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RESEARCH ARTICLE

Zn2+ efflux through lysosomal exocytosis prevents Zn2+-inducedtoxicity

Ira Kukic1, Shannon L. Kelleher2,3,4 and Kirill Kiselyov1,*

ABSTRACT

Zn2+ is an essential micronutrient and an important ionic signal

whose excess, as well as scarcity, is detrimental to cells. Free

cytoplasmic Zn2+ is controlled by a network of Zn2+ transporters and

chelating proteins. Recently, lysosomes became the focus of

studies in Zn2+ transport, as they were shown to play a role in

Zn2+-induced toxicity by serving as Zn2+ sinks that absorb Zn2+ from

the cytoplasm. Here, we investigated the impact of the lysosomal

Zn2+ sink on the net cellular Zn2+ distribution and its role in cell

death. We found that lysosomes played a cytoprotective role during

exposure to extracellular Zn2+. Such a role required lysosomal

acidification and exocytosis. Specifically, we found that the inhibition

of lysosomal acidification using Bafilomycin A1 (Baf) led to a

redistribution of Zn2+ pools and increased apoptosis. Additionally,

the inhibition of lysosomal exocytosis through knockdown (KD) of

the lysosomal SNARE proteins VAMP7 and synaptotagmin VII

(SYT7) suppressed Zn2+ secretion and VAMP7 KD cells had

increased apoptosis. These data show that lysosomes play a

central role in Zn2+ handling, suggesting that there is a new Zn2+

detoxification pathway.

KEY WORDS: Zinc, Golgi, Lysosome, Metallothionein, Exocytosis,

Zn2+ transport, ZnT, Slc30a

INTRODUCTIONCellular Zn2+ dyshomeostasis has been linked to a number of

human pathologies including growth defects (Prasad, 2013),

impaired immune function (Rink and Gabriel, 2000), diabetes(Jansen et al., 2009) and neurodegenerative diseases (Forsleff

et al., 1999; Rulon et al., 2000; Lee et al., 2002; Vinceti et al.,

2002). Regulation of cellular Zn2+ levels involves controlling itsinflux, export and chelation. In general, Zn2+ transport is

regulated by ZnT (also known as solute carrier family 30) andZIP (also known as solute carrier family 39) transporters, and it is

chelated by Zn2+-binding metallothioneins.

In addition to Zn2+ evacuation across the plasma membrane by

the Zn2+ transporter ZnT1 (SLC30A1) (Palmiter and Findley,

1995), Zn2+ is exported from the cytoplasm into the organelles bythe dedicated ZnT transporters such as ZnT6 (SLC30A6) for the

Golgi (Huang et al., 2002), and ZnT2 (SLC30A2) and ZnT4

(SLC30A4) for the lysosome (Palmiter et al., 1996; Huang andGitschier, 1997; Falcon-Perez and Dell’Angelica, 2007;McCormick and Kelleher, 2012). This organellar Zn2+ export

lowers potentially toxic cytoplasmic Zn2+ concentrations inpathophysiological conditions such as neurodegeneration(Kanninen et al., 2013) and breast cancer (Lopez et al., 2011).

Moreover, it provides Zn2+ to organellar processes that require it,such as the maturation of enzymes like the lysosomal acidsphingomyelinase (Schissel et al., 1996), and for the secretion ofZn2+ under normal physiological conditions such as synaptic

transmission (Frederickson and Bush, 2001) and lactation(Kelleher et al., 2009).

The upregulation of Zn2+ chelation and transport machinery

following the activation of the transcription factor MTF-1 by Zn2+

binding (Andrews, 2001) requires time for transcription, translationand protein processing. It is tempting to speculate that Zn2+ export into

organelles serves as a first line of defense to provide temporary Zn2+

storage, giving cells time to upregulate Zn2+ chelators and transporters.Our recent data on the role of lysosomes in Zn2+ handling, as wellas some recently published results suggest that lysosomes play a

role as such Zn2+ sinks, temporarily storing Zn2+ (Hwang et al.,2008; Kukic et al., 2013). In this paper, we sought to delineate therole of lysosomes in protection against Zn2+-induced toxicity.

Zn2+ is transported from the cytoplasm into lysosomes byZnT2 and ZnT4 (Palmiter et al., 1996; Huang and Gitschier,1997; Falcon-Perez and Dell’Angelica, 2007). Zn2+ can also be

delivered to the lysosomes through endocytosis or autophagy(Lee and Koh, 2010; Cho et al., 2012). What happens to Zn2+

absorbed by the lysosomes? A recent series of work from several

laboratories indicate that Zn2+ buildup in the lysosomes is toxic.It leads to lysosomal membrane permeabilization (LMP), to therelease of the lysosomal enzymes such as cathepsins and to celldeath (Hwang et al., 2008; Chung et al., 2009; Lee et al., 2009;

Hwang et al., 2010). As such, the lysosomal Zn2+ accumulationmight constitute a cell death mechanism during normalremodeling of Zn2+-rich tissues, such as the mammary gland

(Kelleher et al., 2011), as well as in pathological conditions. Withthis in mind, we sought to answer whether or not accumulation ofZn2+ in the lysosome is the terminal depot for cellular Zn2+.

