cultured hippocampal neurons using a Seahorse XF96 Extracellular Flux Analyzer (Seahorse Bioscience), an instrument that can simultaneously assess aerobic and anaerobic metabolism in adherent cells cultured in 96-well plates. Cells were washed and preincubated for 30 min in Seahorse assay medium (pH 7.4) containing substrates of interest (30 mM glucose and 10 mM pyruvate). OCR and ECAR were measured at baseline and again after sequential addition of the respiratory inhibitors FCCP (1 M), rotenone (3 M), and oligomycin (2 M), or with veratridine (50 M) to increase neural activity (Lysko et al. 1994). To assess any effects of genotype on cell survival, cells in a subset of wells were also stained with DAPI (4',6diamidino-2-phenylindole) and quantified using Meta-Morph software (Universal Imaging).
Aerobic respiration was also measured in cortical and dopaminergic synaptosomes prepared from ␣syn KO and control mice. Before plating synaptosomes, Seahorse plates were coated with 0.0033% (v/v) polyethyleneimine solution and Geltrex suspension. Synaptosomes were then added to each well (cortical 20 g/well; dopaminergic 40 g/well), and the plates were centrifuged at 3200 ϫ g for 50 mins at 4°C to attach the synaptosomes to the surface. For Seahorse measurements, HBS medium was replaced with Seahorse buffer containing 3.5 mM KCl, 120 mM NaCl, 1.3 mM CaCl 2, 0.4 mM KH 2 PO 4 , 1.2 mM Na 2 SO 4 , 2 mM MgSO 4 , 10 mM TES, 10 mM Na-pyruvate, and 4 mg/ml bovine serum albumin.
VGLUT1-pHluorin fluorescence images were obtained [490/20 excitation (ex), 535/50 emission (em) ( The FRET/donor ratio was calculated for each bouton as described (Xia and Liu, 2001), where FRET ϭ (I FRET Ϫ I CFP ‫ء‬ BT CFP -I YFP ‫ء‬ BT YFP )/I CFP , such that I X is the background-corrected fluorescence intensity measured in a given channel. BT CFP (donor bleed through) and BT YFP (direct excitation of the acceptor) were calculated by expressing CFP and YFP individually and then determining the ratios of I FRET /I CFP and I FRET /I YFP , respectively.

Stereology
Unbiased stereology was used to quantify the number of TH-positive neurons in the SN as described (Gorbatyuk et al. 2010). Sections were visualized using 4x magnification (Olympus BX53 microscope equipped with a motorized stage (Olympus, Center Valley, PA) and a Qimaging 2000R camera (Qimaging, Surrey, BC, Canada)), and the SN was outlined. THϩ cells from every sixth section were counted using the optical fractionator method with a 60x oil objective (Stereo Investigator, MBF Bioscience). The coefficient of error was calculated according to Gundersen and Jensen (Gundersen and Jensen, 1987) and was Ͻ0.1 (Gundersen, m ϭ 1).

Comprehensive Lab Animal Monitoring System Measurements
For metabolic measurements, a Comprehensive Lab Animal Monitoring System (CLAMS, Columbus Instruments) was used to measure the rates of O 2 (VO 2 ) and Negative Results CO 2 consumption (VCO 2 ), the respiratory-exchange ratio (RER; [dot]VCO 2 /VO 2 ), and the activity level of 6-month-old male mice (Millership et al. 2012). These measurements were calculated for both the dark and light cycle for 3 consecutive days. The animals were maintained on a regular chow diet (10% kcal from fat). The body composition (lean and fat mass) of the control and syn KO mice was also analyzed using EchoMRI .

