1. Neuroscience

α-Synuclein plasma membrane localization correlates with cellular phosphatidylinositol polyphosphate levels

  1. Reeba Susan Jacob
  2. Cédric Eichmann
  3. Alessandro Dema
  4. Davide Mercadante
  5. Philipp Selenko  Is a corresponding author
  1. Department of Biological Regulation, Weizmann Institute of Science, Israel
https://doi.org/10.7554/eLife.61951
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  1. Reeba Susan Jacob
  2. Cédric Eichmann
  3. Alessandro Dema
  4. Davide Mercadante
  5. Philipp Selenko
(2021)
α-Synuclein plasma membrane localization correlates with cellular phosphatidylinositol polyphosphate levels
eLife 10:e61951.
https://doi.org/10.7554/eLife.61951
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Abstract

The Parkinson’s disease protein α-synuclein (αSyn) promotes membrane fusion and fission by interacting with various negatively charged phospholipids. Despite postulated roles in endocytosis and exocytosis, plasma membrane (PM) interactions of αSyn are poorly understood. Here, we show that phosphatidylinositol 4,5-bisphosphate (PIP2) and phosphatidylinositol 3,4,5-trisphosphate (PIP3), two highly acidic components of inner PM leaflets, mediate PM localization of endogenous pools of αSyn in A2780, HeLa, SK-MEL-2, and differentiated and undifferentiated neuronal SH-SY5Y cells. We demonstrate that αSyn binds to reconstituted PIP2 membranes in a helical conformation in vitro and that PIP2 synthesizing kinases and hydrolyzing phosphatases reversibly redistribute αSyn in cells. We further delineate that αSyn-PM targeting follows phosphoinositide-3 kinase (PI3K)-dependent changes of cellular PIP2 and PIP3 levels, which collectively suggests that phosphatidylinositol polyphosphates contribute to αSyn’s function(s) at the plasma membrane.

Introduction

Aggregates of human α-synuclein (αSyn) constitute the main components of Lewy body inclusions in Parkinson’s disease (PD) and other synucleinopathies (Goedert et al., 2013). In the brain, αSyn is abundantly found in different types of neurons, where it primarily localizes to presynaptic terminals and regulates synaptic vesicle (SV) clustering and trafficking (Sulzer and Edwards, 2019). Isolated αSyn is disordered in solution, whereas residues 1–100 adopt extended or kinked helical conformations upon binding to membranes containing negatively charged phospholipids (Fusco et al., 2018). Complementary electrostatic contacts between lysine residues within αSyn’s N-terminal KTKEGV-repeats and acidic phospholipid headgroups align these α-helices on respective membrane surfaces (Snead and Eliezer, 2019). Membrane curvature (Middleton and Rhoades, 2010), lipid packing defects (Nuscher et al., 2004; Pranke et al., 2011) and fatty acid compositions (Fortin et al., 2004; Galvagnion, 2017) act as additional determinants for membrane binding. αSyn remodels target membranes (Varkey et al., 2010; Westphal and Chandra, 2013), which likely relates to its biological function(s) in vesicle docking, fusion and fission (Sulzer and Edwards, 2019). Furthermore, αSyn multimerization and aggregation may initiate at membrane surfaces, which holds important ramifications for possible cellular scenarios in PD (Galvagnion, 2017). Early αSyn oligomers bind to and disrupt cellular and reconstituted membranes (Reynolds et al., 2011; Fusco et al., 2017), whereas mature aggregates are found closely associated with membranous cell structures and intact organelles in cellular models of Lewy body inclusions (Mahul-Mellier et al., 2020) and in postmortem brain sections of PD patients (Shahmoradian et al., 2019).

Phosphatidylinositol phosphates (PIPs) are integral components of cell membranes and a universal class of acidic phospholipids with key functions in biology (Balla, 2013). Reversible phosphorylation of their inositol headgroups at positions 3, 4 and 5 generates seven types of PIPs, which act as selective binding sites for folded and disordered PIP-interaction domains (Balla, 2005). In eukaryotic cells, PIPs make up less than 2% of total phospholipids with phosphatidylinositol 4,5-bisphosphate, PI(4,5)P2 or PIP2 hereafter, as the most common species (McLaughlin et al., 2002). PIPs function as core determinants of organelle identity (Di Paolo and De Camilli, 2006). PIP2 is predominantly found at the inner leaflet of the plasma membrane (PM), where it acts as a signaling scaffold and protein-recruitment platform (McLaughlin and Murray, 2005). Carrying a negative net charge of −4 at pH 7 renders it more acidic than other cellular phospholipids such as phosphatidylserine (net charge −1) or phosphatidic acid (net charge −1) (Kooijman et al., 2009). Disordered PIP2-binding domains contain stretches of polybasic residues that establish complementary electrostatic contacts with the negatively charged PIP head groups (McLaughlin et al., 2002) reminiscent of how αSyn KTKEGV-lysines interact with acidic phospholipids (Dettmer, 2018). Indeed, αSyn has been shown to bind to reconstituted PIP2 vesicles in vitro (Narayanan et al., 2005). Phosphatidylinositol 3,4,5-trisphosphate, PI(3,4,5)P3 or PIP3 hereafter, harbors an additional phosphate group, which renders it even more acidic (net charge −5 at pH 7) (Kooijman et al., 2009). The steady-state abundance of PIP3 at the PM is low (Balla, 2013), but local levels increase dynamically in response to cell signaling, especially following phosphatidylinositide-3 kinase (PI3K) activation (Bilanges et al., 2019).

Here, we set out to investigate whether native αSyn interacted with PM PIP2 and PIP3 in mammalian cells. Using confocal and total internal reflection fluorescence (TIRF) microscopy, we show that endogenous αSyn forms discrete foci at the PM of human A2780, HeLa, SK-MEL-2 and neuronal SH-SY5Y cells. The abundance and localization of these foci correlate with pools of PM PIP2 and PIP3. We further delineate high-resolution insights into αSyn interactions with reconstituted PIP2 vesicles by nuclear magnetic resonance (NMR) spectroscopy and establish that αSyn binds PIP2 membranes in its characteristic, helical conformation.

Results

PM localization of endogenous αSyn

To determine the intracellular localization of αSyn, we selected a panel of human cell lines (A2780, HeLa, SH-SY5Y and SK-MEL-2) that expressed low but detectable amounts of the endogenous protein. Confocal immunofluorescence (IF) localization in A2780 cells with an antibody that specifically recognizes αSyn without cross-reacting with its β and γ isoforms (Figure 1—figure supplement 1A) revealed a speckled intracellular distribution with distinct αSyn foci at apical and basal PM regions (Figure 1A). We verified overall antibody specificity by downregulating αSyn expression via siRNA-mediated gene silencing, which established that αSyn foci corresponded to endogenous protein pools (Figure 1B and Figure 1—figure supplement 1B, Figure 1—figure supplement 1—source data 1). To investigate colocalization of αSyn and PIP2, we co-stained A2780 cells with αSyn and PIP2 antibodies, and imaged basal PM planes by IF microscopy (Figure 1C, top panel). In 10–20% of cases, we detected clear superpositions of αSyn and PIP2 signals, which we confirmed by measuring fluorescence intensity profiles over individual cell cross-sections (Figure 1D, top panel). We verified PM colocalization of αSyn with PIP2 in SH-SY5Y cells that we differentiated into dopaminergic-like neurons following a stringent protocol and stimulation with retinoic acid (RA) and brain-derived neurotrophic factor (BDNF) (Encinas et al., 2000; Figure 1—figure supplement 1C, Figure 1—figure supplement 1—source data 2). We found prominent pools of αSyn in expanded structures reminiscent of synaptic boutons along neurites, where they colocalized with PIP2 (Figure 1C, bottom panel, Figure 1D, and Figure 1—figure supplement 1D). These structures also stained positive for the presynaptic V-SNARE component syanptobrevin-2/VAMP2, a known binding partner of αSyn (Burré et al., 2010; Figure 1—figure supplement 1E).

Figure 1 with 2 supplements see all
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Plasma membrane (PM) localization of endogenous α-synuclein (αSyn).

(A) Immunofluorescence detection of endogenous αSyn in A2780 cells by confocal microscopy. PM stained with tetramethylrhodamine-wheat germ agglutinin (WGA) (left panel) or identified via GFP-PLCδ-PH (right panel). Representative apical and basal confocal planes are shown. Scale bars are 2 μm (left) and 10 μm (right). (B) αSyn-PM localization in A2780 cells following control (si NT) and targeted siRNA (si αSyn) knockdown. Phalloidin staining of F-actin marks cell boundaries. Scale bars are 10 μm. (C) Immunofluorescence detection of endogenous αSyn and phosphatidylinositol 4,5-bisphosphate (PIP2) at the PM in A2780 (top) and differentiated SH-SY5Y cells (bottom). Scale bars are 5 μm. (D) Spatially resolved αSyn (green) and PIP2 (red) fluorescence intensity profiles across the dotted lines in the closeup views. Resolved αSyn and PIP2 traces are marked with arrowheads. (E) αSyn-PM localization and quantification after transient green fluorescent protein (GFP) or GFP-PIPKIγ overexpression in A2780 and undifferentiated SH-SY5Y cells. GFP fluorescence identifies transfected cells. Scale bars are 10 μm. Box plots for αSyn immunofluorescence quantification. Data points represent n ~120 cells collected in four independent replicate experiments. Box dimensions represent the 25th and 75th percentiles, whiskers extend to the 5th and 95th percentiles. Data points beyond these values were considered outliers. Significance based on Student’s t tests as ∗∗∗p<0.001. See also Figure 1—source data 1.

Figure 1—source data 1

Raw data of αSyn PM localization upon PIPKinase expression.

https://cdn.elifesciences.org/articles/61951/elife-61951-fig1-data1-v2.xlsx

To test whether changes in cellular PIP2 levels affected αSyn abundance at the PM, we transiently overexpressed green fluorescent protein (GFP)-tagged phosphatidylinositol-4-phosphate 5-kinase PIPKIγ (Krauss et al., 2006). PIPKIγ localizes to the PM via a unique di-lysine motif in its activation loop (Kunz et al., 2000). Upon kinase expression, confirmed by GFP fluorescence, we detected increased amounts of αSyn at the PM of transfected cells (Figure 1E). By contrast, expression of GFP alone did not alter PM levels of αSyn. We obtained similar results in undifferentiated SH-SY5Y and HeLa cells (Figure 1E, Figure 1—source data 1 and Figure 1—figure supplement 2A, Figure 1—figure supplement 2—source data 1) and confirmed that transient PIPKIγ expression did not affect overall αSyn abundance (Figure 1—figure supplement 2B, Figure 1—figure supplement 2—source data 2). These findings suggested that PM localization of endogenous αSyn correlated with cellular PIP2 levels. To better resolve the presence of αSyn at the PM, we resorted to TIRF microscopy. Employing a narrow evanescent field depth of ~50 nm, we detected endogenous αSyn at PM foci in A2780, HeLa, SH-SY5Y and SK-MEL-2 cells, which correlated with total αSyn levels determined by semi-quantitative western blotting (Figure 1—figure supplement 2C, Figure 1—figure supplement 2—source data 3). Importantly, both imaging approaches were targeted toward detecting PM pools of αSyn and did not aim at interrogating cytoplasmic fractions of the endogenous protein.