Alternatively, it is possible that lysosomal Zn2+ dissipates andlysosomes constitute only a temporary Zn2+ storage site. Ourrecently published data suggest that the lysosomal ion channeltransient receptor potential mucolipin 1 (TRPML1, also known as

mucolipin1, encoded by the MCOLN1 gene) is at least partlyresponsible for dissipating lysosomal Zn2+ into the cytoplasm(Kukic et al., 2013). It should be noted that lysosomes fuse with the

plasma membrane through a process involving a specific SNAREcomplex, which includes the VAMP7 protein and synaptotagminVII (SYT7) (Martinez-Arca et al., 2000; Braun et al., 2004; Rao

et al., 2004; Logan et al., 2006; Mollinedo et al., 2006). It has

1The Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA15260, USA. 2The Department of Nutritional Sciences, College of Health andHuman Development, The Pennsylvania State University, University Park, PA16802, USA. 3Department of Surgery, Penn State Hershey Medical Center,Hershey, PA 17033, USA. 4Department of Cellular and Molecular Physiology,Penn State Hershey Medical Center, Hershey, PA 17033, USA.

*Author for correspondence (kiselyov@pitt.edu)

Received 25 October 2013; Accepted 25 April 2014

� 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318

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recently been proposed that such secretion contributes to the

excretion of undigested/indigestible products inside lysosomes(Medina et al., 2011). In the course of the present study, we usedVAMP7 and SYT7 knockdown (KD) to suppress lysosomalsecretion and assess its role in Zn2+ clearance from the cells.

Here, we aimed to establish the functional context of thelysosomal Zn2+ accumulation. Our findings indicate thatlysosomes actively absorb Zn2+ and secrete it across the plasma

membrane, given that suppressing the lysosomal Zn2+ absorptionor secretion causes Zn2+ buildup in the cytoplasm, Golgi andmitochondria, leading to apoptosis.

RESULTSIn order to test the role of the lysosomal Zn2+ sink on cellularZn2+ handling, we blocked the lysosomal H+ pump in HeLa cells

using 1 mM Baf and exposed cells to 100 mM ZnCl2 for 3 hours.The resulting cytoplasmic Zn2+ spikes were measured using live-cell confocal microscopy and FluoZin-3,AM as described

previously (Kukic et al., 2013) Fig. 1A shows that the exposureof Baf-treated cells to Zn2+ caused a significantly higher FluoZin-3,AM response than the exposure of untreated cells to Zn2+.

Although Baf has been shown to decrease cytoplasmic pH,potentially affecting Zn2+ binding to cytoplasmic proteins, orFluoZin-3,AM fluorescence, the magnitude of the observed

effects appear to be incompatible with the quantitativeestimates of changes induced by Baf. Thus, the effect of Baf oncytoplasmic pH appears to be small, within only tenths of pHunits (Heming et al., 1995). The degree of pH change necessary to

cause an effect on Zn2+ handling, by contrast, significantlyexceeds that reported to be caused by Baf. A pH drop below 6.7 isrequired to trigger an increase in intracellular Zn2+ according to

one set of studies (Kiedrowski, 2012), whereas another set hasshown that metallothioneins release Zn2+ only after cytoplasmicpH drops below 5.0 (Jiang et al., 2000). Thus, the increase in

cytoplasmic Zn2+ caused by Baf likely correlates with the loss oflysosomal function, rather than cytosolic pH changes.

We have previously shown that Zn2+ transporters ZnT2 and

ZnT4 colocalize with the lysosomal ion channel TRPML1 in

HeLa cells (Kukic et al., 2013). We suggested that these

transporters play a role in loading of the lysosomes with Zn2+.In order to test this assumption, we performed ZnT2 and ZnT4

Fig. 1. Inhibition of lysosomal Zn2+ sinkfunction by Baf increases cytoplasmic Zn2+

levels. (A) Confocal images of HeLa cells treatedwith 100 mM ZnCl2 and/or 1 mM Baf for 3 hoursthen loaded with FluoZin-3,AM and LysoTracker.Note disappearance of LysoTracker staining,indicative of lysosomal deacidification, and anincrease in FluoZin-3,AM staining intensity,indicative of increased Zn2+. (B) Confocal imagesof control or ZnT siRNA transfected HeLa cells(48 hours post-transfection) treated with 100 mMZnCl2 for 3 hours then loaded with FluoZin-3,AM.Images represent at least three separateexperiments and at least three images percondition in each experiment. Scale bars: 20 mm.

Fig. 2. Inhibition of lysosomal Zn2+ sink function by Baf increases thetranscriptional response of Zn2+-responsive genes. qRT-PCR results ofMT2a (A) and ZnT1 (B) mRNA, shown as percentage of DMSO-only-treated(no Zn2+) cells. RNA was isolated from HeLa cells treated for 3 hours witheither DMSO or 1 mM Baf alone or with 100 mM ZnCl2. Results aremean6s.e.m.; *P,0.05; **P,0.001.

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KD using siRNA as described previously, and tested the resulting

changes in Zn2+ handling using FluoZin-3,AM. Fig. 1B showsthat ZnT2 and ZnT4 KD increased cytoplasmic Zn2+ levelsobserved in these cells after a 3-hour long treatment with 100 mM

ZnCl2. These results are in agreement with the previously

published data on the dependence of ZnTs activity on the acidic

environment of the lysosomes (Chao and Fu, 2004; Ohana et al.,2009) for Zn2+ binding and transporting activity.

The upregulation of MTF-1-dependent, Zn2+-responsive genes

such as metallothionein 2A (MT2a) and ZnT1 (Saydam et al.,2002) indicates elevated cytoplasmic Zn2+. MT2a mRNA wasused previously in our studies of the role of TRPML1 in Zn2+

handling. We measured the expression of the mRNA of these

genes using qRT-PCR (Fig. 2). An increase in MT2a and ZnT1mRNA responses to Zn2+ in cells treated with Baf is evident.With MT2a mRNA levels in DMSO-treated (no Zn2+) cells set as

100%, MT2a mRNA levels were 816.3%9673.61 (in cellstreated with DMSO and Zn2+ (n54; P,0.001), and1174.61%694.20 (mean6s.e.m., n54; P,0.05 relative to

DMSO-only, and DMSO plus Zn2+ controls) in cells treatedwith Baf and Zn2+ (Fig. 2A). ZnT1 mRNA increased to323.43%623.32 (n54; P,0.001) in cells treated with DMSOplus Zn2+, and to 552.09%640.46 (n54; P,0.05 relative to

DMSO-only, and DMSO plus Zn2+ controls) of control values incells treated with Baf plus Zn2+ (Fig. 2B). In addition toconfirming the elevated cytoplasmic Zn2+ levels due to Baf, as

assessed by FluoZin-3,AM in Fig. 1A, these data also corroborateMTF-1 activation due to cytoplasmic Zn2+ buildup.

In addition to increasing cytoplasmic Zn2+, suppression of the

lysosomal function caused redistribution of Zn2+ storage pools.Concentration of FluoZin-3 fluorescence in intracellularinclusions was noted in Baf-treated cells exposed to Zn2+. At

least some of these inclusions were positive for the Golgi markerGalT–mCherry (Fig. 3A). Under these conditions, Zn2+ alsoaccumulated in the mitochondria, which was shown using the

Fig. 3. Inhibition of lysosomal Zn2+ sink function by Bafredistributes cellular Zn2+ pools to the Golgi and themitochondria. (A) Confocal images of GalT–mCherry-transfected HeLa cells treated with 100 mM ZnCl2 and/or 1 mMBaf for 3 hours, then loaded with FluoZin-3,AM. The black-and-white panes show overlap between the green and the redchannels, obtained using the RG2B function of ImageJ.(B) Confocal images of HeLa cells treated with 100 mM ZnCl2and/or 1 mM Baf for 3 hours, then loaded with RhodZin-3,AM.Plot profiles on the right show intensity profiles of RhodZin-3,AMfluorescence recorded along the lines indicated in thecorresponding image. Note increased RhodZin-3,AMflorescence with Baf-treated cells. Images represent at leastthree separate experiments and at least three images percondition in each experiment. Scale bars: 20 mm.

Fig. 4. Inhibition of lysosomal function leads to increased cell deathupon high Zn2+ exposure. (A) Caspase-3 (Cas3) activity assay showingincreased Cas3 activation upon a 48-hour 100 mM ZnCl2 and 1 mM Baftreatment in HeLa cells. A 3-hour long exposure to 1 mM staurosporine wasused as a positive control, which increased Cas3 activity by796.72%6109.06 (n53, P,0.001). Cas3 activity is shown as AMCfluorescence and a percentage of untreated controls. Results aremean6s.e.m.; *P,0.05. (B) Flow cytometry data showing increased AnnVstaining of cells treated with 100 mM ZnCl2 and/or 1 mM Baf for 48 hours.Data represent three experiments.

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mitochondrial Zn2+ dye RhodZin-3,AM, whose signal wasbrighter in Baf-treated than in control Zn2+-treated cells(Fig. 3B). Therefore, suppression of lysosomal function leads to

the loss of Zn2+-buffering capacity and to a spike in cytoplasmicZn2+ when cells are exposed to Zn2+. In the absence of Zn2+

buffering by the lysosomes, Zn2+ is redirected to other organelles.

As previously shown, high Zn2+ is toxic to the cells owing toits buildup in the cytoplasm and in the organelles (Medvedevaet al., 2009). If the lysosomes are a Zn2+-buffering sink, then

inhibiting that function should result in cell death. Our findings inFig. 4A support this: although HeLa cells were fairly resistantto the effects of 100 mM ZnCl2 or 1 mM Baf alone, theircombination caused pronounced caspase-3 (Cas3) activation,

indicative of apoptosis. An exposure of cells to 100 mM ZnCl2for 48 hours increased Cas3 activity by 43.90%614.25(mean6s.e.m., n53; P,0.05) and exposure to 1 mM Baf

increased Cas3 activity by 179.5%668.04 (n53, P,0.05). Acombination of Zn2+- and Baf-exposure increased Cas3 activityby 387.87%636.70 (n53, P,0.001 relative to untreated control),

suggesting that Baf enhances the pro-apoptotic effects of Zn2+.Given that Baf is conventionally used to block the lysosomal H+

pump, we think that the simplest interpretation of these data are

as diminished cytoprotective capacity of the lysosomal Zn2+ sinkin Baf-treated cells.

As a complimentary cell death assay, Annexin V (AnnV) andPropidium Iodide (PI) staining were analyzed using flow

cytometry. Fig. 4B and supplementary material Fig. S1 show anincrease in the number of AnnV-positive, apoptotic cells whencells are treated for 48 hours with Zn2+ plus Baf. Taken together,

these data show that suppression of the lysosomal Zn2+ sinkfacilitates the apoptotic cell death caused by Zn2+ exposure. Theysuggest that lysosomes play a crucial role in buffering

cytoplasmic Zn2+ and in its detoxification. A loss of such a roleexposes cells to the pro-apoptotic effects of Zn2+.

Zn2+ toxicity has been linked to LMP, resulting in cell deathunder some conditions (Hwang et al., 2008; Chung et al., 2009;

Lee et al., 2009; Hwang et al., 2010). Why is the lysosomal Zn2+

sink cytoprotective under some, but not other conditions of Zn2+

exposure? We propose that the switch between pro- and anti-apoptotic effects of the Zn2+ sink is dictated by the rate of Zn2+

absorption by the lysosomes and/or the rate of its clearance fromthe lysosomes. If the rate of Zn2+ clearance exceeds the rate ofits sequestration into the lysosomes, then the Zn2+ sink is

cytoprotective. A rate of sequestration exceeding the rate ofclearance leads to LMP and cell death (see model in Fig. 5). Zn2+

clearance might occur through a Zn2+ leak into the cytoplasm [as

suggested by our recent publication (Kukic et al., 2013)], orthrough its secretion mediated by lysosomal fusion with theplasma membrane. The latter has recently gained a lot of attentionas detoxification mechanism in the lysosomal storage diseases

(Fraldi et al., 2010; Medina et al., 2011; Palmieri et al., 2011;Decressac et al., 2013; Pastore et al., 2013). The next set ofexperiments tested the role of lysosomes in Zn2+ secretion.

We suppressed lysosomal secretion by using siRNAs againsttwo lysosomal SNARE components, VAMP7 and SYT7(Martinez-Arca et al., 2000; Braun et al., 2004; Rao et al.,

2004; Logan et al., 2006; Mollinedo et al., 2006; Flannery et al.,2010). Fig. 6A shows that VAMP7 siRNA reduced VAMP7mRNA to 27.76%63.99 of control siRNA levels (mean6s.e.m.,

n54, P,0.001), whereas Fig. 7A shows that SYT7 siRNAreduced SYT7 mRNA to 30.12%64.11 of control siRNA levels(n53, P,0.001). VAMP7 mRNA levels were not altered byeither a 3- or 48-hour exposure to 100 mM ZnCl2 (I. K.,

unpublished observation; SYT7 dependence on Zn2+ was nottested). VAMP7 KD was confirmed by examining protein levelsthrough western blotting confirming that VAMP7 siRNA reduced

VAMP7 protein expression to 30.21%66.82 of control siRNAlevels (n55, P,0.001) (Fig. 6B). VAMP7 and SYT7 KD wereassociated with changes in the lysosomal numbers and

organization. There was an increase in total lysosomal numbersin VAMP7 KD cells (supplementary material Fig. S2), althoughthere was a high degree of cell-to-cell variation with that metric.Both VAMP7 and SYT7 KD also caused clustering of lysosomes

in large structures, which is compatible with the role of these

Fig. 5. A model of lysosomal Zn2+ sink and its role in Zn2+ detoxification. (A) Under normal conditions, Zn2+ enters the cells through ion channels andZIPs and is (1) recognized by the Zn2+-sensitive transcription factor MTF-1 that, once Zn2+-bound, translocates to the nucleus to upregulate ZnT1 and MT2agene expression. (2) Zn2+ is transported out of the cytoplasm and into the lysosome through ZnT2 and ZnT4. Zn2+ that builds up in the lysosomal Zn2+ sink isthen secreted across the plasma membrane (3) through a VAMP7-dependent mechanism. This prevents toxic Zn2+ buildup in the cytoplasm and otherorganelles (4). (B) Suppression of Zn2+ absorption by the lysosomal Zn2+ sink (5), leads to toxic levels of Zn2+ buildup in the cytoplasm as well as otherorganelles (6), such as the Golgi and mitochondria.

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SNARE proteins in membrane traffic. Examples of suchclustering and its statistical analysis are shown in Figs 6, 7;

supplementary material Fig. S2.VAMP7 and SYT7 KD were functionally significant as they

caused a loss of secreted activity of the lysosomal enzyme b-hexosaminidase (b-hex), a common marker of lysosomal

exocytosis. In these experiments, lysosomal secretion wasinitiated by treatment with 1 mM of the Ca2+ ionophoreionomycin (Ion) (Fig. 6C; Fig. 7B). After 30 minutes, control

KD cells secreted 18.80%60.66 of their total b-hex contentwithout stimulation, whereas VAMP7 KD cells secreted14.09%61.60 of their total b-hex content (mean6s.e.m.,

Fig. 6C). Once stimulated with ionomycin, however, controlKD cells secreted 32.57%62.36 of their total b-hex content,whereas VAMP7 KD cells only secreted 18.87%61.57 of their

total b-hex content (n53, P,0.001) (Fig. 6C). Similarly, SYT7KD was also functionally significant, decreasing lysosomalexocytosis compared to control KD. Fig. 7B shows that after

30 minutes, ionomycin-treated control KD cells secreted35.57%61.36 of their total b-hex content, whereas SYT7 KD

cells secreted 20.29%61.65 of their total b-hex content (n53,P,0.001).

