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Intracellular sphingosine releases calcium from lysosomes

  1. Doris Höglinger
  2. Per Haberkant
  3. Auxiliadora Aguilera-Romero
  4. Howard Riezman
  5. Forbes D Porter
  6. Frances M Platt
  7. Antony Galione
  8. Carsten Schultz  Is a corresponding author
  1. European Molecular Biology Laboratory, Germany
  2. University of Geneva, Switzerland
  3. National Institutes of Health, United States
  4. University of Oxford, United Kingdom
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Cite this article as: eLife 2015;4:e10616 doi: 10.7554/eLife.10616

Abstract

To elucidate new functions of sphingosine (Sph), we demonstrate that the spontaneous elevation of intracellular Sph levels via caged Sph leads to a significant and transient calcium release from acidic stores that is independent of sphingosine 1-phosphate, extracellular and ER calcium levels. This photo-induced Sph-driven calcium release requires the two-pore channel 1 (TPC1) residing on endosomes and lysosomes. Further, uncaging of Sph leads to the translocation of the autophagy-relevant transcription factor EB (TFEB) to the nucleus specifically after lysosomal calcium release. We confirm that Sph accumulates in late endosomes and lysosomes of cells derived from Niemann-Pick disease type C (NPC) patients and demonstrate a greatly reduced calcium release upon Sph uncaging. We conclude that sphingosine is a positive regulator of calcium release from acidic stores and that understanding the interplay between Sph homeostasis, calcium signaling and autophagy will be crucial in developing new therapies for lipid storage disorders such as NPC.

https://doi.org/10.7554/eLife.10616.001

eLife digest

Sphingosine is a small fat molecule that has been suggested to act as a signal inside cells. Individuals with a rare neurodegenerative disease called Niemann-Pick disease type C accumulate sphingosine and other fat molecules in cell compartments called lysosomes. Intriguingly, this fat accumulation is accompanied by an altered movement of calcium ions in and out of lysosomes.

In healthy cells, an increase in calcium ion levels can trigger a process called autophagy, in which proteins and other cell components are destroyed in a controlled manner. This is thought to be caused by the release of calcium ions from lysosomes, which stimulates a protein called TFEB to move into the nucleus of the cell to activate genes involved in autophagy. Two proteins on the surface of lysosomes called TPC1 and TPC2 are believed to act as channels that can release calcium ions from lysosomes. However, it was not clear how sphingosine could disrupt calcium ion movements in patients with Niemann-Pick disease type C.

Here, Hoeglinger et al. have used a new approach to understand how calcium ions and sphingosine are linked in both healthy and diseased cells. The experiments use a form of sphingosine called “caged sphingosine” that is only activated when it is exposed to a flash of light, which makes it possible to increase the levels of this molecule in cells in a precise way. Hoeglinger et al. found that sphingosine triggered the release of calcium ions from lysosomes. This release required the TPC1 protein and resulted in TFEB moving into the cell nucleus.

Further experiments confirm that sphingosine accumulates in the lysosomes of cells taken from patients with Niemann-Pick disease type C. In these cells, the activation of caged sphingosine resulted in a much smaller release of calcium ions from lysosomes than that observed in healthy cells. Together, Hoeglinger et al.’s findings show that sphingosine acts as a signal to trigger the release of calcium ions from lysosomes, which in turn promotes autophagy. The next challenge is to find out exactly how sphingosine opens the calcium ion channels.

https://doi.org/10.7554/eLife.10616.002

Introduction

Sphingosine (Sph) and other small sphingolipids such as ceramide (Cer) and sphingosine 1-phosphate (S1P) are bioactive lipids, which play crucial roles in central cellular processes such as apoptosis, cell growth, differentiation and senescence (Zheng et al., 2006; Bartke and Hannun, 2009; Hannun and Obeid, 2008). While Cer and S1P have been extensively studied and reviewed (Futerman and Hannun, 2004; Spiegel and Milstien, 2003; Alvarez et al., 2007; Pyne and Pyne, 2010; Arana et al., 2010), only limited information on the signaling mechanisms or cellular targets of Sph is available (Jefferson and Schulman, 1988; Smith et al., 1997; Jarvis et al., 1997; Chang et al., 2001). In cells, Sph can be generated in lysosomes, the plasma membrane, the endoplasmic reticulum (ER) as well as the Golgi complex by hydrolysis of Cer through the action of ceramidases (Bernardo et al., 1995; Hwang et al., 2005; Sun et al., 2008; Xu, 2006; Mao et al., 2001). In a physiological context, Sph is reported to mediate cell growth arrest and apoptosis (Cuvillier, 2002; Suzuki et al., 2004) and is involved in the pathophysiology of the lysosomal storage disorder Niemann-Pick disease type C (NPC). In this progressive neurodegenerative disease, which is caused by mutations in the lysosomal proteins NPC1 or NPC2, Sph accumulates alongside other lipids such as cholesterol, sphingomyelin, and glycosphingolipids (te Vruchte et al., 2004). Specifically, Sph storage is the first detectable biochemical change following inactivation of NPC1 leading to chronically lowered calcium concentration in the acidic compartment, as well as subsequent secondary lipid storage (Lloyd-Evans et al., 2008). On the other hand, the calcium release from acidic stores was recently monitored by a genetically encoded calcium indicator directly fused to endolysosomes which showed reduced calcium release from NPC patient cells (Shen et al., 2012) in agreement with previous studies (Lloyd-Evans et al., 2008) but found that the calcium content in the lysosomes remains unchanged and that the calcium release is blocked by the accumulated lipids. This is in contrast to previous studies showing that the defect in calcium levels in NPC disease is likely due to a store filling defect and that intraluminal calcium levels were reduced using an intra-lysosomal calcium probe (Lloyd-Evans et al., 2008). Further research is necessary to fully elucidate the interplay between calcium uptake and release from the endolysosomes and how this mechanism is deregulated in the NPC disease.

Cells use calcium as a versatile tool for modulating intracellular signaling by increasing the intracellular free calcium concentration [Ca2+]i either globally or locally. Second messengers such as myo-inositol 1,4,5-trisphosphate (IP3) or cyclic ADP-ribose (cADPR) open well-characterized calcium channels in the endoplasmic reticulum (ER) or the muscle sarcomere, the main intracellular calcium store (Berridge et al., 2003; Fliegert et al., 2007). The intraluminal calcium concentrations in the ER varies from 100–800 µM (Burdakov et al., 2005). Since the endosomal/lysosomal system was identified as an important source of intracellular calcium (Haller et al., 1996), the luminal calcium concentrations of acidic vesicles was determined to range between 400–600 µM (Christensen et al., 2002). The mechanisms responsible for Ca2+filling and release from these acidic stores are not yet fully understood. The main players involved in lysosomal calcium release and signaling are transient receptor potential (TRP) channels such as mucolipin 1 (TRPML1) (Pryor et al., 2006) and two two-pore channels (TPC1 and TPC2), which are thought to be Ca2+ channels activated by nicotinic acid adenine dinucleotide phosphate (NAADP), Mg2+or phosphatidylinositol 3,5-bisphosphate (PI(3,5)P2) (Brailoiu et al., 2009; Wang et al., 2012; Ruas et al., 2014; Brailoiu et al., 2010; Rybalchenko et al., 2012; Pitt et al., 2010; Pitt et al., 2014; Jha et al., 2014; Ruas et al., 2015; Jentsch et al., 2015). A very recent study showed that lysosomal calcium signaling regulates autophagy through the actions of the phosphatase calcineurin and the transcription factor EB (TFEB) (Medina et al., 2015) which places the lysosome at the center of this very important signaling hub and underlines the importance of understanding the regulation of lysosomal calcium signaling.

In this work, we investigate the role of Sph in intracellular calcium signaling by employing photoactivatable (“caged”) Sph. Caged lipids are biologically inactive as long as they are covalently attached to certain photolabile groups (“cage groups”), which prevent recognition and metabolism inside cells. The active species is then released inside cells by short irradiation with a flash of light, which induces a cleavage reaction in the cage group (Höglinger et al., 2014). In this manner, it is possible to rapidly elevate the intracellular concentration of the lipid with precise spatiotemporal resolution. It is currently difficult to know the physiological and spatial distribution of sphingosine generated under physiological conditions, particularly inside the lysosome where the majority of sphingosine is generated via the action of acidic ceramidases. This tool therefore provides a very useful system with which to probe sphingosine cell biology in greater detail.

We discovered that uncaging Sph in a variety of cell types leads to an immediate and transient increase in cytosolic calcium, which is not released from the ER but from acidic stores. We further pinpoint the endosomal/lysosomal channel TPC1 to be a main contributor to this response and demonstrate that Sph-induced calcium release leads to the nuclear translocation of TFEB. We also show that the calcium release in NPC patient cells is reduced while the Sph levels in the lysosomal compartments are increased. These findings will be helpful in elucidating the pathogenic cascade of NPC and will generally further our understanding of the function of the acidic compartments in calcium homeostasis.

Results and discussion

Design and synthesis of caged Sph and caged dhSph

We synthesized caged variants of Sph by chemically attaching two different cage groups to the amino functionality of Sph (Figure 1): R1 is a diethylamino-coumarin group, which allows for very fast, sub-second uncaging kinetics (Schade et al., 1999; Hagen et al., 2003) as well as visualization of cages lipids via its fluorescent properties. R2 is a well-established, but non-fluorescent 4,5-dimethoxy-2-nitrobenzyl (NB) group (Il'ichev et al., 2004). In order to exclude potential artifacts caused by irradiation or cleavage of the cage groups, we also synthesized a negative control compound, coumarin-caged dihydrosphingosine (dhSph), a naturally occurring lipid that is structurally very closely related to Sph.