αSyn binds reconstituted PIP2 vesicles

To test whether αSyn directly bound PIP2 membranes under physiological salt and pH conditions (150 mM, pH 7.0), we added N-terminally acetylated, 15N isotope-labeled αSyn to reconstituted PIP2 vesicles. Circular dichroism (CD) spectroscopy revealed characteristic helical signatures (Davidson et al., 1998; Jo et al., 2000; Figure 2A), whereas NMR experiments confirmed site-selective line broadening of N-terminal residues 1–100, confirming membrane binding (Bodner et al., 2009; Dikiy and Eliezer, 2014; Figure 2B and Figure 2—figure supplement 1, Figure 2—figure supplement 1—source data 1). In line with these observations, we detected remodeled PIP2 vesicles by negative-stain transmission electron microscopy (EM), manifested by tubular extrusions emanating from reconstituted specimens and agreeing with published findings on other membrane systems (Varkey et al., 2010; Westphal and Chandra, 2013; Figure 2A). Together, these results established that residues 1–100 of αSyn interacted with PIP2 vesicles in helical conformations that imposed membrane remodeling, whereas its 40 C-terminal residues did not engage in membrane binding and remained flexible and disordered. To gain further insights into αSyn-PIP2 interactions, we reconstituted phosphatidylcholine (PC)-PIP2 vesicles (100 nm diameter) at a fixed molar ratio of 13:1 (PC:PIP2) (Figure 2C). We added increasing amounts of these PC-PIP2 vesicles to αSyn and measured CD and dynamic light scattering (DLS) spectra of the resulting mixtures. Up to a ~50-fold molar excess of lipid to protein, αSyn interacted with PC-PIP2 vesicles in a helical conformation without disrupting the monodisperse nature of the specimens, that is, without membrane remodeling (Figure 2C and Figure 2—figure supplement 2A). In parallel, we performed NMR experiments on these samples and measured intensity changes of αSyn resonances in a residue-resolved manner (Figure 2D and Figure 2—figure supplement 2B, Figure 2—figure supplement 2—source data 1). Analyzing signal intensity ratios (I/I0) of unbound (I0) and PC-PIP2-bound αSyn (I), we found that residues 1–10 constituted the primary interaction sites, whereas residues 11–100 displayed progressively weaker membrane contacts. In agreement with our experiments on PIP2-only vesicles, we detected no contributions by C-terminal αSyn residues. These findings confirmed the tri-segmental nature of αSyn-PIP2 interactions and the importance of anchoring contacts by N-terminal αSyn residues, similar to other membrane systems (Bodner et al., 2009; Fusco et al., 2014; Fusco et al., 2016). To further validate our conclusions, we performed NMR experiments with mutant forms of αSyn in which we deleted residues 1–4 (ΔN) (Bartels et al., 2010), substituted Phe4 and Tyr39 with alanine (F4A-Y39A) (Lokappa et al., 2014), or oxidized αSyn Met1, Met5, Met116 and Met123 to methionine-sulfoxides (MetOx) (Maltsev et al., 2013; Figure 2—figure supplement 3A). In line with earlier reports, we did not observe binding to PC-PIP2 vesicles for any of these variants. Our results corroborated that PC-PIP2 interactions strongly depended on intact N-terminal αSyn residues, with critical contributions by Phe4 and Tyr39, and requiring Met1 and Met5 in their reduced states.

Figure 2 with 4 supplements see all
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α-Synuclein (αSyn) binding to reconstituted phosphatidylinositol 4,5-bisphosphate (PIP2) vesicles.

(A) Circular dichroism (CD) spectrum and negative-stain electron micrograph of αSyn-bound PIP2 vesicles (100%). Scale bar is 100 nm. (B) Overlay of 2D 1H-15N nuclear magnetic resonance (NMR) spectra of isolated αSyn in buffer (black) and bound to PIP2 vesicles (green). Remaining signals of C-terminal αSyn residues are labeled. (C) CD spectra of αSyn bound to phosphatidylcholine (PC)-PIP2 vesicles at increasing lipid-to-protein ratios (inset) and negative-stain electron micrograph of the αSyn:PC-PIP2 (1:50 protein:PIP2) sample. Scale bar is 100 nm. (D) NMR signal intensity ratios of bound (I) over unbound (I0) αSyn in the presence of different amounts of PC-PIP2 vesicles (equivalent to (C)). Only residues 1–40 are shown. (E) I/I0 of PC-PIP2 bound αSyn at 1:50 (green) and after addition of phospholipase C (PLC) (dark gray) and Ca2+ (light gray). Selected regions of 2D 1H-15N NMR spectra of PC-PIP2 bound αSyn (left, green) and in the presence of PLC (top right, dark gray) and Ca2+ (bottom right, light gray). Release of N-terminal αSyn residues from vesicles and reappearance of corresponding NMR signals are indicated for Asp2 (D2) as an example. (F) Hydrodynamic diameters of αSyn-bound PC-PIP2 vesicles before (green) and after PLC (dark gray) and Ca2+ (light gray) addition by dynamic light scattering. Errors were calculated based on measurements on three independent replicate samples.

In contrast to other lipids, PIPs offer attractive means to regulate the reversibility of αSyn-membrane interactions. Different charge states of PIPs can be generated from phosphatidylinositol (PI) precursors by action of PIP kinases and phosphatases (Matteis and Godi, 2004), or via PIP conversion by lipases such as phospholipase C (PLC) to produce soluble inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (Berridge and Irvine, 1984; Figure 2—figure supplement 3B). To investigate the reversibility of αSyn-PIP2 interactions, we prepared PC-PIP2 vesicles bound to 15N isotope-labeled αSyn to which we added catalytic amounts of unlabeled PLC. We reasoned that PLC will progressively hydrolyze PIP2 binding sites and, concomitantly, release αSyn. In turn, we expected to observe an increase of αSyn NMR signals corresponding to the fraction of accumulating, unbound protein molecules. Indeed, we detected the recovery of αSyn NMR signals upon PLC addition (Figure 2E and Figure 2—figure supplement 3C, Figure 2—figure supplement 3—source data 1). Next, we asked whether αSyn binding to PC-PIP2 vesicles was sensitive to calcium, a competitive inhibitor of many protein-PIP2 interactions (Bilkova et al., 2017). Whereas overall binding was greatly reduced, we found that the first 10 residues of αSyn displayed residual anchoring contacts with PC-PIP2 vesicles even at high (2.5 mM) calcium concentrations (Figure 2E and Figure 2—figure supplement 4A, Figure 2—figure supplement 4—source data 1), confirming earlier results on the stability of αSyn PC-PIP2 vesicle interactions in the presence of calcium (Narayanan et al., 2005). Notably, DLS measurements showed that hydrodynamic diameters of PC-PIP2 vesicles expanded upon PLC treatment and in the presence of calcium, irrespective of whether αSyn was bound (Figure 2F and Figure 2—figure supplement 4B). This further suggested that vesicle remodeling and concomitant curvature reductions did not abolish αSyn interactions. Finally, we sought to determine whether electrostatic interactions with acidic PIP headgroups alone mediated αSyn binding. To this end, we added a fourfold molar excess of free inositol polyphosphate (IP6) to 15N isotope-labeled αSyn. Surprisingly, we did not detect binding of αSyn to this highly negatively charged entity (Figure 2—figure supplement 4C), which insinuated that αSyn interactions with PIP-containing membranes required additional lipid contributions besides headgroup contacts.

αSyn-PM localization correlates with changes in PIP2-PIP3 levels

Following these results, we asked whether reversible αSyn-PIP2 interactions were present in cells. To answer this question, we transiently overexpressed different PM-targeted PIP phosphatases in A2780 cells and quantified PM localization of endogenous αSyn by confocal IF microscopy (Figure 3A, Figure 3—source data 1). Specifically, we expressed MTM1-mCherry-CAAX, which hydrolyzes PI(3)P to yield PI, INPP5E-mCherry-CAAX to produce PI(4)P from PIP2 and PTEN-mCherry-CAAX to create PIP2 from PI(3,4,5)P3, as described (Posor et al., 2013). In agreement with our hypothesis, only the conversion of PIP2 to PI(4)P by INPP5E led to a marked reduction of endogenous αSyn at the PM (Figure 3A). Together with earlier kinase results, these findings corroborated that PM localization of cellular αSyn was modulated by PIP2-specific enzymes. Next, we asked whether signaling-dependent activation of PI3K and concomitant accumulations of the even more negatively charged PIP3 (Bilanges et al., 2019) led to dynamic changes of αSyn abundance at the PM. To this end, we employed histamine stimulation of SK-MEL-2 cells that we transiently co-transfected with histamine 1 receptor 1 (H1R) and a PH-domain GFP-fusion construct of the general receptor of phosphoinositides 1 (GRP1) that specifically interacts with membrane PIP3 (Kavran et al., 1998). Because histamine-mediated PI3K activation also induces time-dependent secondary effects including PIP2 hydrolysis by PLC (Saheki et al., 2016), we monitored αSyn localization and PIP2-PIP3 levels in a time-resolved fashion by fixing SK-MEL-2 cells at 40, 85, 120, and 240 s after histamine addition (Figure 3B, Figure 3—source data 2). After 40 s, we observed an initial increase of PIP2 and PIP3 levels at the PM, which was mirrored by greater pools of endogenous αSyn at basal membrane regions. While PIP2 levels dropped at intermediate time points (85–120 s), likely due to PLC-mediated PIP2 hydrolysis, PIP3 concentrations were highest at 85 s and leveled off more slowly (120–240 s). Interestingly, PM-αSyn followed the observed PIP3 behavior in a remarkably similar manner. At later time points (240 s), we noted a significant redistribution of cellular PIP2 and PIP3 pools toward the edges of SK-MEL-2 cells, coinciding with the accumulation of bundled actin fibers and in line with expected PI3K-signaling-dependent rearrangements of the cytoskeleton (Bilanges et al., 2019). αSyn colocalization with these peripheral PIP2-PIP3 speckles was significantly higher than at earlier time points (Figure 3B and Figure 3—figure supplement 1A). We independently confirmed these results with single time point measurements by TIRF microscopy (Figure 3—figure supplement 1B, Figure 3—figure supplement 1—source data 1). To investigate whether other PI3K pathways caused similar effects, we stimulated SK-MEL-2 with insulin, which triggers PI3K activation via receptor tyrosine kinase signaling (Ruderman et al., 1990). We verified that SK-MEL-2 cells endogenously expressed the insulin-like growth factor-1 receptor β (IGF-1 Rβ) by western blotting (Figure 3—figure supplement 1C, Figure 3—figure supplement 1—source data 2). In support of our hypothesis, we measured increased αSyn-PM localization by TIRF microscopy upon insulin stimulation for 10 min (Figure 3—figure supplement 1D, Figure 3—figure supplement 1—source data 3). Given the short exposure times to histamine and insulin in these experiments, we reasoned that observed PM accumulations likely reflected enhanced recruitment of existing αSyn pools rather than de novo protein synthesis and PM targeting, thus providing further evidence that αSyn abundance at the PM correlated with signaling-dependent changes of PIP2 and PIP3 levels.

