Cytokines and chemokines are produced and secreted by a broad range of immune cells including macrophages. Remarkably, little is known about how these inflammatory mediators are released from the various immune cells. Here, the endolysosomal cation channel TRPML2 is shown to play a direct role in chemokine trafficking and secretion from murine macrophages. To demonstrate acute and direct involvement of TRPML2 in these processes, the first isoform-selective TRPML2 channel agonist was generated, ML2-SA1. ML2-SA1 was not only found to directly stimulate release of the chemokine CCL2 from macrophages but also to stimulate macrophage migration, thus mimicking CCL2 function. Endogenous TRPML2 is expressed in early/recycling endosomes as demonstrated by endolysosomal patch-clamp experimentation and ML2-SA1 promotes trafficking through early/recycling endosomes, suggesting CCL2 being transported and secreted via this pathway. These data provide a direct link between TRPML2 activation, CCL2 release and stimulation of macrophage migration in the innate immune response.https://doi.org/10.7554/eLife.39720.001
Cytokines/chemokines are released from a wide range of immune cells such as macrophages, B- and T-lymphocytes, neutrophils, mast cells and dendritic cells. They are essential for intercellular communication in both innate and adaptive immunity. Remarkably, our knowledge of the function of cytokines/chemokines in immunity is much more advanced than our knowledge about how they are packaged and secreted from immune cells. Understanding how innate immune cells release cytokines/chemokines is important, as these factors are indispensable for communication between immune but also with non-immune cells to coordinate inflammatory responses (Lacy and Stow, 2011). Importantly, secretion pathways vary between different cell types. Macrophages for example lack typical secretory granules (Lacy and Stow, 2011). Thus, macrophage cytokine/chemokine release is mediated either by direct transport to the cell surface from the trans-Golgi network (TGN) (e.g. IL-10), by transport via recycling endosomes (RE) to the cell surface (e.g. TNF-α, IL-6, IL-10) (Manderson et al., 2007; Murray and Stow, 2014), or via late endosomes/lysosomes (LE/LY), for example IL-1β (Andrei et al., 1999; Lopez-Castejon and Brough, 2011).
We show here that the endolysosomal calcium-permeable cation channel TRPML2 plays a direct role in chemokine secretion, thereby modulating the inflammatory response. Expression of TRPML2 in different immune cells and tissues has been demonstrated by several groups (Cuajungco et al., 2016; Valadez and Cuajungco, 2015; García-Añoveros and Wiwatpanit, 2014; Sun et al., 2015). On the subcellular level, TRPML2 has been shown to be expressed primarily on RE and LE/LY by immunocytochemistry experiments (Sun et al., 2015; Venkatachalam et al., 2006; Karacsonyi et al., 2007). However, functional expression of TRPML2 in different intracellular vesicles and organelles has not been confirmed yet by direct and selective patch-clamp analysis, that is patch-clamping of RE, EE (early endosomes), LE/LY, or other endolysosomal vesicles. Furthermore, it remains unclear whether direct and selective stimulation of TRPML2 leads to an increase in cytokine/chemokine release from macrophages, and which intracellular trafficking pathways mediate the release of these cytokines/chemokines.
One important impediment for the investigation of different endogenous TRPML-like currents and their functional impact on secretion, endolysosomal trafficking, or autophagy, is the lack of isoform-selective agonists. Development of such agonists would allow demonstration of the TRPML isoform-specific contribution towards observed phenomena, for example chemokine secretion. Currently available TRPML channel agonists belong to different chemotypes, including benzenesulfonamides (e.g. SN-1- or SF-21-type), thiophenesulfonamides (e.g. SF-22-type, including MK6-83), isoindolediones (e.g. SF-51-type, including ML-SA1), isoxazolines (e.g. SN-2-type) and others (Grimm et al., 2010; Yamaguchi and Muallem, 2010; Grimm et al., 2012b; Shen et al., 2012). Efficacy, potency and selectivity of these compounds can vary between species. Furthermore, none of the currently available TRPML agonists is selective for TRPML1 or TRPML2. ML-SA1 for example activates TRPML1 and TRPML3 in mouse, while it activates all three human isoforms (Shen et al., 2012; Grimm, 2016). MK6-83 activates TRPML1 and TRPML3 in both mouse and human (Grimm, 2016; Chen et al., 2014). The putative endogenous TRPML channel activator PI(3,5)P2 activates all three TRPML channel isoforms in both species and, in addition, also activates the endolysosomal cation channels TPC1 and TPC2 (Chen et al., 2014; Wang et al., 2012; Cang et al., 2013; Grimm et al., 2014). Through systematic chemical modification of known lead structures we have now generated the first isoform-selective TRPML2 channel agonist, ML2-SA1.
We demonstrate that ML2-SA1 activates TRPML2 in EE and LE/LY as well as in Rab11+ and Tf+/TfR+ (transferrin/transferrin receptor) vesicles. In macrophages, LPS (lipopolysaccharide) exposure leads to a strong upregulation of TRPML2 expression, while TRPML1 and TRPML3 expression levels remain unaffected by LPS (Sun et al., 2015). Importantly, activation by ML2-SA1 was not observed in macrophages without LPS treatment which express TRPML2 only at very low levels, further confirming specificity of the compound. We also show that direct activation of TRPML2 by ML2-SA1 results in an increased release of the chemokine CCL2 from LPS-stimulated WT macrophages, while TRPML2-/- macrophages show no release increase, suggesting that TRPML2 channel activity is directly linked to CCL2 trafficking and secretion. We further provide evidence that CCL2 is released via the early/recycling endosomal pathway but not via LE/LY. Finally, we show that stimulation with ML2-SA1 promotes macrophage migration, one of the major physiological functions of the chemoattractant CCL2, one synonym of which is monocyte chemoattractant protein 1 (MCP-1).
With the aim to further improve the characteristics of existing TRPML channel agonists, we generated more than 80 novel derivatives of recently reported lead activators of TRPML channels which had been originally identified by random screening of the MLSMR small molecule library (Scripps Research Institute Molecular Screening Center) (Grimm et al., 2010). Here, novel derivatives of the lead compounds SN-2 and ML-SA1, a SF-51 analogue (Grimm et al., 2010; Shen et al., 2012; Grimm, 2016; Chen et al., 2014) were evaluated for their efficacy, potency, and selectivity profiles, respectively.
