Lysosomes are organelles responsible for the breakdown and recycling of cellular machinery. Dysfunctional lysosomes give rise to lysosomal storage disorders as well as common neurodegenerative diseases. Here, we use a DNA-based, fluorescent chloride reporter to measure lysosomal chloride in Caenorhabditis elegans as well as murine and human cell culture models of lysosomal diseases. We find that the lysosome is highly enriched in chloride, and that chloride reduction correlates directly with a loss in the degradative function of the lysosome. In nematodes and mammalian cell culture models of diverse lysosomal disorders, where previously only lysosomal pH dysregulation has been described, massive reduction of lumenal chloride is observed that is ~103 fold greater than the accompanying pH change. Reducing chloride within the lysosome impacts Ca2+ release from the lysosome and impedes the activity of specific lysosomal enzymes indicating a broader role for chloride in lysosomal function.https://doi.org/10.7554/eLife.28862.001
In cells, worn out proteins and other unnecessary materials are sent to small compartments called lysosomes to be broken down and recycled. Lysosomes contain many different proteins including some that break down waste material into recyclable fragments and others that transport the fragments out of the lysosome. If any of these proteins do not work, waste products build up and cause disease. There are around 70 such lysosomal storage diseases, each arising from a different lysosomal protein not working correctly.
A recently developed “nanodevice” called Clensor can measure the levels of chloride ions inside cells. Clensor is constructed from DNA, and its fluorescence changes when it detects chloride ions. Although chloride ions have many biological roles, chloride ion levels had not been measured inside a living organism. Now, Chakraborty et al. – including some of the researchers who developed Clensor – have used this nanodevice to examine chloride ion levels in the lysosomes of the roundworm Caenorhabditis elegans. This revealed that the lysosomes contain high levels of chloride ions. Furthermore, reducing the amount of chloride in the lysosomes made them worse at breaking down waste.
Do lysosomes affected by lysosome storage diseases also contain low levels of chloride ions? To find out, Chakraborty et al. used Clensor to study C. elegans worms and mouse and human cells whose lysosomes accumulate waste products. In all these cases, the levels of chloride in the diseased lysosomes were much lower than normal. This had a number of effects on how the lysosomes worked, such as reducing the activity of key lysosomal proteins.
Chakraborty et al. also found that Clensor can be used to distinguish between different lysosomal storage diseases. This means that in the future, Clensor (or similar methods that directly measure chloride ion levels in lysosomes) may be useful not just for research purposes. They may also be valuable for diagnosing lysosomal storage diseases early in infancy that, if left undiagnosed, are fatal.https://doi.org/10.7554/eLife.28862.002
Chloride is the most abundant, soluble anion in the body. Cytosolic chloride can be as low as ~45 mM, while extracellular chloride is ~110 mM (Treharne et al., 2006), (Sonawane et al., 2002). Chloride concentration values thus span a wide range and yet, in each compartment, it is quite tightly regulated (Sonawane and Verkman, 2003). For example, in early endosomes it is ~40 mM, late endosomes it is ~70 mM and lysosomes it is ~108 mM (Hara-Chikuma et al., 2005; Saha et al., 2015; Sonawane et al., 2002). Chloride levels are stringently regulated by chloride channels such as cystic fibrosis transmembrane regulator (CFTR), the CLC family of channels or calcium activated chloride channels, and their dysregulation is directly linked to several diseases including cystic fibrosis, myotonia, epilepsy, hyperekplexia or deafness (Planells-Cases and Jentsch, 2009). Chloride is largely considered to function as a counter ion only to balance changes in cation fluxes related to signaling (Scott and Gruenberg, 2011). In one form, this balancing function serves to reset the membrane potential of depolarized neurons through the operation of plasma membrane resident chloride channels/exchangers (Chen, 2005). In another form, it serves to continuously facilitate organelle acidification, through the operation of intracellular chloride channels (Stauber and Jentsch, 2013). Despite its importance in cell function, intracellular chloride has never been visualized or quantitated in vivo.
DNA nanotechnology has offered creative, functional imaging solutions to quantitate second messengers as well as image organelles in real time in living cells and in genetic model organisms (Bhatia et al., 2016; Chakraborty et al., 2016; Krishnan and Bathe, 2012; Surana et al., 2015). Here, using a previously developed, pH-independent, DNA-based fluorescent chloride reporter called Clensor, we have made the first measure of chloride in a live multicellular organism, creating in vivo chloride maps of lysosomes in C. elegans.
Our investigations reveal that lysosomal chloride levels in vivo are even higher than extracellular chloride levels. Others and we have shown that lysosomes have the highest lumenal acidity and the highest lumenal chloride , among all endocytic organelles (Saha et al., 2015; Weinert et al., 2010). Although lumenal acidity has been shown to be critical to the degradative function of the lysosome (Appelqvist et al., 2013; Eskelinen et al., 2003), the necessity for such high lysosomal chloride is unknown. In fact, in many lysosomal storage disorders, lumenal hypoacidification compromises the degradative function of the lysosome leading to the toxic build-up of cellular cargo targeted to the lysosome for removal, resulting in lethality (Guha et al., 2014). Lysosomal storage disorders (LSDs) are a diverse collection of ~70 different rare, genetic diseases that arise due to dysfunctional lysosomes (Samie and Xu, 2014). Dysfunction in turn arises from mutations that compromise protein transport into the lysosome, the function of lysosomal enzymes, or lysosomal membrane integrity (Futerman and van Meer, 2004). Importantly, for a sub-set of lysosomal disorders like osteopetrosis or neuronal ceroid lipofuscinoses (NCL), lysosomal hypoacidification is not observed (Kasper et al., 2005). Both these conditions result from a loss of function of the lysosomal H+-Cl- exchange transporter CLC-7 (Kasper et al., 2005). In both mice and flies, lysosomal pH is normal, yet both mice and flies were badly affected (Poët et al., 2006; Weinert et al., 2010).
The lysosome performs multiple functions due to its highly fusogenic nature. It fuses with the plasma membrane to bring about plasma membrane repair as well as lysosomal exocytosis, it fuses with the autophagosome to bring about autophagy, it is involved in nutrient sensing and it fuses with endocytic cargo to bring about cargo degradation (Appelqvist et al., 2013; Xu and Ren, 2015). To understand which, if any, of these functions is affected by chloride dysregulation, we chose to study genes related to osteopetrosis in the versatile genetic model organism Caenorhabditis elegans. By leveraging the DNA scaffold of Clensor as a natural substrate along with its ability to quantitate chloride, we could simultaneously probe the degradative capacity of the lysosome in vivo and then in cultured mammalian cells. Our findings reveal that depleting lysosomal chloride showed a direct correlation with loss of the degradative function of the lysosome. We found that lowering lysosomal chloride also reduced the level of Ca2+ released from the lysosome. We also observed that reduction of lysosomal chloride inhibited the activity of specific lysosomal enzymes such as cathepsin C and arylsulfatase B. The role of chloride in defective lysosomal degradation has been hypothesized in the past (Stauber and Jentsch, 2013; Wartosch and Stauber, 2010; Wartosch et al., 2009), and our studies provide the first mechanistic proof of a broader role for chloride in lysosome function.
