2.3 Chemicals .......................................................................................... 204

2.4 Lysosome Patch-Clamp Recording ........................................................ 204

3. Methods ............................................................................................................ 204

3.3.1 Isolation of enlarged lysosomes..............................

3.3.2 Whole-lysosome patch clamping ............................

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CHAPTER............................. 2113.3.3 Other patch configurations...................................

4. Discussion............................................................................

5. Summary ..............................................................................

Acknowledgments ......................................................................

References ................................................................................Methods in Cell Biology, Volume 126, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.10

2015 Elsevier Inc. All rights reserved.......................... 206

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........................... 2113.1 Cell Culture ........................................................................................ 204

3.2 Pipettes and Solutions......................................................................... 204

3.3 Lysosome Patch-Clamp Recording ...............................Lysosomeelectrophysiology 10

Xi Z. Zhong, Xian-Ping Dong1

Department of Physiology and Biophysics, Dalhousie University, Halifax, Nova Scotia, Canada1Corresponding author: E-mail: xpdong@dal.ca

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....................................... 2011.2.3 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 ............................................................................................. 203.022 197

lysosomal ion channels. This technique will expand our understanding of the nature of

lysosomal storage diseases (Lloyd-Evans & Platt, 2011; Luzio et al., 2000; Luzio,

198 CHAPTER 10 Lysosome electrophysiologyPryor, & Bright, 2007).

1.1 LYSOSOME ION CHANNELSAn important feature of the lysosome is an acidic luminal pH (pHw4e5) that en-sures lysosomal hydrolases to function properly. The acidic luminal pH is estab-lished by the vacuolar type H-ATPase, a well-studied H transporter present onlysosomal membranes (Lloyd-Evans & Platt, 2011; Luzio et al., 2000; Luzio, Pryor,et al., 2007; Mindell, 2012). Although H transport has been the most extensivelystudied ion movement across lysosomal membranes, recent studies have also indi-cated that lysosomal membranes are permeable to many other ions, includingNa, K, and Cl (Cang et al., 2013; Cang, Bekele, & Ren, 2014). Advances inmodern cell biology and physiological techniques, together with classical geneticand biochemical approaches, have allowed us to identify a plethora of ion transportproteins in lysosomal membranes (Figure 1), including transient receptor potentialmucolipin 1 (TRPML1) (Cheng, Shen, Samie, & Xu, 2010; Dong et al., 2008,lysosomes and lysosome-related diseases.

INTRODUCTION

1. LYSOSOMELysosomes are specialized acidic intracellular organelles containing acid hydrolasesthat are capable of breaking down macromolecules. The organelles act as wastedisposal systems of the cell by digesting materials that are taken up either fromthe extracellular environment through endocytosis/phagocytosis, or from intracel-lular components of the cell through autophagy. Deficiency in lysosomal acid hydro-lases has been associated with a group of inherited metabolic disorders termedAbstractThe physiology and functions of ion channels have been major topics of interest inbiomedical research. Patch clamping is one of the most powerful techniques used in thestudy of ion channels and has been widely applied to the investigation of electricalproperties of ion channels on the plasma membrane in a variety of cells. A number of ionchannels have been found in intracellular lysosomal membranes. However, their prop-erties had been difficult to study due to the lack of a direct patch-clamping methodologyon lysosomal membranes. Past attempts to record lysosomal channels that were forced toexpress on the plasma membrane or reconstituted into lipid bilayers have largelygenerated inconclusive and conflicting results. Recently, a novel lysosome patch-clamping technique has been developed, making it possible to examine lysosomalchannels under near physiological conditions. This chapter provides a detailed descrip-tion of this technique, which has been successfully applied in several studies concerning

1. Lysosome 1992010; 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

FIGURE 1 Ion channels and transporters on lysosome membranes.

The currently known ion channels and transporters on lysosome membranes are listed.

TRPML1, transient receptor potential mucolipin 1; TRPM2, transient receptor potential

melastatin 2; P2X4, purinergic P2X receptor subtype 4; TPC1, two pore channel 1; TPC2, two

pore channel 2; ClC, ClC family of chloride channels (Cl/H exchanger); H-ATPase,proton-pump ATPase.(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; Weinertet al., 2010) (Figure 1). Interestingly, in addition to lysosomal enzymes, deficiencyin lysosomal ion homeostasis and ion transport has also been associated with lyso-somal 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 intracellularchannels primarily localized in endosomes and lysosomes. The predicted structure ofTRPML proteins includes six transmembrane domains and a putative pore region, similarto that of voltage-gated channels (Nilius et al., 2007; Ramsey et al., 2006). Mutations inthe human TRPML1 gene cause mucolipidosis type IV disease (ML4), a devastating pe-diatric neurodegenerative disease with motor impairment, mental retardation, and iron-deficiency 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 signif-icant Ca2 and Fe2 permeabilities (Bach, 2005). Impaired TRPML1-mediatedCa2/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 ofvoltage-gated channels. TRPM2 has been shown to function as a lysosomal Ca2-release channel activated by intracellular adenosine diphosphateeribose in

200 CHAPTER 10 Lysosome electrophysiologylysosomal 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 lysosomalCa2 release triggered by the second messenger, nicotinic acid adenine dinucleotidephosphate (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 TPC1and TPC2 are in fact highly Na-selective channels with very limited Ca2 perme-ability (Cang et al., 2013, 2014; Wang et al., 2012).

ClCs: ClCs Cl channels (Cl/H exchangers) have functions both on theplasma membrane (ClC-1, -2, -Ka, -Kb) and on intracellular membranes of theendocytotic-lysosomal pathway (ClC3 through ClC7). Plasma membrane ClC chan-nels are known to play a role in the stabilization of membrane potential, transepithe-lial transport, and cell volume regulation, whereas endosomal/lysosomal ClCchannels are thought to provide an electric shunt for the efficient pumping of theH-ATPase. Because ClC3eClC7 primarily reside on the membranes of intracel-lular organelles, their electrophysiological properties and modulations are muchless clear. Most recently, ClC3, ClC4, ClC5, and ClC7 were proposed to be antiport-ers 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).

1.2 METHODS FOR STUDYING LYSOSOMAL ION CHANNELS1.2.1 Methods to study lysosomal channel localizationOne step of characterizing the lysosomal channels is to identify their intracellular lo-calizations. Fluorescent proteins fused to the target proteins provide a useful tool tovirtualize protein localization in live cells. A number of commonly used fluorescentproteins are available with specific colors, for example, GFP (green), YFP (yellow),and RFP/mCherry/DsRed (red) (Ibraheem & Campbell, 2010; Shaner, Steinbach, &Tsien, 2005; Zhang, Campbell, Ting, & Tsien, 2002). Heterologous expression ofGFP fused-TRPML1 revealed that TRPML1 is specifically localized in late endo-somes and lysosomes in a variety of cells (Dong et al., 2008). Because overexpres-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 anddendritic cell maturation and chemotaxis.

P2X4: P2X4 receptor belongs to the purinergic receptor family. It opens inresponse to adenosine triphosphate (ATP) binding at the extracytosolic side (Khakh& North, 2012). In addition to its actions on the plasma membrane, a recent studysuggests that P2X4 is also localized in lysosomal membranes (Qureshi et al.,2007). Lysosomal P2X4 can cycle from the lysosome to phagosome or to the plasmamembrane in response to a variety of stimuli. We recently demonstrated that lyso-somal P2X4 is minimally activated at acidic luminal pH. However, alkalization oflysosome dramatically increases P2X4 channel activity, which may contribute tosion might cause an artificial accumulation of the proteins in cellular compartments,

1. Lysosome 201and because fluorescent proteins could potentially affect the localization of endog-enous proteins (Kim, Soyombo, Tjon-Kon-Sang, So, & Muallem, 2009; Song, Day-alu, Matthews, & Scharenberg, 2006; Venkatachalam, Hofmann, & Montell, 2006),additional approaches are needed to validate the results. Immunostaining is oftenemployed to examine protein localization without interference by heterologousoverexpression. For example, endogenous P2X4 has been detected in lysosomesby immunofluorescent staining (Huang et al., 2014; Qureshi et al., 2007).

