Cells in our body communicate by releasing compounds called transmitters that carry signals from one cell to the next. Packages called vesicles store transmitters within the signaling cell. When the cell needs to send a signal, the vesicles fuse with the cell's membrane and release their cargo. For many signaling processes, such as those used by neurons, this fusion is regulated, fast, and coupled to the signal that the cell receives to activate release. Specialized molecular machines made up of proteins and fatty acid molecules called signaling lipids enable this to happen.

One signaling lipid called PI(4,5)P₂ (short for phosphatidylinositol 4,5-bisphosphate) is essential for vesicle fusion as well as for other processes in cells. It interacts with several proteins that help it control fusion and the release of transmitter. While it is possible to study the role of these proteins using genetic tools to inactivate them, the signaling lipids are more difficult to manipulate. Existing methods result in slow changes in PI(4,5)P₂ levels, making it hard to directly attribute later changes to PI(4,5)P₂.

Walter, Müller, Tawfik et al. developed a new method to measure how PI(4,5)P₂ affects transmitter release in living mammalian cells, which causes a rapid increase in PI(4,5)P₂ levels. The method uses a chemical compound called “caged PI(4,5)P₂” that can be loaded into cells but remains undetected until ultraviolet light is shone on it. The ultraviolet light uncages the compound, generating active PI(4,5)P₂ in less than one second. Walter et al. found that when they uncaged PI(4,5)P₂ in this way, the amount of transmitter released by cells increased. Combining this with genetic tools, it was possible to investigate which proteins of the release machinery were required for this effect.

The results suggest that two different types of proteins that interact with PI(4,5)P₂ are needed: one must bind PI(4,5)P₂ to carry out its role and the other helps PI(4,5)P₂ accumulate at the site of vesicle fusion. The new method also allowed Walter et al. to show that a fast increase in PI(4,5)P₂ triggers a subset of vesicles to fuse very rapidly. This shows that PI(4,5)P₂ rapidly regulates the release of transmitter. Caged PI(4,5)P₂ will be useful to study other processes in cells that need PI(4,5)P₂, helping scientists understand more about how signaling lipids control many different events at cellular membranes.

Signal transduction between cells depends on the regulated exocytosis of vesicles to liberate neurotransmitters, neuropeptides and hormones. Neuronal exocytosis is driven by an evolutionarily conserved machinery that targets vesicles to the plasma membrane, attaches them to the membrane (sometimes referred to as vesicle ‘docking’), and molecularly matures (‘primes’) them to a fusion-competent state. Primed vesicles reside in the so-called Readily Releasable Pool (RRP) whose fusion triggering leads to transmitter/hormone release. The assembly of the thermodynamically stable neuronal SNARE complex formed by the vesicular SNARE protein VAMP2/synaptobrevin-2 and the plasma membrane SNAREs syntaxin-1 and SNAP-25 provides energy needed for vesicle fusion (Jahn and Fasshauer, 2012). Indeed, SNARE proteins are already required for membrane attachment and priming (de Wit et al., 2009; Imig et al., 2014; Walter et al., 2010), and during fusion they continue to influence fusion pore properties through their transmembrane domains (Chang et al., 2015; Dhara et al., 2016; Fang and Lindau, 2014). The protein families Unc18 (sec-1) and Unc13 interact with the neuronal SNAREs and are required for membrane attachment, priming and fusion (Rizo and Südhof, 2012). The Ca²⁺-dependent activator protein for secretion (CAPS) is another priming factor in PC12-cells, chromaffin cells and neurons (Jockusch et al., 2007; Liu et al., 2008; Grishanin et al., 2004), which interacts with SNAREs to stimulate their assembly (James et al., 2009). A function of CAPS in membrane attachment of vesicles was also described in synapses of C.elegans, hippocampal neurons and PC12 cells (Imig et al., 2014; Kabachinski et al., 2016; Zhou et al., 2007), but not in mouse chromaffin cells (Liu et al., 2008). Vesicle fusion is temporally linked to electrical stimulation of the cell by voltage gated Ca²⁺ channels that activate upon depolarization. In mouse chromaffin cells, the resulting increase in intracellular Ca²⁺ concentration is sensed by the vesicular Ca²⁺ binding proteins synaptotagmin-1, and −7 (Schonn et al., 2008), which interact with the SNAREs (Zhou et al., 2015; Schupp et al., 2016), and trigger vesicle fusion (Südhof, 2013).

The lipid bilayer of the plasma membrane – apart from taking part in the vesicle to plasma membrane merger - contains a variety of signaling lipids that can regulate exocytosis, most notably phosphatidylinositols (PIs) and diacylglycerols (DAGs). Phosphatidylinositols constitute a family of lipids, which can be phosphorylated on one or more positions of the inositol headgroup, giving rise to specific signals on cell membrane compartments (Di Paolo and De Camilli, 2006; Balla, 2013). PI(4,5)P₂ is the major phosphatidylinositol in the inner leaflet of the plasma membrane where it plays multiple essential roles in cell motility, actin cytoskeleton organization, ion channel activity, and vesicle exocytosis (Di Paolo and De Camilli, 2006; Martin, 2012).