In this study we use two DNA nanodevices, called the I-switch and Clensor, to fluorescently quantitate pH and chloride respectively (Modi et al., 2009; Saha et al., 2015). The I-switch is composed of two DNA oligonucleotides. One of these can form an i-motif, which is an unusual DNA structure formed by protonated cytosines (Gehring et al., 1993). In the I-switch, intrastrand i-motif formation is used to bring about a pH-dependent conformational change, that leverages fluorescence resonance energy transfer (FRET) to create a ratiometric fluorescent pH reporter. (Figure 1—figure supplement 2)
The DNA-based chloride sensor, Clensor, is composed of three modules: a sensing module, a normalizing module and a targeting module (Figure 1a) (Saha et al., 2015; Prakash et al., 2016). The sensing module is a 12 base long peptide nucleic acid (PNA) oligomer conjugated to a fluorescent, chloride-sensitive molecule 10,10′-Bis[3-carboxypropyl]−9,9′-biacridinium dinitrate (BAC), (Figure 1a) (Sonawane et al., 2002). The normalizing module is a 38 nt DNA sequence bearing an Alexa 647 fluorophore that is insensitive to Clˉ. The targeting module is a 26 nt double stranded DNA domain that targets it to the lysosome via the endolysosomal pathway by engaging the scavenger receptor or ALBR pathway. In physiological environments, BAC specifically undergoes collisional quenching by Clˉ, thus lowering its fluorescence intensity (G) linearly with increasing Clˉ concentrations. In contrast, the fluorescence intensity of Alexa 647 (R) remains constant (Figure 1b). This results in R/G ratios of Clensor emission intensities varying linearly with [Clˉ] over the entire physiological regime of [Clˉ]. Since the response of Clensor is insensitive to pH changes, it enables the quantitation of lumenal chloride in organelles of living cells regardless of their lumenal pH (Saha et al., 2015).
Coelomocytes of C. elegans are known to endocytose foreign substances injected in the body cavity (Fares and Greenwald, 2001). The polyanionic phosphate backbone of DNA can be co-opted to target it to scavenger receptors and thereby label organelles on the endolysosomal pathway in tissue macrophages and coelomocytes in C. elegans (Figure 1c and d) (Bhatia et al., 2011; Modi et al., 2009; Saha et al., 2015; Surana et al., 2011). Alexa 647 labelled I-switch (I4cLY) and Clensor were each injected in the pseudocoelom of 1-day-old adult worms expressing pmyo-3::ssGFP. In these worms, soluble GFP synthesized in muscles and secreted into the pseudocoelom is actively internalized by the coelomocytes resulting in GFP labeling of the coelomocytes (Fares and Greenwald, 2001). After 1 hr, both devices quantitatively colocalize with GFP indicating that they specifically mark endosomes in coelomocytes (Figure 1e and Figure 1—figure supplement 1c). Endocytic uptake of DNA nanodevices was performed in the presence of 30 equivalents of maleylated bovine serum albumin (mBSA), a well-known competitor for the anionic ligand binding receptor (ALBR) pathway (Gough and Gordon, 2000). Coelomocyte labeling by I4cLYor Clensor were both efficiently competed out by mBSA indicating that both reporters were internalized by ALBRs and trafficked along the endolysosomal pathway (Figure 1—figure supplement 1b) (Surana et al., 2011).
Next, the functionality of I4cLY and Clensor were assessed in vivo. To generate an in vivo calibration curve for the I-switch I4cLY, coelomocytes labeled with I4cLY were clamped at various pH values between pH 4 and 7.5 as described previously and in the supporting information (Surana et al., 2011). This indicated that, as expected, the I-switch showed in vitro and in vivo performance characteristics that were extremely well matched (Figure 1—figure supplement 2b–e). To assess the in vivo functionality of Clensor, a standard Clˉ calibration profile was generated by clamping the lumenal [Clˉ] to that of an externally added buffer containing known [Clˉ] as described previously for cultured cells (Saha et al., 2015). Endosomes of coelomocytes were labeled with Clensor and fluorescence images were acquired in the BAC channel (G) as well as Alexa 647 channel (R) as described (see Materials and methods), from which were obtained R/G ratios of every endosome clamped at a specific [Clˉ] (Figure 1b). Endosomal R/G ratios showed a linear dependence on [Clˉ] with ~2 fold change in R/G values from 5 mM to 80 mM [Clˉ] (Figure 1f and g). This is very well matched with its in vitro fold change in R/G over the same regime of [Clˉ].
Before performing quantitative chloride imaging in various mutant nematodes, we checked whether lysosomal targeting of Clensor and the I-switch were preserved in a variety of genetic backgrounds of our interest. Clensor was injected into LMP-1::GFP worms treated with RNAi against specific lysosomal storage disorder (LSD)-related genes or genes linked to osteopetrosis. We observed significant colocalization (>74%) of Clensor with LMP-1-GFP labeled lysosomes in these coelomocytes (Figure 3—figure supplement 2b and c). Given that both Clensor and the I-switch robustly labeled lysosomes of coelomocytes in wild type worms (N2), mutants and RNAi knockdowns of a range of LSD-related genes, we explored whether these devices could report on alterations, if any, in the lumenal ionicity in these lysosomes, and thereby possibly report on lysosome dysfunction.
As an initial study, we focused on C. elegans nematodes in which genes related to osteopetrosis are mutated. Osteopetrosis results from non-functional osteoclasts that lead to increased bone mass and density due to a failure in bone resorption (Sobacchi et al., 2013). In humans, osteopetrosis results from mutations in a lysosomal chloride-proton antiporter CLCN7, and its auxiliary factor OSTM1 (Figure 2a) (Kornak et al., 2001; Lange et al., 2006). It also results from mutations in TCIRG1, which is the a3 subunit of a lysosomal V-ATPase (Kornak et al., 2000) and SNX-10, a sorting nexin implicated in lysosome transport to form the ruffled border of osteoclasts, which is critical for osteoclast function (Aker et al., 2012) (Figure 2a). Lysosomes of CLC7 knockout mice show normal lumenal pH, yet the mice manifest osteopetrosis as well as neurodegeneration, indicating that despite the apparently normal lumenal milieu, the organelle is still dysfunctional (Kasper et al., 2005). The C. elegans homologs for these genes are clh-6 (CLCN7), F42A8.3 (ostm-1; OSTM1), unc-32 (TCIRG1) and snx-3 (SNX10) (Figure 2a).
Clensor was injected into N2, clh-6 and unc-32 mutants and RNAi knockdowns of ostm-1 and snx-3. Chloride concentrations in the lysosomes of each genetic background at 60 min post injection were obtained (Figure 2b,c and Figure 2—figure supplement 1). N2 worms showed a chloride concentration of ~75 mM. Knocking down clh-6 and ostm-1 resulted in a dramatic decrease of lysosomal chloride to ~45 mM due to the loss of function in chloride transport. Lumenal pH in the lysosomes of these mutants was normal, consistent with findings in both flies and mice (Saha et al., 2015; Weinert et al., 2010). As a control, knocking down a plasma membrane resident CLC channel such as clh-4 showed no effect on either lysosomal chloride or pH (Schriever et al., 1999). unc-32c is a non-functional mutant of the V-ATPase a sub-unit, while unc-32f is a hypomorph (Pujol et al., 2001). Interestingly, a clear inverse correlation with unc-32 functionality was obtained when comparing their lysosomal chloride levels i.e.,~55 mM and ~65 mM for unc-32c and unc-32f respectively. Importantly, snx-3 knockdowns showed lysosomal chloride levels that mirrored those of wild type lysosomes. In all genetic backgrounds, we observed that lysosomal chloride concentrations showed no correlation with lysosome morphology (Figure 3—figure supplement 1d).
Dead and necrotic bone cells release their endogenous chromatin extracellularly - thus duplex DNA constitutes cellular debris and is physiologically relevant cargo for degradation in the lysosome of phagocytic cells (Elmore, 2007; Luo and Loison, 2008). Coelomocytes are phagocytic cells of C. elegans, and thus, the half-life of Clensor or I4cLY in these cells constitutes a direct measure of the degradative capacity of the lysosome (Tahseen, 2009). We used a previously established assay to measure the half-life of I-switches in lysosomes (Surana et al., 2013). Worms were injected with 500 nM I4cLY and the fluorescence intensity obtained in 10 cells at each indicated time point was quantitated as a function of time. The I-switch I4cLY had a half-life of ~6 hr in normal lysosomes, which nearly doubled when either clh-6 or ostm-1 were knocked down (Figure 2d and Figure 2—figure supplement 2). Both unc-32c and unc-32f mutants showed near-normal lysosome degradation capacity, inversely correlated with their lysosomal chloride values (Figure 2d and Figure 2—figure supplement 2).
