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10
Lysosome electrophysiology
Xi Z. Zhong, Xian-Ping Dong1 Department of Physiology and Biophysics, Dalhousie University, Halifax, Nova Scotia, Canada 1
Corresponding author: E-mail: [email protected]
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 Ca2þ 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
Methods in Cell Biology, Volume 126, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.10.022 © 2015 Elsevier Inc. All rights reserved.
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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).
1.1 LYSOSOME ION CHANNELS An important feature of the lysosome is an acidic luminal pH (pH w4e5) that ensures lysosomal hydrolases to function properly. The acidic luminal pH is established by the vacuolar type Hþ-ATPase, a well-studied Hþ transporter present on lysosomal membranes (Lloyd-Evans & Platt, 2011; Luzio et al., 2000; Luzio, Pryor, et al., 2007; Mindell, 2012). Although Hþ transport has been the most extensively studied ion movement across lysosomal membranes, recent studies have also indicated that lysosomal membranes are permeable to many other ions, including Naþ, Kþ, and Cl (Cang et al., 2013; Cang, Bekele, & Ren, 2014). Advances in modern cell biology and physiological techniques, together with classical genetic and biochemical approaches, have allowed us to identify a plethora of ion transport proteins in lysosomal membranes (Figure 1), including transient receptor potential mucolipin 1 (TRPML1) (Cheng, Shen, Samie, & Xu, 2010; Dong et al., 2008,
1. Lysosome
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/Hþ exchanger); Hþ-ATPase, proton-pump ATPase.
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/Hþ 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 Ca2þ and Fe2þ permeabilities (Bach, 2005). Impaired TRPML1-mediated Ca2þ/Fe2þ 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 Ca2þrelease 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 Ca2þ 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, Xu’s group at the University of Michigan and others have suggested that TPC1 and TPC2 are in fact highly Naþ-selective channels with very limited Ca2þ permeability (Cang et al., 2013, 2014; Wang et al., 2012). ClCs: ClCs Cl channels (Cl/Hþ 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 Hþ-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 Hþ, rather than ion channels (Accardi & Miller, 2004; Graves et al., 2008; Jentsch, 2007; Weinert et al., 2010).
1.2 METHODS FOR STUDYING LYSOSOMAL ION CHANNELS 1.2.1 Methods to study lysosomal channel localization One step of characterizing the lysosomal channels is to identify their intracellular localizations. Fluorescent proteins fused to the target proteins provide a useful tool to virtualize protein localization in live cells. A number of commonly used fluorescent proteins 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 of GFP fused-TRPML1 revealed that TRPML1 is specifically localized in late endosomes and lysosomes in a variety of cells (Dong et al., 2008). Because overexpression might cause an artificial accumulation of the proteins in cellular compartments,
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.
1.2.2 Methods to study lysosomal Ca2þ channels Ca2þ plays an indispensable role in a variety of intracellular processes. To accomplish their functions, lysosomes also frequently fuse with the plasma membrane and other cellular membranes such as endosomes, autophagosomes, and phagosomes. As with the synaptic vesicle fusion with the plasma membrane, lysosome membrane fusion with other membranes is also Ca2þ-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 the lysosome itself (and/or other organelles) is the main Ca2þ source for membrane fusion processes (Morgan et al., 2011; Pryor et al., 2000). Indeed, lysosomes are emerging as important intracellular Ca2þ stores with luminal [Ca2þ] of approximately 0.5 mM (Christensen, Myers, & Swanson, 2002). Abnormal lysosomal Ca2þ hemostasis is associated with numerous lysosomal storage diseases (LloydEvans et al., 2010; Luzio, Pryor, et al., 2007). In the study of lysosomal Ca2þ-permeable channels, Ca2þ imaging provides a direct way to evaluate channel-mediated Ca2þ release/uptake. Two distinct types of Ca2þ sensors are available: small molecular fluorescent Ca2þ indicator dyes (Grynkiewicz, Poenie, & Tsien, 1985; Takahashi, Camacho, