Lysosomes shape Ins(1,4,5)P₃-evoked Ca²⁺ signals by selectively sequestering Ca²⁺ released from the endoplasmic reticulum

Summary

Most intracellular Ca²⁺ signals result from opening of Ca²⁺ channels in the plasma membrane or endoplasmic reticulum (ER), and they are reversed by active transport across these membranes or by shuttling Ca²⁺ into mitochondria. Ca²⁺ channels in lysosomes contribute to endo-lysosomal trafficking and Ca²⁺ signalling, but the role of lysosomal Ca²⁺ uptake in Ca²⁺ signalling is unexplored. Inhibition of lysosomal Ca²⁺ uptake by dissipating the H⁺ gradient (using bafilomycin A₁), perforating lysosomal membranes (using glycyl-L-phenylalanine 2-naphthylamide) or lysosome fusion (using vacuolin) increased the Ca²⁺ signals evoked by receptors that stimulate inositol 1,4,5-trisphosphate [Ins(1,4,5)P₃] formation. Bafilomycin A₁ amplified the Ca²⁺ signals evoked by photolysis of caged Ins(1,4,5)P₃ or by inhibition of ER Ca²⁺ pumps, and it slowed recovery from them. Ca²⁺ signals evoked by store-operated Ca²⁺ entry were unaffected by bafilomycin A₁. Video-imaging with total internal reflection fluorescence microscopy revealed that lysosomes were motile and remained intimately associated with the ER. Close association of lysosomes with the ER allows them selectively to accumulate Ca²⁺ released by Ins(1,4,5)P₃ receptors.

Introduction

Because Ca²⁺ can be rapidly moved across biological membranes to generate spatially organized increases in cytosolic free Ca²⁺ concentration ([Ca²⁺]_i) (Smith and Parker, 2009), it is a versatile and ubiquitous intracellular messenger (Berridge et al., 2003; Rizzuto and Pozzan, 2006). Most Ca²⁺ signals are generated by regulated opening of Ca²⁺-permeable channels within either the plasma membrane (PM) or the membranes of intracellular organelles, most commonly the endoplasmic reticulum (ER). Within the latter, inositol 1,4,5-trisphosphate receptors [Ins(1,4,5)P₃R] are almost invariably expressed (Foskett et al., 2007). In addition, the ER often expresses ryanodine receptors (RyR) and a variety of other Ca²⁺-permeable channels (Taylor and Dale, 2012). Regulated opening of these channels allows Ca²⁺ to flow rapidly into the cytosol. This, together with the high concentration of cytosolic Ca²⁺ buffers, allows an open Ca²⁺ channel to generate a local, but substantial, increase in [Ca²⁺]_i that decays within about a hundred nanometres of the channel and persists only for as long as the channel is open (Shuai and Parker, 2005). These local microdomains of Ca²⁺ around the mouths of open channels are the fundamental units of Ca²⁺ signalling. They allow different channels to direct Ca²⁺ to different effector systems (Rizzuto and Pozzan, 2006). They allow Ca²⁺-mediated communication between Ca²⁺ channels to facilitate regenerative growth of Ca²⁺ signals by Ca²⁺-induced Ca²⁺ release, usually via Ins(1,4,5)P₃R (Calcraft et al., 2009; Rahman et al., 2009; Smith and Parker, 2009) or RyR (Brailoiu et al., 2010; Cheng and Lederer, 2008). And where Ca²⁺ channels are close to another membrane, they allow Ca²⁺ to be selectively directed to proteins that remove Ca²⁺ from the cytosol. The Ca²⁺ pump of the ER, for example, can sequester Ca²⁺ entering via store-operated Ca²⁺ entry (SOCE) (Parekh and Putney, 2005); and mitochondria can selectively accumulate Ca²⁺ released by Ins(1,4,5)P₃R (Csordás et al., 2010) or entering via SOCE (Parekh and Putney, 2005). These intimate relationships between Ca²⁺ channels and proteins that sequester, release or respond to Ca²⁺ endow the fundamental units of Ca²⁺ signalling, the local increase in [Ca²⁺]_i evoked by opening of a single channel, with the versatility needed to regulate almost every cellular activity (Berridge et al., 2003).

The membranes that have attracted most attention in the context of Ca²⁺ signalling are the PM, the ER and the inner mitochondrial membrane. All three membranes express proteins that remove Ca²⁺ from the cytosol and so terminate Ca²⁺ signals. Mitochondrial Ca²⁺ uptake additionally links cytosolic Ca²⁺ signals to oxidative phosphorylation (Jouaville et al., 1999), mitochondrial motility (Yi et al., 2004) and cell death (Szalai et al., 1999). The ER and PM express a wide variety of Ca²⁺ channels that can generate cytosolic Ca²⁺ signals (Taylor and Dale, 2012; Taylor et al., 2009). The recent identification of regulated Ca²⁺ channels in lysosomes, notably the two-pore channels (TPC) gated by nicotinic acid dinucleotide phosphate (NAADP) (Calcraft et al., 2009) and various transient receptor potential (TRP) channels (Dong et al., 2010), has suggested that they too may contribute to cytosolic Ca²⁺ signalling. Lysosomes are the main degradative compartment of the cell, they contain substantial amounts of Ca²⁺ (Christensen et al., 2002; Morgan et al., 2011), membrane trafficking within the endo-lysosomal pathway is regulated by Ca²⁺ (Luzio et al., 2010), and dysfunctional lysosomes cause devastating lysosomal storage diseases (Vellodi, 2005), some of which are associated with defective lysosomal Ca²⁺ regulation (Lloyd-Evans and Platt, 2011; Morgan et al., 2011; Pereira et al., 2010). There is, therefore, a growing recognition that lysosomal Ca²⁺ channels contribute to the genesis of cytosolic Ca²⁺ signals and to regulation of endo-lysosomal membrane trafficking. However, the mechanism of Ca²⁺ uptake by mammalian lysosomes is not resolved (Lloyd-Evans and Platt, 2011) and nor has the contribution of lysosomal Ca²⁺ uptake to the shaping of Ca²⁺ signals evoked by conventional Ca²⁺ signalling pathways been established. Here, we show that intimate and dynamic associations between lysosomes and the ER allow lysosomes selectively to accumulate Ca²⁺ released from the ER. Lysosomes thereby shape the cytosolic Ca²⁺ signals evoked by activation of Ins(1,4,5)P₃R without affecting those evoked by SOCE.

Results

Dissipating the lysosomal H⁺ gradient enhances carbachol-evoked Ca²⁺ signals

Activation of endogenous muscarinic receptors in human embryonic kidney (HEK) cells with carbachol (CCh) stimulates phospholipase C (PLC), formation of Ins(1,4,5)P₃, release of Ca²⁺ from intracellular stores, and stimulation of Ca²⁺ entry (Tovey et al., 2008). The initial release of Ca²⁺ from intracellular stores is entirely mediated by Ins(1,4,5)P₃R and unaffected by inhibition of RyR or TPC (supplementary material Fig. S1).

Ca²⁺ accumulation by acidic organelles, including lysosomes, requires a transmembrane H⁺ gradient. It is probably mediated by a Ca²⁺/H⁺ exchanger or indirectly via a Ca²⁺/Na⁺ exchanger. The identities of these exchangers in mammalian cells are unknown (Klemper, 1985; Morgan et al., 2011). We used bafilomycin A₁ to inhibit the vacuolar type H⁺-ATPase (V-ATPase) (Yoshimori et al., 1991), dissipate the H⁺ gradient and so prevent Ca²⁺ accumulation by acidic organelles (Fig. 1A). LysoTracker Red, a weak base that accumulates in acidic organelles (Bucci et al., 2000), substantially colocalised with a GFP-tagged lysosomal membrane protein (lysosomal-associated membrane protein 1; LAMP1–GFP) (Fukuda, 1991) transiently expressed in HEK cells (supplementary material Fig. S2). Bafilomycin A₁ (1 µM, 1 h) almost abolished staining with LysoTracker Red without affecting the distribution of LAMP1–mCherry (supplementary material Fig. S3). This demonstrates that lysosomes are the major acidic organelles in HEK cells, and that bafilomycin A₁ discharges their H⁺ gradient without affecting their distribution.