Figure 3 with 1 supplement see all
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Reversible α-synuclein (αSyn)-plasma membrane (PM) localization.

(A) Representative immunofluorescence localization of αSyn at basal A2780 PM planes by confocal microscopy. Cells transiently expressing PM-targeted, mCherry-tagged phosphatidylinositol phosphate (PIP) phosphatases, with mCherry fluorescence indicating successful transfection and phosphatase expression. Mutant, phosphatase-inactive INPP4A-mCherry-CAAX serves as the negative control (mCherry-ctrl, first panel). Scale bar is 10 μm. Box plots of αSyn immunofluorescence quantifications are shown on the right. Approximately 120 data points were collected per cell in four independent replicate experiments. Box dimensions represent the 25th and 75th percentiles, whiskers extend to the 5th and 95th percentiles. Data points beyond these values were considered outliers. Significance based on analysis of variance (ANOVA) tests with Bonferroni’s post-tests as NS >0.05; ∗p<0.05; ∗∗∗p<0.001. See also Figure 3—source data 1. (B) Time-course experiments following histamine stimulation of SK-MEL-2 cells transiently expressing histamine 1 receptor (H1R) and GRP1-PH. Immunofluorescence detection of endogenous phosphatidylinositol 4,5-bisphosphate (PIP2) and αSyn by confocal microscopy of basal PM regions. GRP1-PH GFP-signals report on the presence of PIP3. Phalloidin staining of F-actin marks cell boundaries. Scale bar is 10 μm. Box plots represent data points collected per cell (n ~ 80) from a single experiment, but representative of three independent experiments with similar results. Significance based on ANOVA tests with Bonferroni’s post-tests as NS >0.05; ∗p<0.05; ∗∗p<0.01; ∗∗∗p<0.001. See also Figure 3—source data 1.

Figure 3—source data 1

Quantification of αSyn PM localization following PIP phosphatase expression.

https://cdn.elifesciences.org/articles/61951/elife-61951-fig3-data1-v2.xlsx
Figure 3—source data 2

Quantification of αSyn PM localization after histamine stimulation.

https://cdn.elifesciences.org/articles/61951/elife-61951-fig3-data2-v2.xlsx

Discussion

Our results establish that clusters of endogenous αSyn are found at the PM of human A2780, HeLa, SK-MEL-2 and SH-SY5Y cells, where their abundance correlates with PIP2 levels (Figure 1). Specifically, we show that targeted overexpression of the PIP2-generating kinase PIPKIγ increases αSyn at the PM (Figure 1E), whereas the PIP2-specific phosphatase INPP5E reduces the amount of PM αSyn (Figure 3A). We further demonstrate that PIP3-dependent histamine and insulin signaling redistributes αSyn to the PM (Figure 3B and Figure 3—figure supplement 1), which collectively suggests that changes in PM PIP2 and PIP3 levels affect intracellular αSyn localization in a dynamic and reversible manner. Aiming for a stringent analysis, we investigated αSyn-PM interactions at strictly native, endogenous protein levels and intentionally refrained from transient or stable overexpression to not confound our analysis with non-physiological off-target effects. Moreover, we chose to study αSyn in an unaltered sequence context, that is, without modifying the protein with fluorescent dyes or fusion moieties. These requirements precluded live-cell imaging experiments to determine PM-localization kinetics, although histamine and insulin stimulation experiments suggest that endogenous αSyn pools redistribute readily. While we cannot rule out that additional secondary protein–protein interactions contribute to PM targeting, we demonstrate that αSyn directly interacts with reconstituted PIP2 vesicles in vitro (Figure 2). Importantly, the biophysical characteristics of these interactions are indistinguishable from other previously identified, negatively charged membrane systems with primary contacts by N-terminal αSyn residues 1–10 and progressively weaker interactions along residues 11–100. The last 40, C-terminal residues of αSyn are not involved in PIP2 membrane binding, similar to all other reconstituted vesicular or planar lipid surface interactions studied thus far (Bodner et al., 2009; Fusco et al., 2014; Perrin et al., 2000). Based on the known preferences for negatively charged phospholipids, PIP2 and PIP3 constitute intuitive αSyn binding partners. Not only because of their highly acidic nature (Kooijman et al., 2009), but also because of their acyl chain compositions containing saturated stearic-(18:0) and polyunsaturated arachidonic acids (20:4), the latter conferring ‘shallow’ lipid packing defects (Bigay and Antonny, 2012) that are ideally suited to accommodate αSyn’s helical conformation(s) (Pranke et al., 2011; Pinot et al., 2014). Thus, from a biophysical point of view, phosphatidylinositol polyphosphates satisfy many of the known requirements for efficient αSyn membrane binding. From a biological point of view, PIPs are ubiquitously expressed and stringently required for exocytosis and endocytosis, especially in neurons, where highly abundant PIP2 and PIP3pools (up to ~6 mol%) mark SV uptake and release sites (James et al., 2008). Multiple PIP-binding proteins mediate key steps in SV transmission and recycling (Di Paolo et al., 2004; Milosevic et al., 2005) and, although, αSyn has been implicated in synaptic exocytosis and endocytosis, its role(s) in these processes is ill defined (Huang et al., 2019).

A2780, HeLa, SH-SY5Y and SK-MEL-2 cells are poor surrogates for primary neurons and discussing our results in relation to possible scenarios at the synapse is futile. Endogenous levels of αSyn in the tested cell lines are low, particularly in comparison to presynaptic boutons, where αSyn concentrations reach up to 50 μM (Wilhelm et al., 2014). Similarly, the abundance of PIP2 and PIP3 is much smaller than at presynaptic terminals (James et al., 2008). Hence, αSyn-PIP scenarios in the tested cell lines and in synaptic boutons are at opposite ends of protein and lipid concentration scales. Nonetheless, we believe that key conclusions of our study are generally valid, especially because correlated localizations are equally prominent in non-neuronal cells. The affinity of αSyn to PIP2 vesicles has been reported to be in the low μM range (Narayanan and Scarlata, 2001), similar to most other reconstituted membrane systems containing negatively charged phospholipids (Middleton and Rhoades, 2010; Narayanan et al., 2005; Bodner et al., 2009; Dikiy and Eliezer, 2014; Fusco et al., 2014). In comparison, average dissociation constants for canonical PIP-binding scaffolds such as PH, C2, FYVE, and ENTH domains vary between μM and mM (Balla, 2005; McLaughlin et al., 2002). By contrast, disordered polybasic PIP-binding motifs target negatively charged membranes with much weaker affinities and in a non-discriminatory fashion based on complementary electrostatic interactions (McLaughlin and Murray, 2005). αSyn-PIP binding may define a third class of interactions that is comparable in strength to folded protein domains, but driven, to large parts, by electrostatic contacts similar to those of polybasic motifs (Galvagnion, 2017). Based on these affinity considerations, we speculate that αSyn may successfully compete for cellular PIP2-PIP3 binding sites with other proteins, particularly when their abundance is in a comparable range. For binding scenarios at presynaptic terminals, this is likely the case.

Our findings are additionally supported by recent data showing that intracellular αSyn concentrations directly influenced cellular PIP2 levels and that protein reduction diminished PIP2 abundance, whereas αSyn overexpression increased PIP2 synthesis and produced significantly elongated axons in primary cortical neurons (Schechter et al., 2020a). Conspicuously, these effects depended on αSyn’s ability to interact with membranes and were absent in a membrane-binding-deficient mutant (i.e., K10E-K12E) (Schechter et al., 2020a). Because PM expansions require dedicated cycles of endocytosis and exocytosis (Pfenninger, 2009), αSyn-PIP interactions may contribute to both types of processes, as has been suggested earlier (Lautenschläger et al., 2017). PM-specific αSyn-lipid interactions were additionally confirmed by ‘unroofing’ experiments in related SK-MEL-28 cells (Kaur and Lee, 2020), where endogenous protein pools colocalized with members of the exocytosis machinery including the known αSyn binding partners Rab3A (Chen et al., 2013) and synaptobrevin-2/VAMP2 (Burré et al., 2010). Two other studies implicated αSyn and αSyn-PIP2 interactions in clathrin assembly and clathrin-mediated endocytosis, respectively (Vargas et al., 2020; Schechter et al., 2020b), which further strengthens the notion that phosphatidylinositol polyphosphates contribute to αSyn functions at the PM.