To measure the effect of suppressing lysosomal exocytosis onZn2+ secretion, we incubated control and VAMP7, or SYT7 KD

cells with 100 mM ZnCl2 for 3 hours. Next, cells were placed innormal medium (no Zn2+) and Zn2+ secretion was analyzed usingFluoZin-3 fluorescence as described previously (Kukic et al.,

2013). Both VAMP7 (Fig. 6D) and SYT7 (Fig. 7C) KDsignificantly reduced Zn2+ secretion. Within 30 minutes ofsecretion, VAMP7 KD cells were only able to secrete

51.79%619.26 (n53, P,0.05) of the amount of Zn2+ secretedby the control KD cells (100%) (Fig. 6D), indicating that VAMP7is necessary for Zn2+ secretion. Similarly, SYT7 KD cells were

only able to secrete 58.9062.87% (n53, P,0.05) of the amountof Zn2+ secreted by the control KD cells at 30 minutes, whichwas taken as 100% (Fig. 7C). This inhibition of Zn2+ secretion in

Fig. 6. Inhibition of lysosomal exocytosis throughVAMP7 KD inhibits Zn2+ secretion. (A) qRT-PCR resultsconfirming VAMP7 KD by siRNA. (B) Quantification ofwestern blot results confirming VAMP7 KD. Insert shows arepresentative blot (of five) of endogenous VAMP7 andactin levels under with either control or VAMP7 siRNA.(C) b-hexosaminidase activity assay for lysosomalexocytosis. Results are shown as the percentage of totalb-hexosaminidase secreted after either 10 or 30 minutes;1 mM ionomycin was used to stimulate lysosomalexocytosis. (D) Zn2+ secretion assay using cell-impermeant FluoZin-3 tetrapotassium salt. Results areshown as the percentage of the maximum fluorescencevalue recorded in the control+Zn2+ samples after30 minutes, which were set at 100%. Results aremean6s.e.m.; *P,0.05, **P,0.001.

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VAMP7 and SYT7 KD cells was also corroborated by theelevated cellular Zn2+ levels, as seen by assessing FluoZin-3,AM

staining using confocal microscopy (Fig. 7D).

The effects of suppressing lysosomal secretion on Zn2+-induced cell death were analyzed using the AnnV and PI assay

described above. This assay revealed significant upregulation ofZn2+-induced cell death in VAMP7 KD cells treated with 100 mMZnCl2 for 48 hours, compared to control KD cells undergoing thesame treatment (Fig. 8; supplementary material Fig. S1). In

summary, the data described here show that the Zn2+ sinkintegrates Zn2+ absorption from the cytoplasm with its secretionthrough lysosomal exocytosis. Both aspects of the Zn2+ sink

function are new and necessary for Zn2+ detoxification.Finally, because lysosomal biogenesis and exocytosis are

regulated by TFEB and related factors (Sardiello et al., 2009;

Martina et al., 2014), TFEB should have an impact on lysosomalZn2+ clearance. Indeed, Fig. 9 shows that TFEB overexpression,a common protocol used to study its function, increases both

lysosomal exocytosis and Zn2+ secretion. Fig. 9A shows that after30 minutes, mock (empty vector)-transfected cells secreted13.95%60.67 of their total b-hex content, while TFEB-transfected cells secreted 23.11%63.75 (mean6s.e.m.; n53,

P,0.05) of their total b-hex content. Once stimulated withionomycin, mock-transfected cells secreted 20.30%61.47 (n53,P,0.01) of their total b-hex content, whereas TFEB-transfected

cells secreted 38.65%61.98 (n53, P,0.001) of their total b-hexcontent. TFEB also increased Zn2+ secretion. Fig. 9B shows thatafter 15 minutes, control KD cells were able to secrete

76.70%61.37 of secretable Zn2+ in our assay, whereas TFEB-transfected cells were able to secrete 104.52%610.12 (n53,P,0.05) of secretable Zn2+. This increase in Zn2+ secretion was

dependent on the lysosomal fusion machinery as TFEB cDNA

Fig. 7. Inhibition of lysosomal exocytosisthrough SYT7 KD inhibits Zn2+ secretion.(A) qRT-PCR results confirming SYT7 KD bysiRNA. (B) b-hexosaminidase activity assay forlysosomal exocytosis. Results are shown as thepercentage of total b-hexosaminidase secretedover 10 or 30 minutes; 1 mM ionomycin was usedto stimulate lysosomal exocytosis. (C) Zn2+

secretion assay using cell-impermeant FluoZin-3tetrapotassium salt. Results are shown as percentof the maximum fluorescence value recorded inthe control+Zn2+ samples at 30 minutes, whichwere taken as 100%. Results are mean6s.e.m.;*P,0.05, **P,0.001. (D) Confocal images ofcontrol, VAMP7 and SYT7 KD cells untreated ortreated for 3 hours with 100 mM ZnCl2 and loadedwith FluoZin-3,AM (green) and LysoTracker (red).Scale bars: 20 mm.

Fig. 8. Inhibition of lysosomal exocytosis through VAMP7 KD increasescell death in Zn2+-treated cells. Flow cytometry data of AnnV- and PI-stained control and VAMP7-KD cells treated for 48 hours with 100 mM ZnCl2,showing increased Annexin V fluorescence in VAMP7- Zn2+ treated cells(gray solid line) compared to control- Zn2+ treated cells (solid black line).

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and SYT7 siRNA transfected cells had similar levels of secretion

to control cells (71.70%63.09). To our knowledge, this is thefirst evidence linking increased Zn2+ secretion by TFEB andlysosomal exocytosis.

