This study was carried out in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocol was approved by the Institutional Animal Care and Use Committee of the University of California, San Francisco (protocol number: AN109510). All efforts were made to minimize suffering. LRRK1 KO mice were generated by the Michael J. Fox Foundation and both LRRK1 and 2 KO (Lin et al., 2009) mice obtained from Jackson Labs. Due to reduced survival on the C57Bl/6 background, heterozygous LRRK1 mice on this background were crossed with FVB mice to produce LRRK1 KO mice. LRRK1 KO and WT littermate controls were generated from heterozygous LRRK1 parents. We used LRRK1 heterozygous/LRRK2 KO breeding pairs to produce LRRK1 KO/LRRK2 KO mice and LRRK1 WT/LRRK2 KO littermates as control. Animals were genotyped by PCR.

To produce mouse hippocampal cultures, postnatal day 0 hippocampus was digested in papain (20 U/ml, Worthington), washed, triturated and plated on glass coverslips pre-coated with poly-L-lysine at a density of 700 cells/mm². Neurons were plated in minimal essential media containing 5% FBS, B27 supplement (Gibco), 21 mm glucose, 2 mM glutamax and Mito+ serum extender (VWR). Twenty-four hours after plating, 75% of the medium was removed and replaced with medium containing Neurobasal, B27 supplement, and 2 mM glutamax. After 7 days in culture, one third of the medium was exchanged, and 4 μM cytosine arabinoside added to prevent glial proliferation. Neurons were transfected using calcium phosphate at 6–7 days in vitro (DIV). To produce mixed cortical/striatal neuron cultures, the medial ganglionic eminence and cortex were isolated from embryonic day 16 mouse embryos, digested in trypsin (2.5%, Gibco) and DNase I (10 mg/ml, Roche), washed and triturated. Striatal neurons were transfected by electroporation (Amaxa), mixed with untransfected cortical neurons (3:2 striatal:cortical) and plated on glass coverslips pre-coated with poly-L-lysine at a density of 1100 cells/mm² in Neurobasal medium containing B27 supplement and 2 mM glutamax. Medium was exchanged and cytosine arabinoside added as described for hippocampal cultures.

Hippocampal neurons were transfected with the pH-sensitive GFP variant superecliptic pHluorin, targeted to the lumenal domain of synaptic vesicles by insertion into the large lumenal loop of vesicular glutamate transporter 1 (VGLUT1-pHluorin; Voglmaier et al., 2006). Striatal neurons were transfected with the vesicular GABA transporter (VGAT) tagged with pHluorin at the lumenal/extracellular C-terminus (VGAT-pHluorin; Santos et al., 2013). Neurons were imaged between DIV 14–17, as previously described (Nemani et al., 2010). Briefly, coverslips containing neurons were placed in an RC-21BRFS imaging chamber containing bipolar stimulating electrodes (Warner Instruments) in closed configuration. Neurons were continuously perfused (1 ml/min) at room temperature with Tyrode’s solution (in mM: 119 NaCl, 2.5 KCl, 2 CaCl₂, 2 MgCl₂, 30 glucose and 25 HEPES, pH 7.4) containing glutamate receptor antagonists 6-cyano-7 nitroquinoxaline-2, 3-dione (CNQX; 10 μM) and D, L-2-amino-5-phosphonovaleric acid (APV; 50 μM) and stimulated using trains of 1 ms current pulses to yield fields of 5–10 V/cm. We usually delivered a train of 600 stimuli at 10 Hz for 60 s, but in some experiments we delivered shorter trains at alternate frequencies, as indicated in the text. NH₄Cl buffer (in mM: 69 NaCl, 2.5 KCl, 2 MgCl₂, 2 CaCl₂, 50 NH₄Cl, 30 glucose, 25 HEPES, pH 7.4) was used to reveal total pHluorin fluorescence. Images were collected under epifluorescence illumination using a Nikon TE300 inverted microscope and xenon lamp with ET470/40 nm excitation and ET525/50 nm emission bandpass filters, a 63× 1.2 N.A. water objective, and a EM-CCD camera (Photometrics QuantEM:512SC) controlled by Metamorph software.

Initial rate of fluorescence increase was calculated by determining the slope of a line through the data point immediately prior to stimulation and the first two points during stimulation. To specifically assess exocytosis, we stimulated neurons at 10 or 20 Hz for 150 s in the presence of bafilomycin (0.6 μM, diluted from 1000× stock solutions in DMSO). Initial exocytic rate in bafilomycin was calculated by determining the slope of a line through the data points in the first 10 s after start of stimulation, when the increase in fluorescence is linear.

Endocytosis rate was calculated by normalizing curves to the peak response at the end of stimulation and fitting a single exponential decay curve to determine the decay constant, τ.

For destaining experiments, neurons were loaded by stimulation at 10 Hz for 2 min in 15 μM FM4–64 (ThermoFisher) in Tyrode’s solution, allowed to recover for 2 min, washed briefly with Tyrode’s solution containing 1 mM Advasep-7 (Biotium) followed by a longer wash with Tyrode’s solution, and images were collected as above using ET 470/40 nm excitation and 650 longpass emission filters. Curves were normalized to the initial peak fluorescence and fitted with a single exponential decay curve to determine the time constant τ, reflecting the loss of dye and by inference, the rate of exocytosis.