Signaling lipids are recognized by specific protein motifs, particularly C1, C2 and pleckstrin homology (PH) domains (Martin, 2015). One of the best characterized signaling lipid interactions with exocytosis proteins is that of the synaptotagmin-1 (syt-1) C2B-domain with PI(4,5)P₂ (Schiavo et al., 1996; Honigmann et al., 2013). The syt-1 C2B domain binds Ca²⁺, and is essential for triggering of vesicle fusion (Mackler et al., 2002). In vitro, PI(4,5)P₂ binding to the C2B domain markedly increases the affinity for Ca²⁺ (van den Bogaart et al., 2012; Li et al., 2006). Thus, interactions of syt-1 with synaptic PI(4,5)P₂ ensure that the C2B domain interacts with the plasma membrane (Bai et al., 2004) and aid the triggering of vesicle fusion by bringing its Ca²⁺-affinity into the physiological range. All known Munc13 isoforms contain C1- and C2 domains that regulate exocytosis. The DAG analog phorbolester binds to the Munc13 C1 domain to strongly enhance exocytosis (Rhee et al., 2002; Basu et al., 2007). Membrane binding of Munc13 can be further augmented by PI(4,5)P₂ binding to the neighboring C2B domain, which also influences the fusion probability of synaptic vesicles (Shin et al., 2010). CAPS contains a PH domain which binds PI(4,5)P₂ and is essential for vesicle priming (Nguyen Truong et al., 2014; Loyet et al., 1998; Kabachinski et al., 2014). Because different species of signaling lipids may regulate different essential exocytosis proteins, systematic investigation of the relevant interactions for exocytosis is needed.

The most successful approaches to tease apart molecular components relevant for neurosecretion have been to mutate proteins of the release machinery or to regulate their expression. This is not directly possible for signaling lipids; instead, enzymes of lipid metabolism have been targeted. Early experiments in permeabilized bovine adrenal chromaffin cells using bacterial phospholipase C (PLC) showed that secretion depends on PI(4,5)P₂ at an upstream ATP-dependent priming step (Eberhard et al., 1990). Exocytosis was further found to require the PI(4,5)P₂ synthesizing enzyme phosphatidylinositol-4-phosphate 5-kinase (Hay et al., 1995; Gong et al., 2005). Moreover, experiments using fast capacitance measurements showed that overexpression of PI(4,5)P₂ generating or degrading enzymes increased or decreased the RRP, respectively (Gong et al., 2005; Milosevic et al., 2005). Even though the initial experiments indicated that PLC, which produces DAG at the expense of PI(4,5)P₂, inhibits rather than stimulates secretion (Eberhard et al., 1990; Hay et al., 1995), later experiments showed that phorbolesters – assumed to mimic DAG – strongly stimulate secretion when added to naïve cells (Smith et al., 1998). The phorbol ester effect has since been studied in a number of cell types, and has been found to rely on the activation of two priming factors Munc13 (via its C1 domain)(Rhee et al., 2002; Betz et al., 1998) and Munc18-1 (via protein kinase C phosphorylation)(Wierda et al., 2007; Genc et al., 2014; Nili et al., 2006). In the presence of PLC activity, experiments to increase or decrease the levels of PI(4,5)P₂ might cause correlative changes in DAG. The same concern exists for the conversion of PI(4,5)P₂ to PI(3,4,5)P₃, which might have profound effects on exocytosis in spite of being present in low abundance (Khuong et al., 2013). Even the fastest existing techniques to manipulate PI(4,5)P₂ levels (by voltage, light, or chemical dimerization)(Murata et al., 2005; Suh et al., 2010; Idevall-Hagren et al., 2012) operate on similar speeds as PI(4,5)P₂ metabolism and vesicle priming (i.e. tens of seconds)(Voets et al., 1999), making it a general concern how to tease apart the effect of PI(4,5)P₂ from that of its metabolites including DAG and other phosphatidylinositols.

To manipulate cellular PI(4,5)P₂ levels rapidly and distinguish its function from those of its metabolites in fast cellular reactions we here developed and characterized a new chemical tool: caged, membrane permeant PI(4,5)P₂. We show that our compound is taken up into living cells and verify that its UV-uncaging generates physiologically active PI(4,5)P₂ with sub-second temporal precision. Uncaging induced the re-distribution of proteins containing PI(4,5)P₂ binding motifs and locally increased actin-levels. Capacitance measurements in chromaffin cells showed that following PI(4,5)P₂ uncaging exocytosis is enhanced, and the RRP increased, which we demonstrate is specific to PI(4,5)P₂ by contrasting the effects of DAG-uncaging. Systematic investigation of the relevant effector proteins revealed a requirement for the potentiation on syt-1 and Munc13-2, but not on CAPS. These results suggest two distinguishable types of PI(4,5)P₂ effector proteins: ones that require stoichiometric PI(4,5)P₂-binding to exert their function, and ones that function in the local enrichment of PI(4,5)P₂ at the vesicle fusion site. Finally, making full use of the rapid uncaging kinetics, we investigate the immediate effects of increasing the levels of signaling lipids on exocytosis and discover that PI(4,5)P₂ but not DAG uncaging induces the rapid exocytosis of few vesicles from the RRP. Our data provide an example of how caged lipid compounds can be used to tease apart relevant interactions in fast biological reactions like neurosecretion.

