(2015) Lysosome Electrophysiology
(2015) Lysosome Electrophysiology
(2015) Lysosome Electrophysiology
10
Lysosome
electrophysiology
CHAPTER OUTLINE
Introduction ............................................................................................................ 198
1. Lysosome........................................................................................................... 198
1.1 Lysosome Ion Channels ....................................................................... 198
1.2 Methods for Studying Lysosomal Ion Channels....................................... 200
1.2.1 Methods to study lysosomal channel localization ............................... 200
1.2.2 Methods to study lysosomal Ca2 channels....................................... 201
1.2.3 Studying lysosomal channels in plasma membrane or in artificial
membranes using patch clamping .................................................... 202
1.2.4 Study of lysosomal channels in lysosomes using lysosome
patch clamping ................................................................................ 202
2. Materials........................................................................................................... 203
2.1 Cell Culture ........................................................................................ 203
2.2 Pipettes ............................................................................................. 203
2.3 Chemicals .......................................................................................... 204
2.4 Lysosome Patch-Clamp Recording ........................................................ 204
3. Methods ............................................................................................................ 204
3.1 Cell Culture ........................................................................................ 204
3.2 Pipettes and Solutions......................................................................... 204
3.3 Lysosome Patch-Clamp Recording ........................................................ 206
3.3.1 Isolation of enlarged lysosomes ......................................................... 206
3.3.2 Whole-lysosome patch clamping ....................................................... 206
3.3.3 Other patch configurations................................................................ 208
4. Discussion ......................................................................................................... 210
5. Summary ........................................................................................................... 211
Acknowledgments ................................................................................................... 211
References ............................................................................................................. 211
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Abstract
The physiology and functions of ion channels have been major topics of interest in
biomedical research. Patch clamping is one of the most powerful techniques used in the
study of ion channels and has been widely applied to the investigation of electrical
properties of ion channels on the plasma membrane in a variety of cells. A number of ion
channels have been found in intracellular lysosomal membranes. However, their properties had been difficult to study due to the lack of a direct patch-clamping methodology
on lysosomal membranes. Past attempts to record lysosomal channels that were forced to
express on the plasma membrane or reconstituted into lipid bilayers have largely
generated inconclusive and conflicting results. Recently, a novel lysosome patchclamping technique has been developed, making it possible to examine lysosomal
channels under near physiological conditions. This chapter provides a detailed description of this technique, which has been successfully applied in several studies concerning
lysosomal ion channels. This technique will expand our understanding of the nature of
lysosomes and lysosome-related diseases.
INTRODUCTION
1. LYSOSOME
Lysosomes are specialized acidic intracellular organelles containing acid hydrolases
that are capable of breaking down macromolecules. The organelles act as waste
disposal systems of the cell by digesting materials that are taken up either from
the extracellular environment through endocytosis/phagocytosis, or from intracellular components of the cell through autophagy. Deficiency in lysosomal acid hydrolases has been associated with a group of inherited metabolic disorders termed
lysosomal storage diseases (Lloyd-Evans & Platt, 2011; Luzio et al., 2000; Luzio,
Pryor, & Bright, 2007).
1. Lysosome
2010; Shen, Wang, & Xu, 2011), transient receptor potential melastatin 2 (TRPM2)
(Lange et al., 2009; Sumoza-Toledo et al., 2011), P2X4 purinoceptor (Huang et al.,
2014; Qureshi, Paramasivam, Yu, & Murrell-Lagnado, 2007), two-pore channel 1
(TPC1) (Brailoiu et al., 2009; Cang et al., 2014), TPC2 (Calcraft et al., 2009;
Cang et al., 2013; Wang et al., 2012), and ClC chloride channels (Cl/H
exchanger) (Graves, Curran, Smith, & Mindell, 2008; Jentsch, 2007; Weinert
et al., 2010) (Figure 1). Interestingly, in addition to lysosomal enzymes, deficiency
in lysosomal ion homeostasis and ion transport has also been associated with lysosomal storage diseases (Dong et al., 2008; Lloyd-Evans et al., 2008).
TRPML1: TRPML proteins belong to the TRP family (Nilius, Owsianik, Voets, &
Peters, 2007; Ramsey, Delling, & Clapham, 2006). They form a family of intracellular
channels primarily localized in endosomes and lysosomes. The predicted structure of
TRPML proteins includes six transmembrane domains and a putative pore region, similar
to that of voltage-gated channels (Nilius et al., 2007; Ramsey et al., 2006). Mutations in
the human TRPML1 gene cause mucolipidosis type IV disease (ML4), a devastating pediatric neurodegenerative disease with motor impairment, mental retardation, and irondeficiency anemia (Bassi et al., 2000; Dong et al., 2008; Sun et al., 2000). Recently,
TRPML1 was demonstrated to be a lysosomal nonselective cation channel, with significant Ca2 and Fe2 permeabilities (Bach, 2005). Impaired TRPML1-mediated
Ca2/Fe2 release from lysosomes may underlie ML4 phenotypes (Dong et al., 2008).