We first synthesized and tested >50 chemically modified versions of the TRPML3 activator SN-2 (Figure 1; Figure 1—figure supplement 1; Supplementary file 1). These modifications comprise systematic variations of the substitution pattern of the aryl ring, variations of the aliphatic norbornane ring system, aromatisation of the isoxazoline to an isoxazole fragment, introduction of polar substituents, as well as replacement of the isoxazol(in)e ring by other heterocycles. Crucial steps in these syntheses were Huisgen-type 1,3-dipolar cycloaddition reactions of norbornene (for the closer analogues) and other alkenes with nitrile oxides (Jawalekar et al., 2011; Huisgen, 1963) and related 1,3-dipoles. Related aromatic isoxazole analogues were prepared via cycloaddition of nitrile oxides with ketone enolates (Vitale and Scilimati, 2013) or enamines (Fos et al., 1992). General synthesis strategies for these modifications are shown in Figure 1A.
Derivatives of SF-51/ML-SA1 (Figure 1—figure supplement 2; Figure 1—figure supplement 3; Supplementary file 1) were synthesized by combining appropriate amine building blocks (partially hydrogenated quinolines and other cyclic and open-chain analogues) with N-acyl spacers and imide/lactam-type residues following standard procedures (Figure 1—figure supplement 3A).
Following synthesis, we initially tested the compounds in HEK293 cells transiently transfected with human TRPML1, TRPML2, or TRPML3 (C-terminally fused to YFP) by using the fura-2 calcium imaging technique. When expressed in HEK293 cells, TRPML2 and TRPML3 but not TRPML1 substantially localize at the plasma membrane besides endolysosomes as described previously (Grimm et al., 2010), enabling standard fura-2 calcium imaging experimentation. To evaluate effects on TRPML1, a plasma membrane variant with mutated lysosomal targeting sequences in the N- and C-termini (TRPML1(NC)) was used as reported previously (Grimm et al., 2010).
The majority of the SN-2 and SF-51/ML-SA1 derivatives were either inactive, non-selective like ML-SA1, or selective for TRPML3 like SN-2 (Figure 1B; Figure 1—figure supplement 3B). A subset of molecules however displayed a strong preference for TRPML2: ML2-SA1 (=EVP-22), a derivative of SN-2, as well as derivatives of SF-51/ML-SA1: EVP-198, EVP-207 and EVP-209. The latter three SF-51/ML-SA1 derivatives however showed lower efficacy compared to ML2-SA1 (Figure 1B; Figure 1—figure supplement 3B).
In endolysosomal patch-clamp experiments using transiently transfected HEK293 cells, we investigated TRPML2 channel activity in wortmannin/latrunculin B (Wort./Lat.B)-enlarged EE (Chen et al., 2017a), in YM201636-enlarged LE/LY (Chen et al., 2017a), as well as in vacuolin-enlarged Rab11+ and TfR+ organelles (Figure 2; Figure 2—figure supplement 1). In LE/LY, both ML2-SA1 (Figure 2B; Figure 2—figure supplement 1A) and PI(3,5)P2 (Figure 2—figure supplement 1A) evoked TRPML2 activation while no or very little activation was detectable for TRPML1 and TRPML3. In contrast, the latter ones were robustly activated by ML-SA1 as a positive control (Figure 2C–E). The time course for activation of TRPML2 in LE/LY patch-clamp experiments and the relative Ca2+ permeability are shown in Figure 2G and Figure 2—figure supplement 1B. In addition to LE/LY, TRPML2 channel activity was also detectable in EE after stimulation with ML2-SA1 (Figure 2H and K). In order to patch-clamp discrete populations of vesicles involved in early/recycling endosomal trafficking, cells were transfected with fluorophore-tagged Rab11 or TfR, and enlarged with vacuolin. ML2-SA1 elicited significant currents in Rab11+ and in TfR+ vesicles (Figure 2I–K).
Furthermore, the effect of luminal pH on TRPML2 channel activity was evaluated (Figure 2—figure supplement 1A and C). TRPML2 activity (stimulated with PI(3,5)P2 or with ML2-SA1) increases with increasing, that is less acidic luminal pH. This differs from TRPML1 which shows maximal activity in highly acidic luminal pH (Chen et al., 2017a; Dong et al., 2010). These findings argue TRPML2 channel function is adapted to vesicles of only slightly acidic or neutral pH such as EE/RE rather than highly acidic LE/LY. The strong colocalization between TfR or Rab11 with TRPML2 confirms an important functional role of TRPML2 in RE (Figure 2—figure supplement 2A–B).
In summary, ML2-SA1 was found to be a potent and efficacious activator of both hTRPML2 and mTRPML2. The calculated EC50 values for human and mouse TRPML2 were 1.24 ± 0.12 μM and 2.38 ± 0.01 μM, respectively (Figure 2F; Figure 2—figure supplement 3A). ML2-SA1 shows high selectivity over h/mTRPML1 and h/mTRPML3 in both calcium imaging and endolysosomal patch-clamp experiments and it does not activate TPC1 nor TPC2 (Figure 1; Figure 2; Figure 2—figure supplement 3B–F).
Several recent papers have provided in-depth information on the structures of TRPML1 and TRPML3 channels (Schmiege et al., 2017; Chen et al., 2017b; Hirschi et al., 2017). Schmiege et al., 2017 found that a hydrophobic cavity created by I468 and F465 of PH1 (pore helix 1), F428, C429, V432 and Y436 of S5, F505 and F513 of S6, and Y499 and Y507 of S6 in the neighboring subunit, tightly accommodates ML-SA1 (Figure 3A). In a molecular modeling approach using these recently published structures of TRPML1 and TRPML3 as a basis, we simulated the binding of ML-SA1 as well as ML2-SA1 to hTRPML1 and hTRPML2 (Figure 3; Figure 3—figure supplement 1). Complete 3D models of the open conformation of hTRPML1 and hTRPML2 were constructed and used for ligand docking analysis. Amino acids differing between hTRPML1 and hTRPML2 are colored green (Figure 3B–D). Based on this model, ML2-SA1 (both enantiomers are described, one in Figure 3—figure supplement 1) is predicted to bind to the same binding pocket as ML-SA1 as observed in the cryo-EM structure of hTRPML1 (Figure 3A–B). Six amino acids (A422, A424, G425, A453, V460, and I498) in this pocket are unique to hTRPML2 (highlighted in green; Figure 3C–D). The orientation of ML2-SA1 in the binding pocket of hTRPML2 with the highest docking score is shown in Figure 3C. The dichlorophenyl ring shows favorable π-stacking interaction with F502 whereas the polar isoxazole ring is located near the side chain OH-groups of Y428 and Y496. The hydrophobic norbornane ring is interacting with G425 and Y428. Other possible orientations of ML2-SA1 binding to hTRPML2 are shown in Figure 3—figure supplement 1C–D). The observed binding mode of ML2-SA1 at hTRPML1 is different and appears to be energetically less favorable compared to hTRPML2 due to the observed amino acid substitutions in the predicted binding cavity (Figure 3D). We subsequently replaced each of the six amino acids that are unique to the predicted hTRPML2 binding pocket with the respective amino acids of hTRPML1. We analysed these mutant isoforms first in calcium imaging experiments where we found the strongest reduction of the ML2-SA1 effect in G425A (Figure 3E). In the next step, we performed endolysosomal patch-clamp experiments with this mutant. Mutation of G425 to alanine was found to selectively abrogate the effect of ML2-SA1, while ML-SA1 was still able to activate G425A to a degree not significantly different from WT (Figure 3E–F). G425 is close to the norbornane ring of ML2-SA1 (minimum distance 3.6 Å) docked to hTRPML2 and substitution to alanine is unfavorable for this binding mode (Figure 3C). The experimental data corroborate binding of ML2-SA1 to the ML-SA1 binding pocket and confirm a critical role of G425 in mediating ML2-SA1 selectivity.