In this context, data from snx-3 and unc-32f mutants support that high lysosomal chloride is critical to the degradation function of the lysosome. In humans, SNX10 is thought to be responsible for the vesicular sorting of V-ATPase from the Golgi or for its targeting to the ruffled border (Aker et al., 2012). Non-functional SNX10 can thus be considered a 'secondary V-ATPase deficiency', phenocopying a V-ATPase deficiency and showing osteoclasts without ruffled borders due to defective lysosomal transport (Aker et al., 2012). Importantly, lysosomal pH in snx-3 knockdowns was compromised by 0.3 pH units, while that in unc-32 knockdowns was compromised by 0.2 pH units (Figure 2—figure supplement 1) (Chen et al., 2012). Yet both these genetic backgrounds showed completely normal lysosomal degradation capacity, that is consistent with their normal lumenal chloride levels, rather than their defective pH levels. This further supports that high lysosomal chloride is a sensitive correlate of the degradative function of the lysosome.
Since lysosomal chloride dysregulation correlated with a loss of degradative ability of the lysosome, we wondered whether the converse was true, i.e., whether lysosomes known to be defective in degradation as seen in lysosomal storage disorders, showed depleted chloride levels. Given that in higher organisms such as mice and humans, high acidity has also been shown to be essential for proper lysosome function (Mindell, 2012), we measured both lysosomal pH and lysosomal chloride in C. elegans mutants and RNAi knockdowns for a range of genes that are known to cause lysosomal storage disorders. These included a selection of diseases due to dysfunctional enzymes that metabolize sugar derivatives, such as mannose and glycosaminoglycans, as well as lipids such as sphingomyelin and glucosylceramide. Lysosomal pH and chloride measurements were made with I4cLY and Clensor respectively, in each genetic background at 60 min post injection (Figure 3a and b). We found that in C. elegans mutants for Gaucher’s disease, Batten disease, different forms of NCL, MPS VI and Niemann Pick A/B disease, lysosomal chloride levels were severely compromised (Figure 3a and b). Dysfunctional lysosomes showed three types of ion profiles, those where either lysosomal acidity or chloride levels were reduced, and those where both lysosomal acidity and chloride were reduced. The magnitude of proton dysregulation in these defective lysosomes ranged between 1.9–2.8 µM. However, the magnitude of lysosomal chloride showed a stark drop, decreasing by 19–34 mM in most mutants. Importantly, in mammalian cell culture models for many of these diseases example for Gaucher's disease, NCL, MPS VI, etc., only pH dysregulation has been reported (Bach et al., 1999; Holopainen et al., 2001; Sillence, 2013). Yet we find that in C. elegans models of these diseases that chloride levels are highly compromised. Chloride decreases by nearly three orders of magnitude more than proton decrease, and the percentage changes of both ions are similar.
To check whether such chloride decrease is observed also in higher organisms, we made pH and chloride measurements in mammalian cell culture models of two relatively common lysosomal storage disorders. Macrophages are a convenient cell culture system to study lysosomal storage disorders as they can be isolated from blood samples and have a lifetime of 3 weeks in culture (Vincent et al., 1992). We re-created two widely used murine and human cell culture models of Gaucher’s disease by inhibiting β-glucosidase with its well-known inhibitor conduritol β epoxide (CBE) in murine and human macrophages namely, J774A.1 and THP-1 cells respectively (Hein et al., 2013, 2007; Schueler et al., 2004). We also recreated common mammalian cell culture models of Niemann-Pick A/B disease by inhibiting acid sphinogomyelinase (SMPD1) in J774A.1 and THP-1 cells with a widely used inhibitor amitriptyline hydrochloride (AH) (Aldo et al., 2013; Jones et al., 2008). First we confirmed that Clensor and our DNA-based pH reporter localized exclusively in lysosomes. In both cell lines, DNA nanodevices (500 nM) were uptaken from the extracellular milieu by the scavenger receptors, followed the endolysosomal pathway and showed quantitative colocalization with lysosomes that were pre-labelled with TMR-Dextran (Figure 4—figure supplement 3a and b). In-cell calibration curves of both pH (Figure 4—figure supplement 1) and chloride reporters (Figure 4a) were well matched with their in vitro calibration profiles, indicating that both sensor integrity and performance were quantitatively preserved at the time of making lysosomal pH and chloride measurements in these cells. Both human and murine lysosomes in normal macrophages showed chloride concentrations close to ~118 mM, revealing that lysosomes have the highest chloride levels compared to any other endocytic organelle (Saha et al., 2015; Sonawane et al., 2002). This is nearly 10–15% higher than even extracellular chloride concentrations, which reaches only up to 105–110 mM (Arosio and Ratto, 2014).
Treating J774A.1 cells and THP-1 cells with a global chloride ion channel blocker, such as NPPB (5-Nitro-2-(3-phenylpropylamino) benzoic acid), lowered lysosomal chloride concentrations to 104 and 106 mM respectively, indicating that Clensor was capable of measuring pharmacologically induced lysosomal chloride changes, if any, in these cells. In Gaucher’s cell culture models, murine and human cells showed a substantial decrease in lysosomal chloride to ~101 mM and ~92 mM respectively. This is a drop of 15–25 mM (13‒21% change) chloride, as compared to a drop of ~10 µM in lysosomal proton concentrations. In Niemann-Pick A/B cell culture models, murine and human macrophages showed an even more dramatic decrease in lysosomal chloride to ~77 mM and ~86 mM respectively. This is also a substantial decrease of 30–40 mM (25‒34% change) chloride, as compared to a drop of ~9 µM in lysosomal proton concentrations. On average in these four cell culture models, we find that the magnitude of chloride concentration decrease is at least 3 orders of magnitude greater than proton decrease, indicating that lysosome dysfunction is easily and sensitively reflected in its lumenal chloride concentrations. A Niemann Pick C cell culture model using the inhibitor U18666A recapitulated our findings in nematode models, where only lysosomal pH, but not Cl-, was altered (Figure 4—figure supplement 5)
The ClC family protein CLC-7 is expressed mainly in the late endosomes and lysosomes (Graves et al., 2008; Jentsch, 2007). The loss of either ClC-7 or its β-subunit Ostm1 does not affect lysosomal pH in any way, yet leads to osteopetrosis, resulting in increased bone mass, and severe degeneration of the brain and retina (Lange et al., 2006). Along with our studies in nematodes, this reveals a role for high chloride in lysosome function that is beyond that of a mere counter-ion in the lysosome. We therefore probed whether it could indirectly affect lysosomal function by affecting lysosomal Ca2+ (Luzio et al., 2007; Rodr?guez et al., 1997; Shen et al., 2012). We also considered the possibility that lysosomal chloride could exert a direct effect, where its reduction could impede the function of lysosomal enzymes thus affecting its degradative capacity (Baccino et al., 1975; Cigic and Pain, 1999; Maurus et al., 2005; Wartosch and Stauber, 2010) (Figure 5a).