To achieve fast elevation of PI(4,5)P₂ levels on the relevant timescale for exocytosis we developed photoactivatable (caged), membrane-permeant PI(4,5)P₂ derivatives (Figure 1). Optical uncaging is uniquely suited to increase the levels of signaling molecules non-invasively with high temporal precision (Höglinger et al., 2014). Synthesis was based on a commercially available enantiomerically pure precursor (Figure 1a). Because the hydroxyl and phosphate groups on the inositol ring make PI(4,5)P₂ a highly charged molecule it cannot pass across the cellular plasma membrane. To make the molecule membrane permeant, these groups were equipped with protective groups of acetoxymethyl (AM) esters and butyrates (Bt), respectively (as detailed in Figure 1a legend and Methods). Once inside cells, these protective groups are removed by endogenous carboxyesterases (Schultz, 2003). Similar approaches were successfully applied to other phosphoinositides previously (Laketa et al., 2014; Mentel et al., 2011). Photosensitivity was achieved by the addition of a photocleavable cage designed to interfere with biological functions at the phosphate residues in positions 4 or 5 of the inositol ring. The coumarin caging group was chosen for its extraordinarily fast release kinetics as well as its intrinsic fluorescence, which allows verification of cellular uptake. The resulting intermediates 4a and 4b (Figure 1a) were subsequently coupled to either a dioctanoylglycerol (compound 11, legend to Figure 1) or a stearoyl-arachidonoylglycerol (compound 14, legend to Figure 1) bearing phosphoramidite reagent to form the fully protected caged PI(4,5)P₂ intermediates (Figure 1). One-pot deprotection and alkylation with AM bromide gave the caged, membrane-permeant PI(4,5)P₂ derivatives (1a,b; 2a,b) in 12% and 10% overall yield, respectively (Figure 1a–b).

Figure 1

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We first validated that UV uncaging activated lipid-protein interactions with known PI(4,5)P₂ binding domains. For this, we directly diluted our compound in an imaging buffer (to a final concentration of 20 µM) and made use of the fact that if the solution was not heavily mixed by vortexing, some of the lipid formed micelles clearly visible on the bottom of the coverslip in the light microscope. The solution furthermore contained a reconstituted fusion protein of the PI(4,5)P₂-binding PH domain of PLC-δ₁ linked to EGFP. Illumination in the TIRF field was used to limit light excitation to the surface of the glass coverslip. When EGFP was excited, the presence of PH-EGFP was visible as background in the solution of the TIRF field and some of the PH-EGFP was enriched on the micelles. Following UV-uncaging with a 405 nm laser in the TIRF field, the EGFP signal at these positions was greatly increased, indicative of PH-EGFP recruitment to the micelles from the surrounding solution (Figure 2a). To confirm that the micelles were indeed composed of cg-PI(4,5)P₂, we investigated the images during the irradiation with 405 nm light, which revealed their fluorescence, confirming the presence of the coumarin group (Figure 2a). These results demonstrate that UV-cleavage of the coumarin cage activates the compound for interactions with proteins bearing PI(4,5)P₂ binding motifs.

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To verify that the protective groups synthesized on our compound enabled cellular uptake, we investigated its cellular distribution making use of the intrinsic fluorescence of the coumarin cage. Human Embryonic Kidney (HEK) cells were first loaded with a membrane labelling dye excitable with infrared light (CellMask). Cells were then loaded with caged (cg) PI(4,5)P₂ by incubation for 30 min at 37°C with 20 µM of our compound (2a,b, diluted from a 20 mM DMSO-stock) in the presence of 0.02% Pluronic (prepared with heavy vortexing) to facilitate membrane passage. Cells receiving identical treatment but without the compound served as controls. To assess the localization of cg-PI(4,5)P₂ quantitatively, cells were imaged on a spinning disc confocal microscope and fluorescence line profiles obtained from many cells that bordered open extracellular space (as opposed to ones in contact with other HEK cells). The line profiles were then aligned to local fluorescence maxima of the CellMask signal indicating the position of the plasma membrane. Subsequently, line profiles from all cells were averaged (Figure 2b). We saw that average coumarin fluorescence increased inside the plasma membrane, demonstrating cellular uptake. Moreover, coumarin fluorescence was clearly observed at the position of the plasma membrane; however, it was also present inside the cell, possibly on endosomes. This is not surprising, as the coumarin group is expected (and, indeed, intended) to block interactions with PI(4,5)P₂-binding proteins. This includes those proteins that usually establish a strict pattern of phosphoinositide composition on distinct cellular organelles. Therefore, it is an unavoidable side effect of using caged lipids that their distribution will be broader than the native lipid. Inevitably, visualization of the coumarin fluorescence by its excitation leads to its uncaging. This could be shown by continuous imaging which significantly reduced the coumarin fluorescence at the location of the plasma membrane (bar graph Figure 2b). As expected, neither the fluorescence gradient across the membrane nor the decrease in intensity at the plasma membrane was observed in control cells (Figure 2b), indicating that our compound is taken up into cells, present at the plasma membrane and uncaged there.