Fig. 1.

Inhibition of H⁺ accumulation by acidic organelles exaggerates the increase in [Ca²⁺]_i evoked by receptors that stimulate phospholipase C. (A) Populations of HEK cells in Ca²⁺-free HBS in 96-well plates were stimulated with a maximal concentration of CCh (1 mM), with (red) or without (black) pre-incubation with bafilomycin A₁ (Baf A₁; 1 µM, 1 h). Baf A₁, an inhibitor of the V-ATPase, dissipates H⁺ gradients across the membranes of acidic organelles and thereby diminishes their ability to accumulate Ca²⁺ by an unidentified pathway that may be a Ca²⁺/H⁺ exchanger (inset). Results are means ± s.e.m. for three wells from a single experiment, each typical of at least three similar experiments. (B) Summary results (means ± s.e.m., n = 3 independent experiments) from experiments similar to (A) show concentration-dependent effects of CCh on Ca²⁺ release with and without pre-incubation with bafilomycin A₁. (C) Single-cell analyses in Ca²⁺-free HBS show the peak increase in [Ca²⁺]_i evoked by CCh (1 mM) with and without pre-incubation with bafilomycin A₁ (1 µM, 1 h). Results are means ± s.e.m. from four coverslips, with 50 cells analysed on each. *P<0.05. (D) Intracellular pH (pH_i) was measured in populations of HEK cells with (solid bars) or without (open bars) pre-incubation with bafilomycin A₁ (1 µM, 1 h), vacuolin (10 µM, 1 h) or GPN (200 µM, 10 min) with cathepsin inhibitor 1 (CI; 10 µM, 10 min). Each treatment is paired with its solvent control (0.1–1.1% DMSO). Results are means ± s.e.m. from three independent experiments. (E) COS-7 cells in HBS were stimulated with ATP (10 µM) with or without pre-incubation with bafilomycin A₁ (1 µM, 1 h). Results show means for at least 40 cells from two coverslips. (F,G) Single-cell analyses of COS-7 cells in HBS show the peak increase in [Ca²⁺]_i evoked by ATP (F) and the basal [Ca²⁺]_i (G) with and without pre-incubation with bafilomycin A₁ (1 µM, 1 h). Results are means ± s.e.m. from three coverslips, with ≥30 cells on each analysed. *P<0.05.

Pre-incubation of HEK cells with bafilomycin A₁ caused the increase in [Ca²⁺]_i evoked by a maximally effective concentration of CCh to increase by 1.7±0.1-fold without affecting the sensitivity to CCh (pEC₅₀ = 4.3±0.2 and 4.5±0.2 for control and bafilomycin A₁-treated cells, respectively; where pEC₅₀ is the −log of the half-maximally effective concentration; Fig. 1A,B). Bafilomycin A₁ also caused a small, but significant (P = 0.0035), increase in the [Ca²⁺]_i of unstimulated cells from 30±2 nM to 46±2 nM (n = 5). This modest effect of bafilomycin A₁ is probably due largely to basal activity of Ins(1,4,5)P₃R since it was abolished by inhibition of PLC (supplementary material Fig. S1E). Analyses of single cells confirmed the results from cell populations. The number of cells responding to a maximal concentration of CCh was similar for control and bafilomycin A₁-treated cells (98±1% and 96±3%, respectively) and the increase in [Ca²⁺]_i was 1.9±0.3-fold greater after treatment with bafilomycin A₁ (Fig. 1C; supplementary material Table S1).

Stimulation of HEK cells with a submaximal concentration of CCh (50 µM) evoked sustained Ca²⁺ oscillations, typical of the responses of many cells to physiological stimuli (supplementary material Fig. S4A,B). Bafilomycin A₁ had no effect on the number of cells in which CCh evoked Ca²⁺ oscillations (38±9% and 50±6% in control and bafilomycin-treated cells, respectively), but it increased the amplitude of the Ca²⁺ spikes and reduced their frequency (supplementary material Fig. S4C,D).

The affinities of Ca²⁺ indicators and other Ca²⁺-binding sites for Ca²⁺ and of Ins(1,4,5)P₃R for Ins(1,4,5)P₃ are pH sensitive. We were therefore concerned that bafilomycin A₁ might, via effects on cytosolic pH (pH_i), affect the responses observed. Using SNARF-5F to measure pH_i, we established that neither bafilomycin A₁ nor any of the other inhibitors used affected pH_i (Fig. 1D).

These results establish that the increase in [Ca²⁺]_i evoked by CCh in HEK cells is potentiated by dissipating the lysosomal H⁺ gradient. Similar results were obtained with COS-7 cells, where activation of P2Y receptors by ATP evoked Ca²⁺ release from intracellular stores. Bafilomycin A₁ caused the peak Ca²⁺ signal evoked by ATP to increase by 1.5±0.1 fold (Fig. 1E,F). However, in COS-7 cells, the exaggerated response to ATP occurred without an increase in basal [Ca²⁺]_i (Fig. 1G).

Perforating lysosomal membranes potentiates the increase in [Ca²⁺]_i evoked by CCh

Glycyl-L-phenylalanine 2-naphthylamide (GPN) allows selective disruption of lysosomes because its cleavage by the lysosomal enzyme, cathepsin C, causes osmotic swelling and thereby perforation of lysosomal membranes (Berg et al., 1994) (Fig. 2A). Treatment of HEK cells with GPN abolished staining with LysoTracker Red without affecting the distribution of LAMP1–mCherry (Fig. 2B) or pH_i (Fig. 1D). GPN caused the peak Ca²⁺ signal evoked by a maximal concentration of CCh in Ca²⁺-free HEPES-buffered saline (HBS) to increase by 1.5±0.1-fold (Fig. 2C), without affecting the sensitivity to CCh (pEC₅₀ = 4.68±0.03 and 4.64±0.06 for control and GPN-treated cells; Fig. 2D). GPN also caused a modest increase in basal [Ca²⁺]_i (22±4 nM and 42±5 nM in control and GPN-treated cells, P = 0.025; Fig. 2C). To minimize any non-specific effects arising from release of lysosomal contents into the cytosol, we reduced the period of incubation with GPN from 30 to 10 min and included cathepsin inhibitor 1 (10 µM), which inhibits many lysosomal proteases, but not the cathepsin C that cleaves GPN (Demuth et al., 1996). The effects of GPN on CCh-evoked Ca²⁺ signals were unaffected by this revised protocol: the peak [Ca²⁺]_i signal was increased by 1.6±0.2-fold, with no effect on the sensitivity to CCh (pEC₅₀ = 4.5±0.1 and 4.3±0.2 in control cells and those treated with GPN and cathepsin inhibitor 1; Fig. 2E).

Fig. 2.

Perforation of lysosomal membranes potentiates the increase in [Ca²⁺]_i evoked by CCh. (A) GPN is hydrolysed by the lysosomal enzyme, cathepsin C, causing osmotic disruption of lysosomal membranes. (B) TIRFM images of HEK cells stained with LysoTracker Red (50 nM, 1 h; upper panels) or transfected with LAMP1–mCherry (lower panels) followed by incubation with or without GPN (200 µM, 10 min). Differential interference contrast (DIC) images are shown alongside the corresponding fluorescence images, each typical of those from three different coverslips. Scale bars: 20 µm in all images. (C) Populations of HEK cells in Ca²⁺-free HBS were stimulated with CCh (1 mM) with or without pre-incubation with GPN (200 µM, 30 min). Results are means ± s.e.m. for five wells from two experiments, each typical of at least three similar experiments. (D,E) Summary results (means ± s.e.m. from three independent experiments) show effects of GPN alone (200 µM, 30 min; D) or GPN (200 µM, 10 min) with cathepsin inhibitor 1 (10 µM, 10 min; E) on the concentration-dependent effects of CCh on the Ca²⁺ signals evoked in Ca²⁺-free HBS.