In macrophages significant TRPML2 channel expression is found only after stimulation with LPS, as demonstrated previously by qRT-PCR and western blot analysis (Sun et al., 2015). We confirmed this finding by qRT-PCR and endolysosomal patch-clamping, revealing that only after several hours of LPS treatment, robust endogenous TRPML2 channel expression and activity were detectable (Figure 4A–G). In LPS-stimulated bone marrow-derived macrophages (BMDMΦ) ML2-SA1-induced currents were detectable in Tf-Alexa555 loaded, vacuolin-enlarged vesicles, while no significant TRPML2 channel activity could be detected in non-LPS stimulated BMDMΦ Tf+ vesicles (Figure 4A–B). Currents measured in BMDMΦ LE/LY with ML2-SA1 after LPS-stimulation were smaller than currents measured in Tf+ loaded vesicles (Figure 4C–D). In contrast, in LE/LY isolated from alveolar macrophages (AMΦ), TRPML2 currents elicited with ML2-SA1 were larger on average than in BMDMΦ (Figure 4E–F). These data confirm that ML2-SA1 elicits robust TRPML2 currents in endogenously expressing cells.
To evaluate effects of the novel TRPML2 channel agonist on chemokine secretion from macrophages, we performed experiments based on the results recently provided by Sun et al. (2015) (Figure 5A). We found that incubation with ML2-SA1 significantly increased secretion of the chemokine CCL2 from BMDMΦ, both after 4 hr and 8 hr of LPS treatment (Figure 5A). Importantly, ML2-SA1 did not induce CCL2 secretion in unstimulated BMDMΦ. Furthermore, CCL2 secretion was severely reduced in TRPML2-/- BMDMΦ and ML2-SA1 showed no further increase of CCL2 secretion in the TRPML2-/- BMDMΦ, corroborating the specificity of the agonist (Figure 5A). To characterise the pathway of ML2-SA1-induced CCL2 secretion from macrophages, we performed lysosomal exocytosis and Tf trafficking experiments to distinguish between LE/LY and EE/RE as possible secretion routes. Lysosomal exocytosis experiments revealed no significant effect of ML2-SA1 on lysosomal enzyme (beta-hexosaminidase) release (Figure 5B). In accordance with this, ML2-SA1 application did not result in translocation of LAMP1 to the plasma membrane (Figure 5C), arguing against LE/LY being involved in CCL2 secretion in BMDMΦ. These findings are supported by the LE/LY environment being less favorable for TRPML2 activity as outlined above. More favorable conditions are found in EE/RE compartments (less acid to neutral pH). In line with this, ML2-SA1 application resulted in a significant enhancement of Tf trafficking and recycling through EE/RE (Figure 5D–E). Taken together, these data argue for a TRPML2-dependent trafficking route of CCL2 from Golgi to EE/RE (Figure 5F).
To assess effects of ML2-SA1 on cell migration, we performed migration assays in a modified Boyden chamber setup (Figure 6—figure supplement 1). BMDMΦ in the presence or absence of LPS were seeded in the lower compartment of the chamber and exposed to different concentrations of ML2-SA1. LPS-stimulated, ML2-SA1 pre-treated BMDMΦ were able to significantly increase migration of untreated BMDMΦ through the transwell chamber, while LPS-stimulated BMDMΦ without ML2-SA1 pre-treatment (only DMSO) were not able to alter migration properties of untreated BMDMΦ. (Figure 6A). This is in accordance with the enhanced release of CCL2 by ML2-SA1, which serves as chemoattractant for untreated BMDMΦ. To exclude a chemotactic effect of the compounds themselves, we used a classical Boyden chamber setup without cells in the lower compartment. Yet, neither LPS nor ML2-SA1 alone were able to significantly enhance BMDMΦ migration, while supplementation of recombinant CCL2 led to a substantial increase in BMDMΦ migration (Figure 6B; Figure 6—figure supplement 2A–B). Overall, these data suggest that ML2-SA1 is able to induce CCL2 secretion selectively in TRPML2-expressing macrophages, thus serving as chemoattractant to recruit more macrophages.
We describe here a novel, isoform-selective activator of the TRPML2 channel and describe how TRPML2 activation enhances endosomal trafficking to induce inflammatory mediator release in LPS-stimulated macrophages. Until now, selective activators for TRPML2 had not been available. In an effort to identify such selective activators we synthesized >80 chemical compounds by systematic variation of the known lead structures SN-2 and SF-51/ML-SA1 (Grimm et al., 2010; Shen et al., 2012), generating a library of analogues of sufficient size to deduce structure-activity relationships. In the ML-SA1 series, improved TRPML2 activation was achieved by modification of the length of the acyl spacer, but the resulting selective activators were of only intermediate efficacy and potency. By contrast, the activator ML2-SA1 from the series of norbornene-derived isoxazolines (based on SN-2) is characterized by high TRPML2 subtype selectivity as well as high efficacy and potency, rendering this new small molecule a valuable compound for future studies on this ion channel (Supplementary file 2). Molecular modeling data support specific binding of ML2-SA1 to the pore region of the channel, as observed for ML-SA1. The binding orientation of ML-SA1 at hTRPML2 was found to be similar to the experimentally observed binding to hTRPML1 (Schmiege et al., 2017) which is in agreement with nonselective activation. In contrast, the binding orientation of docked ML2-SA1 at hTRPML1 differs from that found for hTRPML2, suggesting a plausible rationale for its selectivity. In an experimental approach where we investigated the functional consequences of point mutations in hTRPML2 with the endolysosomal patch-clamp technique we found that in mutant G425A activation by ML2-SA1 is selectively lost, while activation by ML-SA1 is preserved, indicating that this amino acid is highly critical for the selective effect of ML2-SA1 on TRPML2.
