ß 2014. Published by The Company of Biologists Ltd | Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
RESEARCH ARTICLE
Zn2+ efflux through lysosomal exocytosis prevents Zn2+-induced
toxicity
ABSTRACT
2+
Zn 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
INTRODUCTION
Cellular 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 its
influx, export and chelation. In general, Zn2+ transport is
regulated by ZnT (also known as solute carrier family 30) and
ZIP (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 by
the dedicated ZnT transporters such as ZnT6 (SLC30A6) for the
1
The Department of Biological Sciences, University of Pittsburgh, Pittsburgh, PA
15260, USA. 2The Department of Nutritional Sciences, College of Health and
Human Development, The Pennsylvania State University, University Park, PA
16802, 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 ([email protected])
Received 25 October 2013; Accepted 25 April 2014
3094
Golgi (Huang et al., 2002), and ZnT2 (SLC30A2) and ZnT4
(SLC30A4) for the lysosome (Palmiter et al., 1996; Huang and
Gitschier, 1997; Falcón-Pérez and Dell’Angelica, 2007;
McCormick and Kelleher, 2012). This organellar Zn2+ export
lowers potentially toxic cytoplasmic Zn2+ concentrations in
pathophysiological 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 acid
sphingomyelinase (Schissel et al., 1996), and for the secretion of
Zn2+ 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, translation
and 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 well
as 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 the
role of lysosomes in protection against Zn2+-induced toxicity.
Zn2+ is transported from the cytoplasm into lysosomes by
ZnT2 and ZnT4 (Palmiter et al., 1996; Huang and Gitschier,
1997; Falcón-Pérez 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 the
release of the lysosomal enzymes such as cathepsins and to cell
death (Hwang et al., 2008; Chung et al., 2009; Lee et al., 2009;
Hwang et al., 2010). As such, the lysosomal Zn2+ accumulation
might constitute a cell death mechanism during normal
remodeling of Zn2+-rich tissues, such as the mammary gland
(Kelleher et al., 2011), as well as in pathological conditions. With
this in mind, we sought to answer whether or not accumulation of
Zn2+ in the lysosome is the terminal depot for cellular Zn2+.
Alternatively, it is possible that lysosomal Zn2+ dissipates and
lysosomes constitute only a temporary Zn2+ storage site. Our
recently published data suggest that the lysosomal ion channel
transient receptor potential mucolipin 1 (TRPML1, also known as
mucolipin1, encoded by the MCOLN1 gene) is at least partly
responsible 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 SNARE
complex, which includes the VAMP7 protein and synaptotagmin
VII (SYT7) (Martinez-Arca et al., 2000; Braun et al., 2004; Rao
et al., 2004; Logan et al., 2006; Mollinedo et al., 2006). It has
Journal of Cell Science
Ira Kukic1, Shannon L. Kelleher2,3,4 and Kirill Kiselyov1,*
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
Fig. 1. Inhibition of lysosomal Zn2+ sink
function by Baf increases cytoplasmic Zn2+
levels. (A) Confocal images of HeLa cells treated
with 100 mM ZnCl2 and/or 1 mM Baf for 3 hours
then loaded with FluoZin-3,AM and LysoTracker.
Note disappearance of LysoTracker staining,
indicative of lysosomal deacidification, and an
increase in FluoZin-3,AM staining intensity,
indicative of increased Zn2+. (B) Confocal images
of control or ZnT siRNA transfected HeLa cells
(48 hours post-transfection) treated with 100 mM
ZnCl2 for 3 hours then loaded with FluoZin-3,AM.
Images represent at least three separate
experiments and at least three images per
condition in each experiment. Scale bars: 20 mm.
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 used
VAMP7 and SYT7 knockdown (KD) to suppress lysosomal
secretion and assess its role in Zn2+ clearance from the cells.
Here, we aimed to establish the functional context of the
lysosomal Zn2+ accumulation. Our findings indicate that
lysosomes actively absorb Zn2+ and secrete it across the plasma
membrane, given that suppressing the lysosomal Zn2+ absorption
or secretion causes Zn2+ buildup in the cytoplasm, Golgi and
mitochondria, leading to apoptosis.
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
In order to test the role of the lysosomal Zn2+ sink on cellular
Zn2+ 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 livecell confocal microscopy and FluoZin-3,AM as described
previously (Kukic et al., 2013) Fig. 1A shows that the exposure
of Baf-treated cells to Zn2+ caused a significantly higher FluoZin3,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, or
FluoZin-3,AM fluorescence, the magnitude of the observed
effects appear to be incompatible with the quantitative
estimates of changes induced by Baf. Thus, the effect of Baf on
cytoplasmic pH appears to be small, within only tenths of pH
units (Heming et al., 1995). The degree of pH change necessary to
cause an effect on Zn2+ handling, by contrast, significantly
exceeds that reported to be caused by Baf. A pH drop below 6.7 is
required to trigger an increase in intracellular Zn2+ according to
one set of studies (Kiedrowski, 2012), whereas another set has
shown that metallothioneins release Zn2+ only after cytoplasmic
pH drops below 5.0 (Jiang et al., 2000). Thus, the increase in
cytoplasmic Zn2+ caused by Baf likely correlates with the loss of
lysosomal function, rather than cytosolic pH changes.
We have previously shown that Zn2+ transporters ZnT2 and
ZnT4 colocalize with the lysosomal ion channel TRPML1 in
Fig. 2. Inhibition of lysosomal Zn2+ sink function by Baf increases the
transcriptional response of Zn2+-responsive genes. qRT-PCR results of
MT2a (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 with
either DMSO or 1 mM Baf alone or with 100 mM ZnCl2. Results are
mean6s.e.m.; *P,0.05; **P,0.001.
3095
Journal of Cell Science
RESULTS
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
KD using siRNA as described previously, and tested the resulting
changes in Zn2+ handling using FluoZin-3,AM. Fig. 1B shows
that ZnT2 and ZnT4 KD increased cytoplasmic Zn2+ levels
observed in these cells after a 3-hour long treatment with 100 mM
ZnCl2. These results are in agreement with the previously
Fig. 4. Inhibition of lysosomal function leads to increased cell death
upon high Zn2+ exposure. (A) Caspase-3 (Cas3) activity assay showing
increased Cas3 activation upon a 48-hour 100 mM ZnCl2 and 1 mM Baf
treatment in HeLa cells. A 3-hour long exposure to 1 mM staurosporine was
used as a positive control, which increased Cas3 activity by
796.72%6109.06 (n53, P,0.001). Cas3 activity is shown as AMC
fluorescence and a percentage of untreated controls. Results are
mean6s.e.m.; *P,0.05. (B) Flow cytometry data showing increased AnnV
staining of cells treated with 100 mM ZnCl2 and/or 1 mM Baf for 48 hours.
Data represent three experiments.
3096
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 was
used 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 ZnT1
mRNA 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 cells
treated with DMSO and Zn2+ (n54; P,0.001), and
1174.61%694.20 (mean6s.e.m., n54; P,0.05 relative to
DMSO-only, and DMSO plus Zn2+ controls) in cells treated
with Baf and Zn2+ (Fig. 2A). ZnT1 mRNA increased to
323.43%623.32 (n54; P,0.001) in cells treated with DMSO
plus Zn2+, and to 552.09%640.46 (n54; P,0.05 relative to
DMSO-only, and DMSO plus Zn2+ controls) of control values in
cells treated with Baf plus Zn2+ (Fig. 2B). In addition to
confirming the elevated cytoplasmic Zn2+ levels due to Baf, as
assessed by FluoZin-3,AM in Fig. 1A, these data also corroborate
MTF-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 intracellular
inclusions was noted in Baf-treated cells exposed to Zn2+. At
least some of these inclusions were positive for the Golgi marker
GalT–mCherry (Fig. 3A). Under these conditions, Zn2+ also
accumulated in the mitochondria, which was shown using the
Journal of Cell Science
Fig. 3. Inhibition of lysosomal Zn2+ sink function by Baf
redistributes cellular Zn2+ pools to the Golgi and the
mitochondria. (A) Confocal images of GalT–mCherrytransfected HeLa cells treated with 100 mM ZnCl2 and/or 1 mM
Baf for 3 hours, then loaded with FluoZin-3,AM. The black-andwhite panes show overlap between the green and the red
channels, obtained using the RG2B function of ImageJ.
(B) Confocal images of HeLa cells treated with 100 mM ZnCl2
and/or 1 mM Baf for 3 hours, then loaded with RhodZin-3,AM.
Plot profiles on the right show intensity profiles of RhodZin-3,AM
fluorescence recorded along the lines indicated in the
corresponding image. Note increased RhodZin-3,AM
florescence with Baf-treated cells. Images represent at least
three separate experiments and at least three images per
condition in each experiment. Scale bars: 20 mm.
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
mitochondrial Zn2+ dye RhodZin-3,AM, whose signal was
brighter 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 cytoplasmic
Zn2+ 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 to
its buildup in the cytoplasm and in the organelles (Medvedeva
et al., 2009). If the lysosomes are a Zn2+-buffering sink, then
inhibiting that function should result in cell death. Our findings in
Fig. 4A support this: although HeLa cells were fairly resistant
to the effects of 100 mM ZnCl2 or 1 mM Baf alone, their
combination caused pronounced caspase-3 (Cas3) activation,
indicative of apoptosis. An exposure of cells to 100 mM ZnCl2
for 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). A
combination of Zn2+- and Baf-exposure increased Cas3 activity
by 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+ sink
in Baf-treated cells.
As a complimentary cell death assay, Annexin V (AnnV) and
Propidium Iodide (PI) staining were analyzed using flow
cytometry. Fig. 4B and supplementary material Fig. S1 show an
increase in the number of AnnV-positive, apoptotic cells when
cells are treated for 48 hours with Zn2+ plus Baf. Taken together,
these data show that suppression of the lysosomal Zn2+ sink
facilitates the apoptotic cell death caused by Zn2+ exposure. They
suggest that lysosomes play a crucial role in buffering
cytoplasmic Zn2+ and in its detoxification. A loss of such a role
exposes cells to the pro-apoptotic effects of Zn2+.
Zn2+ toxicity has been linked to LMP, resulting in cell death
under 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 antiapoptotic effects of the Zn2+ sink is dictated by the rate of Zn2+
absorption by the lysosomes and/or the rate of its clearance from
the lysosomes. If the rate of Zn2+ clearance exceeds the rate of
its sequestration into the lysosomes, then the Zn2+ sink is
cytoprotective. A rate of sequestration exceeding the rate of
clearance 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)], or
through its secretion mediated by lysosomal fusion with the
plasma membrane. The latter has recently gained a lot of attention
as 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 of
experiments tested the role of lysosomes in Zn2+ secretion.
We suppressed lysosomal secretion by using siRNAs against
two 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 VAMP7
mRNA to 27.76%63.99 of control siRNA levels (mean6s.e.m.,
n54, P,0.001), whereas Fig. 7A shows that SYT7 siRNA
reduced SYT7 mRNA to 30.12%64.11 of control siRNA levels
(n53, P,0.001). VAMP7 mRNA levels were not altered by
either a 3- or 48-hour exposure to 100 mM ZnCl2 (I. K.,
unpublished observation; SYT7 dependence on Zn2+ was not
tested). VAMP7 KD was confirmed by examining protein levels
through western blotting confirming that VAMP7 siRNA reduced
VAMP7 protein expression to 30.21%66.82 of control siRNA
levels (n55, P,0.001) (Fig. 6B). VAMP7 and SYT7 KD were
associated with changes in the lysosomal numbers and
organization. There was an increase in total lysosomal numbers
in VAMP7 KD cells (supplementary material Fig. S2), although
there 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
3097
Journal of Cell Science
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 and
ZIPs and is (1) recognized by the Zn2+-sensitive transcription factor MTF-1 that, once Zn2+-bound, translocates to the nucleus to upregulate ZnT1 and MT2a
gene 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 is
then secreted across the plasma membrane (3) through a VAMP7-dependent mechanism. This prevents toxic Zn2+ buildup in the cytoplasm and other
organelles (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 other
organelles (6), such as the Golgi and mitochondria.
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
SNARE proteins in membrane traffic. Examples of such
clustering 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 bhexosaminidase (b-hex), a common marker of lysosomal
exocytosis. In these experiments, lysosomal secretion was
initiated by treatment with 1 mM of the Ca2+ ionophore
ionomycin (Ion) (Fig. 6C; Fig. 7B). After 30 minutes, control
KD cells secreted 18.80%60.66 of their total b-hex content
without stimulation, whereas VAMP7 KD cells secreted
14.09%61.60 of their total b-hex content (mean6s.e.m.,
Fig. 6C). Once stimulated with ionomycin, however, control
KD 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, SYT7
KD was also functionally significant, decreasing lysosomal
exocytosis compared to control KD. Fig. 7B shows that after
3098
30 minutes, ionomycin-treated control KD cells secreted
35.57%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 on
Zn2+ secretion, we incubated control and VAMP7, or SYT7 KD
cells with 100 mM ZnCl2 for 3 hours. Next, cells were placed in
normal medium (no Zn2+) and Zn2+ secretion was analyzed using
FluoZin-3 fluorescence as described previously (Kukic et al.,
2013). Both VAMP7 (Fig. 6D) and SYT7 (Fig. 7C) KD
significantly reduced Zn2+ secretion. Within 30 minutes of
secretion, VAMP7 KD cells were only able to secrete
51.79%619.26 (n53, P,0.05) of the amount of Zn2+ secreted
by the control KD cells (100%) (Fig. 6D), indicating that VAMP7
is necessary for Zn2+ secretion. Similarly, SYT7 KD cells were
only able to secrete 58.9062.87% (n53, P,0.05) of the amount
of Zn2+ secreted by the control KD cells at 30 minutes, which
was taken as 100% (Fig. 7C). This inhibition of Zn2+ secretion in
Journal of Cell Science
Fig. 6. Inhibition of lysosomal exocytosis through
VAMP7 KD inhibits Zn2+ secretion. (A) qRT-PCR results
confirming VAMP7 KD by siRNA. (B) Quantification of
western blot results confirming VAMP7 KD. Insert shows a
representative blot (of five) of endogenous VAMP7 and
actin levels under with either control or VAMP7 siRNA.
(C) b-hexosaminidase activity assay for lysosomal
exocytosis. Results are shown as the percentage of total
b-hexosaminidase secreted after either 10 or 30 minutes;
1 mM ionomycin was used to stimulate lysosomal
exocytosis. (D) Zn2+ secretion assay using cellimpermeant FluoZin-3 tetrapotassium salt. Results are
shown as the percentage of the maximum fluorescence
value recorded in the control+Zn2+ samples after
30 minutes, which were set at 100%. Results are
mean6s.e.m.; *P,0.05, **P,0.001.