Materials and methods

Key resources table
Reagent type
(species) or resource		DesignationSource or referenceIdentifiersAdditional information
Cell line (Homo sapiens)A2780Sigma-AldrichCat# 93112519
RRID:CVCL_0134
Cell line (Homo sapiens)HeLaSigma-AldrichCat# 93021013
RRID:CVCL_0030
Cell line (Homo sapiens)SH-SY5YSigma-AldrichCat# 94030304
RRID:CVCL_0019
Cell line (Homo sapiens)SK-MEL-2Dr. Ronit Sharon
(Hebrew University, Israel) Schechter et al., 2020a
Strain, strain background (Escherichia coli)BL21 (DE3) StarThermo Fisher ScientificCat# C601003Chemically Competent Cells
AntibodyAnti-αSyn
(mouse monoclonal)		Santa CruzCat# sc69977
RRID:AB_1118910		IF (1:200)
WB (1:100)
AntibodyAnti-αSyn MJFR1
(rabbit monoclonal)		AbcamCat# ab138501
RRID:AB_2537217		WB (1:10,000)
AntibodyAnti-PI(4,5)P2
(mouse monoclonal)		Echelon BiosciencesCat# Z-P045
RRID:AB_427225		IF (1:100)
AntibodyAnti-VAMP2
(rabbit monoclonal)		Cell SignallingCat# 13508
RRID:AB_2798240		IF (1:200)
AntibodyAnti-beta actin
(mouse monoclonal)		AbcamCat# ab6276
RRID:AB_2223210		WB (1:5000)
AntibodyAnti-IGF-I Receptor ß
(D23H3) (rabbit monoclonal)		Cell SignallingCat # 9750
RRID:AB_10950969		WB (1:1000)
AntibodyAnti-mouse IgG Alexa 647 conjugated
(goat polyclonal)		AbcamCat# ab150119
RRID:AB_2811129		IF (1:1000)
AntibodyAnti-rabbit IgG Alexa 555 conjugated
(donkey polyclonal)		InvitrogenCat# A-31572
RRID:AB_162543		IF (1:1000)
AntibodyAnti-mouse IgG
HRP-conjugated
(goat polyclonal)		Sigma-AldrichCat# A9917
RRID:AB_258476		WB (1:10,000)
AntibodyAnti-rabbit IgG
HRP-conjugated
(goat polyclonal)		Jackson
Immuno Research Laboratories		Cat# 111-035-003
RRID:AB_2313567		WB (1:5000)
Recombinant DNA reagentEGFP-PLCδ1-PHDr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Várnai and Balla, 1998		PH domain, binds PI(4,5)P2 at PM
Recombinant DNA reagentEGFP-tagged phosphatidylinositol 4-phosphate 5-kinase type Iγ (PIPKIγ)Dr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Krauss et al., 2006		PIP kinase, creates PI(4,5)P2 at PM
Recombinant DNA reagentEGFP-tagged
PIPKIγ
D316A
(mutated)		This paper Krauss et al., 2006PIP kinase, inactive
Recombinant DNA reagentEGFP-tagged
PIPKIγ
K188A
(mutated)		This paper Krauss et al., 2006PIP kinase, inactive
Recombinant DNA reagentMTM1-mCherry-CAAXDr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Posor et al., 2013		PIP phosphatase, acts on PI(3)P Targeted to PM
Recombinant DNA reagentINPP5E-mCherry-CAAXDr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Posor et al., 2013		PIP phosphatase, acts on PI(4,5)P2Targeted to PM
Recombinant DNA reagentPTEN-mCherry-CAAXDr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Posor et al., 2013		PIP phosphatase, acts on PI(3,4,5)P3Targeted to PM
Recombinant DNA reagentINPP4A-mCherry-CAAX (mutated)Dr. Volker Haucke
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany) Posor et al., 2013		PIP phosphatase inactive
Targeted to PM
Recombinant DNA reagentHuman histamine 1 receptor (H1R)Dr. Ronit Sharon
(Hebrew University, Israel) Kumar et al., 2017		Human histamine 1 receptor
Recombinant DNA reagentGRP1-PH pEGFP-C1Addgene Kavran et al., 1998Plasmid# 71378
RRID:Addgene_71378		PH domain binds PI(3,4,5)P3
Sequence-based reagentPIPKIγ D316A_FwThis paperPCR primer
(forward)		GTTTCAAGATCATGGCCTACAGCCTGCTGC
Sequence-based reagentPIPKIγ D316A_RvThis paperPCR primer
(reverse)		GCAGCAGGCTGTAGGCCATGATCTTGAAAC
Sequence-based reagentPIPKIγ K188A_FwThis paperPCR primer
(forward)		GTTCATCATCGCCACCGTCATGCACAAGGAGG
Sequence-based reagentPIPKIγ K188A_RvThis paperPCR primer
(reverse)		TCGTCGTCGCTGGTGACG
Peptide, recombinant proteinN-terminally acetylated αSynThis paper Theillet et al., 2016Purified from E. coli BL21 (DE3) Star
Peptide, recombinant proteinN-terminally truncated (ΔN) αSynThis paper Theillet et al., 2016Purified from E. coli BL21 (DE3) Star
Peptide, recombinant proteinN-terminally acetylated αSyn (F4A-Y39A) mutatedThis paper Theillet et al., 2016Purified from E. coli BL21 (DE3) Star
Commercial assay or kitQ5 Site-Directed Mutagenesis KitNew England BioLabsCat# E0554S
Commercial assay or kitBCA protein quantification kitThermo FisherCat# 23227
Commercial assay or kitSuperSignal West Pico PLUS Chemiluminescent SubstrateThermo FisherCat# 34579
Chemical compound, drugAll-trans retinoic acidSigma-AldrichCat# R2625
Chemical compound, drugRecombinant human/murine/rat BDNFPeprotechCat# 450-02
Software, algorithmImage Analysis
FIJI		imagej.net/Fiji Schindelin et al., 2012RRID:SCR_002285
Software, algorithmMulti-dimensional NMR data processing PROSADr. Peter Güntert
Goethe-University Frankfurt am Main, Germany
Güntert et al., 1992
Software, algorithmComputer-aided NMR resonance assignment CARAcara.nmr.chPhD thesis Rochus Keller
ETH Nr. 15947
OthersLipofectamine 3000Thermo Fisher ScientificCat# L3000015
OthersTransIT-X2Mirus BioCat# MIR 6000
OthersDOPCAvanti Polar LipidsCat# 850375
OthersBrain PI(4,5)P2Avanti Polar LipidsCat# 840046
OthersIP6Dr. Dorothea Fiedler
(Leibniz Institute of Molecular Pharmacology,
FMP-Berlin, Germany)		In-house synthesis
OthersPLC from Clostridium perfringens (Clostridium welchii)Sigma-AldrichCat# 9001-86-9

Mammalian cell lines and growth media

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All cells lines used in this study are described in the Key resources table. Cells were grown in humidified 5% (v/v) CO2 incubators at 37 °C in the following media supplemented with 10% (v/v) fetal bovine serum: RPMI 1640 (A2780), low glucose Dulbecco's modified eagle medium (DMEM)(HeLa), DMEM-Ham’s F-12 (SH-SY5Y), and minimum essential medium (MEM) with 1% non-essential amino acids and 2 mM glutamine (SK-MEL-2). Cells were split at 70–80% confluence with a passage number below 20 for all experiments. All cell lines were routinely tested for being mycoplasma free.

Transient cell transfections

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A2780 cells were seeded on fibronectin (Sigma-Aldrich, USA) coated 18 mm cover slips in 12-well plates at a density of 3 × 105 cells. Cells were transfected using Lipofectamine 3000 (Thermo Fisher, USA) according to the manufacturer's instructions. Undifferentiated SH-SY5Y and SK-MEL-2 cells were seeded on 18 mm coverslips at a density of 2 × 105 cells and transfected using TransIT-X2 (Mirus Bio, USA) according to the manufacturer's instructions. Details of transfection plasmids and mutagenesis primers are provided in the Key Resources Table. Kinase-inactive PIPKIγ mutants were generated with the Q5 site-directed mutagenesis kit (New England BioLabs, USA). Mutant PIPKIγ was confirmed by DNA sequencing. 1 μg of plasmids was used in all cases. Following transfection, cells were grown for 24 hr before analysis.

siRNA knockdown experiments

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Commercial siRNA mixtures against human αSyn (Dharmacon, USA, ON-TARGET plus human SNCA, cat.# L-002000-00-0005) and a non-targeted control (cat.# D-001810-10-05) were used. A2780 cells were seeded at a density of 6 × 105 cells and transfected with 1.7 μg of the respective siRNA mixtures using Lipofectamine 3000 according to the manufacturer’s instructions. After transfection, cells were grown for 48 hr before analysis.

SH-SY5Y differentiation

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Stringent SH-SY5Y differentiation was performed according to Encinas et al., 2000. In short, cells were seeded at a density of 2 × 105 in collagen-coated six-well plates. Twenty-four hours after seeding, cells were pre-differentiated with 10 μM of all trans RA in growth medium for 5 days. Subsequently, cells were cultured in serum-free medium supplemented with 50 ng/mL BDNF (Peprotech, Israel) for 7 days to obtain terminal neuronal differentiation. During the entire process, growth media were exchanged every 2–3 days.

Immunofluorescence

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For IF imaging of endogenous αSyn and expressed PIP-kinase/phosphatases, cells were washed 3 × 5 min with PBS and fixed in 4% (w/v) paraformaldehyde (PFA) for 15 min at room temperature (RT). For PM staining with 5 μg/mL Alexa Fluor 350/tetramethylrhodamine conjugated to wheat germ agglutinin (WGA) (Invitrogen, USA), cells were fixed and washed with PBS before application for 10 min at RT. Excess dye was washed off with PBS. For antibody staining, cells were permeabilized with 0.5% saponin in PBS for 10 min and blocked with 5% (w/v) bovine serum albumin (Sigma-Aldrich, USA) in PBS for 30 min. After blocking, cells were incubated with anti-αSyn antibody for 90 min at RT. After washing 3 × 5 min with PBS, cover slips were incubated with Alexa Fluor-tagged secondary antibody for 45 min at RT. Before confocal microscopy, cover slips were mounted with Immu-Mount (Thermo Fisher, USA), after 3 × 5 min PBS washes. IF detection of PI(4,5)P2 at the PM was performed according to Hammond et al., 2009) with slight modifications. A2780 and SK-MEL-2 cells were cultured on fibronectin-coated coverslips and pre-extracted in PHEM buffer (60 mM PIPES (piperazine-N,N′-bis(2-ethanesulfonic acid), 25 mM HEPES (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid), 5 mM EGTA (ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid), 1 mM MgCl2) to remove the majority of soluble cytoplasmic proteins. Cells were fixed with 4% PFA and 0.2% glutaraldehyde in PHEM buffer for 15 min at RT. All post-fixation steps until mounting were carried out at 4 °C. Washes were performed with ice-cold PIPES buffer (20 mM PIPES, pH 6.8, 137 mM NaCl, 2.7 mM KCl) to minimize damage to endogenous PIP moieties. Following fixation, cells were washed thrice in PIPES buffer containing 50 mM NH4Cl and subsequently blocked and permeabilized in PIPES buffer supplemented with 5% ‘normal goat serum’ and 0.5% saponin for 30 min. Post blocking, cells were incubated with anti-PI(4,5)P2 and anti-αSyn antibodies for 60 min, washed thrice, and incubated with Alexa Fluor 647 secondary antibody for 45 min. Before confocal microscopy, cover slips were mounted with Immu-Mount (Thermo Fisher, USA) after 3 × 5 min PIPES buffer washes. All primary and secondary antibody details are provided in the Key Resources Table.

Confocal microscopy

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Confocal microscopy imaging was performed on a Nikon spinning disk confocal microscope with an oil ×60 objective and additional ×1.5 magnification. Four channels in five optical sections from the basal PM plane were acquired with excitation wavelengths of 405 (blue, 50% laser power, for WGA), 488 (green, 20% for GFP), 568 (red, 20% for mCherry), and 647 (far-red, 20% for goat anti-mouse) with 200 ms exposure times. At least 25 images per biological replicate were collected and 3–4 replicates per experiment were analyzed.