Sun et al. (2015) have recently shown that the levels of TRPML2 are strongly upregulated in macrophages upon TLR4 (toll-like receptor) activation (Supplementary file 2). Thus, treatment with LPS was found to lead to TRPML2 upregulation in, for example microglia, peritoneal macrophages, bone marrow derived macrophages, or alveolar macrophages (Sun et al., 2015). The authors further found that the translation and secretion of several chemokines such as CCL2 was reduced in TRPML2-/- mice, and concluded that TRPML2 might play a role in the regulation of trafficking and/or secretion of these chemokines. However, it remained unclear whether TRPML2 is directly involved in these processes and whether activation of TRPML2 channel activity would show increased release.
Here, we present data strongly supporting a direct involvement of TRPML2, as direct stimulation of TRPML2 with ML2-SA1 leads to an increase in CCL2 secretion from macrophages. Using the specific TRPML2 agonist and the TRPML2-/- knockout mouse model as control, we demonstrate a positive relationship between TRPML2 activity and CCL2 secretion. Using the endolysosomal patch-clamp technique we demonstrate that TRPML2 is present in LE/LY and EE as well as in Rab11+ and TfR+/Tf+ vesicles (Supplementary file 2). However, early endosomes including RE provide more favorable activation conditions for TRPML2 than LE/LY due to their less acidic/neutral luminal pH. In accordance with this, TRPML2 currents elicited with ML2-SA1 in LE/LY isolated from endogenously expressing BMDMΦ were smaller than currents in Tf+ vesicles. In addition, no evidence was found that ML2-SA1 can promote lysosomal exocytosis, while ionomycin or ML-SA1 were able to increase the release of beta-hexosaminidase as previously reported (Samie et al., 2013). The subcellular distribution of LAMP1 did also not change during ML2-SA1 treatment and no translocation to the PM was observed. In contrast, ML2-SA1 application was found to significantly promote Tf trafficking through the early/recycling endosomal compartment, arguing for a role of TRPML2 in CCL2 release via the early/recycling endosomal pathway (Figure 5F).
Loss of function mutations in the TRPML2-related channel TRPML1 result in lysosomal storage and endolysosomal trafficking defects underlying the neurodegenerative disease mucolipidosis type IV (Bach, 2001; Pryor et al., 2006; Chen et al., 2014). Mechanistically, it was postulated that loss of TRPML1 impairs lysosomal exocytosis (LaPlante et al., 2006). It was also suggested that TRPML1 is required for lysosomal pH regulation (Soyombo et al., 2006) and for vesicle fusion (Venkatachalam et al., 2013) while, very recently, data have been presented, supporting that TRPML1 may regulate lysosomal fission (Chen et al., 2017a). A further interesting finding has been presented by Park et al. (2016), suggesting that, in secretory cells, a major role for TRPML1 is to guard against unintended, pathological fusion of lysosomes with other intracellular organelles, for example secretory vesicles. TRPML1 has also been attributed to mediate lysosomal trafficking via Ca2+-dependent motor protein recruitment, its activity favoring retrograde lysosomal movement (Vergarajauregui et al., 2009; Li et al., 2016).
Like TRPML1, TRPML3 was also suggested to regulate membrane trafficking. In particular, it was found to regulate trafficking of early endosomes and to affect endocytosis (Kim et al., 2009). Lelouvier and Puertollano (2011) further presented data showing that TRPML3 is required for proper calcium homeostasis in the endosomal pathway and that impairment of TRPML3 function leads to defective endosomal acidification and defective membrane trafficking. Surprisingly, the authors found increased endosomal fusion after depletion of TRPML3. Recently, Miao et al. (2015) showed that TRPML3 activation, upon neutralization of lysosomal pH, mediates efflux of Ca2+ ions from lysosomes, which in turn induces lysosome exocytosis. TRPML3 is normally inactive in highly acidic lysosomes, in contrast to early/recycling endosomes with more neutral pH, but when the pH in the lumen of the lysosome is neutralized, TRPML3 becomes activated, releases Ca2+ into the cytosol, which in turn triggers spontaneous exocytosis of the lysosome and its contents.
TRPML2 has been suggested to play a role in the regulation of the Arf6-associated pathway and, more specifically, in the trafficking of GPI-APs (Karacsonyi et al., 2007). Arf6 has been implicated in the regulation of endocytosis as well as endocytic recycling and cytoskeleton remodeling. More recently, TRPML2 has been found to increase trafficking efficiency of endocytosed viruses (Rinkenberger and Schoggins, 2018). Furthermore, we are showing here that TRPML2 is, like its relatives TRPML1 and 3, Ca2+ permeable (Figure 2—figure supplement 1C).
Taken together, these findings imply that all three TRPML channels can impact intracellular trafficking processes while the mechanisms how they affect trafficking might differ. While it is likely, based on the available data, that the effect of TRPML2 knockout/activation on CCL2 trafficking and release is occurring at the level of EE/RE, it remains to be further established where along this pathway the effect takes place. Possible scenarios might be: fusion of Golgi vesicles with RE, fission from RE, fusion of Golgi vesicles with EE, fission from EE, fusion of EE-derived vesicles with the RE (Figure 5F).
Functionally, we found that ML2-SA1 promotes migration of untreated macrophages towards LPS-treated macrophages. This suggests that TRPML2-dependent CCL2 release is enhancing the inflammatory response by recruiting innate immune cells to the site of inflammation. This is in accordance with the results presented by Sun et al. (2015) who found that macrophage migration is impaired in vivo in the absence of TRPML2.
CCL2 is known to be a key chemokine regulating migration and infiltration of monocytes/macrophages (Deshmane et al., 2009). Since CCL2 is implicated in the pathogenesis of diseases characterized by infiltrates containing macrophages like psoriasis, rheumatoid arthritis, multiple sclerosis, and atherosclerosis (Deshmane et al., 2009; Xia and Sui, 2009; Daly and Rollins, 2003), we postulate that TRPML2 may be an attractive novel target for the treatment of such innate immunity-related inflammatory diseases.