DISCUSSIONAlthough lysosomes are most commonly discussed in terms of

their role in endocytic digestion and absorption, mountingevidence points towards their role in cell death and in signaling(Settembre et al., 2013). Recent evidence indicates that underoxidative stress conditions, Zn2+ accumulates in the lysosomes

and can lead to LMP (Hwang et al., 2008). Furthermore, this Zn2+

dysregulation is mechanistically linked to autophagy (Lee et al.,2009; Yoon et al., 2010; Cho et al., 2012). Considering that

autophagy and oxidative stress play a crucial role in cell deathand neurodegenerative pathologies, it is likely that Zn2+

represents a new target for modulating diseases and a key step

in understanding neuronal cell death. These recent developmentsemphasize the role of lysosomes in metal toxicity. Thus, theaccumulation of Zn2+ and other metals in lysosomes has beenshown to lead to LMP, to the release of lysosomal enzymes into

the cytoplasm, and to cell death. Although pathological aspects ofZn2+ buildup in the lysosomes are undisputable, its physiologicalrole is unclear.

Our recent data suggest that lysosomes play a role of a ‘Zn2+

sink’, working in parallel with the transcriptional regulation ofZn2+ chelation and transport proteins. Cytoplasmic Zn2+ is

gauged by the transcription factor MTF-1, which, upon Zn2+

binding, induces transcription of Zn2+ chelators, such asmetallothioneins, and Zn2+ transporters, such as ZnT1. The

ability to absorb Zn2+ through active transport involving ZnTtransporters makes lysosomes a good candidate for absorbingrapid cytoplasmic Zn2+ spikes. The recently published data on thelysosomal absorption of Zn2+ released from metallothioneins by

H2O2 highlight such a function of lysosomes (Tang et al., 2002;Suntres and Lui, 2006; Hwang et al., 2008; Lee et al., 2009).

Although ZIP transporters such as ZIP8 have been shown to

exist in the lysosomes (Aydemir et al., 2009), their impact on thefunction of the lysosomal Zn2+ sink has not been shown. At thesame time, the ion channel TRPML1, whose dysfunction causes

the lysosomal storage disease mucolipidosis IV (MLIV)(Slaugenhaupt et al., 1999), has been shown to be a component

of the lysosomal Zn2+ transport machinery. Its loss has been

shown to cause Zn2+ buildup in the lysosomes in studies by twogroups (Eichelsdoerfer et al., 2010; Kukic et al., 2013) and itspermeability to Zn2+ by another group (Dong et al., 2010). It

seems reasonable to conclude that TRPML1 is involved intrafficking Zn2+ from lysosomes into the cytoplasm. We proposedthat upon entering the cell, Zn2+ is registered by MTF-1, which

leads to an eventual increase in transcription and translation ofZn2+ chelators and exporters. However, Zn2+ must be rapidlyeliminated from the cytoplasm in order to protect against toxicity.Thus, in parallel, Zn2+ is scavenged by the lysosomes, to be later

released through a TRPML1-dependent mechanism.The data in Fig. 6D, Fig. 7C and Fig. 9B show that lysosomal

secretion is also involved in Zn2+ clearance from the cell. What is

the relationship between the two mechanisms of Zn2+ clearance?We think that their function reflects their ability to respond tochanges in Zn2+ flow. Such changes can be caused by increased

autophagy of Zn2+-rich organelles such as mitochondria orproteins. Interestingly, TRPML1 is upregulated in response toincreased endocytic load in a manner that requires the transcriptionfactor TFEB, whereas VAMP7 remains unchanged (I.K.,

unpublished observation). Based on this, we propose thatTRPML1-driven Zn2+ release is a dynamic response to increasedZn2+ load, whereas VAMP7- and SYT7-dependent secretion is a

constitutive mechanism. We believe that our data clearly show thatZn2+ absorption into the lysosomes, followed by its secretion, is animportant detoxification mechanism. Furthermore, the fact that

suppression of the lysosomal function causes Zn2+ redistributioninto Golgi and mitochondria shows that the lysosomal Zn2+ sinkhas a major impact on the cellular Zn2+ handling.

LMP following Zn2+ exposure has clearly been shown to be acell death pathway (Hwang et al., 2008; Chung et al., 2009; Leeet al., 2009; Hwang et al., 2010). Why would cells employ a Zn2+

detoxification mechanism that effectively kills them? We think

that the answer to this question lies in the relation between thelysosomal Zn2+ absorption rates and the rate of its secretion andrelease. Such a ratio might depend on the tissue, developmental

stage and other factors. We propose that under normal conditions,or upon moderate Zn2+ exposure, the Zn2+ sink limits cytoplasmicZn2+ by absorbing it (model in Fig. 5). Zn2+ is later released into

the cytoplasm (as discussed above), or secreted across the plasmamembrane. However, during the exposure to high (200 mM)

Fig. 9. Enhancing lysosomal exocytosisthough TFEB overexpression increases Zn2+

secretion. (A) b-hexosaminidase activity assayfor lysosomal exocytosis. Results are shown asthe percentage of total b-hexosaminidasesecreted over 10 or 30 minutes; 1 mM ionomycinwas used to stimulate lysosomal exocytosis.(B) Zn2+ secretion assay using cell-impermeantFluoZin-3 tetrapotassium salt. Results are shownas the percentage of the maximum fluorescencevalue recorded in the control+Zn2+ samples at30 minutes, which were taken as 100%. Theresults shown above are after 15 minutes ofsecretion time (chase). Results are mean6s.e.m.;*P,0.05; **P,0.001.