Lysosomes are also intracellular Ca2+ stores with free Ca2+ ranging between ~400–600 µM (Christensen et al., 2002; Lloyd-Evans et al., 2008). The principal Ca2+ channel responsible for lysosomal Ca2+ release is Mucolipin TRP channel 1 (TRPML1). We therefore sought to estimate lysosomal Ca2+ by measuring Ca2+ that is released from the lysosome using two different triggers under conditions of normal and reduced lysosomal Cl-. Glycyl-L-phenylalanine 2-naphthylamide (GPN) is a substrate for Cathepsin C, which when added to cells, gets cleaved in the lysosome to release naphthylamine that is known to compromise the integrity of the lysosomal membrane, leading to a leakage of ions such as Ca2+ into the cytosol (Berg et al., 1994; Jadot et al., 1984; Morgan et al., 2011). This has been used to induce lysosomal Ca2+ release.
The cytosol of J774A.1 cells are labeled with ~3 µM Fura2-AM to ratiometrically image cytosolic Ca2+ elevation upon its release, if at all, from the lysosome. After addition of 400 µM GPN, cells were continuously imaged ratiometrically over 15–20 mins. Shortly after GPN addition, a burst of Ca2+ was observed in the cytosol, corresponding to released lysosomal Ca2+ (Figure 5b). When the same procedure was performed on cells that had been incubated with 50 µM NPPB that reduces lysosomal Cl-, the amount of lysosomal Ca2+ released was significantly reduced (Figure 5b–d) We then performed a second, more targeted way to release lysosomal Ca2+ into the cytosol, by using 20 µM ML-SA1 which specifically binds to and opens the TRPML1 channel on lysosomes (Shen et al., 2012). We found that when lysosomal Cl- was reduced with NPPB, lysosomal Ca2+ release into the cytosol was near negligible (Figure 5c–d). Taken together this indicates that high lysosomal Cl- is necessary for effective lysosomal Ca2+ release, possibly by affect lysosomal Ca2+ accumulation.
We next investigated whether reducing lysosomal chloride directly impacted the activity of any lysosomal enzymes. In vitro enzymology of Cathepsin C, a lysosome-resident serine protease has revealed that increasing Cl- increased its enzymatic activity (Cigic and Pain, 1999; McDonald et al., 1966). Further, the crystal structure of Cathepsin C shows bound chloride ions close to the active site (Cigic and Pain, 1999; Turk et al., 2012). We therefore used GPN cleavage to probe Cathepsin C activity in the lysosome upon reducing Cl- with NPPB. GPN cleavage by Cathepsin C releases naphthylamine which compromises lysosomal membrane integrity leading to proton leakage from the lysosome into the cytosol. This hypoacidifies the lysosomes resulting in reduced LysoTracker labeling as the labeling efficiency of the latter is directly proportional to compartment acidity.
Lysosomes are pre-labeled with TMR-Dextran, and LysoTracker intensities are normalized to the fluorescence intensity of TMR-Dextran, given as G/R. Hypoacidifying lysosomes by addition of 1 mM NH4Cl indeed reduced LysoTracker labeling, as expected (Figure 5e–f). A similar effect was also obtained upon GPN addition. The presence or absence of NPPB showed no change in LysoTracker labeling in cells (Figure 5e–f), indicating that NPPB by itself caused no alteration in lysosomal pH. However, when GPN was added to NPPB treated cells LysoTracker staining was remarkably well preserved (Figure 5e and f) indicating preservation of lysosomal membrane integrity because GPN was no longer effectively cleaved by Cathepsin C when lysosomal Cl- was reduced. Unlike other cathepsins, Cathepsin C does not undergo autoactivation but requires processing by Cathepsin L and Cathepsin S to convert it into active Cathepsin C (Dahl et al., 2001). We measured the activity of the upstream cathepsins such as Cathepsin L using fluorogenic substrates in the presence and absence of NPPB (Figure 5g, Figure 5—figure supplement 1). We observed no effect of chloride levels on Cathepsin L activity. This indicates that low Cathepsin C activity is not due to decreased amounts of mature Cathepsin C in the lysosome, but rather, reduced activity of mature Cathepsin C (Figure 5g, Figure 5—figure supplement 1).
Based on reports suggesting that arylsulfatase B activity was also affected by low chloride (Wojczyk, 1986), we similarly investigated a fluorogenic substrate for arylsulfatase and found that NPPB treatment impeded arylsulfatase cleavage in the lysosome. Taken together, these results suggest that high lysosomal chloride is integral to the activity of key lysosomal enzymes and that reducing lysosomal chloride affects their function.
The lysosome is the most acidic organelle within the cell. This likely confers on it a unique ionic microenvironment, reinforced by its high lumenal chloride, that is critical to its function (Xu and Ren, 2015). Using a DNA-based, fluorescent reporter called Clensor we have been able to create quantitative, spatial maps of chloride in vivo and measured lysosomal chloride. We show that, in C. elegans, lysosomes are highly enriched in chloride and that when lysosomal chloride is depleted, the degradative function of the lysosome is compromised. Intrigued by this finding, we explored the converse: whether lysosomes that had lost their degradative function – as seen in lysosomal storage disorders - showed lower lumenal chloride concentrations. In a host of C. elegans models for various lysosomal storage disorders, we found that this was indeed the case. In fact, the magnitude of change in chloride concentrations far outstrips the change in proton concentrations by at least three orders of magnitude.
To see whether chloride dysregulation correlated with lysosome dysfunction more broadly, we studied murine and human cell culture models of Gaucher’s disease, Niemann-Pick A/B disease and Niemann Pick C. We found that in mammalian cells too, lysosomes are particularly rich in chloride, surpassing even extracellular chloride levels. Importantly, chloride values in all the mammalian cell culture models revealed magnitudes of chloride dysregulation that were similar to that observed in C. elegans.
Our findings suggest more widespread and as yet unknown roles for the single most abundant, soluble physiological anion in regulating lysosome function. Decrease in lysosomal chloride impedes the release of calcium from the lysosome implicating an interplay between these two ions in the lysosome. It is also possible that chloride accumulation could facilitate lysosomal calcium enrichment through the coupled action of multiple ion channels. The ability to quantitate lysosomal chloride enables investigations into the broader mechanistic roles of chloride ions in regulating multiple functions performed by the lysosome. As such, given that chloride dysregulation shows a much higher dynamic range than hypoacidification, quantitative chloride imaging can provide a much more sensitive measure of lysosome dysfunction in model organisms as well as in cultured cells derived from blood samples that can be used in disease diagnoses and screening applications.
All fluorescently labeled oligonucleotides were HPLC-purified and obtained from IBA-GmBh (Germany) and IDT (Coralville, IA, USA). Unlabeled oligonucleotides were purchased from IDT (Coralville, IA, USA). The peptide nucleic acids (PNA) oligomer, P was synthesized using standard solid phase Fmoc chemistry on Nova Syn TGA resin (Novabiochem, Germany) using analytical grade reagents (Applied Biosystems, USA), purified by reverse phase HPLC (Shimadzu, Japan) as previously reported and stored at −20°C until further use (Prakash et al., 2016).
Bovine serum albumin (66 kDalton), nigericin, valinomycin, monensin, chloride ionophore I, Isopropyl β-D-1-thiogalactopyranoside (IPTG), amitriptyline hydrochloride, 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) and conduritol β epoxide (CBE) were obtained from Sigma (USA). LysoTracker Deep Red, TMR-Dextran (10 kDa) and Oregon Green 488 maleimide was obtained from Molecular Probes, Invitrogen (USA). Lysosomal enzyme kits namely lysosomal sulfatase assay kit was purchased from Marker Gene (USA); Magic Red Cathepsin L assay kit from Immunochemistry Technologies. Gly-Phe β-naphthylamide was purchased from Santa Cruz Biotechnology (USA). All other reagents were purchased from Sigma-Aldrich (USA) unless otherwise specified. BSA was maleylated according to a previously published protocol (Haberland and Fogelman, 1985). Trizol was purchased from Invitrogen (U.S.A.).