Whole-LE/LY and whole-EE recordings have been described previously in detail (Chen et al., 2017a; Chen et al., 2017c). In brief, for whole-LE/LY manual patch-clamp recordings, cells were treated with YM201636 (HEK293 cells: 800 nM o/n; macrophages: 800 nM 1 hr). For whole-EE manual patch-clamp recordings, cells were treated with a combination of 200 nM wortmannin and 10 nM latrunculin B (HEK293 cells: 10–15 min). Cells were treated with compounds at 37°C and 5% CO2. YM201636 was obtained from Chemdea (CD0181), wortmannin and latrunculin B from Sigma (W1628 and L5288), and vacuolin from Santa Cruz (sc-216045). Compounds were washed out before patch-clamp experimentation.
For other organelle patch-clamp recordings, HEK293 cells were transfected with the markers Rab11-DsRed or TfR-mCherry, respectively, and treated with 1 μM vacuolin o/n. Since macrophages could not be transfected with standard transfection protocols or by electroporation, cells were loaded with transferrin-Alexa555 and simultaneously treated with vacuolin for 1 hr to enlarge and visualize vesicles for patch-clamp.
Isolation-micropipettes were used to open up the plasma membrane, and push the enlarged vesicle of interest out of the cell. Afterwards, electrode-micropipettes were applied to patch-clamp the isolated vesicles.
Macrophages were used for experiments within 2–10 days after isolation. Mean capacitance values for Rab11+ vesicles isolated from HEK293 cells was 0.7 ± 0.2 (n = 6), for TfR+ vesicles (n = 3) 1.4 ± 0.3 pF, for EE (n = 10) 0.4 ± 0.1 pF, and for LE/LY (n = 51) 1.0 ± 0.2 pF. For LE/LY isolated from primary macrophages it was 0.8 ± 0.1 pF (n = 41), for Tf-loaded vesicles 1.3 ± 0.5 pF (n = 8). Currents were recorded using an EPC-10 patch-clamp amplifier (HEKA, Lambrecht, Germany) and PatchMaster acquisition software (HEKA). Data were digitized at 40 kHz and filtered at 2.8 kHz. Fast and slow capacitive transients were cancelled by the compensation circuit of the EPC-10 amplifier. Recording glass pipettes were polished and had a resistance of 4–8 MΩ. For all experiments, salt-agar bridges were used to connect the reference Ag-AgCl wire to the bath solution to minimize voltage offsets. Liquid junction potential was corrected. For the application of the lipids (A.G. Scientific) or small molecule agonists (ML2-SA1, ML-SA1), cytoplasmic solution was completely exchanged by cytoplasmic solution containing agonist. Unless otherwise stated, cytoplasmic solution contained 140 mM K-MSA, 5 mM KOH, 4 mM NaCl, 0.39 mM CaCl2, 1 mM EGTA and 10 mM HEPES (pH was adjusted with KOH to 7.2). Luminal solution contained 140 mM Na-MSA, 5 mM K-MSA, 2 mM Ca-MSA 2 mM, 1 mM CaCl2, 10 mM HEPES and 10 mM MES (pH was adjusted with NaOH to 7.2). For optimal conditions of TRPML1, luminal pH was adjusted to 4.6 and Na-MSA was used in the luminal solution. For optimal conditions of TRPML2, luminal pH was adjusted to 7.2 and Na-MSA was used in the luminal solution. For optimal conditions of TRPML3, luminal pH was adjusted to 7.2 and K-MSA was applied to replace Na-MSA in the luminal solution. In all experiments, 500 ms voltage ramps from −100 to +100 mV were applied every 5 s, holding potential at 0 mV. The current amplitudes at −100 mV were extracted from individual ramp current recordings. All statistical analysis was done using Origin8 software.
Calcium imaging experiments were performed using fura-2 as described previously (Grimm et al., 2012a). Briefly, HEK293 cells were plated onto glass coverslips, grown over night and transiently transfected with the respected cDNAs using TurboFect transfection reagent (Thermo Scientific). After 24–48 h cells were loaded for 1 hr with the fluorescent indicator fura2-AM (4 µM; Invitrogen) in a standard bath solution (SBS) containing (in mM) 138 NaCl, 6 KCl, 2 MgCl2, 2 CaCl2, 10 HEPES, and 5.5 D-glucose (adjusted to pH 7.4 with NaOH). Cells were washed in SBS for 30 min before measurement. Calcium imaging was performed using a monochromator-based imaging system (Polychrome IV mono-chromator, TILL Photonics).
Analysis of electron density map. The electron density maps for the cryo-electron microscopy structures of hTRPML1 and hTRPML3 in open agonist-bound form (PDB IDs: 5WJ9 and 6AYF, respectively) were downloaded from the Protein Data Bank (PDB; ww.rcsb.org) (Berman et al., 2000) and visualized in PyMOL (The PyMOL Molecular Graphics System, Version 1.7.4 Schrödinger, LLC). Homology modelling of TRPML2. The amino acid sequence of hTRPML2 was retrieved from UniProt (The UniProt Consortium, 2017); Accession number: Q8IZK6-1) and a Blast (Altschul et al., 1990) search using BLOSUM62 matrix was performed against the PDB to find the closest homologues. Subsequently, sequence alignment of hTRPML2 to the top scored template, hTRPML3 (Sequence identity 59%), was conducted in MOE2012.10 (Molecular Operating Environment (MOE), 2016.08; Chemical Computing Group Inc., 1010 Sherbooke St. West, Suite #910, Montreal, QC, Canada, H3A 2R7, 2016) and the alignment file was used to generate the homology model using MODELLER 9.11 (Webb and Sali, 2014). Ligand-bound homology models of hTRPML2 were finally built using the agonist-bound structure of hTRPML3 (PDB ID: 6AYF) and ranked according to their DOPE score (Shen and Sali, 2006). Molecular docking to hTRPML1 and −2. The ligands were prepared for docking using the LigPrep tool as implemented in Schrödinger’s software (Schrödinger Release 2017–1: LigPrep, Schrödinger, LLC, New York, NY, 2017), where the two stereoisomers of ML2-SA1 were generated and energy minimized using the OPLS force field. Conformers of the prepared ligands were calculated with ConfGen using the default settings and allowing minimization of the output conformations.