Bafilomycin A₁ is expected to inhibit acidification of most acidic organelles, but GPN selectively disrupts lysosomes (Berg et al., 1994). It is therefore significant that GPN abolishes staining by LysoTracker Red (Fig. 2B), confirming that lysosomes are the major acidic organelles in HEK cells. Furthermore, the effects of bafilomycin A₁ (Fig. 1) and GPN (Fig. 2) on the Ca²⁺ signals evoked by CCh are similar (supplementary material Table S1), confirming that potentiation of Ca²⁺ signals by bafilomycin A₁ is due to its effects on lysosomes.

Fusion of lysosomes increases the amplitude of CCh-evoked Ca²⁺ signals

Vacuolin, by an unknown mechanism, causes fusion of lysosomes (Fig. 3A) (Huynh and Andrews, 2005). In HEK cells transiently expressing LAMP1–mCherry, vacuolin caused lysosomes to fuse (Fig. 3B) without affecting pH_i (Fig. 1D). Vacuolin caused a 1.4±0.1-fold increase in the peak [Ca²⁺]_i evoked by CCh without affecting the sensitivity to CCh (pEC₅₀ = 4.7±0.1 and 4.9±0.1 for control and vacuolin-treated cells; Fig. 3C,D). Single-cell analyses confirmed that vacuolin had no effect on the number of cells that responded to CCh (93±2% and 91±1% for control and vacuolin-treated cells), but increased the amplitude of the CCh-evoked Ca²⁺ signal by 2.0±0.2-fold (Fig. 3E). Vacuolin had no effect on basal [Ca²⁺]_i in either single cells or cell populations (25±4 nM and 30±2 nM in control and vacuolin-treated cells, P = 0.31, supplementary material Table S1). The combined effects of maximal concentrations of bafilomycin A₁ and vacuolin on the peak Ca²⁺ signals evoked by CCh were no different from the effects of either treatment alone (Fig. 3F). The results with vacuolin are important because vacuolin is not expected to affect lysosomal ion transport mechanisms, but instead to change the surface area to volume ratio of lysosomes and thereby to perturb interactions with other organelles.

Fig. 3.

Fusion of lysosomes enhances the Ca²⁺ signals evoked by CCh. (A) Vacuolin causes fusion of lysosomes. (B) TIRFM images of single HEK cells transiently expressing LAMP1–mCherry, before and after treatment with vacuolin (10 µM, 1 h). Images are typical of results from three different coverslips. (C) Populations of HEK cells in Ca²⁺-free HBS were stimulated with CCh (1 mM) with or without pre-incubation with vacuolin (10 µM, 1 h). Results are means ± s.e.m. for three wells from a single experiment, each typical of at least five similar experiments. (D) Summary results (means ± s.e.m. from five independent experiments) show effects of vacuolin on the concentration-dependent effects of CCh in Ca²⁺-free HBS. (E) Single-cell analyses in Ca²⁺-free HBS show the peak increase in [Ca²⁺]_i evoked by CCh (1 mM) with and without pre-incubation with vacuolin (10 µM, 1 h). Results are means ± s.e.m. from four coverslips, with ∼50 cells on each analysed. (F) Populations of HEK cells in Ca²⁺-free HBS were stimulated with CCh (1 mM) without or with pre-incubation with vacuolin (100 µM, 1 h), bafilomycin A₁ (Baf A₁; 1 µM, 1 h) or both treatments combined. Results are means ± s.e.m. from four independent experiments). *P<0.05.

Because some, though not all, treatments used to manipulate lysosomal activity caused modest increases in basal [Ca²⁺]_i (Fig. 1E; supplementary material Table S1), we were concerned that potentiated responses to Ins(1,4,5)P₃ might result from enhanced Ca²⁺ uptake by the ER. We therefore used a novel low-affinity Ca²⁺ indicator targeted to the ER, CatchER (Tang et al., 2011), to measure directly the free [Ca²⁺] within the ER of COS-7 cells. These are better suited than HEK cells for these studies because COS-7 cells are larger, flatter and their ER is more easily identified. The results demonstrate that under conditions where bafilomycin A₁ increased the peak [Ca²⁺]_i evoked by ATP, it had no effect on the free [Ca²⁺] within the ER (supplementary material Fig. S5). The larger increases in [Ca²⁺]_i evoked by PLC-linked receptors in the presence of bafilomycin A₁ are not therefore due to increased loading of the ER with Ca²⁺.

Three means of perturbing the behaviour of lysosomes, inhibition of the V-ATPase (Fig. 1), disruption of lysosomal membranes (Fig. 2) or modification of lysosomal morphology (Fig. 3), indistinguishably and non-additively (Fig. 3F) potentiate the Ca²⁺ signals evoked by CCh. Because CCh-evoked Ca²⁺ signals are entirely mediated by Ins(1,4,5)P₃R (supplementary material Fig. S1), we examined the effect of bafilomycin A₁ on the Ca²⁺ signals evoked by directly activating Ins(1,4,5)P₃R.

Lysosomes accumulate Ca²⁺ released by direct activation of Ins(1,4,5)P₃ receptors

Flash photolysis of caged Ins(1,4,5)P₃ (ci-Ins(1,4,5)P₃) non-disruptively loaded into cells allows Ins(1,4,5)P₃ to be delivered directly to the cytosol (Dakin and Li, 2007). Photolysis of ci-Ins(1,4,5)P₃ caused transient increases in [Ca²⁺]_i, the amplitudes of which varied between cells (Fig. 4A), probably reflecting differences between cells in loading and/or de-esterification of ci-Ins(1,4,5)P₃/PM. In a large sample (>500 cells), the peak response to photorelease of ci-Ins(1,4,5)P₃ was 1.3±0.1-fold greater in bafilomycin A₁-treated cells (Fig. 4A–C). This confirms that when Ins(1,4,5)P₃R are directly activated by Ins(1,4,5)P₃, Ca²⁺ signals are potentiated by inhibition of the lysosomal V-ATPase. The increase in [Ca²⁺]_i evoked by photolysis of ci-Ins(1,4,5)P₃ recovered with mono-exponential kinetics (Fig. 4B), but recovery was much slower (P = 0.029) in cells treated with bafilomycin A₁ (half-time, t_1/2 = 45.6±7.8 s) than in control cells (t_1/2 = 18.9±1.7 s; Fig. 4D). Bafilomycin A₁ also slowed recovery from CCh-evoked Ca²⁺ signals (supplementary material Fig. S6). The sustained effects of bafilomycin A₁ on the Ca²⁺ signals evoked by CCh or photolysis of ci-Ins(1,4,5)P₃ probably reflect cycling of Ca²⁺ through the ER fuelled by the activity of the Ca²⁺ pump of the ER (SR/ER Ca²⁺-ATPase; SERCA) and sustained activity of Ins(1,4,5)P₃R. These results confirm that bafilomycin A₁ potentiates Ins(1,4,5)P₃- or CCh-evoked Ca²⁺ signals by inhibiting removal of Ca²⁺ from the cytosol through lysosomal uptake systems.

Fig. 4.