Cellular fractionation provides a separation of homogeneous organelles from totalcell lysates by using centrifugation at controlled speeds (Huang et al., 2014; Wanget al., 2012). With the help of specific antibodies, lysosomal ion channel proteinswere detected in the lysosomal-associated membrane protein 1 (Lamp1) positiveheavy fractions by immunoblotting (Huang et al., 2014; Wang et al., 2012; Zeevi,Frumkin, Offen-Glasner, Kogot-Levin, & Bach, 2009). This can be used to validatethe use of fluorescent fusion proteins in the heterologous systems and immunostainingof endogenous proteins for studying subcellular localization of lysosome channels.

1.2.2 Methods to study lysosomal Ca2 channelsCa2 plays an indispensable role in a variety of intracellular processes. To accom-plish their functions, lysosomes also frequently fuse with the plasma membraneand other cellular membranes such as endosomes, autophagosomes, and phago-somes. As with the synaptic vesicle fusion with the plasma membrane, lysosomemembrane fusion with other membranes is also Ca2-dependent (Cheng et al.,2010; Hay, 2007; Lloyd-Evans & Platt, 2011; Luzio, Bright, & Pryor, 2007; Morgan,Platt, Lloyd-Evans, & Galione, 2011; Peters & Mayer, 1998; Piper & Luzio, 2004;Pittman, 2011; Pryor, Mullock, Bright, Gray, & Luzio, 2000). It is believed that thelysosome itself (and/or other organelles) is the main Ca2 source for membranefusion processes (Morgan et al., 2011; Pryor et al., 2000). Indeed, lysosomes areemerging as important intracellular Ca2 stores with luminal [Ca2] of approxi-mately 0.5 mM (Christensen, Myers, & Swanson, 2002). Abnormal lysosomalCa2 hemostasis is associated with numerous lysosomal storage diseases (Lloyd-Evans et al., 2010; Luzio, Pryor, et al., 2007).

In the study of lysosomal Ca2-permeable channels, Ca2 imaging provides adirect way to evaluate channel-mediated Ca2 release/uptake. Two distinct types ofCa2 sensors are available: small molecular fluorescent Ca2 indicator dyes (Grynkie-wicz, Poenie, & Tsien, 1985; Takahashi, Camacho, Lechleiter, & Herman, 1999) andgenetically encoded Ca2 indicators (GECIs) (Demaurex, 2005; McCombs & Palmer,2008). Fura-2 is one of the most widely used fluorescent dyes that permit ratiometricmeasurement of cytosolic Ca2. However, in cases where the channel is also present inthe plasma membrane or other organelles (e.g., endoplasmic reticulum or mitochon-dria membranes), additional approaches are required to exclude the contribution ofCa2 from other sources. GECIs provide a selective way to examine intracellularCa2 signaling because they can be restricted to desired intracellular compartmentsby fusing the construct to organelle-specific targeting motifs. For instance, fusing

2GCaMP3 to the N-terminus of TRPML1 allows the direct measurement of Ca

202 CHAPTER 10 Lysosome electrophysiologyrelease through TRPML1 on lysosomal membranes (Shen et al., 2012). In addition toGCaMP3, other improved variants of GECIs have been developed, for example,GCaMP6 (Chen et al., 2013) and GECO (Zhao et al., 2011). They could be used tostudy lysosomal Ca2 channels activity at higher spatial and temporal resolutions.