Protein preparation. The cryo-electron microscopy structure of the open conformation of hTRPML1 in complex with ML-SA1 (PDB ID: 5WJ9) and the generated hTRPML2 homology model were prepared with Schrödinger’s Protein Preparation Wizard (Schrödinger Release 2017–1: Schrödinger Suite 2017–1 Protein Preparation Wizard; Epik, Schrödinger, LLC, New York, NY, 2016; Impact, Schrödinger, LLC, New York, NY, 2016; Prime, Schrödinger, LLC, New York, NY, 2017): Hydrogen atoms were added and the H-bond network was subsequently optimized. The protonation states at pH 7.0 were predicted using the PROPKA tool in Schrödinger. The structures were finally subjected to a restrained energy minimization step using the OPLS2005 force field (RMSD of the atom displacement for terminating the minimization was 0.3 Å).
The receptor grid preparation for the docking procedure was carried out by assigning the agonist as the centroid of the grid box. The generated ligand conformers were docked into the proteins using Glide (Small-Molecule Drug Discovery Suite 2017–1: Glide, Schrödinger, LLC, New York, NY, 2017) in the Standard Precision mode. A total of 100 poses per ligand conformer were included in the post-docking minimization step and a maximum of 20 docking clusters were output for each ligand. Redocking of the ligand ML-SA1 into the hTRPML1 pocket gave a docking pose with root mean square deviation of 1.22 Å for the top-ranked solution (Figure 3—figure supplement 1A).
For preparation of primary alveolar macrophages (AMΦ), mice were deeply anesthetized and euthanized by exsanguination. Afterwards, the trachea was carefully exposed and cannulated by inserting a 20 gauge catheter (B. Braun, cat. no. 4252110B). AMΦ were harvested by eight consecutive lung lavages with 1 ml of DPBS each. After a centrifugation step, cells were immediately collected and cultured in RPMI 1640 medium supplemented with 10% fetal bovine serum and 1% antibiotics. AMΦ were directly seeded onto 12 mm glass cover slips and used for experiments within 5 days after preparation. Bone marrow-derived macrophages (BMDMΦ) were isolated from femur and tibias of mice. Thus, bones were isolated and bone marrow was flushed with 10 ml PBS using a sterile 25 gauge needle. Cells were obtained by centrifugation, resuspended and subsequently cultured in 10 cm petri dishes in RPMI 1640 medium supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin and 40 ng/mL murine M-CSF (Miltenyi Biotech). Cells were incubated for 5 days, before they were plated onto poly-L-lysine coated cover slips for experiments. All cells were maintained at 37°C in 5% CO2 atmosphere. If necessary, cells were stimulated with 1 µg/mL LPS (Escherichia coli O26:B6, Sigma, L2762) prior to experiments for different time periods as stated in the text. Animals were used under approved animal protocols and University of Munich (LMU) Institutional Animal Care Guidelines.
Cell culture supernatants from WT or TRPML2-/- BMDMΦ were collected at 4 hr or 8 hr following LPS treatment in the presence or absence of TRPML2 agonist (ML2-SA1), and CCL2 was measured using an ELISA kit (BioLegend, 432707), per the manufacturer’s instructions. Cell culture supernatants were diluted ten times for the assay, and 50 μL diluted supernatant was assessed.
RAW264.7 cells were seeded overnight with 0.1 mg/mL of lipopolysaccharide (LPS) (L4391, Sigma). Then, cells were loaded for 20 min at 37°C with transferrin from human serum, Alexa Fluor 546-conjugated (T23364, ThermoFisher) at the concentration of 50 μg/mL in complete medium (DMEM 10% FBS). The analysis of recycling kinetics was performed by chasing for 5, 10, 15 and 20 min in complete media plus 50 μg/mL of unconjugated transferrin (T0665, Sigma) in the presence of either DMSO or ML2-SA1 (30 μM). Before fixation with 4% paraformaldehyde (PFA), non-internalized transferrin was acid-stripped (150 mM NaCl, 0.5% acetic acid in H2O) for 30 s. Images were acquired using a Zeiss LSM 800 with 63x magnification.
RAW264.7 cells were seeded overnight with 0.1 mg/mL of lipopolysaccharide (LPS) (L4391, Sigma). Then, cells were treated with DMSO, calcium ionophore A23187 (C7522, Sigma) or ML2-SA1 for 3 hr. After 3 h cells were collected and stained with LAMP1 antibody (SC-19992, Santa Cruz) in PBS (1% BSA) during agitation for 20 min (4°C). Cells were then collected by centrifugation and resuspended in PBS (1% BSA) with goat anti-rat, Alexa488 (A-11006 ThermoFisher) during agitation for 1 hr (4°C). Finally, cells were washed in PBS and left on ice until FACS analysis. Cells were loaded into the FACS machine using a nozzle of 100 µm and the LAMP1 fluorescence intensity was measured using a 488 nm excitation laser and a FITCH (530/30 nm) emission filter. The threshold was set using DMSO-treated samples, and 1000 events were counted for each condition.
For measurement of lysosomal hexosaminidase enzyme release, bone marrow macrophages were treated with ML2-SA1, ML-SA1 or DMSO in serum-free RPMI medium, concentrations and durations as indicated. Ionomycin was used as control. After treatment, supernatants were collected, centrifuged and incubated with natrium citrate buffer (pH 4.5) and 4-Methylumbelliferyl N-acetyl-β-D-glucosaminide (M1233, Sigma, 1 mM final concentration) for 1.5 hr. Cells were lysed with Triton-X buffer and lysates were processed in parallel. The reaction was stopped by adding glycin buffer to the samples and the turnover of hexosaminidase substrate was detected as fluorescence (Exitation: 365 nm; Emission: 450 nm) using a plate reader (Spectramax ID3, Molecular Devices). The increase in substrate turnover was analyzed as fluorescence increase in supernatants relative to lysates.
Generation of point mutant isoforms of hTRPML2 (encoded by the MCOLN2 gene) was performed as described previously (Grimm et al., 2010) using the QuikChange protocol. The following primers were used to generate hTRPML2 mutant isoforms: A422C forward primer: CTTCGGTTTTGTTGTTGTGCTGGTATGATTTATCTGGG; A422C reverse primer: CCCAGATAAATCATACCAGCACAACAACAAAACCGAAG; A424V forward primer: CGGTTTTGTGCTTGTGTTGGTATGATTTATCTGGGTTACAC; A424V reverse primer: GTGTAACCCAGATAAATCATACCAACACAAGCACAAAACCG; G425A forward primer: CGGTTTTGTGCTTGTGCTGCTATGATTTATCTGGGTTACAC; G425A reverse primer: GTGTAACCCAGATAAATCATAGCAGCACAAGCACAAAACCG; A453S forward primer: CTGAACACAGTTTCTGAGTGTCTGTTTTCTCTGG; A453S reverse primer: CCAGAGAAAACAGACACTCAGAAACTGTGTTCAG; V460I forward primer: TGTCTGTTTTCTCTGATCAACGGTGATGACATG; V460I reverse primer: CATGTCATCACCGTTGATCAGAGAAAACAGACA; I498V forward primer: CCTTCATCAGCCTTTTTATATATATGGTTCTCAGTCTTTTTATTGC; I498V reverse primer: GCAATAAAAAGACTGAGAACCATATATATAAAAAGGCTGATGAAGG.