Figure 1 with 2 supplements see all
Structures of coumarin-caged sphingosine (Sph-Cou), nitrobenzyl-caged sphingosine (Sph-NB) and the negative control, coumarin-caged dihydrosphingosine (dhSph-Cou), respectively.
https://doi.org/10.7554/eLife.10616.003

Stability of Cou-caged Sph in cells

Next, we checked whether the caged Sph was taken up by cells and if it was stable in the cellular environment. To this end, HeLa cells were incubated with 2 µM Sph-Cou for different times and subjected to +/- UV uncaging. In +UV cases, the whole dish of labelled cells was exposed to UV light from a mercury arc source for a comparably long time of 2 min to ensure complete uncaging of all cells. Cellular lipids were then extracted and coumarin-containing lipids were visualized by thin layer chromatographic analysis (TLC, Figure 1—figure supplement 1). We showed that caged Sph was taken up by cells and that its photo-induced cleavage was successful inside the cells. This experiment also demonstrated that caged Sph did not participate in lipid metabolism during up to 60 min of incubation, as no additional fluorescent bands could be detected on the TLC plate. The drawback of this analysis was that only lipids equipped with a coumarin cage could be visualized via TLC. In order to gain a more accurate estimate of the uptake, stability, and successful uncaging of Sph-Cou as well as the negative control compound dhSph, we investigated control HeLa cells, Sph-Cou and dhSph-Cou treated HeLa cells by lipidomic analysis. Figure 1—figure supplement 2 shows the relative amounts of Sph, S1P, dhSph and dhS1P detected in control, non-UV treated (caged) and UV-treated (uncaged) conditions for uncaging Sph-Cou as well as dhSph-Cou. The stability of the caged Sph (and caged dhSph) in the cells was confirmed because comparable levels of sphingoid bases were measured in control and caged conditions, indicating that no cleavage of the carbamate-linked cage group took place. Additionally, incubation with Sph-Cou or dhSph-Cou did not appear to perturb the long chain base homeostasis in the cells as all endogenous sphingoid bases showed similar concentrations compared to control conditions. Under the exaggerated illumination conditions used in the TLC and lipidomics experiments, Sph levels were found to increase 3.4 fold relative to control conditions in cells incubated with Sph-Cou, indicative of successful uncaging. This increase was specific for Sph and was not observed for dhSph (Figure 1—figure supplement 2a). The levels of S1P also increased in UV-treated conditions, albeit only at a minute level (0.2% of Sph). We attributed this increase to ongoing metabolism during and after uncaging reaction. It seems that the long time of illumination as well as time for the following steps (collection of the cells and extraction of the lipids) was long enough for phosphorylation to occur on a very small fraction of Sph (Figure 1—figure supplement 2). The negative control dhSph was also uncaged successfully, as indicated by a 13.9 fold increase of dhSph in uncaged cells compared to control (Figure 1—figure supplement 2b). This greater increase is explained by the fact that the caged lipids were added at the same concentrations, even though the endogenous levels of dhSph are 5 to 8 fold lower compared to Sph (according to our mass spectrometric data). Finally, it is important to consider that the TLC and mass spectrometric experiments were performed to show that the uncaging reaction is successful in a cellular environment. They should not be interpreted as quantitative measures of the amount of released Sph. Very long illumination conditions were necessary to uncage enough cells so that these analyses were made possible. All following live-cell experiments were performed on a dual scanner confocal microscope with local uncaging of a subcellular area in the cell with simultaneous fluorescence recording. The uncaging area and uncaging times could therefore be kept to a minimum. In this manner, a greatly reduced release of Sph as well as minimal phosphorylation can be expected.

Uncaging of Sph results in cytosolic calcium transients

In order to observe the immediate effects of photo-induced Sph release on intracellular calcium signaling in living cells, we loaded HeLa cells with the membrane-permeant calcium indicator Fluo-4/AM (Williams et al., 1999), which reacts to increases in cytosolic calcium concentrations with corresponding increases in fluorescence intensity. We performed uncaging experiments using a confocal microscope by recording Fluo-4 fluorescence before and after brief irradiation (3 s–6 s) of a local circular area (~9 µm2) within the cell. Cells reacted with an immediate (1–5 s), but transient increase in cytosolic calcium after uncaging either variant of caged Sph (Sph-Cou or Sph-NB, Video 1 and Figure 2a). We calculated the mean calcium traces for each lipid and found a pronounced transient increase for both caged Sph variants, whereas the negative control compound dhSph-Cou gave no significant change in calcium levels under exactly the same delivery/uncaging conditions (Figure 2b), indicating that the signal was specifically due to sphingosine. To analyze single cell responses, we calculated the maximum amplitude of each calcium trace and visualized their distribution in a histogram (Figure 2c). We defined individual cells with more than 20% increase over the average baseline fluorescence as responding cells. While all cells responded when Sph-Cou/+UV treated, less than 15% (4 out of 29 cells) responded under control conditions (dhSph-Cou/+UV). Moreover, under control conditions, the few responding cells gave rise to markedly reduced amplitudes.

Video 1
Local uncaging of Sph leads to calcium transients.

Time-lapse movie of HeLa cells loaded with the calcium indicator Fluo-4 and treated with Sph-Cou. Cells were uncaged a t = 7 s for 3 s and the Fluo-4 fluorescence was recorded.

https://doi.org/10.7554/eLife.10616.006
Figure 2 with 2 supplements see all
Local uncaging of Sph leads to calcium transients.

(A) Time-lapse confocal microscopy images of HeLa cells loaded with the calcium indicator Fluo-4 and treated with Sph-Cou. Cells were irradiated within the white circle at t = 7 s for 3 s. (B) Mean Fluo-4 fluorescence traces of cells loaded with compounds Sph-Cou (17 cells), Sph-NB (15 cells), and dhSph-Cou (29 cells), respectively. Uncaging was carried out for 3 s for coumarin-caged compounds and for 6 s for nitrobenzyl-caged sphingosine. The duration of uncaging for each lipid is represented by the color-coded bars. Traces represent mean values with the standard error of the mean plotted as error bars. (C) Histogram showing the distribution of the maximum observed amplitude compared to baseline of each analyzed cell. The vertical line represents the threshold set at 20% amplitude increase for responding cells.

https://doi.org/10.7554/eLife.10616.007

We then addressed the possibility whether S1P as a potential metabolite of the liberated Sph and known agonist of intracellular calcium release (Meyer zu Heringdorf et al., 2003), was essential for the calcium response. We performed uncaging experiments in the presence of two inhibitors of the sphingosine kinases: N,N-dimethylsphingosine (DMS) (Edsall et al., 1998) is a Sph analogue, which acts by competing for the binding of Sph. The second inhibitor, SKI-II acts as a mixed inhibitor of Sph and ATP binding (Lim et al., 2012). When performing uncaging experiments in presence of these inhibitors, we found only slight differences of the calcium response compared to basal conditions (Figure 2—figure supplement 1). These differences might be explained by cell-to-cell variability as well as a general perturbation of the cells when using these inhibitors. We conclude that the observed calcium signals were caused by the rapid, photoinduced Sph release rather than S1P formation. Interestingly, external addition of free Sph to the medium of the cells did not evoke changes in calcium levels (Figure 2—figure supplement 2a). The reasons for this are not fully resolved, but we speculate that Sph is more rapidly metabolized after entering cells than it is delivered. Sufficiently high doses of externally applied Sph (> 5 µM) are then cytotoxic. The rapid increase in Sph after uncaging overcomes this problem and hence leads to calcium signaling. We also added free Sph prior to the uncaging experiment and found again no difference in the calcium response. (Figure 2—figure supplement 2b).

Investigating the origin of the calcium

To determine the source of the released calcium, we inhibited the plasma membrane calcium channels in an unspecific way by adding 5 mM Ni2+ as well as ethylene glycol tetraacetic acid (EGTA) to the medium to complex all extracellular calcium (PM block). This inhibition did not block the Sph-induced calcium signal, although the mean amplitudes were reduced and 16 of 41 cells exhibited amplitudes below the 20% response threshold (Figure 3a and b). The initial (5 s) calcium increase was the same as for control conditions, demonstrating that plasma membrane calcium channels were not initially involved in this response. However, blocking PM channels leads to reduced ER calcium and thereby reduces calcium-induced calcium release from the ER. This likely accounts for the reduced amplitudes and response rates in PM block conditions.

Investigating the source of Sph-induced calcium.

(A) HeLa cells were loaded with 2 µM Sph-Cou in wild-type conditions (41 cells) or in conditions inhibiting plasma membrane channels by removing extracellular calcium with ethylene glycol tetraacetic acid (EGTA) and by inhibition with 5 mM Ni2+ (PM block, 41 cells) or by blocking IP3 receptors at the ER using Xestospongin C at 25 µM (13 cells). Traces represent the mean values with the standard error of the mean plotted as error bars. (B) Histograms showing the distribution of the maximum observed amplitude change compared to baseline of all cells, with a threshold at 20%. (C) HeLa cells were loaded with 2 µM Sph-Cou. After addition of either 10 µM ATP to stimulate release of ER Ca2+ or addition of 200 µM glycyl-L-phenylalanine-beta-naphthylamide (GPN), which leads to release of Ca2+ from the acidic stores through osmotic rupture, uncaging was performed at t = 100 s after the primary calcium transient had passed, as indicated by the bar. (D) Histograms showing the distribution of the maximum observed amplitude of the calcium increase after uncaging. Since these observed effects are second calcium transients, we lowered the threshold for responding cells to 10% amplitude change over baseline.

https://doi.org/10.7554/eLife.10616.010

To investigate the contribution of the ER as the major intracellular calcium store, we inhibited the inositol trisphosphate receptor with Xestospongin C (Gafni et al., 1997). The calcium signals still persisted with the same initial calcium increase and only slightly reduced amplitudes, suggesting that IP3 channels were not directly involved (Figure 3a and b).

As a complementary approach, we stimulated the release of ER calcium by adding ATP and monitored the increase of cytosolic calcium via G-protein coupled receptor stimulation (Figure 3c). After the ATP-induced calcium transient had passed, we uncaged Sph and observed a second increase in calcium. Since the ER had been previously emptied, we reasoned that this new signal must originate from a different intracellular calcium store. Also, this result further strengthens our hypothesis that liberated Sph conversion to S1P, which is known to induce calcium release from the ER (Mattie et al., 1994), is not critical for the effect we observe with caged Sph.

To determine whether the acidic compartment calcium stores are involved, we used glycyl-L-phenylalanine-beta-naphthylamide (GPN), an agent which is hydrolyzed by cathepsin C in lysosomes and causes osmotic lysis of acidic vesicles, consequently releasing their content. When GPN was added to the cells, we observed an increase in cytosolic calcium, but uncaging Sph after this transient had passed failed to induce a second calcium release. Taken together, these results point towards the acidic compartment as the primary source of the Sph-induced calcium signal that we observed.

Investigating the mechanism of acidic compartment calcium release by Sph

To further specify the machinery involved in this Sph-induced calcium efflux from the acidic compartments, we employed mouse embryonic fibroblasts (MEFs) derived from mice with a single knock out of either the endosomal/lysosomal calcium channels TPC1 or TPC2 as well as MEFs from a double knock out mouse (Ruas et al., 2014). In WT MEFs, uncaging of Sph led to an immediate cytosolic calcium increase in the same way as was observed in HeLa cells, indicating that this effect is not cell-type specific (Figure 4a). Due to the different morphology and size of the fibroblasts, the amplitudes of the calcium transients were less pronounced in MEFs than in HeLa cells, since the area of uncaging was kept identical. This prompted us to set the threshold for responders to 10% increase over baseline. TPC2 knockout MEFs also gave rise to calcium transients with comparable amplitudes. Employing TPC1 knockout fibroblasts, however, resulted in a large reduction of the mean calcium response. Under these conditions, only 6 of 25 cells exhibited amplitudes greater than the response threshold and even those were markedly decreased compared to WT conditions (Figure 4a and b). This striking effect shows the specificity of Sph-induced calcium release via the TPC1 channel. To confirm this, we also employed MEF from the TPC1/TPC2 double knockout mouse. Under these conditions, only 9 of 31 cells gave calcium signals with greater amplitudes than the threshold, much like in the TPC1-KO conditions. We conclude that there is a connection between an increase in cellular Sph and the release of calcium from the acidic compartments through the action of TPC1, which is known to be localized to early, recycling, and late endosomes as well as lysosomes (Brailoiu et al., 2009).

Figure 4 with 2 supplements see all
Knock-out studies of two-pore channels.