TRPM2: TRPM2 is another member of the TRP family (Nilius et al., 2007;
Ramsey et al., 2006). It also displays a transmembrane topology similar to that of
voltage-gated channels. TRPM2 has been shown to function as a lysosomal Ca2release channel activated by intracellular adenosine diphosphateeribose in
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pancreatic b-cells (Lange et al., 2009) and dendritic cells (Sumoza-Toledo et al.,
2011). It may play important roles in hydrogen peroxide-induced b cell death and
dendritic cell maturation and chemotaxis.
P2X4: P2X4 receptor belongs to the purinergic receptor family. It opens in
response to adenosine triphosphate (ATP) binding at the extracytosolic side (Khakh
& North, 2012). In addition to its actions on the plasma membrane, a recent study
suggests that P2X4 is also localized in lysosomal membranes (Qureshi et al.,
2007). Lysosomal P2X4 can cycle from the lysosome to phagosome or to the plasma
membrane in response to a variety of stimuli. We recently demonstrated that lysosomal P2X4 is minimally activated at acidic luminal pH. However, alkalization of
lysosome dramatically increases P2X4 channel activity, which may contribute to
lysosomal membrane trafficking (Huang et al., 2014).
TPCs: TPC1 and TPC2 are cation-selective ion channels with two repeats of a
six-transmembrane-domain module. They were proposed to mediate lysosomal
Ca2 release triggered by the second messenger, nicotinic acid adenine dinucleotide
phosphate (Calcraft et al., 2009; Lloyd-Evans, Waller-Evans, Peterneva, & Platt,
2010). By directly performing patch-clamping recordings in enlarged lysosomes,
Xus group at the University of Michigan and others have suggested that TPC1
and TPC2 are in fact highly Na-selective channels with very limited Ca2 permeability (Cang et al., 2013, 2014; Wang et al., 2012).
ClCs: ClCs Cl channels (Cl/H exchangers) have functions both on the
plasma membrane (ClC-1, -2, -Ka, -Kb) and on intracellular membranes of the
endocytotic-lysosomal pathway (ClC3 through ClC7). Plasma membrane ClC channels are known to play a role in the stabilization of membrane potential, transepithelial transport, and cell volume regulation, whereas endosomal/lysosomal ClC
channels are thought to provide an electric shunt for the efficient pumping of the
H-ATPase. Because ClC3eClC7 primarily reside on the membranes of intracellular organelles, their electrophysiological properties and modulations are much
less clear. Most recently, ClC3, ClC4, ClC5, and ClC7 were proposed to be antiporters with a coupling transport ratio of 2 Cl:1 H, rather than ion channels (Accardi
& Miller, 2004; Graves et al., 2008; Jentsch, 2007; Weinert et al., 2010).
1. Lysosome
and because fluorescent proteins could potentially affect the localization of endogenous proteins (Kim, Soyombo, Tjon-Kon-Sang, So, & Muallem, 2009; Song, Dayalu, Matthews, & Scharenberg, 2006; Venkatachalam, Hofmann, & Montell, 2006),
additional approaches are needed to validate the results. Immunostaining is often
employed to examine protein localization without interference by heterologous
overexpression. For example, endogenous P2X4 has been detected in lysosomes
by immunofluorescent staining (Huang et al., 2014; Qureshi et al., 2007).
Cellular fractionation provides a separation of homogeneous organelles from total
cell lysates by using centrifugation at controlled speeds (Huang et al., 2014; Wang
et al., 2012). With the help of specific antibodies, lysosomal ion channel proteins
were detected in the lysosomal-associated membrane protein 1 (Lamp1) positive
heavy fractions by immunoblotting (Huang et al., 2014; Wang et al., 2012; Zeevi,
Frumkin, Offen-Glasner, Kogot-Levin, & Bach, 2009). This can be used to validate
the use of fluorescent fusion proteins in the heterologous systems and immunostaining
of endogenous proteins for studying subcellular localization of lysosome channels.
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2. Materials
2. MATERIALS
2.1 CELL CULTURE
1. Dulbeccos Modified Eagles Medium (DMEM)/F-12 medium (11330, Gibco,
Life Technologies)
2. Fetal bovine serum (FBS) (26140, Gibco, Life Technologies)
3. Trypsineethylenediaminetetraacetic acid (0.05%; 25300, Gibco, Life Technologies)
4. Opti-MEM (31985, Gibco, Life Technologies)
5. Lipofectamine 2000 (11668, Life Technologies)
6. Poly-L-lysine (0.01%; P4832, Sigma)
7. Cell culture dishes (35 mm; 353001, Falcon, Thomas Scientific)
8. Cell culture plates with 24 wells (142475, Nunc, Thomas Scientific)
9. Glass coverslips (12 mm; 121313, Fisher Scientific)
2.2 PIPETTES
1.