ML2-SA1 effects on macrophage migration were assessed by a modified Boyden chamber setup (Figure 6—figure supplement 2). In the modified Boyden chamber setup, BMDMΦ were plated onto poly-L-lysine coated cover slips in a twenty-four well plate (lower compartment) in the presence or absence of 1 µg/ml LPS for 6 hr. After 6 hr, media was replaced with media containing 10 or 30 µM ML2-SA1 or DMSO. 1 × 105 BMDMΦ were placed on top of the transwell chamber (Corning) in media without any compound. Transwell chambers were placed into the twenty-four well plate and incubated for 3 hr at 37°C in 5% CO2 atmosphere. In the classical Boyden chamber approach a twenty-four well plate was filled with media containing either DMSO, 1 µg/ml LPS and DMSO, 1 µg/ml LPS and 30 µM ML2-SA1, or 10 ng/ml CCL2. Transwell chambers were equally prepared and incubated. Migrated cells were fixed and stained with crystal violet/methanol. The top of the transwell chamber was cleaned an images were taken. Cell covered area was determined with ImageJ (NIH, Bethesda, MD).
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Michael L DustinReviewing Editor; University of Oxford, United Kingdom
Richard AldrichSenior Editor; The University of Texas at Austin, United States
In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.
Thank you for submitting your article "Selective agonist of TRPML2 reveals direct role in chemokine release from innate immune cells" for consideration by eLife. Your article has been reviewed by two peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Richard Aldrich as the Senior Editor. The following individual involved in review of your submission has agreed to reveal his identity: Antony Galione (Reviewer #2).
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
TRPML channels are expressed throughout the endolysosomal system and in some cases at the plasma membrane. Of the three isoforms of TRPMLs, TRPML2 is the least well studied. In this manuscript the authors have developed a selective TRPML2 agonist and used this in concert with macrophages from TRPML2-/- mice to demonstrate a functional role for TRPML2 channels in macrophage physiology. The generation of the agonist is an impressive tour de force and only relatively minor revisions are needed regarding the mechanism of effect on chemokine release.
1) Regarding the agonist action on the channel. Can you provide additional data on the time course of the agonist effect on TRPML2 currents? It would be ideal to have this data in a new main figure panel.
2) The Discussion should be amplified or qualified regarding the mechanism for "direct" events that lead to chemokine secretion and chemotaxis. The question is: how, precisely, does the channel play a role in secretion? You are encouraged to provide a scheme and to discuss what is clearly established and what needs further work. How do you interpret the increase in currents induced by the agonist? Is there a change in ion selectivity or open probability? A change in gating mode? Can you discuss how they think TRPML2 activation leads (directly) to secretion? Is it via cytosolic calcium? What are the effectors? Can you elaborate a bit more on possible mechanisms?
3) Regarding ML2-SA1 effects: Please elaborate a bit more on the potentiation of the LPS effects with ML2-SA1. What is known about endogenous activation of these channels? The Boyden migration assays show a small enhancement + ML2-SA1 in the bar chart. The effects look larger in the images. Please comment. Please clarify the relationship between CCL2 release and migration. The relationship appears complex and non-linear. Does ML2-SA1 affect TPCs, the other major cation channel family of the endolysosomal system? More discussion is needed concerning how CCL2 is released from cells. The authors have ruled out lysosomal exocytosis, but more discussion of alternative mechanisms (or even experiments) would be helpful and important.https://doi.org/10.7554/eLife.39720.021
1) Regarding the agonist action on the channel. Can you provide additional data on the time course of the agonist effect on TRPML2 currents? It would be ideal to have this data in a new main figure panel.
We now provide data on the time course of the ML2-SA1 effect on TRPML2 currents as requested (see new Figure 2G).
2) The Discussion should be amplified or qualified regarding the mechanism for "direct" events that lead to chemokine secretion and chemotaxis. The question is: how, precisely, does the channel play a role in secretion? You are encouraged to provide a scheme and to discuss what is clearly established and what needs further work.
We agree with the reviewers and did recognize that the Discussion section needs to be improved, in particular relevant literature in the field is now discussed in more detail alongside suggested effects of TRPML channel activation on endolysosomal trafficking, secretion, fusion/fission and endocytosis. We have added the following section to the Discussion:
“Loss of function mutations in the TRPML2-related channel TRPML1 result in lysosomal storage and endolysosomal trafficking defects underlying the neurodegenerative disease mucolipidosis type IV (Bach, 2001; Pryor et al., 2006; Chen et al., 2014). […] Possible scenarios might be: fusion of Golgi vesicles with RE, fission from RE, fusion of Golgi vesicles with EE, fission from EE, fusion of EE-derived vesicles with the RE.”
To better visualize these possibilities we now provide a new scheme (see new Figure 5F), showing in more detail what the possible release pathways are that TRPML2 might be involved in and what consequences would result from TRPML2 KO or direct activation. It is established that:
TRPML2 is functionally active in LE/LY, EE and TfR+/Rab11+ RE. In BMDMs there is less TRPML2 channel activity in LE/LY than in RE. It is unclear whether TRPML2 is expressed in Golgi-derived vesicles or in vesicles which are derived from EE or RE by fission processes. According to literature there are no secretory vesicles present in macrophages.
CCL2 release/secretion from BMDMs is increased when applying ML2-SA1 and decreased in TRPML2KO cells, suggesting that release depends, at least in part, on TRPML2 activity.
Transferrin trafficking through the EE/RE system is accelerated by ML2-SA1 while lysosomal exocytosis is not affected.
What remains speculative is the exact location of interference with the CCL2 trafficking process and where along the pathway the effect might take place: possible scenarios are, as now pointed out in the Discussion, fusion of Golgi vesicles with RE, fission from RE, fusion of Golgi vesicles with EE, fission from EE, fusion of EE derived vesicles with the RE.
How do you interpret the increase in currents induced by the agonist? Is there a change in ion selectivity or open probability? A change in gating mode?