Neurons were imaged between DIV 14–17 in Tyrode’s solution with 10 μM CNQX and 50 μM APV at 37°C using a Zeiss LSM 510 confocal microscope with 63× oil immersion objective (NA 1.4) and 488 nm argon laser line. The pinhole was set to 2.5 μm to collect light from the entire bouton and images were collected every 400 ms at 1.5% transmission to reduce bleaching. For photobleaching, 40 consecutive scans were performed over 1 s at 100% transmission. The curves for fluorescence recovery after photobleaching (FRAP) were fit to a single exponential to calculate the time constant for recovery, τ.

Acute hippocampal slices were prepared from 3 week old mice. Three-hundred and fifty micrometer hippocampal slices were cut with a vibratome in aerated slicing solution (in mM, 228 sucrose, 2.5 KCl, 1 NaH₂PO₄, 7 MgSO₄, 0.5 CaCl₂, 26 NaHCO₃ and 11 dextrose) at 4°C, transferred to artificial cerebrospinal fluid (aCSF; in mM, 119 NaCl, 2.5 KCl, 1.3 MgSO₄, 2.5 CaCl₂, 26 NaHCO₃, 1 NaH₂PO₄ and 11 dextrose adjusted to 315 mosmol/L and pH 7.4) at 35°C and allowed to recover for a minimum of 1 h. Slices were visualized with an upright infrared differential interference contrast microscope and perfused with standard aCSF at room temperature. Stimulation (100 μs) pulse was delivered with a bipolar metal electrode (MX21AEW; FHC). Synaptic strength was quantified as the initial slope of field potentials recorded with aCSF-filled microelectrodes (1–2 MΩ).

Data are presented as mean ± SEM. For experiments involving multiple measurements among genotypes, we utilized 2-way analysis of variance (ANOVA), followed by post hoc Bonferonni tests. For experiments involving a single comparison between genotypes, we used two-tailed, unpaired t-tests. All statistics were calculated using Graphpad Prism.

To determine whether the loss of LRRK2 affects synaptic transmission, we used field recordings in hippocampal slices. We first examined the input-output curve, normalizing the slope of the field potential to the strength of the fiber volley. Importantly, the loss of LRRK2 does not influence the evoked synaptic response (Figure 1A). We then assessed release probability using the paired-pulse ratio (PPR), which shows no effect of the LRRK2 KO except at the 20 ms interstimulus interval, suggesting little or no effect on neurotransmitter release (Figure 1B).

To characterize the role of LRRK2 in synaptic vesicle cycling, we first used the styryl dye FM4–64. Like FM1–43, the red-shifted FM4–64 can be loaded into recycling synaptic vesicles by stimulation, and the subsequent stimulation-dependent destaining used to monitor synaptic vesicle exocytosis (Fernández-Alfonso and Ryan, 2004). Surprisingly, hippocampal neurons from WT and LRRK2 KO mice show no difference in either the rate or extent of destaining (Figure 2A), suggesting no role for LRRK2 in either synaptic vesicle exocytosis or the size of the recycling pool.

To monitor synaptic vesicle cycling with a more sensitive reporter that also has the potential to assess endocytosis, we used the pH-sensitive GFP variant superecliptic pHluorin, targeted to the lumenal domain of synaptic vesicles by insertion into a lumenal loop of vesicular glutamate transporter 1 (VGLUT1-pHluorin; Voglmaier et al., 2006). Quenched at the low pH of synaptic vesicles, the fluorescence increases with exocytosis and declines with the reacidification that follows endocytosis (Miesenböck et al., 1998). We transfected this reporter into hippocampal neurons from WT and LRRK2 KO mice, and imaged the change in fluorescence in response to stimulation, normalizing to the total pool of VGLUT1-pHluorin revealed by alkalinization in NH₄Cl (ΔF_NH4Cl). We again observed no difference between WT and LRRK2 KO neurons in the response to stimulation, including the initial rate of response or subsequent endocytosis (Figure 2B), with values very similar to those previously reported (Voglmaier et al., 2006; Nemani et al., 2010; Armbruster et al., 2013). However, the initial rate of fluorescence increase reflects the net contribution of both exocytosis and endocytosis. To more specifically assess the impact of LRRK2 on synaptic vesicle exocytosis, we repeated the experiments in the presence of the proton pump inhibitor bafilomycin to prevent the quenching of VGLUT1-pHluorin fluorescence by reacidification of internalized vesicles, so that the response to stimulation reflects only exocytosis (Figure 2C). In the presence of bafilomycin, there was no difference between WT and LRRK2 KO neurons in the rate or extent of exocytosis, which reflects the total pool of recycling vesicles, consistent with the lack of difference in initial rates observed without bafilomycin (Figures 2B,C).