Disruption of lysosomal Ca²⁺ uptake with bafilomycin A₁ exaggerates the increase in [Ca²⁺]_i evoked by direct activation of Ins(1,4,5)P₃R. (A) Typical increases in fluo-4 fluorescence (RFU, relative fluorescent units) from single HEK cells after flash photolysis of ci-Ins(1,4,5)P₃ in control cells and cells treated with bafilomycin A₁ (Baf A₁; 1 µM, 1 h). The UV flash that caused photolysis of ci-Ins(1,4,5)P₃ was delivered at the times shown by arrowheads. (B) Similar experiments to those in A (reported as F/F_o, see Materials and Methods) showing means ± s.e.m. from a single field of at least 40 cells. Each trace is typical of at least three similar experiments. (C) From experiments similar to those shown in B, average peak responses evoked by photolysis of ci-Ins(1,4,5)P₃ in control and bafilomycin A₁-treated cells are expressed as a percentage of the response in control cells. Results are means ± s.e.m. from three independent experiments. At least three regions of each coverslip, each including 30–70 cells, were analysed. (D) Half-times (t_1/2) for rates of recovery of [Ca²⁺]_i in control and bafilomycin A₁-treated cells (1 µM, 1 h). Results are means ± s.e.m. from three independent experiments. At least three regions of each coverslip (n = 1), each including 30–70 cells, were analysed. *P<0.05. Similar analyses of recovery from CCh-evoked Ca²⁺ signals are shown in supplementary material Fig. S6.

Bafilomycin A₁ also potentiated the transient Ca²⁺ signals evoked by inhibiting the SERCA with thapsigargin in nominally Ca²⁺-free HBS (Fig. 5A,B). Similar results (1.9±0.2-fold increase in the peak Ca²⁺ signal, n = 3) were obtained when cyclopiazonic acid (CPA) was used to inhibit the SERCA (supplementary material Fig. S7). The route by which Ca²⁺ leaks from the ER is undefined, but it may include contributions from translocons (Lang et al., 2011) and Ins(1,4,5)P₃R (supplementary material Fig. S1). Both channels have large Ca²⁺ conductances that might be expected to generate the large local increases in [Ca²⁺]_i that we suggest are required for lysosomal Ca²⁺ uptake.

Fig. 5.

Disruption of lysosomal Ca²⁺ uptake with bafilomycin A₁ does not affect SOCE. (A) Transient increase in [Ca²⁺]_i evoked by thapsigargin (1 µM, solid bar) in HEK cells in nominally Ca²⁺-free HBS in control and bafilomycin A₁-treated cells (Baf A₁; 1 µM, 1 h). (B) Summary results show effects of bafilomycin A₁ (1 µM, 1 h) on the peak increase in [Ca²⁺]_i evoked by thapsigargin (1 µM) in nominally Ca²⁺-free HBS. Results are means ± s.e.m. from three independent experiments. (C) Restoration of extracellular Ca²⁺ (30 mM) to cells pre-treated with thapsigargin (1 µM, 15 min) in nominally Ca²⁺-free HBS to deplete intracellular Ca²⁺ stores evokes SOCE. The response is indistinguishable in control cells and cells treated with bafilomycin A₁ (1 µM, 1 h). (D) SOCE after restoration of different concentrations of extracellular Ca²⁺ ([Ca²⁺]_e) to control and bafilomycin A₁-treated cells (1 µM, 1 h). Results are means ± s.e.m. from three independent experiments. (E) TPEN (100 µM, 2 min) in nominally Ca²⁺-free HBS was used to reduce the free [Ca²⁺] within the ER before restoration of the indicated concentrations of extracellular Ca²⁺ to control or bafilomycin A₁-treated cells (1 µM, 1 h). Results show the peak increase in [Ca²⁺]_i detected within 60 s after restoration of extracellular Ca²⁺. (F) Peak increase in [Ca²⁺]_i evoked by CCh (1 mM) in the presence of TPEN (100 µM, 2 min) with and without pre-incubation with bafilomycin A₁ (1 µM, 1 h). Results are means ± s.e.m. from three (E) or four (F) independent experiments. Examples of traces from which the summary results shown in E and F were derived are shown in supplementary material Fig. S8. *P<0.05.

Ca²⁺ signals evoked by store-operated Ca²⁺ entry are unaffected by lysosomes

Our results so far demonstrate that Ca²⁺ released from the ER via Ins(1,4,5)P₃R or leak pathways (after addition of CPA or thapsigargin) can be accumulated by lysosomes. We next assessed whether Ca²⁺ entering the cell across the plasma membrane via SOCE is also accumulated by lysosomes. Restoration of extracellular Ca²⁺ to cells treated with thapsigargin in nominally Ca²⁺-free HBS evoked sustained increases in [Ca²⁺]_i (Fig. 5C). This is consistent with abundant evidence for SOCE in HEK cells (Parekh and Putney, 2005). Manipulating the extracellular [Ca²⁺] allowed the amplitude of the global increase in [Ca²⁺]_i evoked by SOCE to match and exceed (Fig. 5D) the peak Ca²⁺ signals evoked by CCh in Ca²⁺-free HBS (Fig. 1B). Nevertheless, the Ca²⁺ signals evoked by SOCE were entirely insensitive to bafilomycin A₁ (Fig. 5C,D). Similar results were obtained when CPA (100 µM, 15 min) was used to evoke SOCE: the peak Ca²⁺ signals evoked by addition of HBS containing 30 mM Ca²⁺ were 180±40 nM and 203±48 nM in control and bafilomycin A₁-treated cells, respectively (supplementary material Fig. S7). A possible concern is that Ca²⁺ released from the ER after addition of thapsigargin or CPA is accumulated by lysosomes (Fig. 5A,B) and might thereby limit their capacity to accumulate further Ca²⁺ entering the cell via SOCE. To address this issue, we used a membrane-permeant low-affinity Ca²⁺ buffer, TPEN (N,N,N′,N′-tetrakis(2-pyridylmethyl)-1,2-ethylenediamide), to reduce the free [Ca²⁺] within the ER and so activate SOCE without causing Ca²⁺ release (Hofer et al., 1998) (supplementary material Fig. S8). Restoration of extracellular Ca²⁺ to HEK cells treated with TPEN (100 µM, 2 min) stimulated SOCE-mediated increases in [Ca²⁺]_i that were insensitive to bafilomycin A₁ (Fig. 5E). CCh-evoked Ca²⁺ release in these TPEN-treated cells was potentiated by bafilomycin A₁ (Fig. 5F; supplementary material Fig. S8). There are two important conclusions. First, even when Ca²⁺ within the ER is heavily buffered by TPEN, responses to CCh are potentiated by bafilomycin A₁. This reinforces our conclusion (supplementary material Fig. S5) that potentiation of Ins(1,4,5)P₃-evoked Ca²⁺ signals by inhibitors of lysosomes is not due to enhanced Ca²⁺ loading of the ER. Second, under conditions where lysosomes can attenuate the increases in [Ca²⁺]_i evoked by CCh, they have no effect on SOCE-evoked Ca²⁺ signals. We conclude that lysosomes accumulate Ca²⁺ released from the ER (Figs 1–4), but they do not accumulate Ca²⁺ entering the cell via SOCE (Fig. 5).

CCh increases the luminal pH of lysosomes

So far, our analyses of the contribution of lysosomes to shaping Ins(1,4,5)P₃-evoked Ca²⁺ signals have relied on measurements of [Ca²⁺]_i. Complementary analyses of free [Ca²⁺] within lysosomes ([Ca²⁺]_ly) are more challenging. Indicators can be directed to the lysosome lumen by endocytosis, but their affinity for Ca²⁺ is reduced by the low lysosomal pH (∼pH 4) (Christensen et al., 2002; Lloyd-Evans et al., 2008). We assessed the effects of CCh on lysosomal pH (pH_ly) and [Ca²⁺]_ly using pairs of endocytosed dextran-conjugated indicators: an inert marker (Texas Red, TR) and either Oregon Green (OG, to measure pH) or Oregon Green BAPTA (OGB, to measure [Ca²⁺]; Fig. 6).