2.
3.
4.
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2.3 CHEMICALS
All drugs are obtained from Sigma except for those indicated below.
1. Vacuolin-1 (sc-216,045, Santa Cruz Biotechnology)
2. ML-SA1 (4746, Tocris Bioscience)
3. METHODS
3.1 CELL CULTURE
Cells are maintained in DMEM/F-12 medium (DMEM/Nutrient Mixture F-12) supplemented with 10% FBS at 37 C in a 5% CO2 incubator. Cells are transfected at a
density of approximately 80% confluency using Lipofectamine 2000 as per the vendors instructions. To monitor the expression, enhanced green fluorescent protein is
fused to mouse full-length TRPML1 at the N-terminus. At 4e6 h after transfection,
cells are trypsinized and replated onto 12-mm glass coverslips in 24-well culture
plates. The coverslips are precoated with 0.01% poly-L-lysine overnight, rinsed
with water, and air dried prior to use.
Vacuolin-1 (5 mM) stock solution is prepared by adding 1 mg of vacuolin-1 to
465 mL of dimethyl sulfoxide. The vacuolin-1 stock is mixed, divided into 50-mL aliquots in sterilized tubes, and stored in dark at 20 C. The vacuolin-1 stock is
diluted to 1 mM with DMEM/F-12 culture medium before use. Cells are plated
onto coverslips for approximately 2e4 h, and then treated with vacuolin-1 (1 mM)
for >2 h prior to performing patch-clamp recordings.
3. Methods
visual control using a microforge. Fire polishing allows the pipette to form a narrow
tip opening with rounded edges. The polished pipettes typically have a resistance of
approximately 8e13 MU when filled with the pipette solution.
Preparation of pipette and bath solutions depends on the patch-clamp configuration. It is suggested that the environment of lysosome lumen is similar to extracellular space (Wang et al., 2012). For whole-lysosome recording, the pipette
solution (a modified Tyrodes solution), which mimics a typical extracellular environment bathes the luminal surface of isolated enlarged lysosomes; the bath solution which mimics intracellular environment bathes the cytosolic side of the
isolated enlarged lysosomes (Figure 2). The components of bath and pipette solutions also vary with the objectives of the experiments. With respect to TRPML1 recordings, the bath (internal/cytoplasmic) solution contains 140 mM K-gluconate,
4 mM NaCl, 2 mM MgCl2, 1 mM ethylene glycol tetraacetic acid (EGTA),
0.39 mM CaCl2 (free [Ca2]i equals to 100 nM), and 20 mM 4-(2-hydroxyethyl)1-piperazineethanesulfonic acid (HEPES), with the pH adjusted to 7.2 by KOH
and osmolality adjusted to approximately 290 mOsm by sucrose. The pipette
(luminal) solution contains 145 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM
MgCl2, 10 mM glucose, and 20 mM HEPES, with the pH adjusted to 4.6 (to mimic
the acidic environment of lysosomes) by HCl and osmolality adjusted to approximately 310 mOsm by sucrose.
The pipette solution is filtered through a 0.45-mm (diameter) filter. Before
recording, the tip of the pipette is dipped into the pipette solution to avoid bubbles, and then the pipette is backfilled with the pipette solution using a microfill
needle to half full. The remaining bubbles are removed by gently flicking the
pipette.
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3. Methods
Once a whole-lysosome configuration is established, a designed voltage protocol is applied to record the channel of interest. Figure 4(B) shows representative
IeV curves of whole-lysosome currents measured from Cos-1 cells expressing
TRPML1. Currents are elicited by repeated voltage ramps of 400-ms duration
between 140 mV (relative to the lumen which is set at 0 mV) and 140 mV
every 4 s. The small basal TRPML1 currents are significantly enhanced by the
bath perfusion of 10 mM ML-SA1. Figure 4(C) shows the time course of TRPML1
currents measured at 140 mV in response to ML-SA1 stimulation. The inward
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current at negative potentials indicates an efflux of cations moving from the lumen
of lysosomes to the cytosol due to the opening of TRPML1 (Figure 2).
Further, followed by the establishment of whole-lysosome mode, lysosomal
membrane potential can be measured using the current-clamp recording mode
(Cang et al., 2013). Given that the lysosomal membrane potential (Vm) is defined
as Vcytosol Vlumen (Vlumen 0 mV) (Bertl et al., 1992), opening of TRPML1 results in an increase in Vm, that is, Vlumen becomes more negative. Figure 4(D) shows
that the ML SA1-induced activation of TRPML1 (Figure 4(B) and (C)) is accompanied by a depolarization of the lysosome membrane expressing TRPML1.