TIRF microscopy

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For TIRF localization of endogenous αSyn at the PM, A2780, HeLa, SH-SY5Y, and SK-MEL-2 cells were cultured on 18 mm fibronectin-coated coverslips at a density of 2 × 105 cells for 24 hr and fixed with 4% PFA. After fixation, antibody detection was performed as described in the previous section. Coverslips for TIRF imaging were mounted in PBS after immunostaining and imaged on an Andor Dragonfly spinning disk microscope with a TIRF 100×/NA 1.45 oil objective. For TIRF detection of PM-proximal fluorescence signals, evanescent fields were kept at 50 nm in all experiments. Four lasers operating at 405 nm (15% laser power), 488 nm (20% laser power), 561 nm (20% laser power), and 647 nm (20% laser power) were used for fluorophore excitation along with 200 ms exposure time for image acquisition. At least 20 images per biological replicate were collected and three replicates per experiment were analyzed.

Histamine and insulin stimulation

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PI-3 kinase activity was stimulated by either insulin or histamine addition. For insulin stimulation via the endogenously expressed insulin-like growth factor-1 receptor β (IGF-1 Rβ) (Dricu et al., 1999), SK-MEL-2 cells were seeded on coverslips and starved in Hank's balanced salt solution (HBSS) for 18 hr, as described (Gray et al., 1999). 100 nM of insulin was added for 10 min and cells were fixed immediately afterwards. For histamine stimulation, SK-MEL-2 cells were seeded on 18 mm coverslips at a density of 2 × 105, transiently transfected with histamine 1 receptor (H1R) and serum-starved for 3 hr, as described in Mizuguchi et al., 2011. 500 μM of histamine was added and cells were fixed at indicated time points. All cell samples were further processed as previously outlined for TIRF procedures. F-Actin was detected by Phalloidin-Alexa Fluor 405 staining (1:400, Invitrogen, USA) during secondary antibody incubation.

Image analysis and quantification

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Image analysis and quantification were performed in Fiji (Schindelin et al., 2012). For confocal image quantification, focal planes of apical and basal PMs were selected manually. Images were segmented based on GFP signals by automatic thresholding according to Huang and Wang, 1995. Threshold regions were marked as regions of interest (ROIs), copied to the far-red channel (αSyn IF), and fluorescence intensities were determined. In the box plots of Figure 1E, Figure 1—figure supplement 2A and Figure 3A, B, each ROI corresponds to a single cell and is represented as a data point. For TIRF data in Figure 3—figure supplement 1B, D, images were segmented based on Phalloidin signals via automated thresholding using the default algorithm in Fiji (Zonderland et al., 2019). Different than for confocal images in Figures 1 and 3, TIRF ROIs consist of multiple adjacent cells in a single frame that were copied to the far-red channel (αSyn IF). ROIs of less than 2 μm2 in size were excluded. The Fiji particle counting routine was used to determine the number of αSyn puncta in each ROI. The number of cells in each image was determined manually based on cell outlines marked by Phalloidin. In Figure 3—figure supplement 1B, D, data points in box plots were calculated by dividing the number of αSyn puncta per image by the cell count. All box plots depict median values (center lines) with box dimensions representing the 25th and 75th percentiles. Whiskers extend to 1.5 times the interquartile range and depict the 5th and 95th percentiles. Each box plot in Figure 1E, Figure 1—figure supplement 2A and Figure 3A corresponds to 110–120 data points combined from four independent biological replicates. Box plots in Figure 3B contain data points collected per cell (n ~ 80) from a single experiment but representative of three independent experiments with similar results. Box plots in Figure 3—figure supplement 1B, D contain data points from ~120 cells, combined from three independent biological replicates.

Statistical analysis

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For box plots, data points considered ‘outliers’ were determined based on the criteria defined in the Grubbs, 1969 outlier test and omitted. Analysis of variance tests with Bonferroni’s post-tests (Armstrong, 2014; Dunn, 1961) were used to determine the statistical significance of experiments with more than two samples, whereas Student’s t tests were performed to assess statistical differences between samples (Kalpić et al., 2011). Significance is given as NS >0.05; ∗p<0.05; ∗∗p<0.01; ∗∗∗p<0.001. Absolute p values are given in the respective Source data files.

Cell lysate preparation

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Lysates of A2780, HeLa, SH-SY5Y, and SK-MEL-2 cell lines were prepared by detaching ~5–10 million cells with trypsin/EDTA (0.05%/0.02%) and harvested by centrifugation at 130 × g for 5 min at 25 °C. Sedimented cells were washed once with PBS, counted on a haemo-cytometer, and pelleted again by centrifugation. After resuspending cells in PBS with proteinase inhibitor cocktail (Roche, Switzerland), yielding a cell count of 2 × 107 cells/mL, they were lysed by repeated freeze–thaw cycles. Lysates were cleared by centrifugation at 16,000 × g for 30 min. Supernatants were removed, and total protein concentration was measured with a bicinchoninic acid (BCA) assay kit (Thermo Fisher, USA). For western blotting, 25 µg of protein (per lane, Figure 1—figure supplement 1C) or 50 µg of protein (per lane, all other figures) were applied onto SDS-PAGE.

Western blotting

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Cell lysates and recombinant protein samples were boiled in Laemmli buffer for 10 min before SDS-PAGE separation on commercial, precast 4–18% gradient gels (BioRad, USA). Recombinant N-terminally acetylated α-, β-, and γ-Syn were loaded as reference inputs at specified concentrations (see 'Recombinant protein expression and purification'). Proteins were transferred onto polyvinylidene fluoride (PVDF) membranes and fixed with 4% (w/v) PFA in phosphate-buffered saline (PBS) for 1 hr (Lee and Kamitani, 2011). Membranes were washed 2× with PBS, 2× with tris-buffered saline with 0.1% tween 20 (TBST), and blocked in 5% milk-TBST for 1 hr. After blocking, blots were incubated with primary antibodies overnight at 4 °C. Membranes were washed and probed with horseradish peroxidase (HRP)-conjugated secondary antibodies for 1 hr. All antibody details including respective dilutions are provided in the Key Resource Table. Membranes were developed using the SuperSignal West Pico Plus reagent (Thermo Fisher, USA), and luminescence signals were detected on a BioRad Molecular Imager.

Western blot quantification

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Intensities of αSyn and β-actin bands were quantified using the ImageLab software (BioRad, USA). αSyn reference input was used to generate a standard curve. For cell lysate samples, αSyn intensity was normalized according to the β-actin signal and cell lysate concentrations were calculated with respect to αSyn standards. Error bars denote background (noise).

Recombinant protein expression and purification

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15N isotope-labeled, N-terminally acetylated, human αSyn was produced by co-expressing PT7-7 plasmids with yeast N-acetyltransferase complex B (NatB) (Johnson et al., 2010) in Escherichia coli BL21 Star (DE3) cells in M9 minimal medium supplemented with 0.5 g/L of 15NH4Cl (Sigma-Aldrich, USA). Unlabeled α-, β- and γ-Syn were produced in Luria-Bertani (LB) medium. Generation of αSyn mutants ΔN and F4A-Y39A was described previously (Theillet et al., 2016). For recombinant protein purification under non-denaturing conditions, we followed the protocol by Theillet et al., 2016. Purification of αSyn F4A-Y39A was identical to wild-type αSyn. Lacking the N-terminal substrate specificity for NatB, αSyn ΔN was produced in its non-acetylated form and purified as the wild-type protein. Methionine-oxidized 15N isotope-labeled wild-type αSyn was expressed and purified as described (Binolfi et al., 2016). Protein samples were concentrated to 1–1.2 mM in NMR buffer (25 mM sodium phosphate, 150 mM NaCl) at pH 7.0. Protein concentrations were determined spectrophotometrically by UV absorbance measurements at 280 nm with ε = 5960 M−1 cm−1 for α-, β-Syn ΔN, and methionine-oxidized αSyn. For αSyn F4A-Y39A and γ-Syn, ε = 4470 and 1490 M−1 cm−1 were used. Final aliquots of protein stock solutions were snap frozen in liquid nitrogen and stored at −80 °C until use.

Reconstituted PI(4,5)P2 vesicles

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Phospholipids were purchased from Avanti Polar Lipids (USA). Large unilamellar vesicles (LUVs, 100 nm) were prepared from 100% brain (porcine) PIP2. A thin lipid film was formed in a glass vial by gently drying 1 mg of PIP2 in chloroform-methanol under a stream of nitrogen. To remove residual traces of organic solvent, the lipid film was placed under vacuum overnight. 0.5 mL NMR buffer was then added to hydrate the lipid film for 1 hr at RT while agitating. After five freeze–thaw cycles on dry ice and incubation in a water bath at RT, the lipid suspension was sonicated at 4 °C for 20 min at 30% power settings (Bandelin, Germany). Resulting PIP2 LUVs (2 mg/mL) were used immediately. αSyn:PIP2 molar ratios for sample preparations were calculated using a PIP2 lipid mass of 1096 Da. For PIP2 titration experiments, 1-, 5-, 10-, 15-, 20-, and 30-fold molar excess of lipids was added to 60 μM of 15N isotope-labeled, N-terminally acetylated αSyn (total volume 120 μL) and αSyn PIP2 samples were incubated for 45 min at RT before NMR and CD measurements. Following the same procedure, mixed PC-PIP2 (PC:PIP2 mass ratio 9:1) suspensions were prepared using 9 mg of 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC, 786 Da) and 1 mg PIP2. The dried lipid film was hydrated with 0.25 mL NMR buffer. The PC-PIP2 suspension was then extruded through polycarbonate membranes with a pore size of 100 nm according to the manufacturer’s instructions (mini-extruder, Avanti Polar Lipids) and resulting PC-PIP2 LUVs (40 mg/mL, PC:PIP2 molar ratio 13:1) were used immediately. For sample preparations, an approximate PC-PIP2 lipid mass of ~810 Da ((13/14) × 786 Da + (1/14) × 1096 Da) was used to calculate the αSyn:PC-PIP2 molar ratios. 15N isotope-labeled, N-terminally acetylated αSyn (60 μM) was incubated with ~80-, ~170-, ~340-, and ~680-fold molar excess of total PC-PIP2 lipids. αSyn PC-PIP2 samples (total volume 120 μL) were incubated for 45 min at RT before CD, NMR, and DLS experiments. Synthetic inositol hexaphosphate (IP6) was kindly provided by Dr. Dorothea Fiedler, Department of Chemical Biology, Leibniz Institute of Molecular Pharmacology (FMP-Berlin). Before NMR measurements, 50 μM αSyn was incubated with 200 μM IP6 in NMR buffer (total volume 120 μL) for 45 min at RT.

Phospholipase C reaction

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PLC was purchased from Sigma-Aldrich (USA) and the lyophilized powder was dissolved in NMR buffer at 1000 units (U)/mL. αSyn PC-PIP2 samples at ~680-fold molar excess of PC-PIP2 lipids (60 μM αSyn, ~40 mM PC-PIP2) were incubated while agitating at 37 °C for 45 min with 10 U of PLC and 1 mM phenylmethylsulfonyl fluoride (PMSF) in a total volume of 120 μL.