LRRK2 may contribute to a presynaptic form of degeneration independent of synaptic vesicle cycling. Since association with membranes can influence the misfolding and aggregation of α-syn (Davidson et al., 1998; Lee et al., 2002), we used FRAP to assess a role for LRRK2 in its mobility, which presumably reflects membrane association. In previous work, we showed that after photobleaching a single presynaptic bouton, α-syn-GFP fluorescence recovers much more rapidly than the fluorescence of integral membrane proteins but more slowly than that of soluble GFP (Figures 3A,B), suggesting that α-syn binds specifically but weakly to presynaptic structures (Fortin et al., 2004; Unni et al., 2010). Consistent with this interpretation, the A30P mutation disrupts presynaptic accumulation of α-syn and accelerates recovery from photobleaching to the rate observed for soluble GFP (Fortin et al., 2004, 2005). We now find that the rate and extent of recovery for α-syn-GFP shows no difference between hippocampal neurons from WT and LRRK2 KO mice (Figures 3A,B). LRRK2 thus does not appear to influence the mobility of α-syn at the presynaptic terminal, and by inference, the association of α-syn with presynaptic membranes.

Since loss of the single LRRK protein in Drosophila impairs synaptic vesicle endocytosis (Matta et al., 2012), we considered the possibility that the two mammalian isoforms might be redundant. We therefore generated double KO (DKO) mice lacking both isoforms, and performed field recordings in hippocampal slices. The input-output curve comparing the slope of the field potential to the strength of the fiber volley in LRRK1/2 DKO slices showed a significant main effect of genotype vs. WT (2-way ANOVA, F_(1,271) = 5.094, p = 0.025), but post hoc tests showed no significant differences at any stimulation intensity (Figure 4A). Consistent with the absence of change in baseline synaptic transmission, release probability assessed using the PPR shows no difference from WT in the LRRK1/2 DKO (Figure 4B). We also examined the LRRK1/2 DKO using cultured hippocampal neurons transfected with VGLUT1-pHluorin. Loss of LRRK1 does not by itself alter the initial rate of fluorescence increase or compensatory post-stimulus endocytosis (Figure 4C). We then examined the effect of LRRK1 inactivation in the LRRK2 KO background (LRRK1WT/LRRK2 KO vs. LRRK1 KO/LRRK2 KO). Loss of both isoforms also failed to alter either the fluorescence increase or decrease after stimulation (Figure 4D), demonstrating that LRRK1 does not compensate for the loss of LRRK2.

Several studies have indicated high levels of native LRRK2 expression in the striatum (Melrose et al., 2006; Higashi et al., 2007; Westerlund et al., 2008; Giesert et al., 2013), which contains predominantly GABA neurons, suggesting that these cells may be more affected by the loss of LRRK2. Thus, we also examined the synaptic vesicle cycle in cultured striatal neurons from WT and LRRK2 KO mice, using the VGAT tagged with pHluorin at the lumenal/extracellular C-terminus (VGAT-pHluorin; Santos et al., 2013). The rate of both fluorescence increase and compensatory endocytosis after stimulation at 10 Hz for 60 s did not significantly change with loss of LRRK2, but endocytosis showed a trend to acceleration (Figure 5A). Previous studies using a shorter train of higher frequency stimulation (20 Hz for 15 s) have suggested that loss of LRRK2 inhibits endocytosis in rat striatal neurons (Arranz et al., 2015). Surprisingly, stimulation of mouse striatal neurons transfected with VGAT-pHluorin at 20 Hz for 15 s shows a significantly slower rate of fluorescence increase in LRRK2 KO than WT neurons, with a clear acceleration of compensatory endocytosis after the stimulus (Figure 5B). To determine whether loss of LRRK2 might also affect exocytosis in striatal neurons, we stimulated at 20 Hz in the presence of bafilomycin. Under these conditions, there was no difference in the rate or extent of exocytosis between WT and LRRK2 KO neurons (Figure 5C), supporting our conclusion that loss of LRRK2 does not directly affect the rate of exocytosis. Since stimulation at 20 Hz seemed to reveal a bigger difference in endocytosis between genotypes, we stimulated with sequential trains of 200 stimuli at 5–40 Hz (Figure 5D). As previously reported for hippocampal neurons, endocytosis was slower at higher stimulus frequencies in both genotypes (F_(3,88) = 6.753, p < 0.0004; Armbruster et al., 2013; Kononenko et al., 2014; Figure 5D). However, LRRK2 genotype affected the rate of endocytosis across all stimulation frequencies (F_(1,88) = 08.703, p = 0.0041). Although fluorescence appears to decay below baseline after 5 Hz stimulation, the baseline is in fact elevated due to incomplete decay after an initial stimulus (data not shown): in striatal neurons, fluorescence often fails to recover fully to the pre-stimulus baseline after a single stimulus train (Figures 5A,B); with subsequent stimulation, the fluorescence decays to the original pre-stimulus baseline (Figure 5D). In contrast to striatal neurons, stimulation of hippocampal neurons transfected with VGLUT1-pHluorin at 20 Hz for 15 s showed no difference between genotype in either the rate of fluorescence increase or post-stimulus endocytosis (Figure 2D). Thus, the effect of LRRK2 on rate of endocytosis depends on neuron identity, not stimulation frequency.