(A) Primary mouse embryonic fibroblasts derived from a wild-type mouse (33 cells) or from two-pore channel 1 (TPC1) knock-out mice (25 cells), two-pore channel 2 (TPC2, 20 cells) knock-out mice or from a double knock-out mouse (DKO, 31 cells) were loaded with 2 µM Sph-Cou and uncaged for 3 s as indicated. Traces represent the mean values with standard errors of the mean plotted as error bars. (B) Histograms showing the distribution of the maximum observed amplitude compared to baseline, with the threshold set to 10% increase over baseline.

https://doi.org/10.7554/eLife.10616.011

We also investigated another important acidic store calcium channel, TRPML1, with respect to Sph-induced calcium efflux. We used a fibroblast cell line derived from a patient suffering from mucolipidosis type IV (MLIV) (Sun, 2000). This disease is characterized by a loss-of-function mutation in the acidic store calcium channel TRPML1 (Bargal et al., 2000). Again, control human fibroblasts gave an immediate calcium response upon release of Sph (Figure 4—figure supplement 1). MLIV patient fibroblasts also responded with an immediate and even higher calcium response suggesting that the observed effect is specific to TPC1 and that a loss of TRPML1 function does not interfere with Sph-induced calcium signaling. In fact, the loss of TRPML1 might even increase the importance of calcium efflux through TPCs as the only remaining way of maintaining acidic compartment calcium concentrations. Since two-pore channels are known to be critically involved in NAADP-dependent calcium signaling (Brailoiu et al., 2009; Tugba Durlu-Kandilci et al., 2010), we also tested the effect of a pharmacological antagonist of NAADP, Ned-19 (Naylor et al., 2009), on Sph-induced calcium release in HeLa cells (Figure 4—figure supplement 2). Ned-19 treated HeLa cells did not exhibit a markedly different calcium response after Sph uncaging, which is in line with previous observations that TPC1 is unresponsive to Ned-19 (Pitt et al., 2014). Several questions still remain on the interface between Sph and TPC1. It is unclear if there is direct interaction or if there are additional, yet unidentified mediators in this response. Also, the interaction with other TPC1 ligands such as NAADP needs to be investigated.

Calcium signaling in Niemann-Pick disease type C

Niemann-Pick disease type C is marked by deregulated calcium homeostasis in the acidic compartments as well as an accumulation of Sph (Lloyd-Evans et al., 2008). We investigated Sph-dependent calcium signaling in this disease by performing uncaging experiments on human fibroblast cell lines derived from a healthy subject (control) and from a patient suffering from NPC1. Control cells reacted with an immediate increase in cytosolic calcium, showing again the broad applicability of these caged compounds across different cell types (Figure 5a and b). In these cells, we also reduced the response threshold to 10% due to the size of the fibroblasts compared to HeLa cells. NPC1 patient cells have been shown to have reduced calcium levels in the acidic stores due to a store filling defect (Lloyd-Evans et al., 2008). Using our setup, we could confirm that calcium transients in NPC1 cells after Sph uncaging exhibited significantly lower amplitudes (Figure 5a and b). This observation corroborated our finding that Sph releases calcium from lysosomes and is consistent with other studies (Lloyd-Evans et al., 2008) showing that in NPC disease these vesicles are filled with less calcium compared to control. It should be mentioned that this is in contrast to other findings in (Shen et al., 2012).

Calcium signaling in NPC disease.

(A) Human fibroblasts derived from healthy subjects (control, 25 cells) or patients with Niemann Pick disease type C (NPC, 31 cells) were loaded with 5 µM Sph-Cou (1) and uncaged for 3s as indicated. Traces represent mean values with the standard error of the mean plotted as error bars. (B) Histograms showing the distribution of the maximum observed amplitude compared to baseline of all cells with the response threshold set to 10% increase over baseline.

https://doi.org/10.7554/eLife.10616.014

Subcellular localization of Sph in NPC disease

Next, we investigated whether Sph is localized to the endosomal/lysosomal compartment in control as well as NPC1 fibroblasts by using a newly reported photoactivatable and clickable version of Sph (pacSph) (Haberkant et al., 2015). This Sph analogue is equipped with two modifications on the hydrophobic tail: a photo-crosslinkable diaziridine moiety and a functionality that allows for click reactions, used for staining Sph in cells with a fluorophore after fixation (Figure 6a). Using pacSph we could, for the first time, visualize the subcellular localization of Sph in individual control and NPC1 fibroblasts. After a short (10 min) pulse of 4 µM pacSph, it was localized in endosomal and lysosomal vesicles in both cell types (Figure 6b) as confirmed by co-localization studies with a LAMP1 antibody (Figure 6—figure supplement 1) and quantification by Pearson’s correlation coefficient (Figure 6c). In NPC fibroblasts, these vesicles were bigger and much more numerous than under control conditions. They also exhibited much brighter overall fluorescence, indicative of higher cellular Sph concentrations. It has already been shown that NPC1 disease fibroblasts exhibit both lower acidic compartment calcium levels as well as an increased Sph storage (Lloyd-Evans et al., 2008).

Figure 6 with 1 supplement see all
Subcellular localization of Sph.

(A) Structure of pacSph. (B) Sph distribution in control and NPC human fibroblasts. Cells were incubated with 4 µM pacSph for 10 min, washed and either immediately photo-crosslinked and fixed (0 min) or incubated for further 10 min in buffer before crosslinking and fixation. Visualization was achieved by clicking Alexa488-azide to the terminal alkyne bond of pacSph. Confocal analysis shows a striking accumulation of Sph in the late endosomes/lysosomes of the NPC fibroblasts. For co-staining with the lysosomal marker LAMP1, see Figure 6—figure supplement 1. (C) Quantification of co-localization analysis by calculating Pearson’s correlation coefficient for >6 cells in each condition.

https://doi.org/10.7554/eLife.10616.015

Our findings strengthen the link between Sph and lysosomal calcium homeostasis. Additionally, we followed the metabolism of pacSph in both cell lines by performing pulse chase experiments, fixing and staining cells at different time points. In control fibroblasts, the endosomal/lysosomal staining almost completely disappeared after a chase of only 10 min, as represented by a drop in the Pearson’s coefficient (Figure 6c). This corresponds to the rapid efflux and transport of Sph to other cellular compartments for further metabolism (Hannun and Obeid, 2008). In contrast, NPC fibroblasts still showed a marked endosomal/lysosomal staining with high fluorescence intensities after 10 min, likely due to failed or diminished efflux from these compartments in the disease. These localization studies further support our hypothesis that elevated endosomal/lysosomal Sph concentrations and reduced calcium levels are closely connected.

Transcription factor EB (TFEB) translocation

Lysosomal calcium release was recently shown to lead to activation of calcineurin and subsequent nuclear translocation of transcription factor EB (TFEB), a master regulator of lysosomal biogenesis and autophagy (Medina et al., 2015). To test if Sph-induced calcium release also reproduces this effect, we performed uncaging experiments in HeLa cells overexpressing TFEB-GFP. Indeed, uncaging Sph and the subsequent calcium release (as monitored by the genetically encoded red calcium sensor, R-GECO (Zhao et al., 2011)) induced rapid translocation of TFEB-GFP to the nucleus (Figure 7a) in 14 out of 19 cells. This was not the case when calcium was released by directly uncaging caged calcium to a similar extent (Figure 7b). Only one in 11 cells showed a slight translocation of TFEB to the nucleus in the 15 min timeframe of the experiment. This further supports the findings that vital cellular signaling pathways are distinctly initiated on the lysosome surface. Medina et al. (Medina et al., 2015) found the mucolipin channel TRPML1 to be involved in activating calcineurin and translocating TFEB, while our findings suggest a contribution of TPC1. This could suggest a certain redundancy in lysosomal calcium release and requires further attention. It can be speculated that reduced calcium levels in NPC patients might fail to properly regulatetranscription through TFEB and that the subsequent differences in lysosomal maintenance and autophagy couldcontribute to the NPC phenotype.

Sph uncaging leads to TFEB translocation to the nucleus.

Time-lapse confocal microscopy images of HeLa cells transfected with TFEB-GFP and R-GECO (a genetically encoded calcium sensor) and loaded with either Sph-Cou or caged calcium (o-nitrophenyl EGTA/AM). Uncaging was performed in a small area within the cell for 3 s for Sph-Cou (lower cell) and 2 s for caged calcium.

https://doi.org/10.7554/eLife.10616.017

Conclusion and outlook

In summary, we have developed a method to increase the concentration of Sph in living cells using light in a spatially and temporally precise way. Thus, we interfered with sphingolipid metabolism minimally and were able to investigate resulting effects of elevated sphingosine levels. Dihydrosphingosine, although structurally very similar, served as a good negative control compound and underlined the high structural specificity of the interacting partners of Sph. We applied these tools to a variety of cell types, ranging from cultured HeLa cells to primary mouse embryonic fibroblasts and human patient fibroblasts and showed that in all those cells, an acute increase in Sph was always followed by an immediate release of calcium from GPN-sensitive acidic stores. We also showed that plasma membrane and ER calcium channels were not initially involved in this calcium response. Furthermore, we could pinpoint TPC1, an endosomal/lysosomal channel of the two-pore channel family to be essential in this newly found signaling pathway. When employing embryonic TPC1 knockout fibroblasts, no calcium increase upon uncaging of Sph was observed. TPC2 knockout fibroblasts, however, reacted similarly to WT fibroblasts, confirming the specific requirement of the TPC1 isoform for Sph induced calcium signaling. The involvement of the other main acidic compartment calcium channel TRPML1 was also tested. Loss-of-function in TRPML1 led to a slight increase in Sph-induced calcium release, which might be explained by some redundancy between TRPML1 and TPCs in the regulation of lysosomal calcium concentrations.

Acidic compartment calcium signaling has recently emerged as an important part of intracellular events such as vesicle fusion and secretion (Patel and Docampo, 2010) as well as induction of autophagy and lysosomal biogenesis (Medina et al., 2015). Our knowledge of the machinery involved and its regulation, however, remains incomplete. The two-pore channels (TPCs) are the subject of current controversies regarding ion selectivity and dependencies on activating ligands. While many reports agree that TPC channels are NAADP-regulated Ca2+ channels (Brailoiu et al., 2009; Brailoiu et al., 2010; Pitt et al., 2010; Calcraft et al., 2009), others indicate a broader ion specificity of TPCs (Wang et al., 2012; Rybalchenko et al., 2012; Peiter et al., 2005) or alternative ligands such as PI(3,5)P2 (Wang et al., 2012; Cang et al., 2013). Another study hints towards a convergent regulation of TPC2 by Mg2+, NAADP, PI(3,5)P2 and two protein kinases (JNK and p38) (Jha et al., 2014). A very recent study confirmed that TPCs conduct both Ca2+ and Na+ ions and that they are activated by NAADP as well as PI(3,5)P2 (Ruas et al., 2015; Jentsch et al., 2015). The biological relevance of two-pore channels was recently highlighted by a report showing that TPCs are crucial to successful infection of host cells by the Ebola virus and that disruption of TPC function leads to halted virus trafficking (Sakurai et al., 2015). Interestingly, the progression of Ebola infection is also dependent on the NPC1 protein, which was shown to be necessary for viral escape from the lysosomal compartment (Herbert, et al., 2015Carette, et al., 2011). These recent findings will certainly encourage further efforts to understand the link between lysosomal calcium signaling and the defects in Niemann-Pick disease. In our study, we could confirm that NPC patient fibroblasts responded with lower amplitudes of the calcium transients after Sph uncaging, which is indicative of a reduced pool of calcium in the acidic compartments of these cells, in accordance with other studies (Lloyd-Evans et al., 2008; Shen et al., 2012).