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
VAMP7 and SYT7 KD cells was also corroborated by the
elevated cellular Zn2+ levels, as seen by assessing FluoZin-3,AM
staining using confocal microscopy (Fig. 7D).
Fig. 8. Inhibition of lysosomal exocytosis through VAMP7 KD increases
cell death in Zn2+-treated cells. Flow cytometry data of AnnV- and PIstained 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).
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 of
Zn2+-induced cell death in VAMP7 KD cells treated with 100 mM
ZnCl2 for 48 hours, compared to control KD cells undergoing the
same treatment (Fig. 8; supplementary material Fig. S1). In
summary, the data described here show that the Zn2+ sink
integrates Zn2+ absorption from the cytoplasm with its secretion
through 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 lysosomal
Zn2+ 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 after
30 minutes, mock (empty vector)-transfected cells secreted
13.95%60.67 of their total b-hex content, while TFEBtransfected cells secreted 23.11%63.75 (mean6s.e.m.; n53,
P,0.05) of their total b-hex content. Once stimulated with
ionomycin, 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-hex
content. TFEB also increased Zn2+ secretion. Fig. 9B shows that
after 15 minutes, control KD cells were able to secrete
76.70%61.37 of secretable Zn2+ in our assay, whereas TFEBtransfected 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
3099
Journal of Cell Science
Fig. 7. Inhibition of lysosomal exocytosis
through SYT7 KD inhibits Zn2+ secretion.
(A) qRT-PCR results confirming SYT7 KD by
siRNA. (B) b-hexosaminidase activity assay for
lysosomal exocytosis. Results are shown as the
percentage of total b-hexosaminidase secreted
over 10 or 30 minutes; 1 mM ionomycin was used
to stimulate lysosomal exocytosis. (C) Zn2+
secretion assay using cell-impermeant FluoZin-3
tetrapotassium salt. Results are shown as percent
of the maximum fluorescence value recorded in
the control+Zn2+ samples at 30 minutes, which
were taken as 100%. Results are mean6s.e.m.;
*P,0.05, **P,0.001. (D) Confocal images of
control, VAMP7 and SYT7 KD cells untreated or
treated for 3 hours with 100 mM ZnCl2 and loaded
with FluoZin-3,AM (green) and LysoTracker (red).
Scale bars: 20 mm.
RESEARCH ARTICLE
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
and SYT7 siRNA transfected cells had similar levels of secretion
to control cells (71.70%63.09). To our knowledge, this is the
first evidence linking increased Zn2+ secretion by TFEB and
lysosomal exocytosis.
DISCUSSION
Although lysosomes are most commonly discussed in terms of
their role in endocytic digestion and absorption, mounting
evidence points towards their role in cell death and in signaling
(Settembre et al., 2013). Recent evidence indicates that under
oxidative 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 death
and 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 developments
emphasize the role of lysosomes in metal toxicity. Thus, the
accumulation of Zn2+ and other metals in lysosomes has been
shown to lead to LMP, to the release of lysosomal enzymes into
the cytoplasm, and to cell death. Although pathological aspects of
Zn2+ buildup in the lysosomes are undisputable, its physiological
role is unclear.
Our recent data suggest that lysosomes play a role of a ‘Zn2+
sink’, working in parallel with the transcriptional regulation of
Zn2+ chelation and transport proteins. Cytoplasmic Zn2+ is
gauged by the transcription factor MTF-1, which, upon Zn2+
binding, induces transcription of Zn2+ chelators, such as
metallothioneins, and Zn2+ transporters, such as ZnT1. The
ability to absorb Zn2+ through active transport involving ZnT
transporters makes lysosomes a good candidate for absorbing
rapid cytoplasmic Zn2+ spikes. The recently published data on the
lysosomal 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 the
function of the lysosomal Zn2+ sink has not been shown. At the
same 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
3100
of the lysosomal Zn2+ transport machinery. Its loss has been
shown to cause Zn2+ buildup in the lysosomes in studies by two
groups (Eichelsdoerfer et al., 2010; Kukic et al., 2013) and its
permeability to Zn2+ by another group (Dong et al., 2010). It
seems reasonable to conclude that TRPML1 is involved in
trafficking Zn2+ from lysosomes into the cytoplasm. We proposed
that upon entering the cell, Zn2+ is registered by MTF-1, which
leads to an eventual increase in transcription and translation of
Zn2+ chelators and exporters. However, Zn2+ must be rapidly
eliminated 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 to
changes in Zn2+ flow. Such changes can be caused by increased
autophagy of Zn2+-rich organelles such as mitochondria or
proteins. Interestingly, TRPML1 is upregulated in response to
increased endocytic load in a manner that requires the transcription
factor TFEB, whereas VAMP7 remains unchanged (I.K.,
unpublished observation). Based on this, we propose that
TRPML1-driven Zn2+ release is a dynamic response to increased
Zn2+ load, whereas VAMP7- and SYT7-dependent secretion is a
constitutive mechanism. We believe that our data clearly show that
Zn2+ absorption into the lysosomes, followed by its secretion, is an
important detoxification mechanism. Furthermore, the fact that
suppression of the lysosomal function causes Zn2+ redistribution
into Golgi and mitochondria shows that the lysosomal Zn2+ sink
has a major impact on the cellular Zn2+ handling.
LMP following Zn2+ exposure has clearly been shown to be a
cell death pathway (Hwang et al., 2008; Chung et al., 2009; Lee
et 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 the
lysosomal Zn2+ absorption rates and the rate of its secretion and
release. 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 cytoplasmic
Zn2+ by absorbing it (model in Fig. 5). Zn2+ is later released into
the cytoplasm (as discussed above), or secreted across the plasma
membrane. However, during the exposure to high (200 mM)
Journal of Cell Science
Fig. 9. Enhancing lysosomal exocytosis
though TFEB overexpression increases Zn2+
secretion. (A) b-hexosaminidase activity assay
for lysosomal exocytosis. Results are shown as
the percentage of total b-hexosaminidase
secreted over 10 or 30 minutes; 1 mM ionomycin
was used to stimulate lysosomal exocytosis.
(B) Zn2+ secretion assay using cell-impermeant
FluoZin-3 tetrapotassium salt. Results are shown
as the percentage of the maximum fluorescence
value recorded in the control+Zn2+ samples at
30 minutes, which were taken as 100%. The
results shown above are after 15 minutes of
secretion time (chase). Results are mean6s.e.m.;
*P,0.05; **P,0.001.
levels of Zn2+, Zn2+ extraction lags, leading to LMP and cell
death (supplementary material Fig. S3). Our data show a role of
lysosomes in transition metal toxicity and identify a novel
detoxification mechanism.
MATERIALS AND METHODS
Cell culture
HeLa 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 transfection
The 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 SigmaAldrich. 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.
Microscopy
Cells 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 qPCR
RNA 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-CAGCAGCTGCACTTGTCCGACGC-39; VAMP7, forward, 59-CCGGACAGACTGAAGCCAT-39, reverse, 59-ATCTGCTCTGTCACCTCCAG-39; SYT7, forward,
59-AAGCGGGTGGAGAAGAAGAA-39, reverse, 59-CGAAGGCGAAGGACTCATTG-39; ZnT1 (SLC30A1), forward, 59-GGGAGCAGCGACATCAACGT-39, reverse, 59-GGGTCTGCGGGGTCCAATT-39; ZnT2
(SLC30A2), forward, 59-GCAATCCGGTCATACACGGGAT-39, reverse,
59-CAGCTCAATGGCCTGCAAGT-3; ZnT4 (SLC30A4), forward, 59CACATACAGCTAATTCCTGGAAGTTCATCT-39, reverse, 59-GCCTGTAACTCTGAAGCTGAATAGTACAT-39; and LAMP1, forward, 59GGACAACACGACGGTGACAAG-39, reverse, 59-GAACTTGCATTCATCCCGAACTGGA-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.
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
b-Hexosamindase activity assay
Untreated 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, SigmaAldrich). 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 4nitrophenol produced per mg of protein (Bradford method). Absorbance
was calibrated with different amounts of 4-nitrophenol (N7660, SigmaAldrich) in 0.1 M citrate buffer.
Zinc secretion assay
Cells 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 assay
Cells 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 analysis
Cells were solubilized for 10 min at room temperature in either a 16
detergent 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 cytometry
Transfected 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,
3101
Journal of Cell Science
RESEARCH ARTICLE
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.
Statistics
Statistical 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.
Acknowledgements
This project used the University of Pittsburgh Cancer Institute Cytometry Facility
that is supported in part by award P30CA047904. We thank Michael Meyer and
Bratislav Janjic for help with flow cytometry data analysis. The GalT mcherry
construct was kindly provided by Ora Weisz and the TFEB construct was kindly
provided by Rosa Puertollano.
Competing interests
The authors declare no competing interests.
Author contributions
I.K. conceptualized, designed, executed and interpreted the data, and prepared
the article. S.L.K. conceptualized and interpreted the data. K.K. conceptualized,
designed and interpreted the data, and prepared the article.
Funding
This work was supported by National Institutes of Health [grant numbers
HD058577 and ES01678 to K.K.]. Deposited in PMC for release after 12 months.
Supplementary material
Supplementary material available online at
http://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.145318/-/DC1
References
Andrews, G. K. (2001). Cellular zinc sensors: MTF-1 regulation of gene
expression. Biometals 14, 223-237.