Loss of ␣syn Does Not Affect Mitochondrial Bioenergetics at the Nerve Terminal
To determine if ␣syn is required for neuronal respiration, we measured how ␣syn KO affects the oxygen consumption rate (OCR, a surrogate of respiration) of E18 hippocampal neuronal cultures with a Seahorse instrument. We examined hippocampal neurons because ␣syn biology has been extensively characterized in hippocampal neurons in culture and in vivo (Greten-Harrison et al. 2010;Nemani et al. 2010;Scott et al. 2010;Volpicelli-Daley et al. 2011), and ␣syn also aggregates in hippocampal neurons in PD (Hall et al. 2014). In addition, in contrast to dopamine (DA) neurons that constitute only a fraction of the total neurons in midbrain cultures, hippocampal neuronal cultures consist primarily of excitatory pyramidal neurons (Beaudoin et al. 2012), facilitating their analysis in bulk assays, such as with the Seahorse. Neuronal cultures were grown in serum-free media to minimize the glial content so that respiration is responsible for most of the OCR signal . We found that ␣syn KO cultures had similar basal and maximal (after treatment with 1 M FCCP) respiration as controls (Fig. 1A, B). In addition, increasing neuronal activity with veratridine (50 M) augmented OCR similarly in both groups (Fig. 1C), suggesting that the respiratory function of ␣syn KO neurons upregulates normally when energy requirements are increased. Nonetheless, there was a small trend for ␣syn KO to have decreased basal OCR in several runs (Figs. 1A-C), and we cannot exclude the possibility that ␣syn KO causes a small impairment in respiration that was below the sensitivity of the assay (see statistical table). Notably, ␣syn KO neurons also had similar basal rates of extracellular acidification (ECAR, a surrogate of glycolysis), which increased similarly after treatment with oligomycin (2 M) (Fig. 1D). These results suggest that ␣syn KO neurons also have normal glycolytic capacity. Notably ␣syn KO also did not affect the total number of surviving cells per well, which was assessed in a subset of wells by DAPI staining (control: 296.9 cells/100 mm 2 ؎ 6.0, ␣syn KO: 305.7 cells/100 mm 2 ؎ 7.3; mean ؎ SE, 30 wells per group).
The aforementioned measurements interrogate the overall respiration of neurons, but they may not be sensitive to changes in specific subcellular compartments, such as the nerve terminal. ␣Syn concentrates at synapses, which have high-energy requirements (Rangaraju et al. 2014;, suggesting that any effects of ␣syn KO on bioenergetics would be most prominent in this compartment. To understand the effect of ␣syn KO on respiration specifically in synapses, we ex-amined the respiration of synaptosomes isolated from the brains of 6-month-old control and ␣syn KO mice. We did not observe differences in the basal or maximal respiration in either cortical or dopaminergic synaptosomes (Fig.  1E, F), although there was a small trend for decreased respiration in the ␣syn KO dopaminergic synaptosomes.
We next examined how ␣syn KO impacted ATP levels at the synapse, which reflect the balance between ATP production and consumption. ␣Syn KO and control neurons from postnatal hippocampi were co-transfected with ATP FRET sensors (Imamura et al. 2009b) and mCherrysynaptophysin to identify synaptic boutons and then cultured for 10 days before imaging. As expected, basal ATP levels were normal with glucose and pyruvate treatment in ␣syn KO neurons ( Fig. 2A). To specifically examine the capacity of ␣syn KO mitochondria to produce ATP, we forced neurons to rely on mitochondria-derived ATP by acutely blocking glycolysis (switching to no-glucose media with glycolytic inhibitors). Under these conditions, the assay sensitively detects decreases in energy caused by either acute pharmacologic or chronic, genetic mitochondrial deficits. Specifically, each comparison was sensitive to an Ϸ10%-20% decrease in ATP FRET signal with 90% power and an alpha of 0.05 (statistical table), while the FRET must decrease by Ϸ40% from baseline to pass the threshold level (corresponding to Ϸ0.8 mM ATP) required for endocytosis . Furthermore, because bioenergetic deficits may only appear when energy requirements increase , we tested if ␣syn KO influences the ability of synapses to maintain ATP levels when their energy requirements are increased (Attwell and Laughlin, 2001). Even when neural activity was augmented with electrical field stimulation (10 Hz‫06ء‬ s) (Nemani et al. 2010), ATP levels decreased similarly with or without ␣syn (Fig. 2B, C).