Figure 8 summarizes our results and illustrates that we pinpointed TPC1 as the mediator of a Sph induced rise in cytosolic calcium. We also correlated a reduced calcium release in fibroblasts from NPC patients with a significant storage of Sph in the late endosomes/lysosomes of these patient cells by using a novel bifunctional Sph probe. Short pulse-chase experiments also indicated lack of efflux from lysosomes in NPC fibroblasts compared to cells from healthy controls.

Schematic summary of findings.

Sph causes calcium release from acidic stores through the action of TPC1. In Niemann-Pick disease type C, lower endolysosomal calcium levels occur together with increased Sph concentration. Stimulating lysosomal calcium release by uncaging Sph in NPC patient cells results in lower calcium amplitudes compared to cells from control subjects.

https://doi.org/10.7554/eLife.10616.018

The question whether the NPC1 protein is directly or indirectly involved in the efflux of lipids, specifically Sph, from the acidic compartments has remained a controversial topic. NPC1 has been shown to be involved in the transport of amines (Kaufmann and Krise, 2008), but conclusive data on the transport of Sph and other potential substrates are absent as there is no direct functional assay for NPC1. A study using radiolabelled sphingolipids to follow metabolism and transport in NPC disease concluded that Sph efflux from endolysosomes is not restricted in NPC (Blom et al., 2012). However, these data were obtained using long, 1–4 h chase times during which the precursor lipid was likely metabolized to a high degree and incorporated into other lipids, leaving only a small fraction of Sph for quantitative analysis. This and other obstacles to the study of lipids affirm the need for new tools, which circumvent the lipid metabolic machinery and give a faithful representation of the subcellular lipid localization. With our tools and shorter chase times in the range of 10–20 min, we observed reduced acidic compartment calcium release as well as increased Sph storage and a failure in Sph efflux in NPC disease cells.

Disrupted autophagy is another hallmark of NPC shared by many other neurodegenerative disorders as neurons are particularly vulnerable to defects along the autophagic pathway (Nixon, 2013). TFEB was shown to up-regulate transcription of genes which support autophagosome formation and lysosomal biogenesis (Settembre et al., 2011) and could therefore be a considered a novel target to promote lysosomal clearance in lipid storage and neurodegenerative disorders. TFEB overexpression has already been shown to be beneficial in attenuating the storage phenotype in several storage disorders such as Batten disease (Medina et al., 2011) or Pompe disease (Spampanato et al., 2013). Similar overexpression strategies showed promising results even in mouse models of more common neurodegenerative disorders, such as Parkinson disease (Decressac et al., 2013), Alzheimer disease (Polito et al., 2014) or Huntington disease (Tsunemi et al., 2012). The regulatory functions of Sph and TFEB are therefore key targets for future investigations of neurodegenerative disorders.

Taken together, we describe intracellular Sph as a new small molecule effector of lysosomal calcium homeostasis, the lysosome as triggerable calcium store via the TPC1 channel, and specific transcription factor translocation as a consequence of lysosomal calcium release. Our results further highlight the importance of elucidating the interplay between lysosomal calcium signaling and Niemann-Pick type C disease. This interplay might also be important for the progression of diseases caused by Ebola or Marburg virus infection. Sph should be considered as a new and important player in this field and understanding the regulation of its signaling will likely further our understanding of these named diseases.

Materials and methods

Materials

Common laboratory chemicals were purchased from commercial sources (Acros, Fluka, Merck, Sigma-Aldrich or VWR) at highest available grade and were used without further purification. D-erythro sphingosine and D-erythro dihydrosphingosine, the sphingosine kinase inhibitor SKI-II, Xestospongin C and GPN were obtained from Biomol (Hamburg, DE). 7-Diethylamino-4-hydroxymethylcoumarin was a kind gift from Rainer Müller (EMBL Heidelberg, DE). Deuterated solvents for NMR analysis were purchased from Deutero (Karlsruhe, DE). Sphingosine kinase inhibitor N-N-dimethylsphingosine was obtained from Sigma-Aldrich. NAADP-antagonist Ned-19 was purchased from Tocris Biosciences (Bristol, UK). The fluorescent calcium indicator Fluo-4-AM and Alexa488-azide were obtained from Life Technologies (Thermo Fisher Scientific, Waltham, USA). pEGFP-N1-TFEB was a gift from Shawn Ferguson (Addgene plasmid # 38119) (Roczniak-Ferguson et al., 2012).

General synthetic procedures

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All chemical reactions were carried out using dry solvents under inert atmosphere. Thin layer chromatography (TLC) was performed on plates of silica gel (Merck, 60 F254) and visualized using UV light (254 nm or 366 nm) or a solution of phosphomolybdic acid in EtOH (10% w/v). HPLC grade solvents for chromatography were obtained from VMR. Preparative column chromatography was carried out using Merck silica gel 60 (grain size 0.063–0.200 nm) under a pressure of <1.5 bar. 1H-NMR spectroscopic measurements were conducted on a 400 MHz Bruker UltraShieldTM spectrometer at 25°C. 13C-NMR measurements were performed on a 500 MHz Bruker UltraShieldTM spectrometer at 25°C and were broadband hydrogen decoupled. Chemical shifts are given in ppm referenced to the residual solvent peak. J values are given in Hz and splitting patterns are designated using s for singlet, d for doublet, t for triplet, q for quartet, m for multiplet and b for broad signal. High-resolution mass spectra were recorded at the Organic Chemistry Institute of the University of Heidelberg

Synthesis of 7-(diethylamino)-coumarin-4-yl)-methyl-chloroformate

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A solution of 7-diethylamino-4-hydroxymethylcoumarin (Schönleber et al., 2002) (48 mg, 194 µmol) in 2 mL dry THF was cooled to 0°C. DIPEA (0.1 µL, 575 µmol) and phosgene (300 µL, 610 µmol) were added dropwise and stirred in the dark for 2 h at 0°C. The reaction mixture was extracted with EtOAc/H2O (1:1, 75 mL), the layers were separated, the organic layer was washed with brine and dried using Na2SO4. The solvent was removed under reduced pressure and the product was dried further under high vacuum conditions. 7-(Diethylamino)-coumarin-4-yl]-methyl chloroformate was used without further purification.

Synthesis of N-(7-(diethylamino)-coumarin -4-yl)-methyl)- ((2R,3S,E)-1,3-dihydroxyoctadec-4-en-2-yl)carbamate (Sph-Cou)

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To a solution of D-erythro-sphingosine (30 mg, 100 µmol) in 2 mL dry THF, TEA (70 µL, 500 µmol) and a solution of 7-(diethylamino)-coumarin-4-yl)-methyl chloroformate (46 mg, 148 µmol) in 1 mL dry THF were added. The mixture was stirred at RT for 1 h in the dark. EtOAc (50 mL) was added to stop the reaction and the mixture was washed twice with citric acid (5% w/v, 25 mL) and twice with saturated NaHCO3. The organic layer was dried with Na2SO4 and the solvent was removed under reduced pressure. The residue was purified by repeated flash chromatography (first column: eluent: DCM/MeOH13:1, second column: eluent: cyclohexane/EtOAc 1:5 (+1% TEA)). The compound 1 was obtained as yellow oil (24 mg, 42 µmol, 42% over two steps) 1H NMR (400 MHz, CDCl3) δ = 7.29 (d, J=8.9, 1H), 6.58 (d, J=8.7, 1H), 6.50 (s, 1H), 6.14 (s, 1H), 5.96 (d, J=8.3, 1H), 5.80 (dd, J=14.6, 7.2, 1H), 5.55 (dd, J=15.2, 6.1, 1H), 5.22 (s, 2H), 4.39 (s, 1H), 4.05–3.96 (m, 1H), 3.76 (d, J=11.6, 1H), 3.68 (q, J=3.8, 1H), 3.40 (q, J=7.0, 3H), 3.06 (s, 1H), 2.05 (dd, J=13.5, 6.6, 2H), 1.41–1.10 (m, 28H), 0.87 (t, J=6.6, 3H). 13C NMR (126 MHz, CDCl3) δ = 162.15, 156.18, 155.60, 150.40, 134.45, 128.70, 126.78, 124.43, 108.98, 106.16, 98.13, 74.75, 74.17, 66.85, 62.21, 61.89, 55.73, 44.96, 32.29, 31.93, 29.70, 29.67, 29.64, 29.51, 29.37, 29.22, 29.10, 22.70, 14.12, 12.40. HRMS for C33H53N2O6+ calculated: 573.39036; found: 573.39027.

Synthesis of N-(2,4-dimethoxy-6-nitrophenyl)-((2R,3S,E)-1,3-dihydroxyoctadec-4-en-2-yl)carbamate (Sph-NB)

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A solution of D-erythro-sphingosine (30 mg, 100 µmol) in 2 mL dry THF and TEA (70 µL, 500 µmol) was stirred and 4,5-Dimethoxy-2-nitrobenzyl-chloroformate (41 mg, 150 µmol) in 2 mL dry THF was added dropwise. The mixture was stirred at RT for 1h in the dark. EtOAc (50 mL) was added and the mixture was washed twice with citric acid (5% w/v, 25 mL) and twice with saturated NaHCO3. The organic layer was dried with Na2SO4 and the solvent was removed under reduced pressure. The residue was purified by flash chromatography (eluent: cyclohexane/EtOAc 1:5 (+1% TEA)). Compound 2 was obtained as colorless oil (51.4 mg, 95 µmol, 95% ) 1H NMR (400 MHz, CDCl3) δ = 7.70 (s, 1H), 7.02 (s, 1H), 5.84–5.75 (m, 2H), 5.55 (d, J=6.0, 1H), 5.51 (s, 2H), 4.38 (s, 1H), 4.01 (s, 1H), 3.98 (s, 3H), 3.95 (s, 3H), 3.73 (d, J=11.5, 1H), 3.67 (s, 1H), 2.39 (s, 2H), 2.12–1.96 (m, 2H), 1.42–1.13 (m, 22H), 0.87 (t, J=6.7, 3H) 13C NMR (101 MHz, CDCl3) δ = 155.99, 134.47, 128.58, 110.07, 108.15, 74.81, 63.78, 62.24, 56.52, 56.50, 56.40, 55.57, 32.28, 31.94, 31.92, 29.71, 29.68, 29.62, 29.48, 29.36, 29.22, 29.08, 22.71, 22.69, 14.13. HRMS for C28H47N2O8Na+ calculated: 561.31519, found: 561.31557.