Fig. 6.

CCh causes an increase in lysosomal pH and [Ca²⁺]. (A) Wide-field fluorescence images of HEK cells loaded with dextran-conjugates of Oregon Green (OG) and Texas Red (TR), and the corresponding DIC image, each representative of at least three coverslips. Scale bar applies to all images. (B) Changes in OG and TR fluorescence evoked by CCh (1 mM) are shown as means ± s.e.m. from 28 regions of interest (ROI) from a single coverslip, representative of at least three independent experiments. (C) Summary results (means ± s.e.m.; nine traces from six independent coverslips) show peaks in OG and TR fluorescence measured within 7 min of addition of either HBS or HBS with CCh (1 mM). (D) pH_i measured in populations of HEK cells before and after stimulation with CCh (1 mM, 2 and 5 min) in the absence and presence of extracellular Ca²⁺. (E) Images similar to those shown in A, but with cells loaded with TR and dextran-conjugated Oregon Green BAPTA (OGB); each representative of at least three coverslips. (F) Changes in OGB and TR fluorescence evoked by CCh (1 mM) are shown as means ± s.e.m. from 38 ROIs from a single coverslip, representative of at least three independent experiments. (G) Summary results (means ± s.e.m.; seven traces from four independent coverslips) show peaks in OGB and TR fluorescence measured within 7 min of addition of either HBS or HBS with CCh (1 mM). *P<0.05.

In populations of HEK cells, CCh (1 mM) evoked a sustained increase in OG fluorescence, without affecting TR fluorescence (Fig. 6B,C). This demonstrates that CCh caused an increase in pH_ly. Parallel measurements of cytosolic pH (pH_i) established that in both the presence and absence of extracellular Ca²⁺, CCh evoked a slow and very modest decrease in pH_i (Fig. 6D). The effect was statistically significant (P = 0.009) only for cells stimulated with CCh for 5 minutes in the presence of extracellular Ca²⁺, when pH_i fell from 7.42±0.02 to 7.34±0.01 (Fig. 6D). These results demonstrate that under conditions where pH_i is either stable or very modestly reduced (Fig. 6D), CCh evokes an increase in pH_ly (Fig. 6B,C). The effect of CCh on pH_ly cannot therefore be mediated by changes in pH_i. It is more likely to be caused by CCh-evoked Ca²⁺ signals and subsequent lysosomal Ca²⁺ uptake by Ca²⁺/H⁺ exchange. This interpretation is consistent with evidence that addition of Ca²⁺ to isolated acidic organelles increases luminal pH (Morgan et al., 2011).

It is more difficult to measure directly the free [Ca²⁺] within lysosomes because the Ca²⁺ affinity of OGB, like all other Ca²⁺ indicators, is pH sensitive. Nevertheless, in similar analyses using TR and OGB, CCh selectively increased the fluorescence of the lysosomal Ca²⁺ indicator (OGB; Fig. 6E–G). These direct measurements of pH_ly and indirect measurement of [Ca²⁺]_ly concur with the analyses of [Ca²⁺]_i (Figs 1–5) by suggesting that lysosomes sequester Ca²⁺ released from intracellular stores.

Lysosomes are motile and associated with ER

We used total internal reflection fluorescence microscopy (TIRFM) to examine the distribution of ER and lysosomes in COS-7 cells by labelling ER with GFP–Ins(1,4,5)P₃R1, GFP–Ins(1,4,5)P₃R3 or GFP–ER, all of which colocalised with co-transfected SERCA–mCherry (supplementary material Fig. S9). Lysosomes, identified with either LysoTracker Red or LAMP1–mCherry, were closely associated with the ER (Fig. 7A): 79±3% (n = 7 cells) of lysosomes were close enough to the ER that their separation could not be resolved by TIRFM. As reported for other cells (Matteoni and Kreis, 1987), lysosomes were remarkably motile within COS-7 cells (supplementary material Movie 1).

Fig. 7.

Close association of dynamic lysosomes with ER. (A) TIRFM images of live COS-7 cells transfected with GFP–ER, GFP–Ins(1,4,5)P₃R1 (GFP-IP₃R1) or GFP–Ins(1,4,5)P₃R3 (GFP–IP₃R3) (each shown in green) and co-labelled with either LysoTracker Red (50 nM, 1 h) or co-transfected with LAMP1-mCherry (red). The right-hand column shows magnified images of the boxed regions. The colocalization of lysosome and ER markers was statistically significant (P<0.05) with R_coloc = 0.59±0.1 (LAMP1–mCherry and GFP–ER, n = 5 cells), 0.92±0.003 (LAMP1–mCherry and GFP–Ins(1,4,5)P₃R1, n = 3 cells), 0.82±0.04 (LAMP1–mCherry and GFP–Ins(1,4,5)P₃R3; n = 3 cells), 0.70±0.02 (LysoTracker Red with GFP–ER, n = 3 cells), 0.74±0.06 (LysoTracker Red and GFP–Ins(1,4,5)P₃R1; n = 3 cells) and 0.80±0.009 (LysoTracker Red and GFP–Ins(1,4,5)P₃R3; n = 3 cells). (B) Time-lapse TIRFM images showing a live COS-7 cell co-transfected with GFP–ER (green) and LAMP1–mCherry (red). The interval between each image (1–9) is 16.1 s (every 50th frame of a high-acquisition rate video where individual lysosomes could be tracked accurately). Brightness and contrast were auto-adjusted for each image (Cell∧R software) to mitigate effects of bleaching. Downward arrows indicate a lysosome moving along the ER, and upward arrows indicate a lysosome residing at the leading edge of an extending ER tubule. The complete sequence is shown in supplementary material Movie 1.

Video-rate imaging of COS-7 cells co-transfected with GFP–ER and LAMP1–mCherry allowed tracking of individual lysosomes and showed that lysosomes maintain their association with the ER over prolonged periods (often for the entire 2-minute recording) as they move around the cell (Fig. 7B; supplementary material Movie 1). This movement often occurred along ER tubules, and occasionally lysosomes localised at the tips of ER tubules and moved concomitantly with ER tubule extension, further supporting a close spatial relationship between ER and lysosomes.

Discussion

Lysosomes accumulate Ca²⁺ released from the ER

Disruption of lysosomal Ca²⁺ uptake mechanisms by preventing luminal acidification (Figs 1, 4) or by perforation of lysosomal membranes (Fig. 2), or changing the morphology of lysosomes and so their relationships with other organelles (Fig. 3) exaggerates the cytosolic Ca²⁺ signals evoked by Ins(1,4,5)P₃. This occurs whether Ins(1,4,5)P₃ is delivered by activation of endogenous receptors in the PM (Figs 1–3) or by photolysis of ci-Ins(1,4,5)P₃ (Fig. 4), and it occurs in both HEK and COS-7 cells. Despite the increased Ca²⁺ signals, there was no change in CCh sensitivity, suggesting that the treatments affected neither the intracellular concentrations of Ins(1,4,5)P₃ nor the sensitivity of Ins(1,4,5)P₃R. The decrease in frequency, but increase in amplitude, of CCh-evoked Ca²⁺ spikes after bafilomycin A₁ treatment (supplementary material Fig. S4) is also inconsistent with a simple increase in either Ins(1,4,5)P₃ concentration or Ins(1,4,5)P₃R sensitivity because either would be expected to increase the frequency of Ca²⁺ oscillations. Furthermore, CCh-evoked Ca²⁺ release is accompanied by a sustained increase in lysosomal pH and perhaps lysosomal [Ca²⁺] (Fig. 6), consistent with sequestration of Ca²⁺ by a lysosomal Ca²⁺/H⁺ exchanger.