We next validated that UV uncaging liberated physiologically active PI(4,5)P₂ in living cells. For this, we transfected HEK cells with the PLC-δ₁-PH-EGFP construct and looked for a possible recruitment of EGFP fluorescence to the plasma membrane by TIRF microscopy upon UV-uncaging. However, we found that even before uncaging, EGFP fluorescence intensities were very high and did not increase further upon UV-uncaging, which we attribute to saturation of the sensor due to relatively high plasma membrane levels of PI(4,5)P₂ already at rest (data not shown). To circumvent this problem, we co-transfected COS-7 cells with a plasma membrane targeted 5-phosphatase which degrades PI(4,5)P₂ (Posor et al., 2013). Upon UV-uncaging in cells loaded with cg-PI(4,5)P₂ we found a small, but highly significant increase of EGFP fluorescence at the cell’s footprint in line with the liberation of PI(4,5)P₂ at the plasma membrane, while no such effect was observed in control cells, which were also subjected to UV-light, but not loaded with cg-PI(4,5)P₂ (Figure 2c). The relatively small effect size may be caused by the 5-phosphatase activity which likely rapidly degrades uncaged PI(4,5)P₂ at the plasma membrane before it can be detected by the PLC-δ₁-PH-EGFP sensor which is why this experiment may underestimate the amount of liberated PI(4,5)P₂.

We also investigated the behavior of our compound in cells were PI(4,5)P₂ was not constitutively depleted, but where degradation was acutely induced pharmacologically. Endogenous PI(4,5)P₂ cause the quantitative plasma membrane binding of the high-affinity PLC-δ₁-PH-RFP sensor in tsA-201 cells (Figure 2d). By simultaneously expressing M₁ muscarinic receptors in these cells, we could acutely degrade PI(4,5)P₂ by the application of oxotremorine-M (Oxo-M) which activates M₁ receptors to stimulate PLC. This rapidly decreases PI(4,5)P₂ levels selectively at the plasma membrane and reliably induced the relocalization of PLC-δ₁-PH-RFP sensor from the plasma membrane to the cytosol in control cells (Figure 2d), indicating near complete plasma membrane PI(4,5)P₂ breakdown. Because this assay monitors the sensor’s dissociation it may be better suited to monitor PI(4,5)P₂ liberation close to the sensor’s initial location. We therefore combined OxoM-treatment with PI(4,5)P₂-uncaging, which prevented PLC-δ₁-PH-RFP membrane dissociation (Figure 2d), in line with substantial PI(4,5)P₂ release at the plasma membrane overruling PLC activity. Uncaging itself did not interfere with PI(4,5)P₂ breakdown, because DAG was still produced (validated by parallel imaging with a DAG biosensor)(Figure 2—figure supplement 1b,c).

To verify that UV-uncaging of our compound activated PI(4,5)P₂-dependent cellular responses, we investigated effects of cg-PI(4,5)P₂ uncaging on actin bundles, for whose polymerization a pivotal role of PI(4,5)P₂ is firmly established (Di Paolo and De Camilli, 2006; Rohatgi et al., 1999). Actin bundles were visualized by TIRF microscopy in the footprints of HEK cells expressing the actin marker Lifeact-RFP (Figure 3a). HEK cells loaded with cg-PI(4,5)P₂ were compared to non-loaded cells. Following the measurement of baseline fluorescence (five frames) in the RFP channel, cells were exposed to TIRF illumination with UV light (405 nm laser). This lead to a significant and specific increase in lifeact-RFP in cells loaded with cg-PI(4,5)P₂ (Figure 3a), in line with PI(4,5)P₂ uncaging causing actin accumulation near the plasma membrane.

Figure 3

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Because we eventually wanted to investigate the physiological effects of PI(4,5)P₂ in adrenal chromaffin cells, we next studied whether PI(4,5)P₂ uncaging would also increase PI(4,5)P₂ levels at their plasma membrane. As it is a distinct advantage of our compound that cellular PI(4,5)P₂ can be rapidly and specifically increased by light without interfering with basic PI(4,5)P₂ metabolism, we wanted to verify PI(4,5)P₂ uncaging without the prior manipulation of its resting levels. For this we employed a different biosensor harboring the lower affinity PH-domain of PLCδ₄ (Kabachinski et al., 2014; Lee et al., 2004). Chromaffin cells were infected with a virus expressing PLCδ₄-PH-EGFP, and incubated with caged PI(4,5)P₂ (2a,b). Experiments were performed on the same microscope setup later used for electrophysiological recordings. Uncaging cg-PI(4,5)P₂ by a 1–2 ms light pulse from a Xenon flash bulb caused rapid translocation of the sensor towards the periphery of the cell, in line with PI(4,5)P₂ release in the plasma membrane (Figure 3b). In cells not incubated with cg-PI(4,5)P₂, no translocation was observed. Ongoing imaging of the PLCδ₄-PH-EGFP allowed us to estimate the time constant of the [PI(4,5)P₂] relaxation to ~25 s (Figure 3b). A second flash caused markedly less translocation, suggesting that most of the cg-PI(4,5)P₂ had already been uncaged. In this experiment, the relatively modest recruitment amplitude of to the plasma membrane is probably affected by the widespread intracellular localization of cg-PI(4,5)P₂ (Figure 2b), because uncaging likely also increases PI(4,5)P₂ on intracellular membranes. However, the fact that PH-domains overall relocalize to the plasma membrane (Figures 2c and 3b) indicate that PI(4,5)P₂ is uncaged there.