Synthesis of N-(7-(diethylamino)-coumarin-4-yl)-methyl)-((2R,3S)-1,3-dihydroxyoctadecan-2-yl)carbamate (dhSph-Cou)

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To a solution of D-erythro-dihydrosphingosine (10 mg, 33 µmol) in 1 mL dry THF, DIPEA (23 µL, 230 µmol) and a solution of [7-(diethylamino)-coumarin-4-yl)-methyl chloroformate (15 mg, 50 µmol) in 0,5 mL dry THF were added. The mixture was stirred at RT for 1.5 h in the dark. EtOAc (20 mL) was added to stop the reaction and the mixture was washed twice with citric acid (5% w/v, 10 mL) and twice with saturated NaHCO3. The organic layer was dried with Na2SO4 and the solvent was removed under reduced pressure. The residue was purified by repeated flash chromatography (first column: eluent: DCM/MeOH 13:1, second column: eluent: cyclohexane/EtOAc 1:5). Compound 3 was obtained as yellow oil (14.5 mg, 25 µmol, 76% over two steps) 1H NMR (400 MHz, CDCl3) δ = 7.27 (d, J=8.9 Hz, 1H), 6.56 (dd, J=9.0, 2.3, 1H), 6.49 (d, J=2.3, 1H), 6.14 (s, 1H), 6.08 (d, J=8.4, 1H), 5.22 (s, 2H), 4.07 (dd, J=11.4, 2.3, 1H), 3.82 (d, J=11.7, 2H), 3.66 (s, solvent THF), 3.62 (dd, J=8.1, 3.3, 1H), 3.40 (q, J=7.0, 4H),2,30 (t, J=7.6, solvent), 1.67–1.45 (m, 4H), 1.36–1.13 (m, 30H), 0.87 (t, J=6.7, 3H) 13C NMR (126 MHz, CDCl3) δ = 162.34, 156.17, 155.52, 150.60, 130.02, 129.77, 124.41, 108.83, 106.12, 105.90, 97.90, 74.36, 62.24, 61.83, 55.13, 51.44, 44.82, 34.55, 31.94, 29.71, 29.67, 29.61, 29.59, 29.37, 27.23, 25.99, 24.96, 22.70, 14.12, 12.43. HRMS for C33H55N2O6+ calculated: 575.40601; found: 575.40626.

Cell culture and transfection

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HeLa cells (LGC-ATCC, No. CCL-2, authenticated by STR profiling) were grown in DMEM (1 g/L Glutamate, Gibco) with 10% FBS (Gibco) and 1% Primocin (Invivogen), regularly tested for mycoplasma (using Lookout Mycoplasma PCR detection kit (Sigma, MP0035)) and only used when negative. Primary MEF from Tpcn1-/-, Tpcn2-/- and double knock out mice were characterized previously (Ruas et al., 2015) and only used for 3-4 passages. They were not screened for mycoplasma. Human patient fibroblasts from MLIV patients were obtained from the Coriell Institute (MLIV patient cells: GM02525, authenticated by Nucleoside Phosphorylase Isoenzyme Electrophoresis, control cells: GM05399, authenticated by Chromosome Analysis) and checked for mycoplasma using the Lookout Mycoplasma PCR detection kit (Sigma, MP0035). In case of contamination, Mycoplasma Removal Agent (AbD Serotec, BUF035) was used. NPC1 cells were provided by Dr. Forbes D. Porter (NIHCD, USA). They are primary cultures from an NPC1 patient enrolled in the NIH program. The cells were not screened for mycoplasma, and used for 3–4 passages only. The culture medium for all mentioned fibroblasts consisted of DMEM (4.5 g/L glutatmate, Gibco) with 10% FBS (Gibco), 1% penicillin-streptomycin and 1% L-glutamine (Gibco). Cells were seeded in 8-well Lab-TekTM microscope dishes 48–72 h prior to the experiment. If needed, cells were transfected in OPTIMEM (Gibco) with maximal 200 ng DNA per well and Fugene HD (Promega) according to the manufacturer’s instructions. For the duration of the experiment, the medium of the cells was changed to imaging buffer (20 mM HEPES, 115 mM NaCl, 1.2 mM MgCl2, 1.2 mM K2HPO4, 1.8 mM CaCl2 and 10 mM glucose).

Cell loading with caged compounds

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Each Labtek-well was loaded with 100 µl of a 5 µM Fluo-4 AM (Molecular Probes®) solution in imaging buffer for 30 min at 37°C. The caged lipids were added to a final concentration of 2 µM 15 min prior to the experiment. Immediately prior to the experiment, the cells were washed and kept in imaging buffer.

TLC analyses

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Cells were grown to ~85% confluency in 6 cm dishes and pulsed with indicated concentrations of Sph-Cou in imaging buffer for the indicated times. Under +UV conditions, the cells were irradiated by a UV mercury arc source (Newport) equipped with a 400 nm highpass filter at a distance of 35 cm for 2 min. Cells were washed with PBS and scraped off in 300 µl PBS. The lipids were extracted according to a protocol published by Thiele et al. (Thiele et al., 2012). In brief, 600 µl methanol and 150 µl chloroform was added to the cells, vortexed and centrifuged at 14 000 rpm for 3 min. The supernatant was mixed with 300 µl chloroform and 600 µl acetic acid (0.1% in water). The aqueous phase was discarded and the organic phase was dried on a speedvac at 30°C for 15 min. The dried lipids were redissolved in 30 µl chloroform and applied on a 10 x 10 cm HPTLC Silica 60 glass plate (VWR) using the automatic Camaq system. TLC plates were developed using first CHCl3/MeOH/H2O/AcOH 65:25:4:1 for 6 cm and then cyclohexane/ethylacetate 1:1 for 9 cm. Fluorescently labeled lipids were visualized using a geldoc system.

Lipidomic analyses

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HeLa cells were grown in 6 cm dishes (Nunc) to 85–95% confluency and labelled with 2 µM Sph-Cou or 2 µM dhSph-Cou in imaging buffer or only in imaging buffer for 15 min. After washing with PBS, cells were transferred onto an ice block and UV-irradiated with > 400 nm light on a 450–1,000 W high-pressure mercury lamp (Newport, #66924, series #1166) for 2 min and then immediately pelleted (by centrifugation with 3000 rpm for 1 min) and snap-frozen in liquid nitrogen. Long chain base extraction and analysis was performed as follows. Cell pellets were resuspended in extraction solvent EtOH/H2O/diethylether/pyridine (15:15:5:1) and ammonium hydroxide (2.1.10−3 N). Cells were extracted by vortexing at 4°C for 10 min and incubation on ice for 20 min. Cell debris was pelleted by centrifugation at 20000 x g for 2 min at 4°C. The extraction was repeated one more time without the incubation on ice. Supernatants were combined and dried under vacuum in a Centrivap (Labconco, Kansas City, USA). The extract was resuspended in borate buffer (200 mM boric acid pH8.8, 10 mM tris(2-carboxyethyl)-phosphine, 10 mM ascorbic acid and 33.7 µM 15N13C-valine) and derivatized by reaction for 15 min at 55°C with 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate (AQC, 2.85 mg/ml in acetonitrile). The AQC reagent was synthesized according to Cohen et al (Cohen and Michaud, 1993). Samples were analyzed after overnight incubation at 24°C using a reverse-phase C18 column (HPLC EC 100/2 Nucleoshell RP-18 2.7 µm) on an Accela system high-performance liquid chromatography (ThermoFisher Scientific, Waltham, MA), coupled to a TSQ Vantage (ThermoFisher Scientific, Waltham, USA). MRM-MS was used to identify and quantify lipid species. The relative amounts of long chain bases were normalized to a standard for the derivatization process (15N13C-valine) and internal standards added before extraction (C17-sphingosine, C17-sphinganine, C17-sphingosine-1-phosphate, C17-sphinganine-1-phosphate).

Confocal time-lapse live cell microscopy and photoactivation

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Fluorescence change of the calcium indicators were captured on a dual scanner confocal laser scanning microscope (Olympus Fluoview 1200) with a 63x oil objective using excitation at 488 nm and emission settings between 500–550 nm at an interval of 1 s per frame. For monitoring the red calcium dye R-GECO, the excitation was at 559 nm and emission between 580–650 nm. A baseline of 10 s was captured before photoactivation. Intracellular photoactivation was performed using the tornado function of the Olympus software with a circular region of interest (10 pixel units diameter, 8.9 µm2). For uncaging at 405 nm, we used 50% laser intensity for 3 s at 2 µs per pixel. Nitroveratroyl-uncaging at 375 nm was carried out using 100% laser intensity for 5 s at 2 µs per pixel.

Data analysis

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The acquired time lapse series were analyzed with the Fiji software (W. Rasband, NIH, USA) using the FluoQ macro (Stein et al., 2013) set to the following parameters:

Background subtraction method: Mean of an interactively selected ROI Noise reduction / smoothing method: None Threshold method: Interactively with ImageJ’s built-in threshold window ROI segmentation: Semi-automatically with binary mask modification Calculate amplitude changes: Using maximum observed amplitude change

The maximum amplitude values were calculated by subjecting the raw traces to a central moving average. The maximal amplitude xresponsemax of these smoothed traces was used to calculate the amplitude change in percent %xmax according to following formula:

%xmax=xresponsemaxx¯baseline|x¯baseline|*100

The resulting intensity series/amplitude values represent mean values of whole cells and were further analyzed using R software (Development Core Team, R, 2014).

Sph visualization and co-localization

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Cells were seeded onto 11mm coverslips placed in wells of a 24-well plate and labeled with 4 µM pacSph in imaging buffer for 10 min. Cells were washed, overlaid with 1 mL imaging buffer and UV-irradiated on ice for 2.5 min using a 450–1,000 W high-pressure mercury lamp (Newport, #66924, series #1166) equipped with a glass filter to remove wavelengths below 345 nm (Newport, #20CGA-345), operated at 1,000 W. Cells were immediately fixed with MeOH at -20°C for 20 min. Not crosslinked lipids were extracted by washing 3x with 1 mL of CHCl3/MeOH/AcOH 10:55:0.75 (v/v) at RT. Cells were then incubated with 50 µl of click mixture (1 mM ascorbic acid, 100 μM TBTA, 1 mM CuSO4 and 2 μM Alexa 488 azide in PBS) for 1 h at RT in the dark. Cells were then washed with PBS and incubated with 50 µl of primary antibody (rabbit α-LAMP1, Cell Signaling, 1:100 in PBS supplemented with 4% BSA and 0.02% Triton) overnight at 4°C. Coverslips were again washed in PBS and incubated with secondary antibody (α-rabbit conjugated to AlexaFluor555, Cell Signaling, 1:800) for 1 h, washed and mounted in DAPI-containing mounting medium (Vectashield, Vector Laboratories, Inc. Burlingame, CA 94010, #H-1200). Microscopy images were captured at RT using a confocal laser scanning microscope (Zeiss LSM780) with a 63x oil objective. Settings were as follows: DAPI-channel (405 nm excitation (ex), 409–475 nm emmission (em); green channel: 488 nm ex, 489–550 nm em; red channel: 561 nm ex, 569–655 nm em). Images were further processed using imageJ (http://rsb.info.nih.gov/ij/).

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Decision letter

  1. Suzanne R Pfeffer
    Reviewing Editor; Stanford University School of Medicine, United States

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled "Sphingosine releases calcium from lysosomes via the two-pore channel 1" for peer review at eLife. Your submission has been favorably evaluated by Vivek Malhotra (Senior Editor), a Reviewing Editor, and two reviewers.

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

The reviewers find the work potentially very exciting, but three issues need to be addressed. If you can address the first point convincingly, the reviewers felt that the story would be very important.