1.2.3 Studying lysosomal channels in plasma membrane or in artificialmembranes using patch clamping

The patch-clamp technique allows high-resolution, low noise measurement of theionic currents flowing through the cell membrane (Neher & Sakmann, 1976). It isknown as the most powerful approach in the study of ion channels behaviors, forexample, the ion selectivity, channel kinetics, and gating. Different configurationscan be achieved to record the electrical activity of channels from a section of thecell membrane (known as patch) or the whole cell (Hamill, Marty, Neher, Sakmann,& Sigworth, 1981). For cell-attached mode, the patched membrane adheres tightlyto the pipette, which maintains the intact membrane and intracellular environment.The whole-cell mode is achieved by rupturing the patch formed in the cell-attachedmode through applying a quick suction or a pulse of voltage. It allows recording ofthe whole-cell current at an applied voltage (voltage clamp), or recording of thechanges in the membrane potential where the current is kept constant (currentclamp). The inside-out mode is achieved by pulling the pipette from the cell-attached mode so that the cytosolic side of the membrane is exposed to bath solution.Withdrawing the pipette from whole-cell configuration establishes the outside-outmode, where the outside of the membrane is exposed to the bath solution.

Because of intracellular localization and the relatively small size of vesicles, itwas not feasible to directly measure the electrical activity of lysosomal channelsin the past. Alternative approaches had to be employed. For example, by overex-pressing or introducing some mutations, TRPML1 (Dong et al., 2008; Xu, Delling,Li, Dong, & Clapham, 2007), TPC2 (Brailoiu et al., 2010; Jha, Ahuja, Patel, Brai-loiu, & Muallem, 2014; Wang et al., 2012), and ClCs (Jentsch, 2007; Stauber &Jentsch, 2013) can be redirected to the plasma membrane where they can berecorded using the conventional patch-clamping technique.

Many ion channels such as TRPML1 (Zhang, Jin, Yi, & Li, 2009; Zhang & Li,2007), TPC1 (Pitt, Lam, Rietdorf, Galione, & Sitsapesan, 2014), and TPC2 (Brailoiuet al., 2010; Pitt et al., 2010) have also studied in vitro by reconstituting the channelproteins into planar lipid bilayers. A drawback of this approach is that the proteinsare studied in their nonnative membrane. Indeed, several of the channels appear tohave quite different properties when recorded from lipid bilayers and when studiedfrom the organelles, and a large controversy arises when these channels were studiedin the nonnative membranes (Raychowdhury et al., 2004; Soyombo et al., 2006).

1.2.4 Study of lysosomal channels in lysosomes using lysosome patchclamping

Although several ion channels have been shown to be localized in lysosomal

membranes, the study of functions and properties of these lysosomal channels

et al., 2012).

embryonic kidney 293 (HEK293) or Cos-1 cells.

2. Materials 2032. MATERIALS2.1 CELL CULTURE1. 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,LifeTechnologies)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 PIPETTES1. Glass capillaries (1B150F-4, World Precise Instruments)2. Micropipette puller (Flaming/Brown P-97, Sutter Instruments)3. Microforge (e.g., MF-200, World Precise Instruments)4. Microfill needle (e.g., MF28G-5, World Precise Instruments)The size of a lysosome is usually

plemented with 10% FBS at 37 C in a 5% CO2 incubator. Cells are transfected at a

204 CHAPTER 10 Lysosome electrophysiology3.2 PIPETTES AND SOLUTIONSThe pipettes (electrodes) commonly used for whole-lysosome recordings are similarto those for whole-cell recording except for a smaller size of the pipette tip. Pipettesare pulled from thick-walled borosilicate glass capillaries (1.5-mm outer diameter,1.1-mm inner diameter) using a micropipette puller, and then fire polished underdensity of approximately 80% confluency using Lipofectamine 2000 as per the ven-dors instructions. To monitor the expression, enhanced green fluorescent protein isfused 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 cultureplates. The coverslips are precoated with 0.01% poly-L-lysine overnight, rinsedwith water, and air dried prior to use.