Using the cryo-EM technique, the structure of TRPML1 with ML-SA1 has been solved recently (Schmiege et al., 2017) as well as the structure of TRPML3 with ML-SA1 (Zhou et al., 2017). Schmiege et al. demonstrated that ML-SA1 induces a conformational change of pore helix 1 and transmembrane domain 6 (TM6), effectively dilating the lower selectivity filter and lower hydrophobic gate in a manner reminiscent of capsaicin binding TRPV1. Similarly for TRPML3, Zhou et al. demonstrated ML-SA1 binding to induce a conformational change of the pore loop alongside a TM6 rotation, dilating the pore sufficiently to conduct both hydrated monovalent and divalent cations. We have shown that ML2-SA1 very likely binds in the same binding pocket as ML-SA1 (Figure 3). We therefore speculate that ML2-SA1 may also lead to a pore dilation and thus current increase.
Can you discuss how they think TRPML2 activation leads (directly) to secretion? Is it via cytosolic calcium? What are the effectors? Can you elaborate a bit more on possible mechanisms?
See also answer above (2). Calcium has been postulated to be an important trigger of intracellular vesicle fusion and fission processes as well as fusion with the plasma membrane. We have described above the different locations along the endolysosomal pathway where TRPML2 is functionally expressed (patch clamp data) and where and how it might interfere with the different trafficking steps. We hypothesize that it is the calcium release from the organelle into the cytosol which is the relevant process controlled by TRPML2. ML2-SA1 has been shown to increase cytosolic calcium in calcium imaging experiments. To demonstrate direct calcium release via TRPML2 we now provide endolysosomal patch-clamp experiments assessing the calcium permeability of TRPML2 (see new Figure 2—figure supplement 1C).
3) Regarding ML2-SA1 effects: Please elaborate a bit more on the potentiation of the LPS effects with ML2-SA1.
LPS promotes CCL2 secretion as shown in Figure 5A. The effect on CCL2 release mediated by applying LPS alone is partially dependent on TRPML2 as TRPML2KO leads to a significant reduction in CCL2 release. LPS upregulates the expression of TRPML2 (Sun et al., 2015), hence more TRPML2 would become active (stimulated by endogenous ligands present) or available for activation (e.g. through ML2-SA1) after upregulation of TRPML2 expression. It is also likely that ML2-SA1 increases open-probability and/or pore dilation, thus further increasing TRPML2 activity.
What is known about endogenous activation of these channels?
Generally, TRPML channels can be activated by their putative endogenous ligand PI(3,5)P2. PI(3,5)P2 occurs in LE/LY membranes and, to a lesser extent, in early endosomal membranes. Furthermore, TRPML2 is pH regulated (Figure 2—figure supplement 1; low pH decreases channel activity). How and under which circumstances endogenous ligand(s) become available for TRPML2 activation remains unclear.
The Boyden migration assays show a small enhancement + ML2-SA1 in the bar chart. The effects look larger in the images. Please comment.
We reassessed our statistical analysis. We confirm the validity of the statistical analysis but do now present a selection of alternative images, better reflecting the results of the statistical analysis (see new Figure 6 and new Figure 6—figure supplement 2).
Please clarify the relationship between CCL2 release and migration. The relationship appears complex and non-linear.
When we incubate BMDM with CCL2 (10 ng/ml), we see an increase in migration from 20% (control) to 40% (Figure 6—figure supplement 2). When we incubate with ML2-SA1, we see an increase in migration from 20/25% (control) to 35%. We only see an effect in LPS treated cells. In fact, -LPS BMDM migrate in the direction of +LPS BMDM when both are exposed to ML2-SA1 (see scheme, Figure 6—figure supplement 1). We have not performed multiple dose-response measurements. However, the experiments as presented show a clear effect of ML2-SA1 on cell migration, allowing us to conclude that ML2-SA1 can mimic the effect of CCL2.
Does ML2-SA1 affect TPCs, the other major cation channel family of the endolysosomal system?
We now provide endolysosomal patch-clamp data showing that ML2-SA1 has no effect on TPC1 and TPC2 (see new Figure 2—figure supplement 3C-F).
More discussion is needed concerning how CCL2 is released from cells. The authors have ruled out lysosomal exocytosis, but more discussion of alternative mechanisms (or even experiments) would be helpful and important.
We now discuss in more detail possible secretion routes of CCL2 from macrophages. See also answer above (2).https://doi.org/10.7554/eLife.39720.022
- Christian Grimm
- Christian Grimm
- Christian Wahl-Schott
- Martin Biel
- Lesca M Holdt
- Daniel Teupser
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We thank Lars Allmendinger, Yu-Kai Chao, Berit Noack, Martina Stadler, Wolfgang Wilfert, and Christopher Wolf for technical support. This work was supported, in part, by funding of the German Research Foundation (SFB/TRR152 projects P04 to CG, P06 to CW-S., and P12 to MB. as well as SFB1123 project B1 to LMH and DT), the NCL (Neuronal Ceroid Lipofuscinosis) Foundation Award 2016 to CG, and the University of Pennsylvania Orphan Disease Center and the Mucolipidosis IV Foundation Grant MDBR-17–120 ML4 to CG.
Animal experimentation: This study was performed where applicable in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. This study was performed where applicable in strict accordance with the recommendations of the Bavarian Government (ROB; AZ_55.2-1-54-2532-27-2015).
- Richard Aldrich, The University of Texas at Austin, United States
- Michael L Dustin, University of Oxford, United Kingdom
- Received: July 1, 2018
- Accepted: October 26, 2018
- Version of Record published: November 27, 2018 (version 1)
© 2018, Plesch et al.
This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.
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Heterotrimeric G-proteins are signal transducers involved in mediating the action of many natural extracellular stimuli and many therapeutic agents. Non-invasive approaches to manipulate the activity of G-proteins with high precision are crucial to understand their regulation in space and time. Here, we developed LOV2GIVe, an engineered modular protein that allows the activation of heterotrimeric G-proteins with blue light. This optogenetic construct relies on a versatile design that differs from tools previously developed for similar purposes, that is metazoan opsins, which are light-activated G-protein-coupled receptors (GPCRs). Instead, LOV2GIVe consists of the fusion of a G-protein activating peptide derived from a non-GPCR regulator of G-proteins to a small plant protein domain, such that light uncages the G-protein activating module. Targeting LOV2GIVe to cell membranes allowed for light-dependent activation of Gi proteins in different experimental systems. In summary, LOV2GIVe expands the armamentarium and versatility of tools available to manipulate heterotrimeric G-protein activity.
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