1) The main potential novelty is in defining a physiologic role for sph in regulation of calcium release from endolysosomes with consequent signaling and cell regulatory functions. This is highly suggested by this study, but the study relies primarily on a 'pharmacologic' albeit novel and powerful approach using the delivery of caged sph. As such, the role of endogenous sph is not established. In this regard, the 3.5 fold increase in intracellular sph after uncaging may represent a supraphysiologic response (one reviewer was not aware of physiologic conditions where intracellular sph increases by that much). The study would be much stronger if there were ANY additional results to implicate endogenous sph in this mechanism and this pathway.

Moreover, the authors use various approaches but they don't connect them or control for them appropriately. For example, they show that exogenous sph does not work but they don't measure cellular sph after the exogenous sph which is important. If this is at the same level as what they see with caged-sph, then they have a problem.

Similarly, if they are unable to uncage dh-sph (they don't measure free dh-sph in response to uncaging the way they measure sph after uncaging sph), then they cannot use this as a basis to claim high specificity (which is one of the most important points in support of the specificity and therefore significance of the sph effects on calcium.) The finding that dh-sph is not functional is a strong indication of specificity of action; therefore to clinch this point, the authors need to measure dh-sph after its uncaging.

2) There is no evidence that Sph directly interacts with the TPC1 channel-no targets of Sph were identified. They could use click chemistry and show that uncaged Sph is bound to something specific to a calcium transporter or something specific to the lysosome-but this is likely beyond the scope of the present study. Because exogenous Sph itself does not induce Ca2+ release, the Discussion, Conclusions, and the title should be toned down.

3) The effects of TFEB may or may not be related to the specific action of lysosomal sph. They use pac-sph to 'establish' the sub cellular localization of sph, but this compound would resemble more exogenous sph in its pharmacology and therefore should not act on TFEB or calcium. Does the pac-Sph translocate TFEB? If not, then its cellular localization is irrelevant. If it does, then how come 'plain' sph does not act on calcium? Does exogenous sph induce TFEB?

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for choosing to send your work entitled "Sphingosine releases calcium from endolysosomes via the two-pore channel 1" for consideration at eLife. Your full submission has been evaluated by Vivek Malhotra (Senior Editor) and three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the decision was reached after discussions between the reviewers. Based on our discussions and the comments below, we regret to inform you that the reviewers noted a number of issues that would need to be addressed in full before the manuscript could be considered further for presentation in eLife.

Sphingosine (SPH) has been long demonstrated to act as a bioactive lipid to mediate various biological responses, such as cell proliferation inhibition and programmed cell death, but the mechanisms for it to mediate biological responses remain largely unclear. The present manuscript makes an interesting and potentially important observation that SPH releases calcium from endolysosomes via the two-pore channel 1 (TPC1). The authors developed photoactivatable SPH, caged sph, which was taken up by cells and was converted to free SPH upon photoactivation. This allowed the authors to study how free SPH transiently activates the release of calcium from different cellular compartments. The authors provide evidence that free SPH on its own, not through its metabolite sphingosine-1-phosphate, can induce the release of calcium to the cytosol from intracellular calcium stores. More importantly, the authors demonstrate that the calcium release is specific to SPH but not to its naturally occurring analogue dihydrosphingosine that differs from SPH by a single double bond. These findings provide novel insights into the mechanism by which SPH mediates biological responses.

The results are incomplete with respect to biological effects resulting from calcium mobilization by SPH. One would predict that this magnitude of acute calcium release would mimic many of the effects of stimuli that induce downstream effects through calcium mobilization. Providing evidence with respect to some biological effects would significantly improve the manuscript.

It is worrying that sphingosine proper does not cause a Ca2+ signal (Figure 1—figure supplement 2). This really needs clarification. Additionally, the authors state in the Results and Discussion that Ca2+ waves initiate at the point of uncaging and propagate throughout the cell but this is not quantified and this is not clear from the supplemental movie.

The pharmacology of the sphingosine-evoked Ca2+ signals is not so well defined. In Figure 3A, it is unclear exactly how Ca2+ influx was blocked (Ni v EGTA v both). The effect of XeC is modest but activation of TPCs is thought to result in secondary Ca2+ release from the ER. In Figure 3B, GPN appears to induce a Ca2+ signal that is comparable in amplitude to ATP (which the authors ascribe to ER release) which is surprising. Perhaps the GPN response causes ER Ca2+ release. Uncaging after addition of ATP/GPN is complicated by the elevated baseline. Would exogenous ATP fully deplete ER stores of calcium? What about using thapsigargin? There is little discussion of why Ned-19 is ineffective. Please show effects of Ned-19 on NAADP action or some positive control (Figure 4—figure supplement 2).

It is clear in Figure 4 that sphingosine responses are reduced upon TPC1 knockdown but it is not so clear whether this is a direct effect of sphingosine on TPC1.

The authors claim that sphingosine-mediated Ca2+ signals are reduced in NPC (Figure 5). The quantification (Figure 5B) does not reflect this (the proportion of responding cells is only slightly reduced) and it is unclear if there is an effect on the amplitude. The apparent reduction in signal in Figure 5A in NPC is not dissimilar to the effect of Sph kinase block in Figure 2 (FtY720), XeC/PM block in Figure 3 and TPC2 silencing in Figure 4 yet a role for the intended targets in sphingosine action were discounted in Figures 2-5.

The subcellular localisation/pulse chase data in Figure 6 needs quantitation and the authors' claim that sphingosine localizes to the endolysosomal system (Figure 6—figure supplement 1) is not so convincing.

The authors imply that reduced sphingosine-mediated Ca2+ signal in NPC is due to activation of TPC1 by elevated sphingosine levels causing a reduction of Ca2+ levels within the acid Ca2+ store. A previous study (led by co-authors) however concluded that reduced Ca2+ content in the acid Ca2+ store was due to reduced Ca2+ uptake (Lloyd-Evans, 2008). In the Introduction, the authors refer to a study where Ca2+ release from the acidic Ca2+ store was reduced in NPC but in that study (Shen; 2012), lysosomal Ca2+ content was unchanged and rather lipid accumulation was proposed to block Ca2+ release through TRPML. This is a little confusing and potentially misleading. Cause-effect relationships are blurred.

Other Points:

1) The approach to block the conversion of SPH into S1P with DMS and FTY720 is somewhat problematic because DMS is a non-specific SPHK inhibitor and FTY720 is a SPH analogue, and both may mimic SPH in inducing calcium release on their own. Please note that FTY FTY720 is not an inhibitor of SK1. Moreover, the concentrations of DMS and FTY may not be sufficient to displace sph from sphingosine kinase. The authors should use genetic approaches to block the conversion of SPH to S1P in cells

2) Results in Figure 1—figure supplement 1 do not establish metabolic inertness of the caged compound. There is time dependent uptake, the signal is too faint to detect a sizable metabolite in the order of 10-20% or so, and this approach does not evaluate for spontaneous uncaging (the product would not be fluorescent).

3) The authors need to measure levels of sphingosine and metabolites at the earliest time point if they are able to.

4) Results and Discussion; full name of EGTA is not ethylene tetra-acetic acid

5) Figure 3A. It looks like PM block has a significant effect on the sph response? This appears to be of the same magnitude as the loss in NPC. Also, this figure, in contrast to the legend, does not show the effects of EGTA or Ni.

6) SPH can also be quickly converted to ceramides through ceramide synthases or reverse action of ceramidases. The authors should investigate if blocking the conversion of SPH to ceramides can affect the calcium release by uncaged SPH. Also, studies in Farber's disease would be important.

7) The authors should investigate if increasing the generation of endogenous SPH by activating a ceramidase can induce the release of calcium from a calcium store(s).

8) The authors could measure the content of calcium in the endolysosomal system to determine if uncaging SPH indeed empties this calcium store. This will give another line of evidence that SPH releases calcium from this particular cellular compartment.

9) The max. amplitude change comparisons are often hard to compare – would additional kermel density estimations help, or report means where the data are relatively Gaussian?

10) The data on localization of Sph in NPC are hard to interpret (Figure 6). Only one cell is shown, and it appears to include significant ER staining. LAMP1 staining also seems unusual in this one cell compared to other published reports for NPC1 mutant cells. Please add additional examples and consider including an ER marker to strengthen your conclusions.

11) A recent paper in Science showed a connection between NPC1 and TPC proteins in that both seem to be needed for Ebola virus entry. Please cite and speculate how your work relates to that story.

https://doi.org/10.7554/eLife.10616.021

Author response

1) The main potential novelty is in defining a physiologic role for sph in regulation of calcium release from endolysosomes with consequent signaling and cell regulatory functions.

We agree that one of the most exciting findings of our work is a potential role of Sph. Because the subject is calcium release, the reviewers (and also the readers) expect to see an effect similar to IP3-induced calcium release from the ER. We believe that the effect of Sph on intralysosomal calcium levels is more subtle yet not less important. As our results on monitoring TFEB translocation clearly demonstrate, simple elevation of intracellular calcium by uncaging calcium does not induce translocation. The major effect of Sph on calcium homeostasis therefore is not likely to be the cytosolic calcium increase, but rather the emptying of the lysosomal compartment. Consequently, chronically elevated levels of Sph in NPC cells are responsible for lowered lysosomal calcium levels due to their effect on TPC1 channels. The physiological role is therefore only observable in a pathologic situation. We do NOT demonstrate a classical second messenger system where physiological Sph levels are elevated rapidly and a rapid response results.

This is highly suggested by this study, but the study relies primarily on a 'pharmacologic' albeit novel and powerful approach using the delivery of caged sph.

Based on what we said above, we now indeed create a potentially artificial Sph transient. However, we need to do this, because the global elevation of Sph above a certain level (10 µM) is toxic to cells. In NPC but not in WT cells, Sph accumulates (1-2 µM) in the lysosome, but not in other organelles.

As such, the role of endogenous sph is not established. In this regard, the 3.5 fold increase in intracellular sph after uncaging may represent a supraphysiologic response (one reviewer was not aware of physiologic conditions where intracellular sph increases by that much).

Here we run into one of the aforementioned misunderstandings. We failed to make it clear that the 3.5-fold increase in Sph levels after uncaging as measured by mass spectrometry was due to a supramaximal illumination for chemical analysis and higher than that achieved in the cell biology experiments. For the mass spec analysis we had to perform a batch experiment where a 6 cm dish was fully illuminated for 2 min to be able to detect the uncaged species. In the live cell microscopy experiments, we illuminate only a region of 9 µm2 within a single cell for 3 sec, before we start monitoring cytosolic calcium levels in the entire cell. Hence, the increase in Sph levels will much smaller in the physiological experiments (unfortunately, there is no fluorescent sensor available to directly measure Sph levels in cells).

The study would be much stronger if there were ANY additional results to implicate endogenous sph in this mechanism and this pathway.

We would like to state that there is indeed support of our claim that Sph acts as a signaling molecule in lysosomes. In an admittedly correlative study, Fran Platt’s lab showed that increased endogenous levels of Sph correlate well with reduced lysosomal calcium levels in NPC disease (Lloyd-Evans et al. Nat. Med. 2008).

Further, we have invested quite some effort to measure lysosomal calcium release on isolated lysosomes upon addition of “endogenous” Sph. However, after consultation of specialists in the field we learned that this has never been published before. Not surprisingly we failed to set up a robust assay that demonstrates the release of lysosomal calcium after Sph elevation in a lysosomal preparation. Measuring lysosomal calcium directly in living cells is complicated by the fact that all calcium sensors are strongly pH-dependent. Ideally, we would generate a new construct of TPC1 fused to an extralysosomal calcium indicator such as a genetically-encoded calcium indicator. This is subject of current investigations in the Galione lab and needs significant optimization before it will be useful for our purpose.