Vacuolin-1 (5 mM) stock solution is prepared by adding 1 mg of vacuolin-1 to465 mL of dimethyl sulfoxide. The vacuolin-1 stock is mixed, divided into 50-mL al-iquots in sterilized tubes, and stored in dark at 20 C. The vacuolin-1 stock isdiluted to 1 mM with DMEM/F-12 culture medium before use. Cells are platedonto coverslips for approximately 2e4 h, and then treated with vacuolin-1 (1 mM)for >2 h prior to performing patch-clamp recordings.2.4 LYSOSOME PATCH-CLAMP RECORDING1. Microscope, air table, and Faraday cage2. Micromanipulator (e.g., MP225, Shutter Instruments)3. Head stage (Axon CV203BU, Molecular Devices)4. Electrode holder (Axon HL-U, Molecular Devices)5. Perfusion chamber (RC-26Z, Warner Instruments)6. Chamber platform (PH-1, Warner Instruments)7. Patch-clamping amplifier (Axon multiclamp 200B, Molecular Devices)8. Digitizer (Axon digidata 1440, Molecular Devices)9. pClamp 10.0 software (Molecular Devices)

10. Bath perfusion system for fast solution exchange

3. METHODS3.1 CELL CULTURECells are maintained in DMEM/F-12 medium (DMEM/Nutrient Mixture F-12) sup-2.3 CHEMICALSAll 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)

visual control using a microforge. Fire polishing allows the pipette to form a narrowtip opening with rounded edges. The polished pipettes typically have a resistance ofapproximately 8e13 MU when filled with the pipette solution.

Preparation of pipette and bath solutions depends on the patch-clamp configu-ration. It is suggested that the environment of lysosome lumen is similar to extra-cellular space (Wang et al., 2012). For whole-lysosome recording, the pipettesolution (a modified Tyrodes solution), which mimics a typical extracellular envi-ronment bathes the luminal surface of isolated enlarged lysosomes; the bath solu-tion which mimics intracellular environment bathes the cytosolic side of theisolated enlarged lysosomes (Figure 2). The components of bath and pipette solu-tions also vary with the objectives of the experiments. With respect to TRPML1 re-cordings, 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 [Ca

2]i equals to 100 nM), and 20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), with the pH adjusted to 7.2 by KOHand osmolality adjusted to approximately 290 mOsm by sucrose. The pipette(luminal) solution contains 145 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM

3. Methods 205MgCl2, 10 mM glucose, and 20 mM HEPES, with the pH adjusted to 4.6 (to mimicthe acidic environment of lysosomes) by HCl and osmolality adjusted to approxi-mately 310 mOsm by sucrose.

The pipette solution is filtered through a 0.45-mm (diameter) filter. Beforerecording, the tip of the pipette is dipped into the pipette solution to avoid bub-bles, and then the pipette is backfilled with the pipette solution using a microfillneedle to half full. The remaining bubbles are removed by gently flicking thepipette.

FIGURE 2 Illustration of the whole-lysosome recording configuration.

The pipette contains a modified Tyrode solution with pH 4.6, which mimics the typical

lysosomal environment; the bath solution is a standard intracellular solution, which mimics

the intracellular environment. Opening of transient receptor potential mucolipin 1

(TRPML1) leads to an efflux of cations (Na/Ca2), moving from the lumen of lysosome to

the cytosol.

206 CHAPTER 10 Lysosome electrophysiology3.3 LYSOSOME PATCH-CLAMP RECORDINGLysosome patch-clamp recordings are performed on manually isolated enlarged ly-sosomes as previously reported (Dong et al., 2008, 2010; Wang et al., 2012). All ex-periments are conducted at room temperature (w20 C).