We also assessed whether loss of ␣syn affected synaptic vesicle cycling (particularly endocytosis), an ATPconsuming process that is sensitive to decreases in ATP (Rangaraju et al. 2014;. To monitor synaptic vesicle cycling in individual boutons, we used the VGLUT1-pHluorin reporter, which targets a pH-sensitive GFP to the lumen of synaptic vesicles. The pHluorin does not fluoresce in acidified vesicles, but does when synaptic vesicles fuse and expose their contents to the alkaline extracellular environment (Voglmaier et al. 2006;Nemani et al. 2010). Hippocampal neurons expressing VGLUT1-pHluorin were incubated in buffer with 10 mM pyruvate, but without glucose to favor reliance on mitochondria for energy. Even with glycolytic inhibitors, synaptic vesicle cycling (10 Hz‫06ء‬ s, which preferentially targets the recycling pool) was normal in synaptic boutons lacking ␣syn, further supporting that mitochondrial-derived ATP persists at functionally significant levels (Fig. 2D). Because developmental compensation may occur in ␣syn KO neurons, we also examined how knocking down ␣syn with shRNA impacts ATP levels in rat neurons. The shRNA reduced ␣syn expression by ϳ60% based on immunofluorescence (Fig. 2E); however, this decrease did not affect synaptic vesicle cycling (Voglmaier et al. 2006;Nemani et al. 2010) (Fig. 2F).
Increased ␤ and ␥, syn TKO) isoforms (but not loss of ␣syn alone) decreases mouse lifespan, raising the possibility that ␤ and ␥syn compensate to maintain mitochondrial function when ␣syn is lost. However, sustained electrical stimulation (5 Hz‫574ء‬s) decreased ATP levels similarly in syn  for neurons and synaptosomes). A, ␣Syn KO had no effect on the basal or maximal (after FCCP) respiration of hippocampal neurons in medium containing 10 mM pyruvate and 30 mM glucose (compilation of two experiments, n ϭ 13 wells per group). B, Oligomycin and rotenone similarly decrease OCR in ␣syn KO and control groups (compilation of two experiments, n ϭ 10 wells per group). C, Increasing neuronal activity with veratridine similarly increased OCR (compilation of two experiments, n ϭ 6 wells per group), while oligomycin similarly increased ECAR (D; extracellular acidification rate, a surrogate of glycolysis; compilation of two experiments, n ϭ 9 wells per group). E, F, cortical synaptosomes (E) and dopamine (DA) synaptosomes (F, right) isolated from the striatum also had similar basal and maximal rates of respiration (n ϭ 15 wells per group from two experiments for cortical synaptosomes; n ϭ 7-8 wells per group from two experiments for DA synaptosomes). As expected, western blotting (F, left) shows that both control and ␣syn KO DA synaptosomes are enriched in tyrosine hydroxylase (TH). All graphs show mean Ϯ SEM. NS ϭ not significant by two-way ANOVA and Sidak's posthoc test. TKO and control synaptic boutons, suggesting that concurrent loss of isoforms does not affect mitochondriaderived ATP levels at the nerve terminal (Fig. 3A). As ATP levels depend on the balance between energy production (aerobic respiration and glycolysis) and consumption, we also assessed if syn TKO might impact the rate of ATP consumption. However, when energy production was blocked (the respiratory chain was blocked with rotenone (2 M), and external glucose was removed to limit glycolysis), and energy consumption was increased with repetitive electrical stimulation (30 Hz‫5ء‬s every 120s), the stimulus-dependent endo-and exocytic response of syn TKO and control boutons failed at the same rate ( Fig.  3B-D), indicating that ATP levels fell below the threshold level needed to support synaptic vesicle cycling at the same rate . Since the ATP level reflects a balance between ATP production and consumption, these results suggest that the net balance of ATP consumption and any residual ATP produc-tion by glycolysis is also similar in syn TKO and control synaptic boutons.

Lowering ␣syn Does Not Impact Mitochondrial Distribution in Axons
Even when the intrinsic function of individual mitochondria is normal, changes in the distribution of mitochondria could create regions within neurons (especially axons) without sufficient mitochondria to meet energy requirements, leading to energy failure in that region. ␣Syn primarily locates to presynaptic terminals (Jakes et al. 1994;Iwai et al. 1995), and mutant A53T ␣syn decreases the movement and density of mitochondria in axons (Choubey et al. 2011;. To determine if endogenous ␣syn influences the localization of axonal mitochondria to synapses, we used cre-dependent AAVbased viral reporters to visualize mitochondria [mitochondrial-targeted GFP (mitoGFP)] specifically in individual DA neurons and their synapses (mCherry-  4A, B).