Moreover, the authors use various approaches but they don't connect them or control for them appropriately. For example, they show that exogenous sph does not work but they don't measure cellular sph after the exogenous sph which is important. If this is at the same level as what they see with caged-sph, then they have a problem.

We refer to the misunderstanding mentioned above. The levels of Sph measured in mass spec studies after uncaging is not relevant for the live-cell microscopy experiments. It simply served to show that Sph was not released until illumination and that “endogenous” Sph was released. Nevertheless, we tried to measure intracellular Sph levels after addition of exogenous Sph as suggested. However, this is quite challenging as most of the exogenous Sph simply associates with the PM and cannot be washed away. This has been previously seen by the Riezman lab when radiolabeling cells using 3H-sphinganine. This rapid cell association leads to overestimation of the “internal” Sph levels. A crude estimate of how much Sph was taken up by the cells can be gained from measuring S1P-levels, which increased quite slowly in a time-dependent manner (much slower than the calcium signal happens after uncaging Sph, see inserts). We are aware that a quantification of Sph after addition to cells is not useful because excess Sph cannot be washed away. Due to these complications, we decided not to include the data in the manuscript, but we add it for the reviewers only.

Author response image 1
Comparative lipid analysis by mass spectrometry.

Lipid extracts of HeLa cells pulsed with (A). Sph-Cou and then uncaged or (B). upon addition of exogenous Sph were AQC derivatized and measured on a TSQ Vantage mass spectrometer. Values are normalized to the C17 internal standards.

https://doi.org/10.7554/eLife.10616.019

Similarly, if they are unable to uncage dh-sph (they don't measure free dh-sph in response to uncaging the way they measure sph after uncaging sph), then they cannot use this as a basis to claim high specificity (which is one of the most important points in support of the specificity and therefore significance of the sph effects on calcium.) The finding that dh-sph is not functional is a strong indication of specificity of action; therefore to clinch this point, the authors need to measure dh-sph after its uncaging.

Again, this is a point we were not sufficiently clear about: In Figure 2, all controls were performed by uncaging dh-Sph. When we uncaged Sph in Figure 1—figure supplement 2, we also measured dh-Sph (which did not change). We now added to the data set the results of uncaging dh-Sph and measuring dh-Sph and Sph levels (Figure 1—figure supplement 2) and added a paragraph in the Results section.

2) There is no evidence that Sph directly interacts with the TPC1 channel-no targets of Sph were identified. They could use click chemistry and show that uncaged Sph is bound to something specific to a calcium transporter or something specific to the lysosome-but this is likely beyond the scope of the present study. Because exogenous Sph itself does not induce Ca2+ release, the Discussion, Conclusions, and the title should be toned down.

We agree that showing direct interaction of Sph with TPC1 would improve the manuscript and simplify conclusions. However, we currently only have hints pointing to such an interaction. Crosslinking experiments followed by proteomic profiling with pacSph (see Haberkant et al, 2015) revealed TPC1 as a Sph interactor in one experiment out of three (interestingly, TPC2 did not show up at all). Since TPC1 expression levels are quite low and any potential interaction could easily be missed in such a complex experiment, we tried crosslinking on lysosomal purifications and detection by Western Blot. Unfortunately, we could not reliably detect any such interaction. We also tried to overexpress a new TPC1-FLAG construct followed by crosslinking with pacSph. Again, Western Blot experiments were not successful with this construct. We are now considering the possibility that Sph, analogous to NAADP, exerts its function on TPC1 via an adaptor protein. The search for such proteins will unfortunately exceed the scope of this manuscript (as mentioned by the reviewers). However, we believe that the lysosomal interactome of Sph will be a fabulous topic for the future. To reflect these difficulties, we changed the title to “Intracellular sphingosine releases calcium from lysosomes”

3) The effects of TFEB may or may not be related to the specific action of lysosomal sph. They use pac-sph to 'establish' the sub cellular localization of sph, but this compound would resemble more exogenous sph in its pharmacology and therefore should not act on TFEB or calcium.

We acknowledge that there was a mix-up of results and reasoning on our side. We have now changed the first sentence subsection “Subcellular localization of Sph in NPC disease to “Next, we investigated whether Sph is localized to the endosomal/lysosomal compartment in control as well as NPC1 fibroblasts by using a newly reported photoactivatable and clickable version of Sph (pacSph) “We clearly demonstrate that pac-Sph is located predominantly to the lysosome. This is supported now by Haberkant et al. where the metabolism of pac-Sph is shown, albeit in cells that lack the S1P-lyase.

Does the pac-Sph translocate TFEB? If not, then its cellular localization is irrelevant. If it does, then how come 'plain' sph does not act on calcium? Does exogenous sph induce TFEB?

We investigated the effects of pacSph as well as exogenous Sph on TFEB and did not observe any translocation. This is to be expected as both lipids do not release lysosomal calcium. As for using pacSph to probe for lipid localization, we would argue that this is an important tool. Our study (and the related manuscript Haberkant et al, 2015) uses pacSph for the first time to visualize subcellular sphingolipid distribution. Even though pacSph does not induce lysosomal calcium, most probably because it does not reach sufficiently high concentrations in a short period of time, this probe still reports on endogenous Sph localization. This is important for our conclusion that Sph acts on the lysosome.

To demonstrate the difference of elevated Sph levels after uncaging vs addition, we now used a caged version of pac-Sph and showed that it elevates calcium levels after uncaging just as caged-Sph does (see the Author response image 2).

Author response image 2
Uncaging of caged pacSph increases cytosolic calcium.

Mean Fluo-4 fluorescence traces of HeLa cells incubated with Sph-Cou or TFS. Uncaging was performed in a small circular area within the cell for 3 s as indicated by the black bar. The standard error of the mean is plotted as error bars.

https://doi.org/10.7554/eLife.10616.020

We think however that this additional data set inflates the manuscript and provide it only for inspection by the reviewers.

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

The results are incomplete with respect to biological effects resulting from calcium mobilization by SPH. One would predict that this magnitude of acute calcium release would mimic many of the effects of stimuli that induce downstream effects through calcium mobilization. Providing evidence with respect to some biological effects would significantly improve the manuscript.

Following the findings of Medina et al. (Nat Cell Biol 2015), starvation conditions lead to mTOR release from the lysosomal surface as well as lysosomal calcium release, subsequent TFEB dephosphorylation and nuclear translocation. We confirm this now by showing that Sph-induced lysosomal calcium release leads to TFEB translocation (new Figure 7). Interestingly, uncaging calcium in the cytosol by a flash of light does not lead to TFEB translocation despite the same massive signal (new Figure 7). It should be mentioned that many other calcium effects will likely also be induced by lysosomal calcium release, simply because the fast increase following Sph uncaging will induce calcium-induced calcium release from the ER.

It is worrying that sphingosine proper does not cause a Ca2+ signal (Figure 1—figure supplement 2). This really needs clarification.

Unfortunately, the addition of Sph to the extracellular space is cytotoxic and frequently induces apoptosis. Therefore only low micromolar amounts can be added in a live cell experiment which is mentioned in several papers (e.g. Chang, J. Dermat. 2004). We believe that, while the lipid enters properly, fairly rapid metabolism will prevent that sufficient amounts will reach the lysosome. Local uncaging at internal membranes seems to overcome this problem. Sph is now available in sufficient amounts close to its target. Further, we can dose Sph release well by choosing a larger or smaller area or less light. We now discuss this in the main text.

Additionally, the authors state in the Results and Discussion that Ca2+ waves initiate at the point of uncaging and propagate throughout the cell but this is not quantified and this is not clear from the supplemental movie.

To us, it was not surprising that a calcium wave starts from the point of uncaging. This can be explained by the above mentioned calcium-induced calcium release that is known to propagate through the cytosol. In addition, the diffusion or transport of the released Sph will also not be as fast as a water soluble compound would be. Nevertheless, we eliminated the sentence stating that the calcium wave propagates through the cell to avoid confusion.

The pharmacology of the sphingosine-evoked Ca2+ signals is not so well defined. In Figure 3A, it is unclear exactly how Ca2+influx was blocked (Ni v EGTA v both).

We thank the reviewers to spot this shortcoming. We now altered the figure legend accordingly.

The effect of XeC is modest but activation of TPCs is thought to result in secondary Ca2+ release from the ER.

First, it was important to see that XeC was unable to block Ca2+ release. This supported our hypothesis that Sph does not activate release from the ER. The traces shown in Figure 3A further show that the response to Sph uncaging is now reduced which we attribute to the lack of calcium-induced calcium release via the IP3 channel. It should be mentioned that the initial phase of the calcium increase is the same in XeC and control treated cells (and also in PM blocked cells).

In Figure 3B, GPN appears to induce a Ca2+ signal that is comparable in amplitude to ATP (which the authors ascribe to ER release) which is surprising. Perhaps the GPN response causes ER Ca2+ release.

First, it was important to see that XeC was unable to block Ca2+ release. This supported our hypothesis that Sph does not activate release from the ER. The traces shown in Figure 3A further show that the response to Sph uncaging is now reduced which we attribute to the lack of calcium-induced calcium release via the IP3 channel. It should be mentioned that the initial phase of the calcium increase is the same in XeC and control treated cells (and also in PM blocked cells).

Uncaging after addition of ATP/GPN is complicated by the elevated baseline. Would exogenous ATP fully deplete ER stores of calcium?

We tried to give another dose of ATP and most cells did not produce a second calcium transient. Of course, this only controls for ATP dosage. The proper experiment would have been to over-express the adrenergic receptor. Nevertheless, the strong calcium release after ATP will have likely emptied most of the available ER pool. In fact, after ATP addition, Fluo-4 (Kd for Ca2+ = 345 nM) saturates meaning that the intracellular calcium concentration probably exceeds 1 µM. As ATP is not washed away, the ER stores should not refill.

What about using thapsigargin?

We tried thapsigargin once and found no calcium elevation after Sph uncaging. When looking up the literature, we found that thapsigargin-induced calcium mobilization indeed abolishes the effect of GPN, meaning that lysosomal calcium stores are sensitive to thapsigargin and likely its inhibitory effect on ATPases (Sivaramakrishnan et al., J. Cell Sci. 2012).

There is little discussion of why Ned-19 is ineffective. Please show effects of Ned-19 on NAADP action or some positive control (Figure 4—figure supplement 2).

Our mistake: It was shown that Ned-19 is ineffective on TPC1 (but triggers TPC2). We now added a sentence stating this fact and cite Pitt et al., Sci Signal 2014.

It is clear in Figure 4 that sphingosine responses are reduced upon TPC1 knockdown but it is not so clear whether this is a direct effect of sphingosine on TPC1.

We agree with the reviewers that we do not show direct binding of Sph to TPC1 and that this would be very important. But with all respect, this is a major task beyond the scope of this manuscript. We are aiming on showing this interaction in a future collaboration with structural biologists. For the reviewers only: we used a photo-crosslinking and clickable Sph derivative and performed proteomic analysis after cross-linking, clicking on biotin and extracting the lipid-protein conjugates. In one of two replicates, TPC1 was identified while TPC2 was not found. While these data are very preliminary, they support a direct interaction of Sph with TPC1.