3.3.1 Isolation of enlarged lysosomesRemove the glass coverslip that contains vacuolin 1-treated cells from the 24-wellplate and place it in the perfusion chamber. Positively transfected cells are recog-nized by green fluorescence. Mount a pipette (electrode) to the electrode holder,and micromanipulate it to touch the cell containing enlarged lysosomes to bepatched. The patch pipette is pressed against the cell and quickly pulled away toslice the cell membrane. Enlarged lysosomes are allowed to release into therecording chamber by pushing the top of the cell with the same pipette (Figure 3).

3.3.2 Whole-lysosome patch clampingAfter an enlarged lysosome is released into the bath, a new pipette is mounted. Toprevent backflow of the bath solution into the pipette and to prevent the pipettefrom getting plugged with debris, a slight positive pressure is applied to the pipettebefore the pipette is dipped into the bath solution. Manipulate the pipette until its tipis just above the isolated enlarged lysosomes without touching it. Set the holding po-tential at 0 mV, apply a 5-mV voltage test pulse, and zero out the offset potential.Slowly micromanipulate the pipette until the tip reaches the surface of the enlargedlysosomes, and then release the positive pressure. Watch for a reduction of the testpulse-induced current, and apply a slight negative pressure to obtain a tight (gigaohm) seal between the pipette and the lysosome membrane. There are severalways to control the positive or negative pressure at the tip of the pipette. The methodwe commonly use is to apply pressure or suction by mouth from the end of the tubeconnected to the pipette. Notably, the tube connected to the pipette holder must befirmly anchored to the head stage so as to minimize the vibration while applyingpressure or suction.

When a tight seal is formed, a current transient is normally observed. Pipettecapacitance compensation is performed to reduce the transient. In order to achievea whole-lysosome configuration, a quick suction by mouth or a brief voltage pulse isapplied. The successful break-in is verified by the reappearance of capacitance tran-sients (sharp capacitance spike with fast decay kinetics) in response to the 5-mV testpulse (Figure 4(A)). Care must be taken to ensure that the lysosome does not enterlysosome cytoplasmic-side-out patch configuration, which, unfortunately, happensquite often. During the experiment, this can be monitored as a loss of capacitancetransients and a reduction in current noise. However, one should bear in mind thatthe fluid level in the perfusion chamber can also affect the capacitance transients.Because of the ubiquitous expression of TRPML1, alternatively, the detection ofendogenous TRPML1 current induced by PI(3,5)P2 or ML-SA1 (a commonlyused TRPML1 agonist) could be another way to differentiate a whole-lysosome

recording from a patch recording (Dong et al., 2010; Shen et al., 2012).

3. Methods 207Once a whole-lysosome configuration is established, a designed voltage proto-col is applied to record the channel of interest. Figure 4(B) shows representativeIeV curves of whole-lysosome currents measured from Cos-1 cells expressingTRPML1. Currents are elicited by repeated voltage ramps of 400-ms durationbetween 140 mV (relative to the lumen which is set at 0 mV) and 140 mVevery 4 s. The small basal TRPML1 currents are significantly enhanced by thebath perfusion of 10 mM ML-SA1. Figure 4(C) shows the time course of TRPML1currents measured at 140 mV in response to ML-SA1 stimulation. The inward

FIGURE 3 Isolation of enlarged lysosomes.

(A) Two enhanced green fluorescent protein- transient receptor potential mucolipin 1

(EGFP-TRPML1) expressing HEK293 cells pretreated with vacuolin-1. Note the

EGFP-positive enlarged lysosomes inside the cell. (B) A pulling pipette (the lower one)

pressed against the lower cell. An enlarged lysosome is isolated and released into the

recording chamber. The recording is then made on the isolated EGFP-positive enlarged

lysosome using a recording pipette (the upper one), which is filled with Rhodamine B dye for

illustration purpose. (See color plate)

Adopted from Dong et al. (2008).

208 CHAPTER 10 Lysosome electrophysiologycurrent at negative potentials indicates an efflux of cations moving from the lumenof lysosomes to the cytosol due to the opening of TRPML1 (Figure 2).