NMR spectroscopy

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For best comparison of protein reference and αSyn-lipid NMR data, final concentrations of 15N isotope-labeled, N-terminally acetylated αSyn samples were adjusted to 60 μM, supplemented with 5% D2O, and measured in 3 mm (diameter) Shigemi tubes in all cases. NMR experiments were acquired on a Bruker 600 MHz Avance spectrometer equipped with a cryogenically cooled proton-optimized 1H{13C/15N} TCI probe. Reference and αSyn-lipid NMR spectra were acquired with identical spectrometer settings and general acquisition parameters. Specifically, we employed 2D 1H-15N SOFAST HMQC NMR pulse sequences (Schanda et al., 2005) with 512 x 128 complex points for a sweep width of 28.0 ppm (15N) and 16.7 ppm (1H), 128 scans, 60 ms recycling delay at 283 K. Inspection of the highly pH-sensitive His50 (H50) 1H-15N chemical shift indicated that the sample pH changed from 7 to 6.5 during the PLC reaction (Figure 2E). To accurately delineate I/I0 values, we recorded reference NMR spectra at pH 6.5. All NMR spectra were processed with PROSA, zero-filled to four times the number of real points and processed without window function. Visualization and data analysis were carried out in CARA. NMR signal intensity ratios (I/I0) of isolated αSyn (I0) and in the presence of lipids (I) were determined for each residue by extracting maximal signal peak heights in the respective 2D 1H-15N NMR spectra.

CD spectroscopy

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NMR samples of isolated αSyn and αSyn in the presence of lipid vesicles were diluted with NMR buffer to a final protein concentration of 10 μM for CD measurements. CD spectra (200–250 nm) were collected on a Jasco J-720 CD spectropolarimeter in a 1 mm quartz cell at 25 °C. One replicate per sample was recorded. Six scans were averaged and blank samples (without αSyn) were subtracted from protein spectra to calculate the mean residue weight ellipticity (θMRW).

Dynamic light scattering

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DLS measurements were acquired on a Zetasizer Nano ZS (Malvern Instruments, UK) operating at a laser wavelength of 633 nm equipped with a Peltier temperature controller set to 25 °C. Data were collected on all NMR samples containing αSyn, isolated PC-PIP2 vesicles and PC-PIP2 vesicles in the presence of Ca2+ and PLC. Using the Malvern DTS software, mean hydrodynamic diameters were calculated from three replicates of the same sample in the intensity-weighted mode.

Negative-stain electron microscopy

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NMR samples of αSyn at ~30- and ~680-fold molar excess of PIP2 and PC-PIP2 lipids were diluted to a protein concentration of ~10 μM in NMR buffer. 5 μL aliquots were added to glow-discharged carbon-coated copper grids for 1 min. Excess liquid was removed with filter paper and grids were washed twice with H2O before staining with 2% (w/v) uranyl acetate for 15 s. Negative-stain transmission EM images were acquired on a Technai G2 TEM.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting files. Source data files have been provided for Figures 1,2,3 and all figure supplements.

References

    1. Dunn OJ
    (1961) Multiple comparisons among means
    Journal of the American Statistical Association 56:52–64.
    https://doi.org/10.1080/01621459.1961.10482090
  37. Book
    1. Kalpić D
    2. Hlupić N
    3. Lovrić M
    (2011) Student’s t-Tests
    In: Lovric M, editors. International Encyclopedia of Statistical Science. Springer. pp. 1–2.
    https://doi.org/10.1007/978-3-642-04898-2

Decision letter

  1. Axel T Brunger
    Reviewing Editor; Stanford University, United States
  2. Olga Boudker
    Senior Editor; Weill Cornell Medicine, United States
  3. David Eliezer
    Reviewer; Weill Cornell Medical College, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

Acceptance summary:

In this short report it is shown that phosphatidylinositol 4,5-bisphosphate (PIP2) and phosphatidylinositol 3,4,5-trisphosphate (PIP3), two highly acidic components of inner plasma membrane leaflets, mediate plasma membrane localization of endogenous pools of α-synuclein in A2780, HeLa, SH-SY5Y and SK-MEL-2 cells. Moreover, it is shown that PIP2 synthesizing kinases and hydrolyzing phosphatases can redistribute α-synclein in cells. In addition, structural details of the interactions of αSyn with PIP2 containing membranes have been revealed by NMR spectroscopy.

Decision letter after peer review:

Thank you for submitting your article "α-Synuclein plasma membrane localization correlates with cellular phosphatidylinositol polyphosphate levels" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Olga Boudker as the Senior Editor. The following individual involved in review of your submission has agreed to reveal their identity: David Eliezer (Reviewer #2).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

As the editors have judged that your manuscript is of interest, but as described below that additional experiments are required before it is published, we would like to draw your attention to changes in our revision policy that we have made in response to COVID-19 (https://elifesciences.org/articles/57162). First, because many researchers have temporarily lost access to the labs, we will give authors as much time as they need to submit revised manuscripts. We are also offering, if you choose, to post the manuscript to bioRxiv (if it is not already there) along with this decision letter and a formal designation that the manuscript is "in revision at eLife". Please let us know if you would like to pursue this option. (If your work is more suitable for medRxiv, you will need to post the preprint yourself, as the mechanisms for us to do so are still in development.)

Summary:

The normal behavior and function of α-synuclein remain poorly understood, at least in part because the majority of studies of this protein focus on its pathological self-assembly and deposition, processes that are thought to be critical for its role in neurodegeneration. Whether or not the latter is true, an improved understanding of the cell biology of α-synuclein is necessary to clarify when and how this protein deviates from its normal physiological contexts during the course of disease. To that end, understanding the cellular localization of the protein, (which has remained highly controversial) and to explore how it is regulated (which has hardly been studied) are important goals that are addressed by this study. Given the importance of PIPs in membrane trafficking pathways, and the growing consensus that α-synuclein functions in the arena of synaptic vesicle trafficking, uncovering a role for PIPs in attracting α-synuclein to membranes is a significant contribution to our understanding of this protein.

In this short report it is shown that phosphatidylinositol 4,5-bisphosphate (PIP2) and phosphatidylinositol 3,4,5-trisphosphate (PIP3), two highly acidic components of inner PM leaflets, mediate plasma membrane localization of endogenous pools of αSyn in A2780, HeLa, SH-SY5Y and SK-MEL-2 cells. Although it is already known that αSyn binds to PIP2 and other anionic lipids and that this binding induces α-helicity in αSyn this work additionally shows that PIP2 synthesizing kinases and hydrolyzing phosphatases can redistribute αSyn in cells. Moreover, structural details of the interactions of αSyn with PIP2 containing membranes have been revealed by NMR spectroscopy.

Essential revisions:

While overall, there is good evidence that αSyn targets PIP2 at the plasma membrane, the in vivo relevance remains unclear, in particular with neuronal activity-dependent roles of PIP2 in both, synaptic vesicle exo- and endocytosis, and of PIP3 in postsynaptic receptor trafficking. The reviewers are concerned about the cellular systems that were chosen to assess αSyn plasma membrane localization – mostly of non-neuronal origin – which the authors also highlight themselves. The authors have the neuronal SH-SY5Y cell line and produced little data with this line, but most of the experiments presented here were performed in A2780 (ovarian carcinoma), HeLa (cervical cancer) and SK-MEL-2 (skin metastasis) cells. If some of the experiments could be repeated in the neuroblastoma cell line, ideally differentiated into a more neuron-like shape, it would make the paper more compelling.

Figure 1E and Figure 1—figure supplement 1B: The authors describe increased amounts of αSyn at the PM by expression of PIPKIγ. Are total αSyn levels affected, and could this result in an increased pool on the PM? Please test by western blotting.

Figure 1—figure supplement 1D: An overlay or quantification of the localization of αSyn at the PM would be helpful. The overlap in A2780 cells seems to be high, but the overlap in SH-SY5Y and SK-MEL-2 cells is not particularly convincing.

Figure 2C: Please justify the use of 100 nm vesicles. If the PM should be mimicked, the vesicles should have a larger diameter.

Figure 3A: The quantification of αSyn intensity with expression of INPP5E is not reflected by the provided image where there is essentially no αSyn signal. Please provide a more representative image.

Figure 3A: The authors argue that αSyn has a specificity for PIP2 and PIP3, but why then are there no apparent changes between αSyn intensity upon expression of MTM1 and INPP5E (which result in production of π and PI4P, respectively) versus PTEN (which produces PIP2 from PIP3)? In addition, the authors describe that "only the conversion of PIP2 to PI4P by INPP5E led to a marked reduction" but their statistics show significant changes for all three phosphatases compared to mCherry only.

Figure 3: There may be a contribution of coincidental labeling. Could the authors reduce PIP2 and/or PIP3 levels as in panel 3A and repeat their analysis?

Are the observed CD spectra and NMR spectra shown in Figure 2 consistent with residues ~ 1-100 in an helical conformation and the remaining ~ 40 residues being unstructured under high PIP2 to αSyn ratios, similar to the solution NMR structure of micelle-bound αSyn (PDB ID 1XQ8) or similar to the solid state NMR studies of αSyn interactions with lipid bilayers by Fuso et al., Nature Comm, 2014? In other words, are residues 100-140 unstructured in all these studies? Please comment.

The experiments with calcium (Figure 2E) suggest that the first ten residues always interact with PC-PIP2 vesicles, although overall binding was greatly reduced. However, plasma membranes contain anionic lipids that may mitigate the effect of calcium that is observed in these experiments. Please comment.

https://doi.org/10.7554/eLife.61951.sa1

Author response

Essential revisions:

While overall, there is good evidence that αSyn targets PIP2 at the plasma membrane, the in vivo relevance remains unclear, in particular with neuronal activity-dependent roles of PIP2 in both, synaptic vesicle exo- and endocytosis, and of PIP3 in postsynaptic receptor trafficking. The reviewers are concerned about the cellular systems that were chosen to assess αSyn plasma membrane localization – mostly of non-neuronal origin – which the authors also highlight themselves. The authors have the neuronal SH-SY5Y cell line and produced little data with this line, but most of the experiments presented here were performed in A2780 (ovarian carcinoma), HeLa (cervical cancer) and SK-MEL-2 (skin metastasis) cells. If some of the experiments could be repeated in the neuroblastoma cell line, ideally differentiated into a more neuron-like shape, it would make the paper more compelling.

We have now included new figure panels depicting co-localization of endogenous αSyn with PIP2 in fully differentiated SH-SY5Y cells. Specifically, we followed a 14-day differentiation protocol that employs initial stimulation with retinoic acid (RA) for 5 days, followed by brain-derived neurotrophic factor (BDNF) treatment for 7 additional days. The procedure commonly accepted as the most stringent differentiation routine for SH-SY5Y cells1, especially in comparison to differentiation with RA only2. In line with expected results, we observed substantial neurite outgrowth and shrinkage of cell bodies, concomitant with increased expression of endogenous αSyn (new Figure 1—figure supplement 1C). We detected αSyn in cell bodies and neurites, although protein pools were particularly abundant along neurites where they colocalize with PIP2 in expanded structures reminiscent of synaptic boutons (new Figure 1C, D, lower panel). Indeed, co-staining for the vesicular SNARE-component synaptobrevin-2 (Syb2/VAMP2) confirmed that αSynPIP2 and αSyn-Syb2/VAMP2 localize to these structures (new Figure 1—figure supplement 1D, E). As the reviewers are obviously aware, Syb2/VAMP2 is one of the few “broadly accepted” binding partners of αSyn in presynaptic terminals3.