Notably, ␣syn KO mice may have developmental changes that compensate for the loss of ␣syn (Kuhn et al. 2007). Although we did not detect changes in the level of ␤syn in total brain lysates (Fig. 4C), other groups found it upregulated in the midbrain of ␣syn KO mice (Robertson et al. 2004;Thomas et al. 2011) and that compensation could occur independent of changes in expression. To further exclude the possibility of any developmental compensation, we also examined if lowering ␣syn with shRNA in adult mice (Gorbatyuk et al. 2010;Kanaan and Manfredsson, 2012) would impact the axonal localization of mitochondria. Using AAV expressing an shRNA against ␣syn, we lowered ␣syn levels in DA neurons by ϳ60% (Figs 4D-E, 4E-1) (as measured by immunofluorescence), a level of decrease that may not be quite sufficient to produce neuronal loss in rats (Gorbatyuk et al. 2010), and mice may also be more resistant (Benskey et al. 2016b). Consistent with this, there was no significant loss of DA neurons in the ␣syn shRNA group (Fig. 4F), although there was a trend for decreased THϩ counts. This level of ␣syn decrease also did not affect the proportion of boutons containing mitochondria (Fig. 4G). Therefore, ␣syn levels can be significantly lowered in DA neurons without impacting the synaptic targeting of mitochondria. We cannot exclude the possibility that further lowering of ␣syn by shRNA such that the DA neurons die would have disrupted mitochondria, and it would be difficult to attribute such a change specifically to ␣syn lowering as mitochondria are typically disrupted during neuronal death, regardless of the cause. Nonetheless, when considered with the lack of effect in ␣syn KO mice, our data suggest that either ␣syn does not normally impact mitochondrial distribution in DA axons or that other factors compensate for ␣syn loss .
␣Syn is expressed throughout the brain, including in regions that influence respiration and metabolism. ␣Syn is also present at high levels in certain peripheral tissues including in red blood cells, liver and spleen (Kuo and Nussbaum, 2015), and phosphorylated ␣syn accumulates Figure 2. Loss of ␣syn does not affect mitochondrial-derived ATP levels at the nerve terminal. A-C, ATP levels of hippocampal neurons were assessed using an ATP YEMK FRET sensor, and synaptic boutons were identified with mCherry-synaptophysin. Basal ATP levels in Tyrodes buffer containing glucose and pyruvate were identical in neurons isolated from control and ␣syn KO mice (A; n ϭ 14 -15 coverslips, not significant (NS) by unpaired two-tailed t test). Electrical field stimulation (10 Hz‫06ء‬ s, blue lines) in pyruvate buffer without (B) and with (C) 2-deoxyglucose (2DG, 5 mM) and iodoacetate (IAA, 1 mM) to completely block glycolysis reduced ATP levels similarly in neurons in control and ␣syn KO mice (compilation of two experiments, n ϭ 6 -7 coverslips/group with 15-20 boutons/coverslip). NS for ATP level of ␣syn KO versus control groups at corresponding time points. Note that overall ATP levels (control and ␣syn KO) decreased after the first electrical stimulation (B and C, p Ͻ 0.01 for ATP at 10 min versus 9 min pre-stimulation time points), while the acute drop in ATP levels after the second stimulations did not reach significance. D-F, Synaptic transmission at individual boutons was assessed using a pH-sensitive GFP targeted to synaptic vesicles (VGLUT1-pHluorin), again in pyruvate buffer, as well as 2DG and IAA to force reliance on glycolysis. Neither ␣syn KO (D) or shRNA against ␣syn (E, F) affected synaptic vesicle cycling after repeated stimulation (10 Hz‫06ء‬ s, blue lines). Bar graph confirms that shRNA decreased ␣syn levels by immunofluorescence (E) (compilation of three experiments, n ϭ 10 -12 coverslips/group with 10 -15 cells/coverslip). NS for extent of endocytosis [(amplitude endocytosis)/(amplitude exocytosis)] versus respective control by two-way ANOVA and Sidak's posthoc test. All graphs show mean Ϯ SEM.