The essential role of PI(4,5)P₂ for exocytosis was realized more than 25 years ago, through experiments showing that PI(4,5)P₂ depletion inhibits exocytosis from cracked-open adrenal chromaffin cells and PC12 cells (Eberhard et al., 1990; Hay et al., 1995). Later experiments showed that PI(4,5)P₂ delivery to the intracellular compartment via the patch pipette increased the RRP (Milosevic et al., 2005). To investigate the physiological effects of PI(4,5)P₂ uncaging, cg-PI(4,5)P₂ was loaded into the cells (see Materials and methods) and exocytosis was induced with a depolarization protocol to allow Ca²⁺ influx (Voets et al., 1999). Exocytosis was monitored using patch-clamp capacitance measurements, which report on increased plasma membrane area upon vesicle fusion. After a pre-pulse, which elicited indistinguishable exocytosis in both groups (Figure 4a), cells were either subjected to UV-light flashes (PI(4,5)P₂ uncaging group) or not (control group), and a second depolarization protocol was used to assess the effect of uncaging. PI(4,5)P₂ uncaging significantly augmented exocytosis in wildtype cells (Figure 4b,ci). The overall level of exocytosis found in these experiments is similar to previous findings from wild type Bl6 mice (Voets et al., 2001a; Voets et al., 2001b). Note that in these experiments, to protect against any effect of the loading protocol or the cg-PI(4,5)P₂ compound itself, both control and uncaging groups were loaded with the cg-PI(4,5)P₂, but only the uncaging group was exposed to UV-light. Nevertheless, in separate experiments we compared cells loaded with cg-PI(4,5)P₂ to cells exposed to the same loading protocol, but without cg-PI(4,5)P₂, and found that without UV light cg-PI(4,5)P₂ had no effect on depolarization induced exocytosis (Figure 4—figure supplement 1).

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We specifically investigated the role of the fatty acid tail in exocytosis using two different compounds, one containing a short DOG-analog (1a,b) and one containing the natural SAG-chain (2a,b). However, we found similar potentiation in both cases, showing that the inositol headgroup is responsible for the enhanced exocytosis (Figure 4b,ci). Importantly, the augmentation of exocytosis was not due to changes in Ca²⁺ influx during membrane depolarization, as revealed by similar Ca²⁺ currents during the depolarizations (Figure 4cii, Figure 4—figure supplement 2), and unchanged intracellular Ca²⁺ concentration immediately after PI(4,5)P₂ uncaging (Figure 4—figure supplement 3a).

Because the compounds 1a,b and 2a,b elicited similar effects, we pooled both datasets for further analysis. The depolarization protocol included steps of varying duration which can be used to quantify the release of different populations of vesicles undergoing fusion. The first six 10 ms depolarizations release vesicles positioned close to Ca²⁺ channels in the so-called Immediately Releasable Pool (IRP)(Voets et al., 1999), while the following two longer (100 ms) depolarizations (a total of four were given) are assumed to deplete the full Readily Releasable Pool (RRP; the IRP is part of the RRP [Voets et al., 1999]). We found that PI(4,5)P₂ uncaging did not influence the release of the IRP. However, the RRP was approximately doubled by PI(4,5)P₂ uncaging (Figure 5a,b), confirming that increasing PI(4,5)P₂ enhances priming of vesicles into the RRP (Milosevic et al., 2005). In addition, secretion elicited by residual Ca²⁺ in-between depolarizations was enhanced (as seen by steeper slopes of the capacitance increase, Figure 5a, right-hand panels); this could be due to faster priming followed by fusion or increased fusion probability of the remaining vesicles.

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Augmentation of the RRP was noted before in adrenal chromaffin cells following longer-term elevation of PI(4,5)P₂ (Milosevic et al., 2005), but those manipulations might also have elevated the levels of the downstream metabolite DAG. Indeed, Phorbol esters, which are assumed to act as DAG analogues, augment the RRP size in chromaffin cells (Smith et al., 1998). Therefore, we wanted to distinguish between PI(4,5)P₂ vs. DAG requirements for rapid exocytosis augmentation in chromaffin cells. To this end we performed DAG uncaging. Coumarin-caged DAG (Nadler et al., 2013) (cg-DAG, Figure 5b) was infused into cells via the patch pipette. Because both cg-DAG and cg-PI(4,5)P₂ bear the same fluorescent coumarin cage, titration of coumarin fluorescence at different (known) DAG concentrations in the patch pipette allowed us to estimate the cellular concentration of cg-PI(4,5)P₂ in the experiments above. This concentration was not known, since AM-ester loading allows progressive accumulation in the cell with time. Based on comparable fluorescence values in the titration, we estimated that the final intracellular concentration of the caged PI(4,5)P₂ corresponded to ~29 μM (Figure 5c) and therefore performed DAG uncaging using 30 or 45 μM DAG. However, UV-induced DAG uncaging (unlike PI(4,5)P₂ uncaging) failed to potentiate secretion in adrenal chromaffin cells (Figure 5d).