3. Methods
when the pipette is sealed onto the isolated enlarged lysosomes without breaking
into the vacuolar membrane. The luminal-side-out mode is achieved by quickly
withdrawing the pipette from the enlarged lysosomes after forming the
lysosome-attached mode. Therefore, the luminal surface of the enlarged lysosomes
is exposed to the bath solution. Figure 5 shows representative IeV curves of
TRPML1Va (a gain-of-function mutant) currents under lysosome-attached and
luminal-side-out configurations (Dong et al., 2008). Switching from lysosomeattached to luminal-side-out modes induces a decrease in the amplitude of the
currents.
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4. DISCUSSION
Lysosome patch clamping has been a powerful technique to study lysosomal ion
channels. However, the mechanisms of action of vacuolin-1 are still not clear. The
membrane components in the enlarged lysosomes induced by vacuolin-1 could be
different from bona fide lysosomes in intact cells. One concern of this technique
is that vacuolin-1 treatment may affect the channel properties. Given that enlarged
lysosomes are also present in a very small number of nontreated cells, the channel
properties of enlarged lysosomes obtained from cells untreated and treated with
vacuolin-1 were compared. As for TRPML1 (Dong et al., 2008, 2010), TPC1
(Cang et al., 2013; Wang et al., 2012), and P2X4 (Huang et al., 2014), no significant
difference in channel properties was detected for enlarged lysosomes obtained with
or without vacuolin-1 treatment. However, the possibility of a change in properties
induced by vacuolin-1 for other lysosomal ion channels cannot be excluded.
Notably, the lysosome recording is performed on isolated lysosomes. Although
the membrane of lysosomes is intact, the cytosolic environment is altered when
the lysosome is isolated. The loss of cytosolic regulatory factors associated with
lysosomal membranes could be one problem for studying the regulation of lysosomal channels. In this case, regulatory factors should be considered to be included
in the system when doing lysosome patch clamping. For instance, PI(3,5)P2 (an
endolysosome specific PIP2) has been found to be required for the activation of
TRPML1 (Dong et al., 2010) and TPC currents (Cang et al., 2013; Dong et al.,
2010). In addition, cytosolic ATP has been shown to regulate TPC2 currents
(Cang et al., 2013). Similarly, some factors in the lumen should also be taken into
consideration, such as ATP (Huang et al., 2014).
The development of lysosome patch clamping has made it easier to identify
novel lysosome channels (Cang et al., 2014) and to characterize known ones. For
instance, by using this technique, lysosomal membranes have been shown to be
permeable to other ions including Na, K, and Cl (Cang et al., 2013), and a number of lysosomal channels have been well characterized, including TRPML1 (Dong
et al., 2008, 2010), TPC2 (Cang et al., 2013; Wang et al., 2012), and P2X4 (Huang
et al., 2014). However, the regulation of these channels remains largely unclear. We
believe that lysosome patch clamping in combination with other methods may provide a complete insight into the regulation of lysosomal ion channels. Taken TPC2,
for example, it has been shown to be regulated by mammalian target of rapamycin
(mTOR) and be involved in the nutrient-sensing mTOR pathway (Cang et al., 2013).
On the other hand, this technique also represents a unique approach to validate potential drugs that target lysosome channels, which helps find new therapeutic strategies for lysosomal ion channel diseases.
In principle, this technique may be modified for recording other lysosomerelated organelles such as endosomes, phagosomes, autophagosomes, melanosomes,
lytic granules, and many other secretory granules. Indeed, Xus group has successfully recorded the TRPML1 current in phagosomes (Samie et al., 2013). Although
the approach has limitations, it provides a unique method to measure ion transport
References
across lysosomal membranes and allows the characterization of ion channels in lysosomes and lysosome-related organelles.
5. SUMMARY
Similar to the studies of lysosomal enzymes, the study of lysosomal ion transport is
an important aspect in our understanding of lysosomal functions. With the advancement of lysosome patch clamping that allows the direct measurement of lysosomal
channels in their native environment, we expect that more lysosome ion channels
and their regulatory mechanisms will be elucidated in the near future. Since deficiency in lysosomal membrane ion channels and dyshomeostasis of lysosomal
ions have been implicated in a group of lysosomal storage diseases (Cheng et al.,
2010; Lloyd-Evans et al., 2008; Weinert et al., 2010) and classical neurodegenerative diseases (e.g., Alzheimers Disease) (Coen et al., 2012), we believe that this
technical advance will dramatically improve our understanding of basic lysosome
physiology, and their implications in lysosome-related diseases.
ACKNOWLEDGMENTS
Work in the Dong laboratory is funded by DMRF, CIHR grant (MOP-119349), NSHRF Establishment Grant (MED-PRO-2011-7485), and CFI Leaders Opportunity Fund-Funding for
research infrastructure (29291).
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