Negative Results in peripheral tissues in PD (Beach et al. 2010). However, there is very little information on if ␣syn expression in these areas impacts metabolic functions. Because ␣synlowering therapies will likely lower ␣syn levels throughout the brain and in peripheral tissues, we also examined if interactions between ␣syn and mitochondria affect energy metabolism on a whole-body level using metaboliccage analyses and CLAMS (Columbus Instruments). At 6 months of age, ␣syn KO mice had total and lean body masses similar to controls, as assessed by EchoMRI (Fig.  5A). Food consumption (Fig. 5B) and total locomotor activity (Fig. 5C) were also unchanged, although the assessment of activity lacked sensitivity due to high variability, and there was a trend toward less movement in the dark cycle in the ␣syn KO group. However, importantly, ␣syn KO did not affect oxygen consumption (VO 2 ), carbon dioxide production (VCO 2 ), or the respiratory-exchange ratio (RER; [dot]VCO 2 /VO 2 ) in either the light or dark cycle (Fig 5D-F). Taken together, these data strongly suggest that loss of ␣syn does not impact total energy consumption in mice.

Discussion
␣Syn likely plays a central role in the pathogenesis of sporadic PD. Mitochondria are also compromised in PD, so therapeutic lowering of ␣syn will be done in the context of damaged mitochondria. In addition, increased ␣syn disrupts a range of mitochondrial functions, suggesting that decreasing ␣syn also influences mitochondrial function. However, here, we show that loss of ␣syn does not significantly impact the intrinsic bioenergetic function of mitochondria (i.e., respiration and ATP levels) in rodent neurons, even regionally at the synapse where ␣syn concentrates. Loss of ␣syn also fails to influence the local-  )) were scored with regard to synaptic vesicle cycling response at each stimulus burst. The stimulus burst at which the response ЉfailedЉ was recorded, and data were plotted as survival curves. Boutons were scored as failed if stim ⌬F was Ͻ10% of the ⌬F from first stim, or if ⌬F 120s after stimulus was Ͼ33% of the peak Fstim-F0 value (ie endocytic failure). Wt and syn TKO boutons in rotenone without glucose progressively failed to respond at a similar rate (p ϭ 0.21 by Gehan-Breslow-Wilcoxon test). n ϭ 325 (wt) and 340 (TKO) boutons for rotenone/no glucose, 98 (wt) and 93 (TKO) boutons for glucose-containing Tyrode's. The average lifespan of boutons by coverslip in rotenone/no glucose was also similar (wt ϭ 3.75 Ϯ 0.253 and syn TKO ϭ 3.61 Ϯ 0.137). Adeno-associated viruses (AAVs) expressing mitochondria-targeted GFP (mitoGFP; green, to visualize mitochondria) and mCherry-Synaptophysin (red, to visualize synaptic boutons) in DIO constructs (Sohal et al. 2009) that express only in Cre-expressing neurons were coinjected into the substantia nigra pars compacta (SNc) of 3-and 7-month-old DATcre control and ␣syn KO-DATcre mice. Mice were sacrificed one month later at 4 and 8 months of age, respectively. Roughly 60% of control and ␣syn KO synaptic boutons show mitochondria in the caudate putamen (CPu) (n ϭ 3-4 mice per group, where each value is the mean of 18 -21 fields; NS ϭ not significant by two-way ANOVA and Sidak's posthoc test). C, Western blot shows that ␣syn KO mice have similar levels of ␤syn as controls (n ϭ 3 mice per group; ‫ء‬p Ͻ 0.05, ‫‪p‬ءءء‬ Ͻ 0.001 by one-way ANOVA and Dunnet's posthoc test). D-G, AAVs expressing expressing mitoGFP and mCherry-Synaptophysin in DIO constructs were co-injected with shRNA scramble TagBFP or shRNA ␣syn TagBFP into the SNc of 7-month-old Dat icre/wt mice, and brains were harvested 6 weeks later. D-E, shRNA against ␣syn decreased ␣syn immunofluorescence ϳ60% versus shRNA scramble in DA neurons (n ϭ 3-4 mice, 57-86 cells per mouse (␣syn immunofluorescence level of individual cells for each mouse is shown in Fig. 4E-1); ‫ء‬p Ͻ 0.001 by unpaired two-tailed t test), identified by ization of mitochondria in DA axons or disrupt normal energy consumption in the whole body. Thus, our findings suggest that either ␣syn has no significant physiologic impact on mitochondrial bioenergetic function, or that any such functions are fully compensated for when lost or emerge only in the presence of specific stressors.