Further, followed by the establishment of whole-lysosome mode, lysosomalmembrane potential can be measured using the current-clamp recording mode(Cang et al., 2013). Given that the lysosomal membrane potential (Vm) is definedas Vcytosol Vlumen (Vlumen 0 mV) (Bertl et al., 1992), opening of TRPML1 re-sults in an increase in Vm, that is, Vlumen becomes more negative. Figure 4(D) showsthat the ML SA1-induced activation of TRPML1 (Figure 4(B) and (C)) is accompa-nied by a depolarization of the lysosome membrane expressing TRPML1.

3.3.3 Other patch configurationsIn addition to whole-lysosome mode, other patch configurations are also availablefor lysosome patch-clamp recording. The lysosome-attached mode is obtained

FIGURE 4 Whole-lysosome recording of transient receptor potential mucolipin 1 (TRPML1).

(A) Representative current traces before (black) and after (red) break-in responding to a

5-mV test pulse. Note the appearance of capacitance transients after break-in. (B)

Representative IeV curves of whole-lysosome TRPML1 activated by bath perfusion of 10 mM

ML-SA1 (short for Mucolipin Synthetic Agonist 1). (C) Current amplitudes measured at

140 mV are used to plot the time course of activation. (D) The activation of TRPML1 isaccompanied by depolarization (Vlumen becomes more negative) of the lysosome recorded in

the current clamp mode. (See color plate)

when the pipette is sealed onto the isolated enlarged lysosomes without breakinginto the vacuolar membrane. The luminal-side-out mode is achieved by quicklywithdrawing the pipette from the enlarged lysosomes after forming thelysosome-attached mode. Therefore, the luminal surface of the enlarged lysosomesis exposed to the bath solution. Figure 5 shows representative IeV curves ofTRPML1Va (a gain-of-function mutant) currents under lysosome-attached andluminal-side-out configurations (Dong et al., 2008). Switching from lysosome-attached to luminal-side-out modes induces a decrease in the amplitude of thecurrents.

FIGURE 5 Common lysosomal recording configurations in the voltage-clamp mode.

(A) Illustration of lysosome-attached, lysosome luminal-side-out, and whole-lysosome

configurations. The arrows indicate the direction of the transient receptor potential

3. Methods 209mucolipin 1 (TRPML1) inward current recorded at negative potentials (flow of cations

moving out of the lysosomes). (B) Two traces to show the currents of TRPML1Va, a gain-

of-function mutant, under lysosome-attached, and lysosome luminal-side-out

configurations. Due to the pH-dependent activation of TRPML1, switching from the

lysosome-attached (luminal side exposed to pH 4.6) to the luminal-side-out configuration

(luminal side exposed to pH 7.2) resulted in a decrease in the current amplitude of

TRPML1Va. (C) A large whole-lysosome current in a lysosome expressing TRPML1Va. A

Cs-based solution (147 mM Cs-methanesulfonate) was used as the pipette solution forboth configurations. (See color plate)

Adopted from Dong et al. (2008).

210 CHAPTER 10 Lysosome electrophysiology4. DISCUSSIONLysosome patch clamping has been a powerful technique to study lysosomal ionchannels. However, the mechanisms of action of vacuolin-1 are still not clear. Themembrane components in the enlarged lysosomes induced by vacuolin-1 could bedifferent from bona fide lysosomes in intact cells. One concern of this techniqueis that vacuolin-1 treatment may affect the channel properties. Given that enlargedlysosomes are also present in a very small number of nontreated cells, the channelproperties of enlarged lysosomes obtained from cells untreated and treated withvacuolin-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 significantdifference in channel properties was detected for enlarged lysosomes obtained withor without vacuolin-1 treatment. However, the possibility of a change in propertiesinduced by vacuolin-1 for other lysosomal ion channels cannot be excluded.