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levels of Zn2+, Zn2+ extraction lags, leading to LMP and celldeath (supplementary material Fig. S3). Our data show a role of

lysosomes in transition metal toxicity and identify a noveldetoxification mechanism.

MATERIALS AND METHODSCell cultureHeLa cells were maintained in Dulbecco’s modified Eagle’s medium

(DMEM; Sigma-Aldrich, St Louis, MO) supplemented with 10% FBS.

For siRNA and cDNA transfection, medium was changed after 24 hours.

100 mM ZnCl2 was added directly to the medium, containing serum,

24 hours after transfection, for the indicated times. Bafilomycin A1

(#196000, EMD MIllipore, Darmstadt, Germany) was used at 1 mM for

the indicated amount of time.

siRNA-mediated KD and plasmid transfectionThe VAMP7 siRNA (cat. number SASI_Hs01_00197188), SYT7 (cat.

number SASI_Hs01_0047888), ZnT2 siRNA (Cat number SASI_

Hs01_00055662), ZnT4 siRNA (cat. number SASI_Hs00225995), and

MTF-1 siRNA (cat. number SASI_Hs01_00177112) were from Sigma-

Aldrich. Non-targeting control siRNA#1 (Sigma) was used as a negative

control. Cells were transfected using Lipofectamine 2000 (Invitrogen,

Carlsbag, CA) as described by the manufacturer’s protocol using 600 nM

siRNA per 35-mm well (1200 nM siRNA per 35-mm well for efficient

VAMP7 and SYT7 KD). All KDs were confirmed using SYBR-green

based qPCR. For DNA transfections, 2 mg of GalT–mCherry and TFEB

was transfected per 35-mm dish.

MicroscopyCells were seeded on glass coverslips and loaded with dyes for

15 minutes at 37 C in buffer containing, in mM: 150 NaCl, 5 KCl, 1

CaCl2, 1 MgCl2, 10 HEPES pH 7.4, 1 g/l glucose. The loading was

followed by a 15-minute washout in all cases. Lysotracker Red DND-99

and FluoZin-3,AM (F-24195, Invitrogen, Carlsbag, CA) were used at

0.1 mM. Confocal microscopy was performed using Leica TCS SP5 and

BioRad 3000 confocal microscopes. Live cells were treated as above.

Images were analyzed using ImageJ (Bethesda, MD).

Reverse transcriptase and quantitative qPCRRNA was isolated from cells using Trizol (Invitrogen, Carlsbag, CA)

according to the manufacturer’s protocol. cDNA was synthesized using

the GeneAmp RNA PCR system (Applied Biosystems, Carlsbad, CA)

with oligo(dT) priming. qPCR was performed using SYBR green

(Fermentas, Glen Burnie, MD). The amount of cDNA loaded was

normalized to starting RNA concentrations, with a final concentration of

9 ng (40 ng in ZnT experiments) of RNA loaded per experimental well.

Six-point standard curves were generated for each primer using 1:2

dilutions of cDNA. cDNA for the following genes were amplified using

the indicated primers (IDT, Coralville, IA). MT2a, forward,

59AAGTCCCAGCGAACCCGCGT-39, reverse, 59-CAGCAGCTGCAC-

TTGTCCGACGC-39; VAMP7, forward, 59-CCGGACAGACTGAAGC-

CAT-39, reverse, 59-ATCTGCTCTGTCACCTCCAG-39; SYT7, forward,

59-AAGCGGGTGGAGAAGAAGAA-39, reverse, 59-CGAAGGCGAA-

GGACTCATTG-39; ZnT1 (SLC30A1), forward, 59-GGGAGCAGCGA-

CATCAACGT-39, reverse, 59-GGGTCTGCGGGGTCCAATT-39; ZnT2

(SLC30A2), forward, 59-GCAATCCGGTCATACACGGGAT-39, reverse,

59-CAGCTCAATGGCCTGCAAGT-3; ZnT4 (SLC30A4), forward, 59-

CACATACAGCTAATTCCTGGAAGTTCATCT-39, reverse, 59-GCCT-

GTAACTCTGAAGCTGAATAGTACAT-39; and LAMP1, forward, 59-

GGACAACACGACGGTGACAAG-39, reverse, 59-GAACTTGCATTC-

ATCCCGAACTGGA-39. All primers were designed to span exons and

reverse-transcriptase-negative controls were tested to ensure amplification

of cDNA only. qPCR was performed using the standard curve method on

the 7300 Real Time System (Applied Biosystems, Carlsbad, CA).