The authors claim that sphingosine-mediated Ca2+ signals are reduced in NPC (Figure 5). The quantification (Figure 5B) does not reflect this (the proportion of responding cells is only slightly reduced) and it is unclear if there is an effect on the amplitude.

The average responder shows a clearly reduced amplitude. Further, the fraction of cells that respond with less than 50% amplitude change was 28% in control cells and 69% in NPC fibroblasts.

The apparent reduction in signal in Figure 5A in NPC is not dissimilar to the effect of Sph kinase block in Figure 2 (FtY720), XeC/PM block in Figure 3 and TPC2 silencing in Figure 4 yet a role for the intended targets in sphingosine action were discounted in Figures 2-5.

We agree that we show many traces with a reduced calcium response after treatment. Here, however, there is no treatment. We just compare two very similar cell types (healthy vs NPC subject cells). Just because a pharmacological treatment reduces the calcium response, this does not invalidate a comparison of the two cell lines.

The subcellular localisation/pulse chase data in Figure 6 needs quantitation and the authors' claim that sphingosine localizes to the endolysosomal system (Figure 6—figure supplement 1) is not so convincing.

We thank the reviewers for this hint. We now quantified the colocalization by determining Pearson’s correlation coefficients, added a new Figure 6C and expanded the corresponding Figure 6—figure supplement 1.

The authors imply that reduced sphingosine-mediated Ca2+ signal in NPC is due to activation of TPC1 by elevated sphingosine levels causing a reduction of Ca2+ levels within the acid Ca2+ store. A previous study (led by co-authors) however concluded that reduced Ca2+ content in the acid Ca2+ store was due to reduced Ca2+ uptake (Lloyd-Evans, 2008). In the Introduction, the authors refer to a study where Ca2+ release from the acidic Ca2+ store was reduced in NPC but in that study (Shen; 2012), lysosomal Ca2+ content was unchanged and rather lipid accumulation was proposed to block Ca2+ release through TRPML. This is a little confusing and potentially misleading. Cause-effect relationships are blurred.

We agree that the statements are a bit confusing. We now changed the Introduction and made clear in the Discussion that the given statements are part of an ongoing discussion in the field.

Other Points: 1) The approach to block the conversion of SPH into S1P with DMS and FTY720 is somewhat problematic because DMS is a non-specific SPHK inhibitor and FTY720 is a SPH analogue, and both may mimic SPH in inducing calcium release on their own. Please note that FTY FTY720 is not an inhibitor of SK1. Moreover, the concentrations of DMS and FTY may not be sufficient to displace sph from sphingosine kinase. The authors should use genetic approaches to block the conversion of SPH to S1P in cells.

Unfortunately, the use of RNAi against both Sph kinases was unsuccessful, probably because we blocked the main sphingolipid degradation pathway leading to toxic sphingolipid levels (now stated in the text). We then employed the non-sphingosine analog inhibitor SKI-II which gave a similarly small effect on Sph-induced calcium release. The data is incorporated into Figure 2, Figure 2—figure supplement 2.

2) Results in Figure 1—figure supplement 1 do not establish metabolic inertness of the caged compound. There is time dependent uptake, the signal is too faint to detect a sizable metabolite in the order of 10-20% or so, and this approach does not evaluate for spontaneous uncaging (the product would not be fluorescent).

3) The authors need to measure levels of sphingosine and metabolites at the earliest time point if they are able to.

We agree that a thorough control of sphingosine levels before and after uncaging was missing. We now teamed up with the group of Howard Riezman. They performed a lipidomics analysis. Results: incubation with caged Sph does not lead to changes in any of the sphingoid bases (Figure 1—figure supplement 2). After uncaging, Sph increased 3.4-fold while dihydroSph did not.

4) Results and Discussion; full name of EGTA is not ethylene tetra-acetic acid.

Corrected, thank you.

5) Figure 3A. It looks like PM block has a significant effect on the sph response? This appears to be of the same magnitude as the loss in NPC. Also, this figure, in contrast to the legend, does not show the effects of EGTA or Ni.

It is well known that the use of calcium channel blockers reduces the size of the intracellular calcium pool simply because the leaky ER loses calcium that is partially pumped out and cannot be refilled. Therefore, in PM blocked cells, the response will be always reduced.

6) SPH can also be quickly converted to ceramides through ceramide synthases or reverse action of ceramidases. The authors should investigate if blocking the conversion of SPH to ceramides can affect the calcium release by uncaged SPH. Also, studies in Farber's disease would be important.

Although we did not specifically investigate ceramide formation, the Sph kinase inhibitor FTY720 we used also happens to block ceramide synthase activity (Lahiri et al, J. Biol. Chem. 2009). Regarding Farber’s disease: there are around 80 cases reported worldwide and it would be very tough to get cells from such patients. We agree that it would be very interesting to do such experiments.

7) The authors should investigate if increasing the generation of endogenous SPH by activating a ceramidase can induce the release of calcium from a calcium store(s).

We are not aware of a small molecule able to activate ceramidase. Over-expression looks like a dirty experiment as the cell would try to adapt to the higher Sph levels, especially considering the aforementioned toxicity.

8) The authors could measure the content of calcium in the endolysosomal system to determine if uncaging SPH indeed empties this calcium store. This will give another line of evidence that SPH releases calcium from this particular cellular compartment.

This would be a great experiment. Unfortunately, these are also very difficult experiments as the lysosome is quite acidic (and worse, inhomogeneous ‘pH’) and calcium indicators will have altered calcium affinities (and quantification) under these conditions. Genetically encoded calcium sensors cannot be employed because the uncaging procedure would bleach them and hence would prevent quantification.

9) The max. amplitude change comparisons are often hard to compare – would additional kermel density estimations help, or report means where the data are relatively Gaussian?

We agree that the histograms are not very easy to compare. However, once the reader has focused on this form of presentation, it is quite informative. We looked at a Gaussian presentation of the mean. Unfortunately, the cell number available does not suffice to present decent curves.

10) The data on localization of Sph in NPC are hard to interpret (Figure 6). Only one cell is shown, and it appears to include significant ER staining. LAMP1 staining also seems unusual in this one cell compared to other published reports for NPC1 mutant cells. Please add additional examples and consider including an ER marker to strengthen your conclusions.

We agree. We now show at least two cells in all co-localization conditions. Further, we also show a new Figure 6C that provides the Pearson’s coefficient for 6 to 10 cells per co-localization.

11) A recent paper in Science showed a connection between NPC1 and TPC proteins in that both seem to be needed for Ebola virus entry. Please cite and speculate how your work relates to that story.

The mentioned paper came out after we submitted the original manuscript. We now cite this paper and discuss its content in the conclusion section.

https://doi.org/10.7554/eLife.10616.022

Article and author information

Author details

  1. Doris Höglinger

    Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    Contribution
    DH, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  2. Per Haberkant

    Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    Contribution
    PH, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  3. Auxiliadora Aguilera-Romero

    Department of Biochemistry, University of Geneva, Geneva, Switzerland
    Contribution
    AAR, Design, performance and analysis of lipidomics experiments, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  4. Howard Riezman

    Department of Biochemistry, University of Geneva, Geneva, Switzerland
    Contribution
    HR, Design and analysis of lipidomics experiments, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  5. Forbes D Porter

    Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, United States
    Contribution
    FDP, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  6. Frances M Platt

    Department of Pharmacology, University of Oxford, Oxford, United Kingdom
    Contribution
    FMP, Conception and design, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  7. Antony Galione

    Department of Pharmacology, University of Oxford, Oxford, United Kingdom
    Contribution
    AG, Drafting or revising the article, Contributed unpublished essential data or reagents
    Competing interests
    The authors declare that no competing interests exist.
  8. Carsten Schultz

    Cell Biology and Biophysics Unit, European Molecular Biology Laboratory, Heidelberg, Germany
    Contribution
    CS, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    For correspondence
    schultz@embl.de
    Competing interests
    The authors declare that no competing interests exist.

Funding

Seventh Framework Programme (289278 - Sphingonet)

  • Doris Höglinger
  • Howard Riezman
  • Frances M Platt
  • Carsten Schultz

National Center for Competence in Research (Chemical Biology)

  • Auxiliadora Aguilera-Romero
  • Howard Riezman

European Commission (EMBL Interdisciplinary Postdoc Programme)

  • Per Haberkant

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

Acknowledgements

We are grateful for the support by the staff of the Advanced Light Microscopy Facility of the European Molecular Biology Laboratory for maintaining the microscopes used in this study. We thank Frank Stein for helpful suggestions for data analysis with FluoQ. We gratefully acknowledge help from Isabelle Riezman for the long chain base analysis and support from the Swiss National Science Foundation and the NCCR Chemical Biology. F.M.P. is a Royal Society Wolfson Research Merit Award holder. The research leading to these results has received funding from the European Union Seventh Framework Programme (FP7 2007–2013) under grant agreement no 289278 - "Sphingonet".

Reviewing Editor

  1. Suzanne R Pfeffer, Stanford University School of Medicine, United States

Publication history

  1. Received: August 4, 2015
  2. Accepted: November 27, 2015
  3. Accepted Manuscript published: November 27, 2015 (version 1)
  4. Version of Record published: January 27, 2016 (version 2)

Copyright

This is an open-access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

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Further reading

    1. Cell Biology
    Hongyan Hao et al.
    Research Article

    KASH proteins in the outer nuclear membrane comprise the cytoplasmic half of LINC complexes that connect nuclei to the cytoskeleton. Caenorhabditis elegans ANC-1, an ortholog of Nesprin-1/2, contains actin-binding and KASH domains at opposite ends of a long spectrin-like region. Deletion of either the KASH or calponin homology (CH) domains does not completely disrupt nuclear positioning, suggesting neither KASH nor CH domains are essential. Deletions in the spectrin-like region of ANC-1 led to significant defects, but only recapitulated the null phenotype in combination with mutations in the trans-membrane span. In anc-1 mutants, the ER, mitochondria, and lipid droplets were unanchored, moving throughout the cytoplasm. The data presented here support a cytoplasmic integrity model where ANC-1 localizes to the ER membrane and extends into the cytoplasm to position nuclei, ER, mitochondria, and likely other organelles in place.

    1. Cell Biology
    2. Physics of Living Systems
    Manuel Giménez-Andrés et al.
    Research Article

    Numerous proteins target lipid droplets (LDs) through amphipathic helices (AHs). It is generally assumed that AHs insert bulky hydrophobic residues in packing defects at the LD surface. However, this model does not explain the targeting of perilipins, the most abundant and specific amphipathic proteins of LDs, which are weakly hydrophobic. A striking example is Plin4, whose gigantic and repetitive AH lacks bulky hydrophobic residues. Using a range of complementary approaches, we show that Plin4 forms a remarkably immobile and stable protein layer at the surface of cellular or in vitro generated oil droplets, and decreases LD size. Plin4 AH stability on LDs is exquisitely sensitive to the nature and distribution of its polar residues. These results suggest that Plin4 forms stable arrangements of adjacent AHs via polar/electrostatic interactions, reminiscent of the organization of apolipoproteins in lipoprotein particles, thus pointing to a general mechanism of AH stabilization via lateral interactions.