Aydemir, T. B., Liuzzi, J. P., McClellan, S. and Cousins, R. J. (2009). Zinc
transporter ZIP8 (SLC39A8) and zinc influence IFN-gamma expression in
activated human T cells. J. Leukoc. Biol. 86, 337-348.
Braun, V., Fraisier, V., Raposo, G., Hurbain, I., Sibarita, J. B., Chavrier, P.,
Galli, T. and Niedergang, F. (2004). TI-VAMP/VAMP7 is required for optimal
phagocytosis of opsonised particles in macrophages. EMBO J. 23, 4166-4176.
Chao, Y. and Fu, D. (2004). Kinetic study of the antiport mechanism of an
Escherichia coli zinc transporter, ZitB. J. Biol. Chem. 279, 12043-12050.
Cho, K. S., Yoon, Y. H., Choi, J. A., Lee, S. J. and Koh, J. Y. (2012). Induction of
autophagy and cell death by tamoxifen in cultured retinal pigment epithelial and
photoreceptor cells. Invest. Ophthalmol. Vis. Sci. 53, 5344-5353.
Chung, H., Yoon, Y. H., Hwang, J. J., Cho, K. S., Koh, J. Y. and Kim, J. G.
(2009). Ethambutol-induced toxicity is mediated by zinc and lysosomal
membrane permeabilization in cultured retinal cells. Toxicol. Appl. Pharmacol.
235, 163-170.
Decressac, M., Mattsson, B., Weikop, P., Lundblad, M., Jakobsson, J. and
Björklund, A. (2013). TFEB-mediated autophagy rescues midbrain dopamine
neurons from a-synuclein toxicity. Proc. Natl. Acad. Sci. USA 110, E1817E1826.
Dong, X. P., Shen, D., Wang, X., Dawson, T., Li, X., Zhang, Q., Cheng, X.,
Zhang, Y., Weisman, L. S., Delling, M. et al. (2010). PI(3,5)P(2) controls
membrane trafficking by direct activation of mucolipin Ca(2+) release channels
in the endolysosome. Nat. Commun. 1, 38.
Eichelsdoerfer, J. L., Evans, J. A., Slaugenhaupt, S. A. and Cuajungco, M. P.
(2010). Zinc dyshomeostasis is linked with the loss of mucolipidosis IVassociated TRPML1 ion channel. J. Biol. Chem. 285, 34304-34308.
Falcón-Pérez, J. M. and Dell’Angelica, E. C. (2007). Zinc transporter 2
(SLC30A2) can suppress the vesicular zinc defect of adaptor protein 3depleted fibroblasts by promoting zinc accumulation in lysosomes. Exp. Cell
Res. 313, 1473-1483.
Flannery, A. R., Czibener, C. and Andrews, N. W. (2010). Palmitoylationdependent association with CD63 targets the Ca2+ sensor synaptotagmin VII to
lysosomes. J. Cell Biol. 191, 599-613.
Forsleff, L., Schauss, A. G., Bier, I. D. and Stuart, S. (1999). Evidence of
functional zinc deficiency in Parkinson’s disease. J. Altern. Complement. Med.
5, 57-64.
Fraldi, A., Annunziata, F., Lombardi, A., Kaiser, H. J., Medina, D. L.,
Spampanato, C., Fedele, A. O., Polishchuk, R., Sorrentino, N. C., Simons,
K. et al. (2010). Lysosomal fusion and SNARE function are impaired by
cholesterol accumulation in lysosomal storage disorders. EMBO J. 29, 36073620.
3102
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
Frederickson, C. J. and Bush, A. I. (2001). Synaptically released zinc:
physiological functions and pathological effects. Biometals 14, 353-366.
Heming, T. A., Traber, D. L., Hinder, F. and Bidani, A. (1995). Effects of
bafilomycin A1 on cytosolic pH of sheep alveolar and peritoneal macrophages:
evaluation of the pH-regulatory role of plasma membrane V-ATPases. J. Exp.
Biol. 198, 1711-1715.
Huang, L. and Gitschier, J. (1997). A novel gene involved in zinc transport is
deficient in the lethal milk mouse. Nat. Genet. 17, 292-297.
Huang, L., Kirschke, C. P. and Gitschier, J. (2002). Functional characterization
of a novel mammalian zinc transporter, ZnT6. J. Biol. Chem. 277, 2638926395.
Hwang, J. J., Lee, S. J., Kim, T. Y., Cho, J. H. and Koh, J. Y. (2008). Zinc and 4hydroxy-2-nonenal mediate lysosomal membrane permeabilization induced by
H2O2 in cultured hippocampal neurons. J. Neurosci. 28, 3114-3122.
Hwang, J. J., Kim, H. N., Kim, J., Cho, D. H., Kim, M. J., Kim, Y. S., Kim, Y.,
Park, S. J. and Koh, J. Y. (2010). Zinc(II) ion mediates tamoxifen-induced
autophagy and cell death in MCF-7 breast cancer cell line. Biometals 23, 9971013.