Notably, the lysosome recording is performed on isolated lysosomes. Althoughthe membrane of lysosomes is intact, the cytosolic environment is altered whenthe lysosome is isolated. The loss of cytosolic regulatory factors associated withlysosomal membranes could be one problem for studying the regulation of lyso-somal channels. In this case, regulatory factors should be considered to be includedin the system when doing lysosome patch clamping. For instance, PI(3,5)P2 (anendolysosome specific PIP2) has been found to be required for the activation ofTRPML1 (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 intoconsideration, such as ATP (Huang et al., 2014).

The development of lysosome patch clamping has made it easier to identifynovel lysosome channels (Cang et al., 2014) and to characterize known ones. Forinstance, by using this technique, lysosomal membranes have been shown to bepermeable to other ions including Na, K, and Cl (Cang et al., 2013), and a num-ber of lysosomal channels have been well characterized, including TRPML1 (Donget al., 2008, 2010), TPC2 (Cang et al., 2013; Wang et al., 2012), and P2X4 (Huanget al., 2014). However, the regulation of these channels remains largely unclear. Webelieve that lysosome patch clamping in combination with other methods may pro-vide 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 po-tential drugs that target lysosome channels, which helps find new therapeutic strate-gies for lysosomal ion channel diseases.

In principle, this technique may be modified for recording other lysosome-related organelles such as endosomes, phagosomes, autophagosomes, melanosomes,lytic granules, and many other secretory granules. Indeed, Xus group has success-fully recorded the TRPML1 current in phagosomes (Samie et al., 2013). Although

the approach has limitations, it provides a unique method to measure ion transport

Journal of Cell Biology, 186, 201e209.Brailoiu, E., Rahman, T., Churamani, D., Prole, D. L., Brailoiu, G. C., Hooper, R., et al.

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5. SUMMARYSimilar to the studies of lysosomal enzymes, the study of lysosomal ion transport isan important aspect in our understanding of lysosomal functions. With the advance-ment of lysosome patch clamping that allows the direct measurement of lysosomalchannels in their native environment, we expect that more lysosome ion channelsand their regulatory mechanisms will be elucidated in the near future. Since defi-ciency in lysosomal membrane ion channels and dyshomeostasis of lysosomalions 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 neurodegenera-tive diseases (e.g., Alzheimers Disease) (Coen et al., 2012), we believe that thistechnical advance will dramatically improve our understanding of basic lysosomephysiology, and their implications in lysosome-related diseases.

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10. Lysosome electrophysiologyIntroductionIntroduction1. Lysosome1.1 Lysosome Ion Channels1.1 Lysosome Ion Channels1.2 Methods for Studying Lysosomal Ion Channels1.2 Methods for Studying Lysosomal Ion Channels1.2.1 Methods to study lysosomal channel localization1.2.1 Methods to study lysosomal channel localization1.2.2 Methods to study lysosomal Ca2+ channels1.2.2 Methods to study lysosomal Ca2+ channels1.2.3 Studying lysosomal channels in plasma membrane or in artificial membranes using patch clamping1.2.3 Studying lysosomal channels in plasma membrane or in artificial membranes using patch clamping1.2.4 Study of lysosomal channels in lysosomes using lysosome patch clamping1.2.4 Study of lysosomal channels in lysosomes using lysosome patch clamping

2. Materials2.1 Cell Culture2.1 Cell Culture2.2 Pipettes2.2 Pipettes2.3 Chemicals2.3 Chemicals2.4 Lysosome Patch-Clamp Recording2.4 Lysosome Patch-Clamp Recording

3. Methods3.1 Cell Culture3.1 Cell Culture3.2 Pipettes and Solutions3.2 Pipettes and Solutions3.3 Lysosome Patch-Clamp Recording3.3 Lysosome Patch-Clamp Recording3.3.1 Isolation of enlarged lysosomes3.3.1 Isolation of enlarged lysosomes3.3.2 Whole-lysosome patch clamping3.3.2 Whole-lysosome patch clamping3.3.3 Other patch configurations3.3.3 Other patch configurations

4. Discussion5. SummaryAcknowledgmentsReferences