In HEK cells, inhibition of lysosomal Ca²⁺ accumulation with bafilomycin A₁ or GPN was accompanied by a small (<20 nM) increase in basal [Ca²⁺]_i. Nevertheless, we were concerned that these small increases in basal [Ca²⁺]_i might contribute to the increased amplitude of the Ins(1,4,5)P₃-evoked Ca²⁺ signals by enhancing loading of Ca²⁺ stores or by sensitization of Ins(1,4,5)P₃R. The latter is unlikely because none of the treatments affected the sensitivity to CCh (pEC₅₀; Fig. 1B; Fig. 2D,E; Fig 3D; and see above) and they potentiated the Ca²⁺ signals evoked by Ca²⁺ leaks from the ER (Fig. 5A; supplementary material Fig. S7A). Additional evidence establishes that enhanced Ca²⁺ loading of the ER is not the explanation. First, bafilomycin A₁ enhanced the increase in [Ca²⁺]_i evoked by ATP in COS-7 cells without affecting basal [Ca²⁺]_i (Fig. 1E–G) and vacuolin mimicked the effects of bafilomycin A₁ and GPN on CCh-evoked Ca²⁺ signals in HEK cells without affecting basal [Ca²⁺]_i (supplementary material Table S1). Second, direct measurement of luminal [Ca²⁺] within the ER demonstrated that bafilomycin A₁ did not affect Ca²⁺ uptake by the ER (supplementary material Fig. S5), and buffering ER Ca²⁺ with TPEN did not prevent bafilomycin A₁ from potentiating CCh-evoked Ca²⁺ signals (Fig. 5F). Third, after CCh stimulation (supplementary material Fig. S6) or flash photolysis of ci-Ins(1,4,5)P₃ (Fig. 4), [Ca²⁺]_i recovered more slowly in bafilomycin A₁-treated cells, confirming that bafilomycin A₁ inhibits Ca²⁺ removal from the cytosol. Finally, CCh increased pH_ly and perhaps [Ca²⁺]_ly in the absence of inhibitors (Fig. 6), suggesting that Ca²⁺ released via Ins(1,4,5)P₃R is sequestered by lysosomes.

We conclude that the increased amplitude of the Ca²⁺ signals evoked by Ins(1,4,5)P₃ after disruption of lysosomes is due to their diminished ability to sequester Ca²⁺ released from the ER. These observations are consistent with studies where thapsigargin evoked larger cytosolic Ca²⁺ signals (Pereira et al., 2010), and lysosomal Ca²⁺ uptake was diminished (Lloyd-Evans et al., 2008) in cells with lysosomal storage disorders.

ER and lysosomes are intimately associated

CCh evokes Ca²⁺ signals in HEK cells that are initiated by Ins(1,4,5)P₃R and then sustained by SOCE. Although many lysosomes were present in the TIRFM field, suggesting that they lie within ∼100 nm of the PM, they did not sequester Ca²⁺ entering via SOCE even when the global increases in [Ca²⁺]_i exceeded those evoked by Ins(1,4,5)P₃. This demonstrates that lysosomes selectively sequester Ca²⁺ released from the ER. There are several implications of this conclusion. First, the global elevations in [Ca²⁺]_i that occur as Ca²⁺ diffuses away from open channels are insufficient to allow significant stimulation of Ca²⁺ uptake by lysosomes. This suggests that the lysosomal uptake system responsible for these effects has a low affinity for Ca²⁺. The sparse studies of Ca²⁺ uptake by isolated lysosomes differ enormously in their estimates of the affinity (K_m) for Ca²⁺: 108 nM and 5.7 mM for lysosomes isolated from neutrophils (Styrt et al., 1988) and fibroblasts (Lemons and Thoene, 1991), respectively. The significance of the latter study is unclear because the incubations excluded ATP. Second, the lysosomal Ca²⁺ uptake system responsible for the effects observed in our study is unlikely to be a Ca²⁺ pump because pumps are too slow (e.g. turnover number ∼10 s⁻¹ for SERCA) (Lytton et al., 1992) to sequester Ca²⁺ effectively before it diffuses away from an open Ins(1,4,5)P₃R that probably conducts ∼500,000 Ca²⁺ s⁻¹ (Vais et al., 2010). Ca²⁺ exchangers, which have been proposed to mediate Ca²⁺ uptake by lysosomes (Lloyd-Evans and Platt, 2011) and are likely to have higher turnover numbers (up to 5000 s⁻¹ for the NCX1 Na⁺/Ca²⁺ exchanger, for example) (Hilgemann et al., 1991), are better able rapidly to sequester Ca²⁺ released by Ins(1,4,5)P₃R.

The third implication is the need for selective association of lysosomes and ER to allow lysosomes, some of which are no more than ∼100 nm from the PM (Fig. 7; supplementary material Movie 1), to ignore Ca²⁺ signals generated by SOCE, but accumulate Ca²⁺ released by Ins(1,4,5)P₃R. Local increases in [Ca²⁺]_i near Ins(1,4,5)P₃R with their large unitary currents are likely to be much greater (perhaps 50 µM) (Vais et al., 2010) than those near SOCE channels where unitary Ca²⁺ currents are ∼100-times smaller (Zweifach and Lewis, 1993). Ins(1,4,5)P₃R are probably better able than SOCE to generate high local [Ca²⁺]_i and so allow Ca²⁺ sequestration by low-affinity transport pathways. However, these high [Ca²⁺]_i dissipate steeply within ∼100 nm of the open channel (Shuai and Parker, 2005). Selective sequestration of Ca²⁺ by lysosomes probably requires that Ins(1,4,5)P₃R are within ∼100 nm of lysosomes. Our analyses are consistent with such an intimate association. Lysosomes and ER are closely associated and maintain their association during organelle movements (Fig. 7; supplementary material Movie 1).

ER–organelle junctions: a recurring theme in Ca²⁺ signalling

We have shown that dynamic lysosomes and ER are intimately associated (Fig. 7; supplementary material Movie 1), allowing lysosomes selectively to accumulate Ca²⁺ released from the ER. The reciprocal relationship is also important because in many cells NAADP-evoked Ca²⁺ release from lysosomes triggers Ca²⁺ release from the ER by Ca²⁺-induced Ca²⁺ release via Ins(1,4,5)P₃R (Calcraft et al., 2009) or RyR (Brailoiu et al., 2010; Cancela et al., 1999; Kinnear et al., 2008; Lee et al., 1997). Lysosome–ER junctions are reminiscent of those between mitochondria and ER (Fig. 8). The latter are maintained by specific tethering proteins between dynamic organelles (Csordás et al., 2006) and they allow mitochondria, despite the low affinity of their Ca²⁺ uptake pathway, to sequester Ca²⁺ released by Ins(1,4,5)P₃R (Rizzuto et al., 2009). Mitochondrion–ER junctions allow bi-directional interactions between organelles: mitochondria can relieve Ca²⁺ inhibition of Ins(1,4,5)P₃R by rapidly sequestering Ca²⁺ (Olson et al., 2010), while Ca²⁺ provided by ER Ca²⁺ channels allows mitochondrial Ca²⁺ uptake to regulate oxidative phosphorylation (Jouaville et al., 1999), apoptosis (Szalai et al., 1999) and mitochondrial motility (Yi et al., 2004). We suggest a similar bi-directional interplay between ER and lysosomes (Fig. 8). Ca²⁺ release via TPC2 can trigger Ca²⁺ release via RyR or Ins(1,4,5)P₃R in the ER (Brailoiu et al., 2010; Calcraft et al., 2009; Kinnear et al., 2008), and Ca²⁺ release via Ins(1,4,5)P₃R is selectively accumulated by lysosomes. The latter may regulate the behaviour of lysosomes by increasing lysosomal pH; by priming TPC2, which appears to be stimulated by luminal Ca²⁺ (Pitt et al., 2010), to respond to NAADP; and it may regulate endolysosomal trafficking (Luzio et al., 2010). Conversely, lysosomal Ca²⁺ sequestration shapes cytosolic Ca²⁺ signals, including the frequency and amplitude of the Ca²⁺ oscillations typically evoked by physiological stimuli, and by analogy with mitochondria (Olson et al., 2010) lysosomes may modulate feedback regulation of Ins(1,4,5)P₃R gating (Fig. 8).