Next, we sought an independent method to confirm that refilling of the primed vesicle pool (RRP) depends on PI(4,5)P₂, and not on DAG in a similar concentration regime. To this end, we performed double Ca²⁺-uncaging experiments (Figure 5—figure supplement 1). In these experiments no caged lipids were present, but UV uncaging of a photolysable Ca²⁺ chelator allowed the direct triggering of vesicles without voltage depolarization of the cell. Ca²⁺ uncaging increases intracellular Ca²⁺ concentrations to the tens-of micromolar range, which is sufficient to deplete the entire RRP and expected to activate endogenous PLC, leading to PI(4,5)P₂ degradation. Two sequential Ca²⁺-uncaging stimuli were used to assess RRP sizes and thus refilling in cells incubated with a PLC inhibitor, or an inactive control compound. Recovery of the RRP was significantly enhanced by inhibiting PLC (Figure 5—figure supplement 1), indicating that preventing PI(4,5)P₂ degradation enhances vesicle priming. Thus, conversion of endogenous PI(4,5)P₂ to DAG is overall negative for refilling of the RRP, which confirms our findings using caged lipid compounds.

We next sought to identify relevant PI(4,5)P₂ effectors among the molecular release machinery. Syt-1, the Ca²⁺ sensor for rapid exocytosis in chromaffin cells (Voets et al., 2001a), was among the first PI(4,5)P₂-binding presynaptic proteins to be identified (Schiavo et al., 1996; Honigmann et al., 2013; van den Bogaart et al., 2012; Bai et al., 2004). The relevance of PI(4,5)P₂-binding was indicated by mutation, which increased the Ca²⁺ requirements for exocytosis. However, mutations can have other effects than those intended in the experiment. For instance, the same residues in the syt-1 C2B domain interacting with PI(4,5)P₂ were also shown to interact with the neuronal SNARE complex (Zhou et al., 2015). Therefore, and to complement those experiments, we here uncaged PI(4,5)P₂ in syt-1 knockout mice, and found that uncaging did not potentiate exocytosis (Figure 6a). Proper loading of the compound was ensured after the experiment by the intrinsic fluorescence of the coumarin group (Figure 6—figure supplement 1a). Exocytosis from syt-1 KO cells is reduced compared to wild type cells (Voets et al., 2001a), although sizable release – including a small RRP (Mohrmann et al., 2013) – remains. We asked whether the lack of PI(4,5)P₂ augmentation was due to the smaller exocytosis amplitude, rather than the lack of syt-1. To this end, we reanalyzed data, identifying wild type cells with intrinsically low exocytosis amplitude, and syt-1 KO cells with high exocytosis amplitude. However, we still found significant potentiation in WT cells, but not in syt-1 KO cells (Figure 6—figure supplement 1d), suggesting a molecular requirement for syt-1. Neurosecretion is known to depend on the key vesicle priming factor Munc13, and the relevant isoform in chromaffin cells, Munc13-2, harbors a C2-domain (C2B), which displays a strong PI(4,5)P₂-dependence (Shin et al., 2010; Kabachinski et al., 2014). Adrenal chromaffin cells isolated from Munc13-2 knockout mice lacked the capacity of PI(4,5)P₂ uncaging to potentiate exocytosis (Figure 6b). Thus, PI(4,5)P₂ potentiation in chromaffin cells occurs via specific activation of the vesicular release machinery and requires syt-1 and Munc13-2. To identify additional molecular targets, we repeated experiments in knockout mouse cells for the major PI(4,5)P₂ binding proteins CAPS1 and −2. CAPS interacts with PI(4,5)P₂ via a pleckstrin homology domain and loss of this interaction impedes vesicle exocytosis by reducing the number of releasable vesicles (Nguyen Truong et al., 2014). However, uncaging PI(4,5)P₂ in CAPS1 and −2 double knockout mice revealed a similar enhancement of release upon PI(4,5)P₂ uncaging as in wild type cells (Figure 6c), arguing that augmentation of exocytosis observed here occurs independently of CAPS, or bypasses CAPS (see Discussion). Surprisingly, the IRP size was actually reduced by PI(4,5)P₂-uncaging in the Munc13-1 KO, whereas it was (nonsignificantly, p<0.08) increased in the CAPS-1/2 DKO (Figure 6). The implication of this finding is unclear, but Munc13 and CAPS-proteins play distinct roles during priming (Kabachinski et al., 2014; Liu et al., 2010), and if they are both required for the formation of the IRP-vesicles, then the elimination of one or the other might create IRPs with distinct properties, including PI(4,5)P₂-dependence. We conclude that PI(4,5)P₂-dependent activation of exocytosis operates via Munc13-2 and syt-1 to potentiate RRP size.

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Use of cg-PI(4,5)P₂ for the first time allowed investigating the consequences of an abrupt increase in PI(4,5)P₂ abundance on a subsecond timescale. When inspecting the capacitance trace around the first uncaging flash (see Figure 4a for stimulation protocol), we found an abrupt jump in the capacitance, indicating fast fusion of a few (5-10) vesicles (Figure 7a). This jump was observed only with the first uncaging flash, indicating that it is unlikely to be a photo-artifact (Figure 7—figure supplement 1). The release appears specific, because the size of the response strongly correlated with the RRP sizes in these cells (Figure 7b). Furthermore, uncaging of the PI(4,5)P₂ downstream metabolite DAG, bearing the same photolysable coumarin group as PI(4,5)P₂ did not induce any capacitance increase (Figure 7a–c). Finally, the jump was reduced in size – or absent – in cells from Munc13-2, CAPS-1/2 and syt-1 knockout mice, which all have smaller RRP sizes (Liu et al., 2008; Mohrmann et al., 2013; Man et al., 2015) (Figures 5, 6 and 7b). Thus, rapidly increasing PI(4,5)P₂ levels can fuse vesicles. The release of a fraction of the RRP is consistent with the stimulation of some vesicles close to fusion threshold (Yang et al., 2002) whose Ca²⁺-sensitivity may increase further due to the interaction of syt-1 with PI(4,5)P₂ (van den Bogaart et al., 2012;Li et al., 2006 ), leading to increased release probability.