To the best of our knowledge, these are the first reported results of endogenous αSyn-PIP2 and Syb2/VAMP colocalization in fully differentiated SH-SY5Y cells.

We updated the manuscript accordingly:

Abstract: “… endogenous pools of αSyn in A2780, HeLa, SK-MEL-2 and differentiated and undifferentiated neuronal SH-SY5Y cells.”

Results: “We verified PM colocalization of αSyn with PIP2 in SH-SY5Y cells that we differentiated into dopaminergic-like neurons following a stringent protocol and stimulation with retinoic acid (RA) and brain-derived neurotrophic factor (BDNF)1 (Figure 1—figure supplement 1C). We found prominent pools of αSyn in expanded structures reminiscent of synaptic boutons along neurites, where they colocalized with PIP2 (Figure 1C, bottom panel, Figure 1D and Figure 1—figure supplement 1D). These structures also stained positive for the presynaptic V-SNARE component syanptobrevin2/VAMP2, a known binding partner of αSyn3 (Figure 1—figure supplement 1E).”

We also attempted to transfect fully differentiated SH-SY5Y cells with GFP-PIPKIγ to ask whether ectopic expression of the PIP2 kinase triggered localization of higher levels of endogenous αSyn at the PM. This is a challenging experiment in any kind of terminally differentiated cell line. We accomplished this goal after careful evaluation of various transfection agents and procedures. However, we were unable to perform consistent confocal imaging at equivalent planes for cell bodies and neurites given their highly irregular shapes and extended morphologies. Accordingly, we did not quantify αSynfluorescence on a per cell basis in differentiated SH-SY5Y cells, as we had done for A2780 and HeLa cells.

Instead, we extended the analysis of induced αSyn PM localization upon PIPKIγ expression in undifferentiated SH-SY5Y cells, which is now included in the new Figure 1E (right panel). Compared to the original submission, we increased the number of analyzed cells (n > 120) to derive quantitative information about the significance of the observed effect. We are happy to report that we obtained p-values < 0.001 (***) for this and all other cell lines indicating the general significance and reproducibility of the PMtargeting effect upon induced accumulation of PIP2 at the PM. We moved the analysis of HeLa cells to Figure 1—figure supplement 2A.

We updated the manuscript accordingly (see also the next question).

Results: “We obtained similar results in undifferentiated SH-SY5Y and HeLa cells (Figure 1E and Figure 1—figure supplement 2A) and confirmed that transient

PIPKIγ expression did not affect overall αSyn abundance (Figure 1—figure supplement 2B).”

Figure 1E and Figure 1—figure supplement 1B: The authors describe increased amounts of αSyn at the PM by expression of PIPKIγ. Are total αSyn levels affected, and could this result in an increased pool on the PM? Please test by western blotting.

We quantified levels of endogenous αSyn in A2780 and undifferentiated SHSY5Y cells upon expression of wild-type (WT) GFP-PIPKIγ and GFP only. In addition, we included two PIPKIγ mutants (D316A and K188A), which are catalytically inactive but targeted to the PM similar to WT PIPKIγ. As can be appreciated from the Western blots (new Figure 1 —figure supplement 2B), transient expression of any of these constructs did not alter the global levels of endogenous αSyn. We include this analysis in the revised manuscript and updated the text accordingly (see above).

Figure 1—figure supplement 1D: An overlay or quantification of the localization of αSyn at the PM would be helpful. The overlap in A2780 cells seems to be high, but the overlap in SH-SY5Y and SK-MEL-2 cells is not particularly convincing.

There appears to be a misunderstanding as to what the PM-dye in these images shows. This panel confirms that the chosen TIRF fields correctly represent the PM, which is identified by staining with tetramethylrhodamine-WGA (as specified in the figure legend). No co-localization between this dye and αSyn is expected. The dye simply identifies the basal plane of the PM, whereas αSyn signals show pools of endogenous protein at the PM and cell type-specific variations thereof. Brighter patches of PM-dye correspond to preferred incorporation of tetramethylrhodamine-WGA, which does not correlate with (nor represent) PIP2 or αSyn localization. Irrespectively, we selected new image panels to better reflect unrelated WGA and αSyn “staining” in the revised figure (now Figure 1—figure supplement 2C)

Figure 2C: Please justify the use of 100 nm vesicles. If the PM should be mimicked, the vesicles should have a larger diameter.

While we share the reviewer’s concern regarding the (implied) “curvature” aspect of 100 nm vesicles versus larger, giant unilamellar vesicles (GUVs) with a diameter > 1 µm, PIP2-containing GUVs are inherently unstable in the presence of low µM amounts of Ca2+ and rupture within minutes4, thus precluding experiments presented in Figure 2E. Despite these shortcomings, we believe that 100 nm vesicles (i.e. LUVs) represent excellent surrogates for studying αSyn interactions with “planar” membranes.

Considering the geometries of αSyn and target vesicles as spherical in a first approximation, αSyn molecules approaching these membranes display the following properties:

Author response image 1
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Given the known diameter of αSyn in solution (~6 nm5, 6) and that of our reconstituted vesicles (i.e. ~100 nm), the local interaction surface displays minimal curvature over the entire dimensions of the protein (indicated by arrow heads). Because αSyn residues 1-12 act as the primary anchoring sites7, 8, the extension of the corresponding α-helix (~1.6 nm) can be extracted from available experimental data9. As can be appreciated in Author response image 1, curvature is negligible over this range of dimensions and initial contacts between αSyn monomers and 100 nm vesicles or planar membranes are virtually identical on the microscopic level. Even in the fully extended helical conformation, spanning roughly 12 nm10, curvature effects imposed by 100 nm vesicles are small (not taking membrane bending and remodeling activities by αSyn into account11).It is also important to keep in mind that in today’s discussions about αSyn-membrane interactions, “membrane curvature” mostly serves as a proxy for “lipid packing defects”. While it is clear that high curvature induces/exacerbates these defects and enhances αSyn binding, membrane fluidity and composition especially with regard to saturated versus unsaturated fatty acids (FA) act as important additional contributors to packing defects. In this regard, we believe that the poly-unsaturated FA character of most PIPs and induced packing defects exert a stronger influence than curvature itself.

Figure 3A: The quantification of αSyn intensity with expression of INPP5E is not reflected by the provided image where there is essentially no αSyn signal. Please provide a more representative image.

This has been updated and we now provide an image that better reflects residual amounts of endogenous αSyn at the PM upon INPP5E overexpression (new panel in Figure 3A). Specifically, we have chosen a comparative view that shows cells not transfected with INPP5E-mCherry (upper left, lower right corners) together with a transfected one in the same frame (center). The differences in PM-localization levels of endogenous αSyn are readily evident (see also Q6 next).

Figure 3A: The authors argue that αSyn has a specificity for PIP2 and PIP3, but why then are there no apparent changes between αSyn intensity upon expression of MTM1 and INPP5E (which result in production of π and PI4P, respectively) versus PTEN (which produces PIP2 from PIP3)? In addition, the authors describe that "only the conversion of PIP2 to PI4P by INPP5E led to a marked reduction" but their statistics show significant changes for all three phosphatases compared to mCherry only.

We fully agree with the reviewer that our p-value analysis and corresponding statistical significance (** versus ***) were at odds with our statement. As a matter of fact, they were also at odds with what is shown in the whisker blots (of Figure 3A), as mCherryctrl and MTM1, PTEN datasets were obviously too similar to rationalize the low p-values. We revisited the source data file and recalculated p-values. Indeed, we found that the mCherry-ctrl – MTM1 comparison yields a p-value of 0.6297. Therefore, the correct labeling should have been “not significant” (NS).

The p-value for the mCherry-ctrl – PTEN is 0.0198, thus should have been labeled * (p < 0.05). Upon closer inspection, we realized that the significance analysis in this particular case, is dominated by the spread of data points in the control dataset compared to the phosphatase experiments and, therefore, should be considered with caution. By excluding single high-value outliers in the mCherry-ctrl dataset (by adjusting the overall threshold), we learned that the INPP5E experiment displayed the only “robust” value (*** p < 0.001).

We rechecked all other p-values and now include them in the revised source data files.

In summary, we had mislabeled the significance values in our original submission and updated the figure accordingly. Having made those changes, we believe that our initial statement “In agreement with our hypothesis, only the conversion of PIP2 to PI(4)P by INPP5E led to a marked reduction of endogenous αSyn at the PM (Figure 3A).” remains valid. We kept the original wording in the revised versions of the manuscript.

Figure 3: There may be a contribution of coincidental labeling. Could the authors reduce PIP2 and/or PIP3 levels as in panel 3A and repeat their analysis?

Coincidental labeling between PIP2 and PIP3 can be excluded due to the fact that PIP2 detection is antibody-mediated, whereas PIP3 identification is accomplished via transient expression of the GRP1-PH-domain fused to GFP. The affinity of GRP1-PH domain towards PIP3 is 2-3 orders of magnitudes greater than for PIP2, and negligible for other PIPs12. Hence, “PIP3-fluorescence” originates from interaction of a PH domain that specifically recognizes PIP3 only. Technically speaking, no cross-reactivity is possible in such a system. We believe that this notion is corroborated by the differential appearance of PIP2 and PIP3 signals in the time course. At 120 seconds, for instance, endogenous αSyn localization “follows” the PIP3 signal, whereas PIP2 fluorescence is virtually absent (Figure 3B). In that sense, we are confident that our experimental setup cleanly reports on signaling mediated redistributions of cellular αSyn in a manner that is PIP2 and PIP3-dependent.

Are the observed CD spectra and NMR spectra shown in Figure 2 consistent with residues ~ 1-100 in an helical conformation and the remaining ~ 40 residues being unstructured under high PIP2 to αSyn ratios, similar to the solution NMR structure of micelle-bound αSyn (PDB ID 1XQ8) or similar to the solid state NMR studies of αSyn interactions with lipid bilayers by Fuso et al., Nature Comm, 2014? In other words, are residues 100-140 unstructured in all these studies? Please comment.

Yes, based on available NMR spectra (Figure 2B and Figure 2—figure supplement 1A and 2B), it is evident that resonances corresponding to residues 100-140, do not undergo line-broadening and do not display chemical shift changes, i.e., superimpose perfectly with those of disordered monomeric αSyn. This corroborates that C-terminal residues remain dynamic and disordered upon PIP2-vesicle binding, similar to all previously determined αSyn-membrane interactions. To strengthen this notion, we modified the text at several points:

Results: “Together, these results established that residues 1-100 of αSyn interacted with PIP2vesicles in helical conformations that imposed membrane remodeling, whereas C-terminal residues 100-140 did not engage in membrane-binding and remained flexible and disordered.”