All oligonucleotides were ethanol precipitated and quantified by their UV absorbance. For I-switch (I4cLYA488/A647) sample preparation, 5 μM of I4 and I4′ were mixed in equimolar ratios in 20 mM potassium phosphate buffer, pH 5.5 containing 100 mM KCl. The resulting solution was heated to 90°C for 5 min, cooled to the room temperature at 5°C/15 mins and equilibrated at 4°C overnight. Samples were diluted and used within 7 days of annealing. A sample of Clensor was similarly prepared using HPLC purified oligonucleotides and PNA oligomer at a final concentration of 10 μM by mixing D1, D2 and P (see Table S1 for sequence information) in equimolar ratios in 10 mM sodium phosphate buffer, pH 7.2 and annealed as described above. For ImLy, Oregon Green maleimide was first conjugated to the thiol labeled oligonucleotide (Hermanson, 2008). Briefly, to 10 µM thiol labelled oligonucleotide in HEPES pH 7.4, 500 µM of TCEP (tris-carboxyethylphosphine) was added to reduce the disulfide bonds. After 1 hr at room temperature, 50 µM Oregon Green Maleimide was added and the reaction was kept overnight at room temperature. The reaction mixture was purified using an Amicon cutoff membrane filter (3 kDa, Millipore) to remove unreacted dye (Figure 4—figure supplement 1). A sample of ImLy was similarly prepared using HPLC purified oligonucleotides at a final concentration of 5 μM by mixing ImLY OG and ImLYAT647 (see Table S1 for sequence information) in equimolar ratios in 10 mM sodium phosphate buffer, pH 7.2 and annealed as described above. Prior to use, all buffer stock solutions were filtered using 0.22 μm disk filters (Millipore, Germany).
Standard methods were followed for the maintenance of C. elegans. Wild type strain used was the C. elegans isolate from Bristol, strain N2 (Brenner, 1974). Strains used in the study, provided by the Caenorhabditis Genetics Center (CGC), are RRID:WB-STRAIN:RB920 clh-6(ok791), RRID:WB-STRAIN:FF451 unc-32(f131), RRID:WB-STRAIN:CB189 unc-32(e189), RRID:WB-STRAIN:MT7531 ppk-3(n2835), RRID:WB-STRAIN:VC3135 gba-3(gk3287), RRID:WB-STRAIN:VC183 ppt-1(gk139), and RRID:WB-STRAIN:XT7 cln-3.2(gk41) I; cln-3.3(gk118) cln-3.1(pk479). Transgenics used in this study, also provided by the CGC, are RRID:WB-STRAIN:GS1912 arIs37 [pmyo-3::ssGFP], a transgenic strain that expresses ssGFP in the body wall muscles, which is secreted in the pseudocoelom and endocytosed by coelomocytes and RRID:WB-STRAIN:RT258 pwIs50 [lmp-1::GFP + Cb-unc-119(+)], a transgenic strain expressing GFP-tagged lysosomal marker LMP-1. Genes, for which mutants were unavailable, were knocked down using Ahringer library based RNAi methods (Kamath and Ahringer, 2003). The RNAi clones used were: L4440 empty vector control, ncr-1 (clone F02E8.6, Ahringer Library), ostm1 (clone F42A8.3, Ahringer Library), snx-3 (clone W06D4.5, Ahringer Library), manba (clone C33G3.4, Ahringer Library), aman-1 (clone F55D10.1, Ahringer Library), sul-3 (clone C54D2.4, Ahringer Library), gba-3 (clone F11E6.1, Ahringer Library) and asm1 (clone B0252.2, Ahringer Library).
Coelomocyte labeling and competition experiments were carried out with I4cLYA647, and ClensorA647 as described previously by our lab (Surana et al., 2011). For microinjections, the samples were diluted to 100 nM using 1X Medium 1 (150 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH 7.2). Injections were performed, in the dorsal side in the pseudocoelom, just opposite to the vulva, of one-day old wild type hermaphrodites using an Olympus IX53 Simple Inverted Microscope (Olympus Corporation of the Americas, Center Valley, PA) equipped with 40X, 0.6 NA objective, and microinjection setup (Narishige, Japan). Injected worms were mounted on 2.0% agarose pad and anesthetized using 40 mM sodium azide in M9 buffer. In all cases labeling was checked after 1 hr incubation at 22°C.
I4cLYA647 or ClensorA647 sample was diluted to 100 nM using 1X Medium 1 and injected in 10 arIs37 [pmyo-3::ssGFP] hermaphrodites as described previously by our lab (Surana et al., 2011). Imaging and quantification of the number of coelomocytes labeled, after 1 hr of incubation, was carried out on the Leica TCS SP5 II STED laser scanning confocal microscope (Leica Microsystems, Inc., Buffalo Grove, IL) using an Argon ion laser for 488 nm excitation and He-Ne laser for 633 excitation with a set of dichroics, excitation, and emission filters suitable for each fluorophore. Cross talk and bleed-through were measured and found to be negligible between the GFP/Alexa 488/BAC channel and Alexa 647 channel.
Bacteria from the Ahringer RNAi library expressing dsRNA against the relevant gene was fed to worms, and measurements were carried out in one-day old adults of the F1 progeny (Kamath and Ahringer, 2003). RNA knockdown was confirmed by probing mRNA levels of the candidate gene, assayed by RT-PCR. Briefly, total RNA was isolated using the Trizol-chloroform method; 2.5 μg of total RNA was converted to cDNA using oligo-dT primers. 5 μL of the RT reaction was used to set up a PCR using gene-specific primers. Actin mRNA was used as a control. PCR products were separated on a 1.5% agarose-TAE gel. Genes in this study that were knocked down by RNAi correspond to clh-6, ncr-1 and ostm-1 that showed expected 1.1 kb (clh-6); 1.1 kb (ncr-1); 0.9 kb (ostm-1) etc (Figure 1—figure supplement 1).
Fluorescence spectra were measured on a FluoroMax-4 Scanning Spectrofluorometer (Horiba Scientific, Edison, NJ, USA) using previously established protocols (Modi et al., 2009; Saha et al., 2015). Briefly, I4cLYA488/A647 was diluted to 50 nM in 1X pH clamping buffer of desired pH for all in vitro fluorescence experiments. All samples were vortexed and equilibrated for 30 min at room temperature. The samples were excited at 488 nm and emission collected between 505–750 nm. A calibration curve was obtained by plotting the ratio of donor emission intensity (D) at 520 nm and acceptor intensity (A) at 669 nm (for A488/A647) as a function of pH. Mean of D/A from three independent experiments and their s.e.m were plotted for each pH value. For in vitro calibration of ImLy, 50 nM of the sensor is diluted into 1X pH clamping buffer of desired pH. Oregon Green and Atto 647N are excited at 490 nm and 645 nm respectively. Emission spectra for Oregon Green and Atto 647N were collected between 500–550 nm and 650–700 nm respectively. A calibration curve was obtained by plotting the ratio of Oregon Green (G) at 520 nm and Atto 647N (R) at 665 nm (for G/R) as a function of pH. Mean of G/R from three independent experiments and their s.e.m were plotted for each pH value.
For chloride measurements, 10 μM stock of Clensor was diluted to a final concentration of 200 nM using 10 mM sodium phosphate buffer, pH 7.2 and incubated for 30 min at room temperature prior to experiments. BAC and Alexa 647 were excited at 435 nm for BAC and 650 nm for Alexa 647 respectively. Emission spectra of BAC and Alexa 647 were collected between 495–550 nm and 650–700 nm respectively. In order to study the chloride sensitivity of Clensor, final chloride concentrations ranging between 5 mM to 80 mM were achieved by addition of microliter aliquots of 1 M stock of NaCl to 400 μL of sample. Emission intensity of BAC at 505 nm (G) was normalized to emission intensity of Alexa 647 at 670 nm (R). Fold change in R/G ratio was calculated from the ratio of R/G values at two specific values of [Clˉ], either 5 mM and 80 mM or 5 mM and 120 mM as mentioned in the text.