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Here, we developed a photocaged membrane-permeant PI(4,5)P₂ and combined this compound with high time-resolution electrophysiology and genetic manipulations to identify relevant PI(4,5)P₂ effectors in neuroendocrine chromaffin cells. The main PI(4,5)P₂-binding proteins in the secretory pathway are syt-1 (Schiavo et al., 1996; Honigmann et al., 2013; van den Bogaart et al., 2012; Bai et al., 2004), the Ca²⁺-sensor for exocytosis, and Munc13-2 (Shin et al., 2010) and CAPS (Loyet et al., 1998; Kabachinski et al., 2014), two priming proteins, which are responsible for establishing and replenishing the RRP (Man et al., 2015; Liu et al., 2010). Possible effects of PI(4,5)P₂-binding to these proteins in living cells were previously not investigated by altering PI(4,5)P₂ levels, but by using correlative analyses following protein mutation (Li et al., 2006; Shin et al., 2010). However, exactly which of these proteins are acutely activated by PI(4,5)P₂ to facilitate secretion was not clear. By uncaging our compound we could now verify that PI(4,5)P₂ enhances exocytosis and by studying mouse knockouts we could provide mechanistic insight into distinct PI(4,5)P₂-dependent processes in exocytosis:

• Synaptotagmin-1 and Munc13-2 are required for the potentiating effect of PI(4,5)P₂ on exocytosis.

• The priming- and PI(4,5)P₂-binding proteins CAPS-1/2 are not involved in (or are bypassed by, see below) the potentiating effect of PI(4,5)P₂ uncaging.

• Increasing PI(4,5)P₂ triggers the rapid release of a part of the Readily Releasable Pool of vesicles.

PI(4,5)P₂ uncaging specifically potentiated RRP size, but not the size of the IRP, which forms a subpool of the RRP, consisting of vesicles co-localized with Ca²⁺-channels (Voets et al., 1999). One interpretation is that IRP-vesicles are already saturated with PI(4,5)P₂; another possible explanation is that the IRP is limited in size by additional factors (for instance the availability of Ca²⁺-channels), and therefore cannot be further augmented. Based on the different types of effects we observe in our experiments (loss of PI(4,5)P₂-dependent augmentation in syt-1 KO and Munc13-2 KO vs. no effect in CAPS DKO), two different mechanisms of PI(4,5)P₂-binding proteins can be envisioned (Figure 8): the first is specific, and probably stoichiometric, PI(4,5)P₂-binding to support protein function, for instance the ability of Munc13 to stimulate SNARE-complex formation, or to increase the Ca²⁺ binding affinity of syt-1 during secretion triggering. The other potential function is to co-localize the fusion machinery with PI(4,5)P₂-patches in the plasma membrane. The latter, but not the former, function might be bypassed by PI(4,5)P₂-uncaging, which uncovers PI(4,5)P₂ under the vesicle (Figure 8). Thus, lipid uncaging can serve as an exquisite tool to distinguish between these two different functions of PI(4,5)P₂-binding proteins, just as Ca²⁺ uncaging has been instrumental in distinguishing between effects on Ca²⁺-binding to the release machinery itself, and effects on colocalizing vesicles with Ca²⁺ channels (Voets et al., 1999; Wadel et al., 2007). Therefore, the established essential requirement of the CAPS PH-domain for its function in vesicle priming (Nguyen Truong et al., 2014; Kabachinski et al., 2014) can be reconciled with our findings here, if the function of CAPS is to cause local enrichment of PI(4,5)P₂ at the sites of vesicle priming, where it will interact with other exocytotic proteins.

Figure 8

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The use of uncaging made it possible for the first time to investigate the consequences of an acute, millisecond, increase in PI(4,5)P₂ and DAG abundance. We found that PI(4,5)P₂, but not DAG, uncaging caused the rapid fusion of vesicles (Figure 7a,c). The amount of fusion correlated with the RRP size, and this correlation extended to the knockouts tested (Figure 7b). Thus, the acutely fusing vesicles probably constitute a fraction of the RRP. The fact that rapid fusion was only seen when uncaging PI(4,5)P₂, but not DAG, argue that PI(4,5)P₂ is more directly linked to exocytosis triggering in adrenal chromaffin cells, possibly because PI(4,5)P₂ binding to the C2-domains in Munc13-2 and syt-1 directly change the Ca²⁺ affinities of those domains. The fusing vesicles might be members of the ‘Highly Calcium Sensitive Pool’ (HCSP), which fuse at lower Ca²⁺ concentrations than the rest of the RRP vesicles (Yang et al., 2002). Since these vesicles are close to fusion threshold, rapid binding of PI(4,5)P₂ might increase the Ca²⁺-affinity of syt-1 enough that the vesicles fuse due to a rapid increase in Ca²⁺-affinity rather than a rapid increase in Ca²⁺ concentration as would normally be the case.