Fig. 8.

Functional Ca²⁺ junctions: similarities between lysosomes and mitochondria. Mitochondria, via the low-affinity Ca²⁺ uniporter, rapidly accumulate Ca²⁺ when exposed to high local [Ca²⁺]_i around the mouth of open Ca²⁺ channels, such as Ins(1,4,5)P₃R (IP₃R). Our results suggest that lysosomes can also selectively accumulate Ca²⁺, presumably via a low-affinity uptake pathway, from within microdomains of substantially increased [Ca²⁺]_i around open Ins(1,4,5)P₃R. We have shown that lysosomes can thereby shape the Ca²⁺ signals evoked by Ins(1,4,5)P₃R. Mitochondria likewise regulate cytosolic Ca²⁺ signals both locally (by intercepting Ca²⁺-mediated interactions between Ins(1,4,5)P₃R) and more globally. The reverse interaction, where Ca²⁺ release from lysosomes via TPC2, selectively stimulates Ins(1,4,5)P₃R or RyR by Ca²⁺-induced Ca²⁺ release, seems to have no obvious parallel with mitochondria. However, Ca²⁺ uptake by mitochondria triggered by opening of Ins(1,4,5)P₃R regulates mitochondrial activity. We speculate that Ca²⁺ uptake by lysosomes may also regulate their activity by, for example, regulating lysosomal pH and thereby enzyme activity, endolysosomal trafficking and priming TPC2 to respond to NAADP.

Materials and Methods

Materials

Culture media, Lipofectamine 2000, LysoTracker Red DND-99, SNARF-5F/AM, fluo-4/AM, fura-2/AM, dextran-conjugated Texas Red (M_r, 70,000), dextran-conjugated Oregon Green BAPTA 488 (M_r, 10,000), dextran-conjugated Oregon Green 488 (M_r, 10,000) and the Ca²⁺ standard solutions used to calibrate fura-2 fluorescence signals to [Ca²⁺]_i were from Invitrogen (Paisley, UK). Cell culture plastics and 96-well assay plates were from Greiner (Stonehouse, UK). Imaging dishes (35-mm diameter with a 7-mm No. 0 glass insert) were from MatTek Corporation (Ashland, USA) or PAA Laboratories (Yeovil, UK). U73122 (1-[6-[[(17β)-3-methoxyestra-1,3,5(10)-trien-17-yl]amino]hexyl]-1H-pyrrole-2,5-dione) and CPA were from Tocris (Bristol, UK). Bafilomycin A₁ was from AG Scientific (California, USA) or Fluorochem (Hadfield, UK). Glycyl-L-phenylalanine 2-naphthylamide (GPN) was from Bachem (St. Helens, UK). Cathepsin inhibitor 1 was from Calbiochem (Nottingham, UK). BAPTA was from Molekula (Dorset, UK). CCh, ATP, DMSO, foetal bovine serum, poly-L-lysine and vacuolin were from Sigma-Aldrich (Poole, UK). Ionomycin was from MerkEurolab (Nottingham, UK). Thapsigargin was from Alomone Labs (Jerusalem, Israel). Caged cell-permeant Ins(1,4,5)P₃ (ci-Ins(1,4,5)P₃/PM) was from SiChem (Bremen, Germany). NED 19 and TPEN were from Enzo Life Sciences (Exeter, UK). Ryanodine was from Ascent Scientific (Bristol, UK).

Plasmids

Plasmids encoding LAMP1–mCherry and LAMP1–GFP were made by transferring the LAMP1 fragment from LAMP1–tdTomato (a gift from T. Carter, MRC National Institute for Medical Research, London, UK) (Babich et al., 2008) into pmCherry N1 or pAcGFP-N1 (Clontech, Mountain View, USA) respectively, using EcoRI/BamHI restriction sites. A plasmid encoding the ER marker, GFP–ER, in which GFP is attached to sequences that allow insertion into the cytosolic leaflet of the ER membrane, was a gift from V. J. Allan (University of Manchester, UK) (Woźniak et al., 2009). A plasmid encoding SERCA type-1 tagged at its C-terminus with mCherry was prepared by insertion of mCherry from pmCherry-N1 into SERCA-GFP (a gift from J. M. East, University of Southampton, UK) (Newton et al., 2003) using AgeI/NotI restriction sites. Plasmids encoding rat types 1 and 3 Ins(1,4,5)P₃R tagged at their N-termini with GFP have been described previously (Pantazaka and Taylor, 2011). The coding sequences of all plasmids were verified. A plasmid encoding CatchER, an EGFP-based Ca²⁺ indicator targeted to the ER lumen, was generously provided by J. J. Yang (Georgia State University, USA) (Tang et al., 2011).

Cell culture and transfection

HEK 293 cells were cultured at 37°C in Dulbecco's modified Eagle's medium/Ham's F12 with GlutaMAX and foetal bovine serum (10%) in a humidified atmosphere containing 95% air and 5% CO₂. Medium was replaced every third day, and cells were passaged when they reached ∼80% confluence. For experiments with cell populations, cells were seeded (∼2×10⁴ cells/well) into 96-well plates. For single-cell analyses or TIRFM, cells were seeded (1.2×10⁵ cells/well) onto either 22-mm round glass coverslips or glass-bottomed culture dishes pre-coated with 0.01% (w/v) poly-L-lysine. Cells were grown for a further 2–3 days before experiments or transfection. The latter used Lipofectamine 2000 according to the manufacturer's instructions with 1 µg DNA/well for cells in glass-bottomed culture dishes.

COS-7 cells were cultured at 37°C in RPMI 1640 with foetal bovine serum (10%) in a humidified atmosphere containing 95% air and 5% CO₂ (Pantazaka and Taylor, 2011). For TIRFM experiments, COS-7 cells were transiently transfected with plasmids using Neon nucleofection (Invitrogen) with 5 µg DNA/100 µl cells (10⁵ cells). Cells were then plated onto poly-L-lysine-coated glass-bottomed culture dishes and used 1–2 days after transfection.

Measurements of [Ca²⁺]_i

Measurements of [Ca²⁺]_i in HEK cells were performed at 20°C in HBS (Tovey et al., 2008). HBS had the following composition: 135 mM NaCl, 5.9 mM KCl, 1.2 mM MgCl₂, 1.5 mM CaCl₂, 11.5 mM glucose, 11.6 mM HEPES, pH 7.3. Ca²⁺ was omitted from nominally Ca²⁺-free HBS, and replaced by BAPTA (10 mM) in Ca²⁺-free HBS. For cell populations, confluent cells in 96-well plates were loaded with fluo-4/AM (2 µM, 1 h, 20°C), washed and incubated for a further 1 h in HBS. Fluorescence, from which [Ca²⁺]_i was determined, was then measured using a fluorescence plate reader (FlexStation 3, MDS Analytical Devices, Wokingham, UK). Fluo-4 fluorescence was calibrated to [Ca²⁺]_i using the equation: [Ca²⁺]_i = K_D (F−F _min)/(F _max−F), using a K_D for Ca²⁺ of 345 nM. Fluorescence from Ca²⁺-saturated (F_max) and Ca²⁺ free (F_min) fluo-4 was measured in adjacent wells by addition of Triton X-100 (0.1%) with CaCl₂ (10 mM) or BAPTA (10 mM) (Tovey et al., 2008).