Clamping and real time measurement experiments were carried out with I4cLYA488/A647 as described by our lab previously (Modi et al., 2009; Surana et al., 2011). For microinjections, the I-switch sample was diluted to 500 nM using 1X Medium 1. Briefly, worms were incubated at 22˚C for 1 hr post microinjection and then immersed in clamping buffers (120 mM KCl, 5 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 20 mM HEPES) of varying pH, containing 100 µM nigericin and 100 µM monensin. In order to facilitate entry of the buffer into the body, the cuticle was perforated at three regions of the body using a microinjection needle. After 75 mins incubation in the clamping buffer, coelomocytes were imaged using wide field microscopy. Three independent measurements, each with 10 worms, were made for each pH value.
Chloride clamping and real time measurements were carried out using Clensor. Worms were injected with 2 μM of Clensor and incubated at 22˚C for 2 hr. To obtain the chloride calibration profile, the worms were then immersed in the appropriate chloride clamping buffer containing a specific concentration of chloride, 100 μM nigericin, 100 μM valinomycin, 100 μM monensin and 10 μM chloride ionophore I for 45 mins at room temperature. Chloride calibration buffers containing different chloride concentrations were prepared by mixing the 1X chloride positive buffer (120 mM KCl, 20 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH, 7.2) and 1X chloride negative buffer (120 mM KNO3, 20 mM NaNO3, 1 mM Ca(NO3)2, 1 mM Mg(NO3)2, 20 mM HEPES, pH 7.2) in different ratios.
For real-time lysosomal pH or chloride measurements, 10 hermaphrodites were injected with I4cLYA488/A647 or Clensor respectively and incubated at 22°C for 1 hr. Worms were then anaesthetized and imaged on a wide field inverted microscope for pH measurements and confocal microscope for chloride measurements.
Mouse alveolar macrophage J774A.1 cells were a kind gift from Prof Deborah Nelson, Department of Pharmacological and Physiological Sciences, the University of Chicago, cultured in Dulbecco’s Modified Eagle’s Medium/F-12 (1:1) (DMEM-F12) (Invitrogen Corporation,USA) containing 10% heat inactivated Fetal Bovine Serum (FBS) (Invitrogen Corporation, USA). THP-1 monocyte cell line was obtained from late Professor Janet Rowley’s Lab at the University of Chicago. Cells were cultured in RPMI 1640 containing 10% heat-inactivated FBS, 10 mM HEPES, 2 mM glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin, and maintained at 37°C under 5% CO2. All reagents and medium were purchased from (Invitrogen Corporation,USA). THP-1 monocytic cells were differentiated into macrophages in 60 mm dishes containing 3 ml of the RPMI 1640 medium containing 10 nM PMA over 48 hr. These cells are not on the list of commonly misidentified cell lines maintained by the International Cell Line Authentication Committee. The sources of each cell line used in this study are as mentioned above and were used directly by us without additional authentication beyond that provided by the sources. All cells were regularly checked for mycoplasma contamination and were found to be negative for contamination as assayed by DAPI staining.
Chloride clamping and measurements were carried out using Clensor using a previously published protocol from our lab (Saha et al., 2015). J774A.1 and THP-1 cells were pulsed and chased with 2 μM of Clensor. Cells are then fixed with 200 μL 2.5% PFA for 2 min at room temperature, washed three times and retained in 1X PBS. To obtain the intracellular chloride calibration profile, perfusate and endosomal chloride concentrations were equalized by incubating the previously fixed cells in the appropriate chloride clamping buffer containing a specific concentration of chloride, 10 μM nigericin, 10 μM valinomycin, and 10 μM tributyltin chloride (TBT-Cl) for 1 hr at room temperature.
Chloride calibration buffers containing different chloride concentrations were prepared by mixing the 1X chloride positive buffer (120 mM KCl, 20 mM NaCl, 1 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH, 7.2) and 1X - chloride negative buffer (120 mM KNO3, 20 mM NaNO3, 1 mM Ca(NO3)2, 1 mM Mg(NO3)2, 20 mM HEPES, pH 7.2) in different ratios.
For real-time chloride measurements, cells are pulsed with 2 μM of Clensor followed by a 60 min chase. Cells are then washed with 1X PBS and imaged. To see whether Clensor can detect changes in Clˉ accumulation under perturbed conditions, we treated cells with 50 µM NPPB, which is a well-known non-specific Clˉ channel blocker. Cells were labeled with 2 μM Clensor for 30 mins and chased for 30 mins at 37°C. The cells were then chased for 30 mins in media containing 50 µM NPPB and then imaged.
To estimate the chloride accumulation in the lysosomes of Gaucher’s Disease in two cell models that is murine J774A.1 and human THP-1 cells, glucosylceramide storage was induced catalytically inactivating the enzyme acid β-glucosidase, using its well-known inhibitor conduritol β epoxide (CBE) (Grabowski et al., 1986; Schueler et al., 2004). These are both well-documented murine and human cell culture models of Gaucher's disease. Macrophage cells were cultured with 400 µM CBE for 48 hr. Cells were then pulsed and chased with 2 μM Clensor as previously described.
To estimate chloride accumulation in the lysosomes of Niemann Pick A/B disease, the same murine and human cell lines were used, and the activity of acid sphingomyelinase (ASM) in these macrophage cell lines was inhibited using the well-known inhibitor, amitriptyline hydrochloride (Beckmann et al., 2014; Kornhuber et al., 2010). Cells were labeled with 2 μM Clensor for 30 mins and chased for 30 mins at 37°C. The cells were then chased for 30 mins in media containing 10 µM amitriptyline hydrochloride and then imaged.
In cellulo pH clamping and measurement experiments were carried out with ImLy with modifications to protocols described by our lab previously (Modi et al., 2013, 2009). J774A.1 and THP-1 cells were pulsed and chased with 500 nM of ImLy. Cells are then fixed with 200 μL 2.5% PFA for 20 mins at room temperature, washed three times and retained in 1X PBS. To obtain the intracellular pH calibration profile, perfusate and endosomal pH were equalized by incubating the previously fixed cells in the appropriate pH clamping buffer clamping buffers (120 mM KNO3, 5 mM NaNO3, 1 mM Mg(NO3)2, 1 mM Ca(NO3)2, 20 mM HEPES, MES and NaOAc) of varying pH, containing 25 µM nigericin and 25 µM monensin for 30 mins at room temperature.
For real-time pH measurements, cells are pulsed with 500 nM of ImLy followed by a 60 mins chase. Cells are then washed with 1X PBS and imaged. pH measurements in the lysosomes of Gaucher’s Disease and of Niemann Pick A/B disease, in the two cell models that is murine J774A.1 and human THP-1 cells, were carried out similar to the protocol above using 500 nM of ImLy.