Jansen, J., Karges, W. and Rink, L. (2009). Zinc and diabetes – clinical links and
molecular mechanisms. J. Nutr. Biochem. 20, 399-417.
Jiang, L. J., Vasák, M., Vallee, B. L. and Maret, W. (2000). Zinc transfer
potentials of the alpha- and beta-clusters of metallothionein are affected by
domain interactions in the whole molecule. Proc. Natl. Acad. Sci. USA 97, 25032508.
Kanninen, K. M., Grubman, A., Caragounis, A., Duncan, C., Parker, S. J.,
Lidgerwood, G. E., Volitakis, I., Ganio, G., Crouch, P. J. and White, A. R.
(2013). Altered biometal homeostasis is associated with CLN6 mRNA loss in
mouse neuronal ceroid lipofuscinosis. Biol. Open 2, 635-646.
Kelleher, S. L., Seo, Y. A. and Lopez, V. (2009). Mammary gland zinc
metabolism: regulation and dysregulation. Genes Nutr. 4, 83-94.
Kelleher, S. L., McCormick, N. H., Velasquez, V. and Lopez, V. (2011). Zinc in
specialized secretory tissues: roles in the pancreas, prostate, and mammary
gland. Adv. Nutr. 2, 101-111.
Kiedrowski, L. (2012). Cytosolic acidification and intracellular zinc release in
hippocampal neurons. J. Neurochem. 121, 438-450.
Kukic, I., Lee, J. K., Coblentz, J., Kelleher, S. L. and Kiselyov, K. (2013). Zincdependent lysosomal enlargement in TRPML1-deficient cells involves MTF-1
transcription factor and ZnT4 (Slc30a4) transporter. Biochem. J. 451, 155163.
Lee, S. J. and Koh, J. Y. (2010). Roles of zinc and metallothionein-3 in oxidative
stress-induced lysosomal dysfunction, cell death, and autophagy in neurons and
astrocytes. Mol. Brain 3, 30.
Lee, J. Y., Cole, T. B., Palmiter, R. D., Suh, S. W. and Koh, J. Y. (2002).
Contribution by synaptic zinc to the gender-disparate plaque formation in human
Swedish mutant APP transgenic mice. Proc. Natl. Acad. Sci. USA 99, 77057710.
Lee, S. J., Cho, K. S. and Koh, J. Y. (2009). Oxidative injury triggers autophagy in
astrocytes: the role of endogenous zinc. Glia 57, 1351-1361.
Logan, M. R., Lacy, P., Odemuyiwa, S. O., Steward, M., Davoine, F., Kita, H.
and Moqbel, R. (2006). A critical role for vesicle-associated membrane protein7 in exocytosis from human eosinophils and neutrophils. Allergy 61, 777784.
Lopez, V., Foolad, F. and Kelleher, S. L. (2011). ZnT2-overexpression represses
the cytotoxic effects of zinc hyper-accumulation in malignant metallothioneinnull T47D breast tumor cells. Cancer Lett. 304, 41-51.
Martina, J. A., Diab, H. I., Lishu, L., Jeong-A, L., Patange, S., Raben, N. and
Puertollano, R. (2014). The nutrient-responsive transcription factor TFE3
promotes autophagy, lysosomal biogenesis, and clearance of cellular debris.
Sci. Signal. 7, ra9.
Martinez-Arca, S., Alberts, P., Zahraoui, A., Louvard, D. and Galli, T. (2000).
Role of tetanus neurotoxin insensitive vesicle-associated membrane protein (TIVAMP) in vesicular transport mediating neurite outgrowth. J. Cell Biol. 149, 889900.
McCormick, N. H. and Kelleher, S. L. (2012). ZnT4 provides zinc to zincdependent proteins in the trans-Golgi network critical for cell function and Zn
export in mammary epithelial cells. Am. J. Physiol. 303, C291-C297.
Medina, D. L., Fraldi, A., Bouche, V., Annunziata, F., Mansueto, G.,
Spampanato, C., Puri, C., Pignata, A., Martina, J. A., Sardiello, M. et al.
(2011). Transcriptional activation of lysosomal exocytosis promotes cellular
clearance. Dev. Cell 21, 421-430.
Medvedeva, Y. V., Lin, B., Shuttleworth, C. W. and Weiss, J. H. (2009).
Intracellular Zn2+ accumulation contributes to synaptic failure, mitochondrial
depolarization, and cell death in an acute slice oxygen-glucose deprivation
model of ischemia. J. Neurosci. 29, 1105-1114.
Mollinedo, F., Calafat, J., Janssen, H., Martı́n-Martı́n, B., Canchado, J.,
Nabokina, S. M. and Gajate, C. (2006). Combinatorial SNARE complexes
modulate the secretion of cytoplasmic granules in human neutrophils.
J. Immunol. 177, 2831-2841.
Ohana, E., Hoch, E., Keasar, C., Kambe, T., Yifrach, O., Hershfinkel, M. and
Sekler, I. (2009). Identification of the Zn2+ binding site and mode of operation of
a mammalian Zn2+ transporter. J. Biol. Chem. 284, 17677-17686.