For single-cell imaging, confluent cultures of HEK or COS-7 cells on 22-mm round, poly-L-lysine-coated glass coverslips were loaded with fura-2/AM (2 µM, 1 h, 20°C), washed and incubated for a further 1 h in HBS. Fluorescence, detected at >510 nm after alternating excitation at 340 and 380 nm, was detected using an Olympus IX71 inverted fluorescence microscope with a Luca electron multiplying charge-coupled device (EMCCD) camera (Andor Technology, Belfast, UK) with a 40×1.35 NA objective and analysed using MetaFluor software (MDS Analytical Devices, Wokingham, UK). After correction for background fluorescence, fluorescence ratios (F₃₄₀/F₃₈₀) were calibrated to [Ca²⁺]_i using Ca²⁺ standard solutions (Tovey et al., 2008).

Measurement of lysosomal pH and [Ca²⁺]

Almost confluent cultures of HEK cells grown on poly-L-lysine-coated, glass-bottomed culture dishes, were incubated in culture medium with dextran-conjugated Texas Red (TR; 0.1 mg/ml) together with a dextran-conjugate (0.1 mg/ml) of either Oregon Green (OG) or Oregon Green BAPTA (OGB) for 12 h at 37°C, and then for 4 h without the indicators. Cells were then washed with HBS and fluorescence was recorded using an Olympus IX81 microscope with a 40×/1.35 NA objective. Cells were illuminated with a mercury xenon lamp using alternating filter sets: U-MNIBA (Olympus; excitation 470–495 nm, emission 510–550 nm) and LF561A (Semrock; excitation 550–570 nm, emission 580–630 nm) for OG/OGB and TR, respectively. Images were captured at 2-second intervals using an EMCCD camera (Andor iXon 897) and analysed using Cell∧R software (Olympus, Milton Keynes, UK). All records were corrected for background fluorescence determined under identical conditions from cells that had not been loaded with indicators. Fluorescence changes from defined regions of interest (ROI) were then expressed as F/F_o, where F_o and F denote the average fluorescence within the ROI at the start of the experiment (F_o) and at each time point (F).

Measurement of luminal free [Ca²⁺] in the ER

The luminal free [Ca²⁺] of the ER was measured using an ER-targeted Ca²⁺-sensor CatchER (Tang et al., 2011). COS-7 cells in 35-mm imaging dishes were transfected with the CatchER coding sequence in pcDNA3.1 (2 µg/well) using Lipofectamine 2000 (1∶1, DNA/Lipofectamine). After 48 h, cells were imaged using an Olympus IX81 TIRF microscope with 60× or 150× TIRFM objectives. In time-lapse experiments, images were captured using wide-field illumination (488 nm, 200-ms exposure time) with a 60× TIRF objective and an Andor iXon 897 EMCCD camera. Control cells were used to correct images for photobleaching (∼15% over 1 h).

Flash photolysis of caged Ins(1,4,5)P₃

HEK cells grown on poly-L-lysine-coated, glass-bottomed culture dishes were loaded with ci-Ins(1,4,5)P₃/PM (1 µM, 45 min) (Dakin and Li, 2007), and then with fluo-4/AM (2 µM) and ci-Ins(1,4,5)P₃/PM (1 µM) for a further 45 min. After washing and incubation in HBS for a further 45 min, cells were imaged (20°C) using an Olympus IX81 microscope equipped with a 40×/1.35 NA objective. Cells were illuminated with a 488-nm diode-based solid-state laser (Olympus Digital Laser Systems) and emitted fluorescence (500–550 nm) was captured with an EMCCD camera (Andor iXon 897). After 60 s, ci-Ins(1,4,5)P₃ was uncaged by exposing an entire field to three pulses of UV light (<345 nm, ∼1 ms, 3000 µF, 300 V, ∼170 J) delivered within ∼6 s using a JML-C2 xenon flash-lamp (Rapp OptoElectronic GmbH, Hamburg, Germany). Images were acquired at 1-second intervals with an Andor iXon 897 camera (512×512 pixels) and analysed using Cell∧R software (Olympus, Milton Keynes, UK). For these measurements of [Ca²⁺]_i using a non-ratiometric indicator (fluo-4), responses are reported as F/F_o, where F_o is the average fluorescence intensity recorded from a ROI immediately before flash photolysis, and F is the fluorescence intensity from the same region after the flash.

Measurement of cytosolic pH

Cytosolic pH (pH_i) was measured using the fluorescent pH-sensitive indicator, SNARF-5F (Liu et al., 2001). Cells in 96-well plates were incubated with SNARF-5F/AM (2 µM, 30 min), washed with HBS, and fluorescence (excitation at 561 nm, and emission at 580 and 640 nm) was measured using a FlexStation fluorescence plate reader. Fluorescence ratios (F₅₈₀/F₆₄₀) were calibrated to pH_i using standard pH solutions (Owen, 1992).

Total internal reflection fluorescence microscopy

For TIRFM, cells on poly-L-lysine-coated, glass-bottomed culture dishes were imaged using an Olympus IX81 microscope with 150×/1.45 NA or 60×/1.45 NA TIRF objectives. For staining of lysosomes, cells were incubated with LysoTracker Red DND-99 (50 nM, 1 h, 20°C). Cells were illuminated with 488 nm (for GFP) or 561 nm (for mCherry or LysoTracker Red) diode-based solid-state lasers, and images were acquired with an Andor iXon 897 EMCCD camera. With the filters used there was no significant crosstalk between green and red channels. Images were processed using Cell∧R software (Olympus, Milton Keynes, UK).

Analysis

Concentration-effect relationships for each experiment were individually fitted to Hill equations using non-linear curve-fitting (GraphPad Prism, version 5) and the results obtained from each (pEC₅₀, Hill coefficient h, maximal response) were pooled for statistical analysis and presentation.

Rates of recovery of [Ca²⁺]_i after flash photolysis of ci-Ins(1,4,5)P₃ were determined by fitting mono-exponential decay equations (GraphPad Prism, version 5) to the averaged responses of all cells within the field exposed to the UV-flash (30–70 cells/field). At least three such fields were analysed for each coverslip and the half-times (t_1/2) were pooled to produce a single value for each coverslip. Rates of recovery of [Ca²⁺]_i after stimulation of cell populations with CCh were determined by fitting bi-exponential decay equations (GraphPad Prism, version 5). Each determination comprises the average response from at least three wells in a single experiment.

For quantitative analyses of the colocalisation of two fluorophores, Cell∧R software was used to correct for background fluorescence using an area outside the cell and then to define ROI (∼200–400 µm²) within the peripheral cytoplasm that excluded the nucleus and perinuclear area. For each ROI examined (Fig. 7; supplementary material Figs S2, S9), there was a statistically significant (P<0.05) colocalisation of fluorophores defined using the Colocalisation Analysis/Colocalisation Test plugin (ImageJ). This applies the Costes randomization method with 100 iterations and ignores pixels in which there is no fluorescence from either fluorophore (Costes et al., 2004). Colocalisation was then quantified using the Colocalisation Analysis/Colocalisation Threshold plugin (ImageJ). This was applied to threshold images and then calculate Pearson's correlation coefficient (R_coloc), ignoring pixels with intensities below threshold. R_coloc = Σ(R_i−R_m)(G_i−G_m)/√Σ(R_i−R_m)²Σ(G_i−G_m)², where G_i and R_i are the intensities of individual green and red pixels respectively, and G_m and R_m are the mean intensities of green and red pixels. R_coloc = 1 denotes perfect colocalisation.

Student's t-test or 1-way ANOVA was used for statistical analyses with P<0.05 considered significant. All statistical analyses were performed on raw data, although for clarity of presentation some results are shown normalized and with statistical significance shown for the underlying raw data (Fig. 1C,F; Fig. 3E; Fig. 4C).