Wide field microscopy was carried out on IX83 research inverted microscope (Olympus Corporation of the Americas, Center Valley, PA, USA) using a 60X, 1.42 NA, phase contrast oil immersion objective (PLAPON, Olympus Corporation of the Americas, Center Valley, PA, USA) and Evolve Delta 512 EMCCD camera (Photometrics, USA). Filter wheel, shutter and CCD camera were controlled using Metamorph Premier Ver 220.127.116.11 (Molecular Devices, LLC, USA), suitable for the fluorophores used. Images on the same day were acquired under the same acquisition settings. All the images were background subtracted taking mean intensity over an adjacent cell free area. Mean intensity in each endosome was measured in donor (D) and acceptor (A) channels. Alexa 488 channel images (D) were obtained using 480/20 band pass excitation filter, 520/40 band pass emission filter and a 89016- ET - FITC/Cy3/Cy5 dichroic filter. Alexa 647 channel images (A) were obtained using 640/30 band pass excitation filter, 705/72 band pass emission filter and 89016- ET - FITC/Cy3/Cy5 dichroic filter. For FRET channel images were obtained using the 480/20 band pass excitation filter, 705/72 band pass emission filter and 89016- ET - FITC/Cy3/Cy5 dichroic filter. Mean intensity in each endosome was measured in donor and acceptor channels. A ratio of donor to acceptor intensities (D/A) was obtained from these readings. Pseudocolor images were generated by calculating the D/A ratio per pixel. Confocal images were captured with a Leica TCS SP5 II STED laser scanning confocal microscope (Leica Microsystems, Inc., Buffalo Grove, IL, USA) equipped with 63X, 1.4 NA, oil immersion objective. Alexa 488 was excited using an Argon ion laser for 488 nm excitation, Alexa 647 using He-Ne laser for 633 excitation and BAC using Argon ion laser for 458 nm excitation with a set of dichroics, excitation, and emission filters suitable for each fluorophore.
Ratiometric calcium imaging of Fura-2 was carried out on an Olympus IX81 microscope equipped with a 40x objective, NA = 1.2. Excitation of Fura-2 is performed using 340/26 and 380/10 nm excitation filters, equipped with a 455 nm dichroic mirror and a 535/40 nm emission filter. Exposure time was kept at 100 ms for all the imaging experiments to minimize phototoxicity.
Images were analyzed with ImageJ ver 1.49 (NIH, USA). For pH measurements Alexa 488 and Alexa 647 images were overlapped using ImageJ and endosomes showing colocalization were selected for further analysis. Intensity in each endosome was measured in donor (D) and FRET (A) channels and recorded in an OriginPro Sr2 b9.2.272 (OriginLab Corporation, Northampton, MA, USA) file from which D/A ratio of each endosome was obtained. The mean D/A of each distribution were converted to pH according to the intracellular calibration curve. Data was represented as mean pH value ± standard error of the mean. Data for pH clamping experiments was analysed similarly.
For chloride measurements, regions of cells containing lysosomes in each Alexa 647 (R) image were identified and marked in the ROI plugin in ImageJ. The same regions were identified in the BAC (G) image recalling the ROIs and appropriate correction factor for chromatic aberration if necessary. After background subtraction, intensity for each endosome was measured and recorded in an Origin file. A ratio of R to G intensities (R/G) was obtained from these values by dividing the intensity of a given endosome in the R image with the corresponding intensity in the G image. For a given experiment, mean [Clˉ] of an organelle population was determined by converting the mean R/G value of the distribution to [Clˉ] values according to the intracellular calibration profile. Data was presented as mean of this mean [Clˉ] value ± standard error of the mean. Data for chloride clamping experiments was analyzed similarly.
Colocalization of GFP and Alexa 647 was determined by counting the numbers of Alexa 647 positive puncta that colocalize with GFP and representing it as a Pearson's correlation coefficient.
Temporal mapping of I-switch and Clensor was done in 10 worms of pwIs50 [lmp-1::GFP + Cb-unc-119(+)] as previously described by our lab (Surana et al., 2011). Briefly, worms were injected with 500 nM of I4cLYA647 or ClensorA647, incubated at 22°C for 1 hr, and then imaged using Leica TCS SP5 II STED laser scanning confocal microscope (Leica Microsystems, Inc., Buffalo Grove, IL, USA). Colocalization of GFP and I4cLYA647 or ClensorA647 was determined by counting the numbers of Alexa647 positive puncta that colocalize with GFP positive puncta and expressing them as a percentage of the total number of Alexa 647 positive puncta. In order to confirm lysosomal labeling in a given genetic background, the same procedure was performed on the relevant mutant or RNAi knockdown in pwIs50 [lmp-1::GFP + Cb-unc-119(+)].
All pH and chloride clamping experiments (Figure 1b, Figure 1—figure supplement 2, Figure 4—figure supplement 2) were performed in triplicates and the standard error of mean (s.e. m) values are plotted with the number of cells considered being mentioned in each legend. Experiment with murine macrophage, J774A.1 and THP-1 cells (Figure 4) has been performed in triplicates. Ratio of standard error of the mean is calculated for n = 20 cells and n = 10 cells and is plotted in Figure 4d and e respectively. All pH and chloride measurements in C.elegans of indicated genetic backgrounds (Figures 2c and 3c and Figure 2—figure supplement 1c ) were carried out in n = 10 worms and the standard error of mean (s.e.m) values are plotted with the number of cells considered being mentioned in each legend.
Coelomocyte labeling for stability assay were carried out with I4cLYA647, and ClensorA647. For microinjections, the samples were diluted to 500 nM using 1X Medium 1 (150 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH 7.2). Post injection the worms are incubated at 22°C. After requisite time the injected worms are anesthetized in 40 mM sodium azide in M9 buffer and mounted on a glass slide containing 2% agarose pad. Worms were imaged using Olympus IX83 research inverted microscope (Olympus Corporation of the Americas, Center Valley, PA, USA). The average whole cell intensity in the Alexa 647 channel was plotted as a function of time (Figure 2—figure supplement 2)
J774A.1 and THP-1 macrophage cells labeling was carried out using 500 nM ClensorA647 using 1X Medium 1 (150 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 20 mM HEPES, pH 7.2). Cells were pulsed for 30 mins and then chased at 37°C for the indicated time points. The average whole cell intensity in the Alexa 647 channel was plotted as a function of time (Figure 4—figure supplement 3).
Cells were loaded with 3 μM Fura-2 AM in HBSS (137 mM NaCl, 5 mM KCl, 1.4 mM CaCl2, 1 mM MgCl2, 0.25 mM Na2HPO4, 0.44 mM KH2PO4,4.2 mM NaHCO3 and 10 mM glucose) at 37°C for 60 min. Post incubation cells were washed with 1X PBS and then imaged in Low Ca2+ or Zero Ca2+ buffer (145 mM NaCl, 5 mM KCl, 3 mM MgCl2, 10 mM glucose, 1 mM EGTA, and 20 mM HEPES (pH 7.4)). Ca2+ concentration in the nominally free Ca2+ solution is estimated to be 1–10 μM. With 1 mM EGTA, the free Ca2+ concentration is estimated to be <10 nM based on the Maxchelator software (http://maxchelator.stanford.edu/). Florescence was recorded using two different wavelengths (340 and 380 nm) and the ratio (F340/F380) was used to calculate changes in intracellular [Ca2+].
Enzyme activity assays were performed in J774A.1 cells. For Cathepsin C enzyme activity; we used Gly-Phe β-naphthylamide as a substrate. Lysosomes of J774A.1 cells were pre-labeled with TMR-dextran (0.5 mg/mL; G) for 1 hr and chased in complete medium for 16 hr at 37°C. The cells were then labeled with 50 nM LysoTracker in complete medium for 30 mins at 37°C. 50 μM NPPB or 200 μM GPN were then added to the cells and incubated for 30 mins at 37°C. The cells then washed and imaged in HBSS buffer containing either NPPB or GPN. The whole cell intensity ratio (G/R) was plotted to quantify the level of LysoTracker labelling of the endosomes. For Cathepsin L and Aryl Sulfatase enzyme activity Magic Red Cathepsin L assay kit (Immunochemistry Technologies) and Lysosomal sulfatase assay kit (Marker Gene) were used. The experiment was performed using the manufacture’s protocol. Briefly, cells were incubated with 1X Magic Red Cathepsin L assay probe or 200 μM Lysosomal sulfatase assay probe for 4 hr in complete medium. The cells were then labelled with 10 μM Hoechst stain for 10 mins at 37°C after which the cells were washed and imaged. For low chloride containing dishes; cells were preincubated with 50 μM NPPB before the addition of the enzyme probes. The ratio of enzyme substrate whole cell intensity to that of DAPI was used to quantify enzyme activity.