Reactions were run on the following parameters: 2 minutes at 50 C,

10 minutes at 95 C, and 40 cycles at 95 C for 15 seconds followed by 60 C

for 1 minute. All experimental samples were run in triplicate and

normalized to an RPL32 endogenous control.

b-Hexosamindase activity assayUntreated control and transfected cells were washed with 37 C PBS, and

300 ml 37 C PBS with 1 mM CaCl2 was added to each 35-mm dish. For

each sample, 100 ml of the supernatant was incubated with 300 ml 3 mM

4-nitrophenyl N-acetyl-b-D-glucosaminide (N9376, Sigma-Aldrich) for

30 minutes at 37 C in 0.1 M citrate buffer (pH 4.5) (C2488, Sigma-

Aldrich). This volume was replaced with fresh 100 ml of 37 C of PBS

with 1 mM CaCl2 to the culture dish. Samples were collected every after

0, 10 and 30 minutes. Reactions were stopped by adding 650 ml borate

buffer (100 mM boric acid, 75 mM NaCl, 25 mM sodium borate pH 9.8)

and the absorbance was measured in a spectrophotometer at 405 nm. To

determine total cellular content of b-hexosamindase, cells were lysed

with 300 ml 1% Triton X-100 in PBS and, after a 10,000 g spin for

5 minutes at 4 C, 10 ml of the cell extracts were used for the enzyme

activity reaction. Enzyme activity was determined as the amount of 4-

nitrophenol produced per mg of protein (Bradford method). Absorbance

was calibrated with different amounts of 4-nitrophenol (N7660, Sigma-

Aldrich) in 0.1 M citrate buffer.

Zinc secretion assayCells were plated on a 12-well plate and 48 hours after, transfection

pulsed with 100 mM ZnCl2 for 3 hours, washed twice with warm PBS,

and chased in 1 ml DMEM per well. Duplicate time-points were

collected for 0, 5, 15, 30 or 60 minutes and replaced with new 50 ml of

DMEM. For each sample, 50 ml of supernatant was placed in a 96-well

plate. Zn2+ content was measured by incubating the supernatants with

10 mM cell-impermeant FluoZin-3 tetrapotassium salt (F-24194,

Molecular Probes, Invitrogen, Carlsbag, CA) for 15 minutes at 37 C.

The 96-well plate was read using a FujiFilm FLA-5100 fluorescent image

analyzer. After the last time point, cells were washed with PBS, 200 ml

detergent solution was added to lyse the cells and fluorescence was

normalized to total protein in each sample.

Caspase-3 activity assayCells were prepared and measured using the EnzChek Caspase-3 Assay

Kit #1 (E13183, Invitrogen, Carlsbag, CA) following the manufacturer’s

instructions. AMC substrate fluorescence was measured using a

fluorometer at an excitation wavelength of 342 nm and an emission

wavelength of 441 nm.

Western blot analysisCells were solubilized for 10 min at room temperature in either a 16detergent solution (0.5 M EDTA, pH 8.0, 1 M Tris, pH 8.0, 0.4%

deoxycholate, 1% Nonidet P-40 substitute) for (LAMP1) or a 1% CHAPS

in PBS solution (for VAMP7) containing protease inhibitor mixture III

(Calbiochem, Gibbstown, NJ) and centrifuged at 16,000 g for 5 minutes.

The supernatant was collected and protein concentrations were

determined using a Bradford assay. Protein was incubated at 100 C for

5 minutes in sample buffer containing 14% b-mercaptoethanol. Equal

amounts of protein were loaded on a 10% precast Tris-HCl

polyacrylamide gel (Bio-Rad) for each experimental sample. Proteins

were transferred onto PVDF membrane (EMD Millipore, Darmstadt,

Germany) and blocked in 10% nonfat dried milk for 1 hour. The

following primary antibodies were used: monoclonal LAMP1 (sc-20011,

Santa Cruz, Santa Cruz, CA) at 1:1000 dilution, monoclonal VAMP7

(ab36195, Abcam, Cambridge, US) at 1:500, and monoclonal b-actin

(ab6276, Abcam, Cambridge, US) at 1:5000 dilution. Horseradish

peroxidase (HRP)-conjugated goat anti-mouse Ig secondary antibodies

(Amersham Biosciences) were used at 1:20,000 dilutions.

Immunodetection was performed with the Luminata Forte HRP

substrate (EMD Millipore, Darmstadt, Germany). Band densities were

measured using ImageJ (Bethesda, MD).

Flow cytometryTransfected HeLa cells were treated with either 100 mM ZnCl2 for

48 hours for Baf and VAMP7 experiments, or 200 mM ZnCl2 for

12 hours for TFEB experiments. Cells were then washed with PBS,

trypsinized, and counted. 26106 cells were pelleted for each sample,

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washed with PBS and then resuspended in Annexin Binding Buffer from

the Vybrant Apoptosis Assay Kit #3 (V.13242, Molecular Probes). Cells

were then loaded with 1 ml propidium iodide and 2 ml Annexin V 488

and sorted on the Accuri (BD) C6 at the University of Pittsburgh Cancer

Institute Cytometry Facility. Cell sorting was gated to include healthy

and apoptotic cells while excluding debris.

StatisticsStatistical significance was calculated using a one-tailed, unpaired

Student’s t-test with P#0.05 considered significant. Data are presented

as mean6s.e.m.

AcknowledgementsThis project used the University of Pittsburgh Cancer Institute Cytometry Facilitythat is supported in part by award P30CA047904. We thank Michael Meyer andBratislav Janjic for help with flow cytometry data analysis. The GalT mcherryconstruct was kindly provided by Ora Weisz and the TFEB construct was kindlyprovided by Rosa Puertollano.

Competing interestsThe authors declare no competing interests.

Author contributionsI.K. conceptualized, designed, executed and interpreted the data, and preparedthe article. S.L.K. conceptualized and interpreted the data. K.K. conceptualized,designed and interpreted the data, and prepared the article.

FundingThis work was supported by National Institutes of Health [grant numbersHD058577 and ES01678 to K.K.]. Deposited in PMC for release after 12 months.

Supplementary materialSupplementary material available online athttp://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.145318/-/DC1

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