Palmieri, M., Impey, S., Kang, H., di Ronza, A., Pelz, C., Sardiello, M. and
Ballabio, A. (2011). Characterization of the CLEAR network reveals an
integrated control of cellular clearance pathways. Hum. Mol. Genet. 20, 38523866.
Journal of Cell Science
RESEARCH ARTICLE
Palmiter, R. D. and Findley, S. D. (1995). Cloning and functional characterization of a
mammalian zinc transporter that confers resistance to zinc. EMBO J. 14, 639-649.
Palmiter, R. D., Cole, T. B. and Findley, S. D. (1996). ZnT-2, a mammalian
protein that confers resistance to zinc by facilitating vesicular sequestration.
EMBO J. 15, 1784-1791.
Pastore, N., Ballabio, A. and Brunetti-Pierri, N. (2013). Autophagy master
regulator TFEB induces clearance of toxic SERPINA1/a-1-antitrypsin polymers.
Autophagy 9, 1094-1096.
Prasad, A. S. (2013). Discovery of human zinc deficiency: its impact on human
health and disease. Adv. Nutr. 4, 176-190.
Rao, S. K., Huynh, C., Proux-Gillardeaux, V., Galli, T. and Andrews, N. W.
(2004). Identification of SNAREs involved in synaptotagmin VII-regulated
lysosomal exocytosis. J. Biol. Chem. 279, 20471-20479.
Rink, L. and Gabriel, P. (2000). Zinc and the immune system. Proc. Nutr. Soc. 59,
541-552.
Rulon, L. L., Robertson, J. D., Lovell, M. A., Deibel, M. A., Ehmann, W. D. and
Markesber, W. R. (2000). Serum zinc levels and Alzheimer’s disease. Biol.
Trace Elem. Res. 75, 79-85.
Sardiello, M., Palmieri, M., di Ronza, A., Medina, D. L., Valenza, M., Gennarino,
V. A., Di Malta, C., Donaudy, F., Embrione, V., Polishchuk, R. S. et al. (2009).
A gene network regulating lysosomal biogenesis and function. Science 325,
473-477.
Saydam, N., Adams, T. K., Steiner, F., Schaffner, W. and Freedman, J. H.
(2002). Regulation of metallothionein transcription by the metal-responsive
transcription factor MTF-1: identification of signal transduction cascades that
control metal-inducible transcription. J. Biol. Chem. 277, 20438-20445.
Journal of Cell Science (2014) 127, 3094–3103 doi:10.1242/jcs.145318
Schissel, S. L., Schuchman, E. H., Williams, K. J. and Tabas, I. (1996). Zn2+stimulated sphingomyelinase is secreted by many cell types and is a product of
the acid sphingomyelinase gene. J. Biol. Chem. 271, 18431-18436.
Settembre, C., Fraldi, A., Medina, D. L. and Ballabio, A. (2013). Signals from the
lysosome: a control centre for cellular clearance and energy metabolism. Nat.
Rev. Mol. Cell Biol. 14, 283-296.
Slaugenhaupt, S. A., Acierno, J. S., Jr, Helbling, L. A., Bove, C., Goldin, E.,
Bach, G., Schiffmann, R. and Gusella, J. F. (1999). Mapping of the
mucolipidosis type IV gene to chromosome 19p and definition of founder
haplotypes. Am. J. Hum. Genet. 65, 773-778.
Suntres, Z. E. and Lui, E. M. (2006). Antioxidant effect of zinc and zincmetallothionein in the acute cytotoxicity of hydrogen peroxide in Ehrlich ascites
tumour cells. Chem. Biol. Interact. 162, 11-23.
Tang, Z. L., Wasserloos, K. J., Liu, X., Stitt, M. S., Reynolds, I. J., Pitt, B. R.
and St Croix, C. M. (2002). Nitric oxide decreases the sensitivity of pulmonary
endothelial cells to LPS-induced apoptosis in a zinc-dependent fashion. Mol.
Cell. Biochem. 234-235, 211-217.
Vinceti, M., Bergomi, M., Nacci, G., Pietrini, V., Ferrari, A., Fortini, K., Guidetti,
D., Sola, P., Rocchi, E., Mancia, D. et al. (2002). Erythrocyte zinc, copper, and
copper/zinc superoxide dismutase and risk of sporadic amyotrophic lateral
sclerosis: a population-based case-control study. Amyotroph. Lateral Scler.
Other Motor Neuron Disord. 3, 208-214.
Yoon, Y. H., Cho, K. S., Hwang, J. J., Lee, S. J., Choi, J. A. and Koh, J. Y.
(2010). Induction of lysosomal dilatation, arrested autophagy, and cell death by
chloroquine in cultured ARPE-19 cells. Invest. Ophthalmol. Vis. Sci. 51, 60306037.
Journal of Cell Science
RESEARCH ARTICLE
3103