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Suzanne R PfefferReviewing Editor; Stanford University School of Medicine, 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.
[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration and the paper was accepted for publication. The first decision letter after peer review is shown below.]
Thank you for submitting your work entitled "High lumenal chloride in the lysosome is critical for lysosome function" for consideration by eLife. Your article has been favorably evaluated by a Senior Editor and three reviewers, one of whom, Suzanne R Pfeffer (Reviewer #1), is a member of our Board of Reviewing Editors. The following individuals involved in review of your submission have agreed to reveal their identity: Haoxing Xu (Reviewer #2) and Carsten Schultz (Reviewer #3).
Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that this manuscript will not be considered further for publication in eLife. As you will see in the following comments, the reviewers noted a number of strengths in the use of your novel chloride sensor. However, they felt that additional work would be required to bring the work to the level of significance and novelty expected for presentation in eLife.
Quoting a review from Jentsch (2013), exact measurements of lysosomal [Cl−] have not been reported due to the lack of suitable Cl− sensors for the high [Cl−] at low pH. However, from measurements with cells treated with low Cl− and from model calculations, lysosomal [Cl−]lumen is greater than 80 mM. 2010 data strongly point to a crucial role of Cl− transport in organellar physiology that "goes beyond merely providing the electrical shunt for proton pumping by the V-ATPase." Despite maintaining lysosomal conductance and normal lysosomal pH, Clcn7unc/unc mice showed lysosomal storage disease like mice lacking ClC-7. Although various proteins are known to be regulated by Cl−, the mechanism by which (mainly luminal) Cl− affects membrane traffic and organellar function has remained elusive; the 2010 analysis of Clcn5unc and Clcn7unc mice has suggested that luminal anion concentration is important all along the endosomal-lysosomal pathway.
Here, the authors use a novel, DNA based chloride sensor to try more accurately to determine lysosomal chloride levels. Like the previous work from the Jentsch lab, the findings using C. elegans lysosomes indicate a high level of lysosomal chloride. Chloride ion levels are decreased under conditions of lysosomal dysregulation due to mutations in genes that lead to lysosomal storage disorders. Not clear are the mechanisms underlying these observations.
Recently, the lysosome has been shown to represent a significant calcium store, and in worms, CUP-5 is the MCOLN-1 homolog that is important for lysosomal calcium levels and functions. If chloride is not the counterion for protons, it may be important for calcium homeostasis. Given that it is not clear how chloride influences lysosome function in disease states or in normal cells, it would be very important for these authors to clarify the connection between chloride accumulation defects and calcium uptake and regulation. Without this, the findings seem somewhat anecdotal, despite highlighting the recently uncovered importance of chloride ions in lysosome function. No doubt this is an interesting area for further investigation.
Cl- transport across endosomal and lysosomal membranes are known to be crucial for lysosomal acidification and physiology, but the underlying mechanisms are poorly understood, largely due to the lack of the methods that can be used to reliably monitor luminal Cl- levels in live cells. The authors previously reported the development of a DNA-based fluorescent reporter (i.e., Clensor) for Cl-. Compared with protein-based ion-sensing probes, a major advantage for DNA-based probes is their relative insensitivity to pH, which is low in the endosomes and lysosomes. Building on this exciting development in the field, the authors now report the use of Clensor in C. elegans and mammalian phagocytes, providing evidence that a decrease in lysosomal [Cl-] is likely to be a primary pathogenic factor in a number of lysosome storage diseases (LSDs). Overall, the cell biology and imaging experiments in the study were carefully designed, and the results were mostly clean and convincing, and carefully interpreted. Both the use of Clensor and the major conclusions of the paper are of substantial interests to researchers working on lysosome biology, anion channels and transporters, membrane transport, organellar channels and transporters, and LSDs. The manuscript can be improved if following points are taken into consideration.
1) Cl- channels are known to exhibit poor anion selectivity. What is the anion selectivity for Clensor? When lysosomal [Cl-] is reduced in some LSD cells, assuming that lysosome lumen is still electroneutral and iso-osmotic, what are substituted anions? Phosphate is probably an obvious candidate. Therefore, it would be helpful if the authors can show the effects of NO3(-) and PO4(3-) in Clensor in vitro.
2) Lysosomal delivery of Clensor might be slowed down in LSD cells, compared with wild-type (WT) cells. Could trafficking defects contribute to the observed reduction in lysosomal [Cl-]? Given the 40 mM difference in late endosomal vs. lysosomal [Cl-], it is plausible that a block in endosome maturation might affect the [Cl-] measurement. I would assume that the calibration curves and maximal signals are identical for WT and LSD cells. Is that the case?
The work of Krishnan and co-workers describes the use of previously published fluorescent sensors for measuring lysosomal pH and Cl levels. Elegantly, the work was performed in circulating nematode macrophages as well as cultured mammalian cells. This is a very exciting application and the authors observe that a lack of chloride levels correlates with a loss of degrading capacity of the lysosome. The authors follow the hypothesis that the drop-in chloride content is responsible for driving lysosomal function. They tested model conditions for lysosomal storage diseases in C. elegans and found indeed reduced chloride levels under such conditions. However, this is not a prove that the chloride ion levels are instrumental. The question remains if chloride levels are the key component in lysosomal dysfunction or a by-stander effect.
The authors argue that CF patients show lysosomal distress symptoms at the molecular level (changes in enzyme activities) similar to Niemann-Pick patients. What the authors do not consider in their argumentation is that CF patients share very little of the NPC phenotype. There is no early neurodegeneration in CF nor a liver accumulation of lipids such as cholesterol or sphingosine. As a result, the conclusions driven by the present study do not hold. Down to the bare bones, the study reveals very interesting data regarding the chloride ion levels in lysosomes but lacks any mechanistic insight. Unfortunately, there is no consideration (or even mentioning) of the calcium levels in the lysosome that have received quite some attention in the regulation of lysosomal function and signaling lately (for instance, see Medina et al. Nature Cell Biology 17, 288-299 (2015)). Chloride might change driving forces for lysosomal im- or export but a direct switching function through changes in protein conformation as is known for calcium ions would be a large surprise.
For publication in eLife, I would expect a more profound mechanistic insight of what chloride ions are doing in lysosomes. So far, the authors present highly interesting observations in very relevant cells followed by a conclusion section that is in my opinion not coherent with the observed phenotype (in CF). To make this work suitable for eLife, I would expect measuring and/or manipulating calcium levels under conditions where lysosomal chloride levels are high or low, respectively. In addition, the authors could manipulate chloride levels in lysosomes acutely and observe effects spontaneously.
In summary, I suggest to reject this manuscript until the above-mentioned experiment sets have been performed and a mechanistic model is presented.https://doi.org/10.7554/eLife.28862.025
- Yamuna Krishnan
- Yamuna Krishnan
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
The authors thank Professors John Kuriyan, Susan Cotman, Drs A H Rahmathullah and D McEwan for critical comments and valuable suggestions. The authors thank the Integrated Light Microscopy facility at the University of Chicago, the C. elegans Genetic Center (CGC) for strains, Koushika, S, Glotzer, M, and Ausbel, F for Arhinger Library RNAi clones. Data described in the paper are presented in the main text and the supplementary materials. This work was supported by the National Center for Advancing Translational Sciences of the National Institutes of Health through grant no: UL1 TR000430 and U Chicago startup funds to YK. YK is a Brain Research Foundation Fellow.
- Suzanne R Pfeffer, Reviewing Editor, Stanford University School of Medicine, United States
- Received: May 21, 2017
- Accepted: June 23, 2017
- Version of Record published: July 25, 2017 (version 1)
© 2017, Chakraborty et al.
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