Increased ␣syn expression selectively inhibits complex I function (Devi et al. 2008;Chinta et al. 2010;Loeb et al. 2010) or the flux between complex I and III (Ellis et al. 2005;Devi et al. 2008). However, we do not yet understand the precise mechanisms of these effects or if the decrease in complex I function impacts energy production. Insufficient energy could also result from changes in the mass or distribution of mitochondria, even if the mitochondria have normal function. Indeed, increased mutant A53T ␣syn augments Parkin-dependent mitophagy in cortical neurons (Choubey et al. 2011) and the number of mitochondria in autophagosomes in midbrain DA neurons (Chinta et al. 2010). However, we found that decreased ␣syn did not affect the bioenergetic function of mitochondria, including regionally at the synapse, or the mass or distribution of mitochondria in nigrostriatal DA neurons in vivo.
The lack of effect of ␣synKO on bioenergetic function has three potential explanations. The first is that ␣syn KO does actually impair bioenergetic function but our studies failed to detect this due to insufficient sensitivity. Indeed, many of our assays lacked the sensitivity to reliably detect changes less than Ϸ10% -15%, and hence, subtle changes would not have been detected. However, all of the approaches used to assess bioenergetic function have been validated for their sensitivity to detect the effects of acute pharmacologic and chronic genetic inhibitors of respiration, and the uniform lack of significant changes across multiple complementary approaches provides strong evidence that ␣syn KO does not significantly impact bioenergetic function in the paradigms studied. Insufficient sensitivity could also have resulted if we assayed the wrong type of cell or the wrong subcellular compartment. In particular, our study focuses on neurons because ␣syn primarily localizes to neurons normally and accumulates in neurons in PD. Moreover, within neurons, we focused on synapses where most ␣syn KO localizes. To specifically assay nigrostriatal DA neurons we examine DA synaptosomes. However, recognizing the potential for artifact from the antibody bead-based isolation of DA synaptosomes, we also examined cortical synaposomes that were isolated without use of antibodies. Since synaptosomes likely have distinct bioenergetic properties from intact neurons, and even the standard isolation of continued tyrosine hydroxylase (TH), but had no effect on either the number of THϩ neurons as measured by stereology (F; n ϭ 6-8 mice per group; NS ϭ not significant (p ϭ 0.16) by unpaired two-tailed t test) or on the localization of mitochondria to synaptic boutons (G; n ϭ 3-4 mice per group, where each value is the mean of 6 -10 fields; NS ϭ not significant by unpaired two-tailed t test). All graphs show mean Ϯ SEM. Figure 5. ␣syn KO does not impact total body metabolism in mice. A, Body composition was measured using EchoMRI. Control and ␣syn KO had identical lean and fat body mass composition at 6 months of age. B-F, Body metabolism was assessed using a Comprehensive Lab Animal Monitoring System (CLAMS; Columbus Instruments). ␣syn KO and control mice had similar daily food intake (B) and locomotor activity (C). They also had similar VO 2 (D), VCO 2 (E), and respiratory exchange ratio (RER, ratio of VCO 2 produced and VO 2 used) (F) during both the light and dark cycles. n ϭ 6 mice per group; NS ϭ not significant by two-way ANOVA and Sidak's posthoc test. All graphs show mean Ϯ SEM. cortical synaptosomes may introduce artifacts, we assayed mitochondria-derived ATP levels in individual synaptic boutons of live neurons in complementary assays.
Insufficient sensitivity for an effect of decreasing ␣syn on bioenergetics could also have resulted if decreasing ␣syn affects only certain neuron types that we failed to assay. For instance, although we assayed DA neurons when possible, many of our assays focused on hippocampal neurons. However, we hypothesize lowering ␣syn will have similar effects across most neuron types. Indeed, ␣syn is present in neurons throughout the brain and, undoubtedly, has normal functions outside of nigrostriatal DA neurons. (Bendor et al. 2013;Benskey et al. 2016b) Thus, we must understand the impact of lowering ␣syn on nonDA neurons, in addition to DA neurons, since any ␣syn lowering therapy will likely be delivered to the entire brain. Moreover, the fact that human nigrostriatal DA neurons are susceptible to increased ␣syn in familial forms of PD does not mean they will also be more susceptible to ␣syn loss. In fact, it could mean just the opposite, and the underlying mechanisms of toxicity of increasing versus decreasing ␣syn could also be very different. For instance, increased ␣syn in some forms of PD may cause toxicity through a toxic gain of function of ␣syn, but loss of ␣syn could produce toxicity through loss of its normal function. Moreover, in PD, ␣syn accumulates in many nonDA neurons, including hippocampal neurons (e.g., see Hall et al. Brain, 2014), presumably contributing to the many non-motor features of the disease. As such, hippocampal neurons are an important and appropriate system to study normal ␣syn biology, and the wealth of preexisting studies in this neuron type facilitates interpretation of our results. However, the possibility remains that ␣syn-lowering compromises bioenergetics only in certain other neuron types.