A definite advantage of the photocaged approach is that it allows inducing sub-second increases in the phospholipid composition of membranes, which can be used to identify direct effects of a phospholipid before its metabolism takes place. A possible complication is that by shielding the head group, the lipid will no longer be recognized by proteins (enzymes, lipid-shuttling proteins) that establish the cellular pattern of lipid composition between different organelles. Thus, the localization of the caged lipid will likely be broader than for the native lipid. Indeed, investigating the sub-cellular distribution of our cg-PI(4,5)P₂ revealed the uptake in compartments other than the plasma membrane (Figure 2b). However, our data also clearly show its specific uncaging at the plasma membrane, making it a suitable tool to address reactions at the latter (Figures 2 and 3). Moreover, uncaging PI(4,5)P₂ in chromaffin cells led to a specific potentiation of vesicle priming (Figures 4–6), which is consistent with previous findings using enzymatic over/underexpression (Gong et al., 2005; Milosevic et al., 2005), and with the use of a PLC-inhibitor to prevent PI(4,5)P₂ breakdown (Figure 5—figure supplement 1). Furthermore, the effect depended on known PI(4,5)P₂-binding proteins. Thus, although we cannot rule out that PI(4,5)P₂ is liberated elsewhere in the cell, PI(4,5)P₂ uncaging results in valid and specific effects on exocytosis. The wide-spread distribution of our caged compound may also be considered a distinct advantage, because this allows to study the consequences of its focal liberation in regions where PI(4,5)P₂ is sparse.

Collectively, our data demonstrate the power of caged phospholipids to dissect physiological functions of different, but interconvertible, phospholipids. The main power of the approach is that it outpaces the rate of metabolism/interconversion of one lipid into another. Using this method we have dissected the molecular requirement for the potentiating effect of PI(4,5)P₂ on exocytosis, and we have demonstrated a novel, acute effect of uncaged PI(4,5)P₂: to trigger rapid exocytosis. We anticipate that caged lipid second messengers will serve as valuable experimental tools to uncover mechanistic details of fast cellular processes.

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Author details

1. Alexander M Walter

   1. Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark
   2. Leibniz-Forschungsinstitut für Molekulare Pharmakologie, Berlin, Germany

   Contribution

   Investigation, Methodology

   Contributed equally with

   Rainer Müller and Bassam Tawfik

   For correspondence

   awalter@fmp-berlin.de

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5646-4750

2. Rainer Müller

   Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany

   Contribution

   Investigation, Methodology

   Contributed equally with

   Alexander M Walter and Bassam Tawfik

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3464-494X

3. Bassam Tawfik

   Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Investigation, Methodology

   Contributed equally with

   Alexander M Walter and Rainer Müller

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1193-8494

4. Keimpe DB Wierda

   Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Resources, Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8784-9490

5. Paulo S Pinheiro

   Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark

   Contribution

   Formal analysis, Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

6. André Nadler

   1. Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
   2. Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany

   Contribution

   Formal analysis, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

7. Anthony W McCarthy

   Leibniz-Forschungsinstitut für Molekulare Pharmakologie, Berlin, Germany

   Contribution

   Formal analysis, Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-3771-351X

8. Iwona Ziomkiewicz

   1. Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark
   2. Discovery Sciences, AstraZeneca, Cambridge, United Kingdom

   Contribution

   Investigation, Methodology, Writing—review and editing

   Competing interests

   Performed experiments as an employee of University of Copenhagen and is now an employee of AstraZeneca; IZ has no financial investments in AstraZeneca

9. Martin Kruse

   Department of Physiology and Biophysics, School of Medicine, University of Washington, Seattle, United States

   Contribution

   Validation, Investigation, Writing—review and editing

   Competing interests

   No competing interests declared

10. Gregor Reither

    Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany

    Contribution

    Formal analysis, Investigation, Writing—review and editing

    Competing interests

    No competing interests declared

11. Jens Rettig

    Cellular Neurophysiology, Center for Integrative Physiology and Molecular Medicine, Saarland University, Homburg, Germany

    Contribution

    Resources, Supplied CAPS double knockout mice

    Competing interests

    No competing interests declared

12. Martin Lehmann

    Leibniz-Forschungsinstitut für Molekulare Pharmakologie, Berlin, Germany

    Contribution

    Resources, Investigation, Methodology

    Competing interests

    No competing interests declared

13. Volker Haucke

    Leibniz-Forschungsinstitut für Molekulare Pharmakologie, Berlin, Germany

    Contribution

    Resources, Funding acquisition

    Competing interests

    No competing interests declared

14. Bertil Hille

    Department of Physiology and Biophysics, School of Medicine, University of Washington, Seattle, United States

    Contribution

    Conceptualization, Supervision, Funding acquisition, Writing—review and editing

    Competing interests

    No competing interests declared

15. Carsten Schultz

    Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany

    Contribution

    Conceptualization, Resources, Supervision, Funding acquisition, Validation, Methodology, Writing—review and editing

    For correspondence

    schultz@embl.de

    Competing interests

    No competing interests declared

16. Jakob Balslev Sørensen

    Neurosecretion group, Center for Neuroscience, Faculty of Health and Medical Sciences, University of Copenhagen, Copenhagen, Denmark

    Contribution

    Conceptualization, Supervision, Funding acquisition, Writing—original draft, Project administration, Writing—review and editing

    For correspondence

    jakobbs@sund.ku.dk

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0001-5465-3769

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