Discussion: “… these interactions are indistinguishable from other previously identified, negatively charged membrane systems with primary contacts by N-terminal αSyn residues 1-10 and progressively weaker interactions along residues 10-100. Cterminal residues 100-140 are not involved in PIP2 membrane binding, similar to all other reconstituted vesicular or planar lipid surface interactions studied thus far.”

The experiments with calcium (Figure 2E) suggest that the first ten residues always interact with PC-PIP2 vesicles, although overall binding was greatly reduced. However, plasma membranes contain anionic lipids that may mitigate the effect of calcium that is observed in these experiments. Please comment.

Yes, this is exactly what we intended to recapitulate in this experiment. As outlined in Materials and methods, we used 60 µM of isotope-labeled (15N) αSyn and a 50fold mass excess of PC-PIP2 vesicles. In molar terms, this corresponds to ~41.25 mM of total lipids and ~2.95 mM of PIP2 given the molar ratio between DOPC:PIP2 of 13:1. We added Ca2+ to a final concentration of 2.5 mM. However, membrane-embedded PIP2 can sequester up to 3 Ca2+ ions per lipid (3:1)13, which indicates that Ca2+ is present at substoichiometric amounts with respect to direct PIP2 binding, but in large excess to αSyn.

When αSyn binds to membranes in its characteristic extended helical conformation, it coordinates between 20-30 lipid molecules, depending on lipid type and clustering14, 15. If αSyn were to only interact with PIP2 molecules on DOPC:PIP2 vesicles, this would translate into 1.2-1.8 mM of PIP2 (at 60 µM of αSyn) and, accordingly, reduce the concentration of membrane-embedded PIP2 not bound by αSyn to 1.75-1.15 mM. At 3:1 of Ca2+:PIP2 binding, all of the added Ca2+ could coordinate free PIP2 on DOPC:PIP2 vesicles and not displace any of the PIP2-bound αSyn. As our results show, this is clearly not the case, i.e. C-terminal portions of αSyn are effectively displaced by Ca2+.

This strongly argues for a scenario in which αSyn actively clusters PIP2 on the surface of DOPC:PIP2 vesicles (in line with MD simulations 16). These PIP2 clusters likely exhibit higher affinity for Ca2+ than the remaining portions of non-clustered PIP2. As a result, Ca2+ progressively displaces parts of αSyn that exhibit weaker PIP2 interactions (i.e. residues 10100) whereas regions of higher PIP2 affinity are unaffected and remain vesicle-bound (i.e. residues 1-10). In that sense, PIP2 clustering by αSyn is “replaced” by protein-independent divalent metal coordination after the addition of Ca2+17. Thus, our experimental setup effectively recapitulates a competitive binding scenario in which “physiological” amounts of Ca2+ (within the range of synaptic transmission, for example) compete for anionic lipid interactions with membrane-bound proteins-, in our case αSyn-, that are present at substoichiometric concentrations. This is in line with expected, general behaviors of PIP2 micro-domains in biology18.

We did not dwell on this aspect in the original submission because we consider it is outside the scope of the manuscript. We are grateful for being able to communicate it here so that readers can access it in the published Author Response.

References

Encinas M, et al. Sequential treatment of SH-SY5Y cells with retinoic acid and brain-derived neurotrophic factor gives rise to fully differentiated, neurotrophic factor-dependent, human neuron-like cells. J Neurochem 75, 991-1003 (2000).

Lopes FM, et al. Comparison between proliferative and neuron-like SH-SY5Y cells as an in vitro model for Parkinson disease studies. Brain Res 1337, 85-94 (2010).

Burre J, Sharma M, Tsetsenis T, Buchman V, Etherton MR, Sudhof TC. Α-synuclein promotes SNARE-complex assembly in vivo and in vitro. Science 329, 1663-1667 (2010).

Carvalho K, Ramos L, Roy C, Picart C. Giant unilamellar vesicles containing phosphatidylinositol(4,5)bisphosphate: characterization and functionality. Biophys J 95, 4348-4360 (2008).

Paleologou KE, et al.Phosphorylation at Ser-129 but not the phosphomimics S129E/D inhibits the fibrillation of α-synuclein. J Biol Chem 283, 16895-16905 (2008).

Stephens AD, Nespovitaya N, Zacharopoulou M, Kaminski CF, Phillips JJ, Kaminski Schierle GS. Different Structural Conformers of Monomeric α-Synuclein Identified after Lyophilizing and Freezing. Anal Chem 90, 6975-6983 (2018).

Bodner CR, Dobson CM, Bax A. Multiple tight phospholipid-binding modes of α-synuclein revealed by solution NMR spectroscopy. J Mol Biol 390, 775-790 (2009).

Lokappa SB, Suk JE, Balasubramanian A, Samanta S, Situ AJ, Ulmer TS. Sequence and membrane determinants of the random coil-helix transition of α-synuclein. J Mol Biol 426, 2130-2144 (2014).

Fusco G, De Simone A, Arosio P, Vendruscolo M, Veglia G, Dobson CM. Structural Ensembles of Membrane-bound α-Synuclein Reveal the Molecular Determinants of Synaptic Vesicle Affinity. Sci Rep 6, 27125 (2016).

Jao CC, Hegde BG, Chen J, Haworth IS, Langen R. Structure of membrane-bound α-synuclein from site-directed spin labeling and computational refinement. Proc Natl Acad Sci U S A 105, 19666-19671 (2008).

West A, Brummel BE, Braun AR, Rhoades E, Sachs JN. Membrane remodeling and mechanics: Experiments and simulations of α-Synuclein. Biochim Biophys Acta 1858, 1594-1609 (2016).

Kavran JM, et al. Specificity and promiscuity in phosphoinositide binding by pleckstrin homology domains. J Biol Chem 273, 30497-30508 (1998).

Bradley RP, Slochower DR, Janmey PA, Radhakrishnan R. Divalent cations bind to phosphoinositides to induce ion and isomer specific propensities for nano-cluster initiation in bilayer membranes. R Soc Open Sci 7, 192208 (2020).

Galvagnion C, et al. Lipid vesicles trigger α-synuclein aggregation by stimulating primary nucleation. Nat Chem Biol 11, 229-234 (2015).

Middleton ER, Rhoades E. Effects of curvature and composition on α-synuclein binding to lipid vesicles. Biophys J 99, 2279-2288 (2010).

Gambhir A, et al. Electrostatic sequestration of PIP2 on phospholipid membranes by basic/aromatic regions of proteins. Biophys J 86, 2188-2207 (2004).

Wang YH, Slochower DR, Janmey PA. Counterion-mediated cluster formation by polyphosphoinositides. Chem Phys Lipids 182, 38-51 (2014).

Borges-Araujo L, Fernandes F. Structure and Lateral Organization of Phosphatidylinositol 4,5bisphosphate. Molecules 25, (2020).

https://doi.org/10.7554/eLife.61951.sa2

Article and author information

Author details

  1. Reeba Susan Jacob

    Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing - original draft, Project administration, Writing - review and editing
    Contributed equally with
    Cédric Eichmann
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0676-834X

  2. Cédric Eichmann

    Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Conceptualization, Resources, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Contributed equally with
    Reeba Susan Jacob
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-8476-1936

  3. Alessandro Dema

    Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
    Present address
    UCSF, School of Dentistry, Department of Cell and Tissue Biology, San Francisco, United States
    Contribution
    Conceptualization, Resources, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - original draft, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0976-9396

  4. Davide Mercadante

    Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
    Present address
    School of Chemical Sciences, University of Auckland, Auckland, New Zealand
    Contribution
    Conceptualization, Resources, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-6792-7706

  5. Philipp Selenko

    Department of Biological Regulation, Weizmann Institute of Science, Rehovot, Israel
    Contribution
    Conceptualization, Resources, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Methodology, Writing - original draft, Project administration, Writing - review and editing
    For correspondence
    philipp.selenko@weizmann.ac.il
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2590-5899

Funding

European Research Council (647474)

  • Philipp Selenko

Swiss National Science Foundation (P300PA_160979)

  • Cedric Eichmann

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We are grateful to Dr. Peter Schmieder and Monika Beerbaum for excellent maintenance of the NMR infrastructure at the Leibniz Institute of Molecular Pharmacology (FMP Berlin) and Dr. Tali Scherf for NMR infrastructure maintenance at the Weizmann Institute of Science. We thank Dr. Dmytro Puchkov (FMP-Berlin) for assistance with negative-stain electron microscopy and Drs. Michael Krauss and Volker Haucke (FMP-Berlin) for tools and reagents, helpful discussions throughout the project and useful feedback on the manuscript. Dr. Dorothea Fiedler (FMP-Berlin) for sharing aliquots of IP6. We also thank Drs. Martin Lehmann (Cellular imaging, FMP-Berlin) and Yoseph Addadi (Life Sciences Core Facilities, Weizmann Institute of Science) for excellent maintenance of imaging facilities and their support at the respective institutes. We acknowledge the highly valuable input by Drs. Meir Schechter and Ronit Sharon, Hebrew University Jerusalem, especially with regard to time-resolved histamine experiments. We further thank them for kindly providing aliquots of the SK-MEL-2 cell line. We are grateful to Drs. Ori Avinoam, Hagen Hofmann (Weizmann), and Andres Binolfi (CONICET) for carefully reading the manuscript. CE was supported by a Swiss National Science Foundation (SNSF) Advanced -Postdoc.Mobility fellowship P300PA_160979. PS acknowledges funding by the European Research Council (ERC) Consolidator Grant NeuroInCellNMR (647474). TIRF imaging was made possible with the help and support of the de Picciotto Cancer Cell Observatory in memory of Wolfgang and Ruth Lesser. Work in the Selenko laboratory is supported by the Willner Family Foundation.

Senior Editor

  1. Olga Boudker, Weill Cornell Medicine, United States

Reviewing Editor

  1. Axel T Brunger, Stanford University, United States

Reviewer

  1. David Eliezer, Weill Cornell Medical College, United States

Version history

  1. Received: August 10, 2020
  2. Accepted: February 12, 2021
  3. Accepted Manuscript published: February 15, 2021 (version 1)
  4. Version of Record published: March 3, 2021 (version 2)

Copyright

© 2021, Jacob et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Reeba Susan Jacob
  2. Cédric Eichmann
  3. Alessandro Dema
  4. Davide Mercadante
  5. Philipp Selenko
(2021)
α-Synuclein plasma membrane localization correlates with cellular phosphatidylinositol polyphosphate levels
eLife 10:e61951.
https://doi.org/10.7554/eLife.61951

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