A second possibility for a lack of effect of decreasing ␣syn on bioenergetic function is that ␣syn may normally interact with mitochondria and influence respiration, but the effects of ␣syn loss may be compensated for by other factors, such as ␤syn, which has similar, albeit less, potent effects on mitochondrial morphology when overexpressed Taschenberger et al. 2013). Indeed, the three syn isoforms can compensate for each other, because ␣syn, ␤syn, and ␥syn single-KO mice and ␣syn/␤syn double-KO mice have normal lifespans (Chandra et al. 2004), but triple syn-KO mice die early (Greten-Harrison et al. 2010). As evidence against this possibility, however, we found no effect of synTKO on mitochondria-derived ATP levels at the nerve terminal or on the rate of ATP consumption. In apparent contradiction, Ludtmann et al. (Ludtmann et al. 2016) recently reported that synTKO neurons have impaired bioenergetic function, suggesting that other syn isoforms compensate for certain ␣syn-effects on mitochondria. The reasons for this discrepancy are unclear, but could reflect differences between subcellular compartments. Specifically, our studies on synTKO neurons focused on changes at the nerve terminal where ␣syn and ␤syn accumulate. They also observed ATP changes using a mitochondriatargeted ATP sensor, while we examined cytosolic ATP levels, raising the possibility that synuclein isoforms might specifically alter ATP levels in the mitochondria. However, other methodological differences including the use of permeabilized versus intact neurons and the use of Mg 2ϩ homeostasis (versus our use of synaptic vesicle cycling) to assay for ATP consumption may also underlie some of the differences, and will require additional experimentation to resolve.
A third possibility is that ␣syn normally has minimal interactions with mitochondria and little effect on bioenergetic function. Although increased ␣syn disrupts mitochondrial morphology and function, these effects may reflect direct toxicity from the interaction of ␣syn oligomers with mitochondria (van Rooijen et al. 2009;Nakamura, 2013;Luth et al. 2014), and they may not occur under normal conditions. Also, while endogenous ␣syn can influence mitochondrial morphology (Kamp et al. 2010;Norris et al. 2015), and disrupt mitochondrial protein import (Di Maio et al. 2016), these changes may not be sufficiently robust to compromise respiration under basal conditions, although may be more prominent under pathologic conditions as in PD. Furthermore, although ␣syn KO mice resist MPTP and other mitochondrial toxins (Dauer et al. 2002;Klivenyi et al. 2006), and decreasing endogenous ␣syn levels protects against rotenone (Zharikov et al. 2015), the mechanism of these effects may be independent of bioenergetic function and the other parameters studied here.
Importantly, our study does not exclude the notion that lowering ␣syn may impact other mitochondrial functions, such as mitochondrial Ca 2ϩ import and buffering (Cali et al. 2012;Guardia-Laguarta et al. 2014), reactive oxygen species production, and lipid metabolism (Ellis et al. 2005;Cole et al. 2008;Nunnari and Suomalainen, 2012). Nonetheless, significant changes in any of these parameters would likely affect bioenergetic function and mitochondrial morphology, suggesting that any such changes would likely be subtle.
Our findings show that ␣syn can be safely lowered in mice without affecting mitochondrial bioenergetics. We believe that these studies suggest that therapeutically lowering ␣syn is unlikely to further disrupt mitochondrial bioenergetic function in PD. These results will need to be established in humans, especially if intended for therapies for PD patients, which will require many years. Moreover, lowering ␣syn could also produce toxicity through nonmitochondrial functions, such as disrupting synaptic vesicle release. Considering the rapid development of ␣synlowering therapies, we can expect to gain new insights into the safety and biological impact of ␣syn-lowering therapies over the coming decade.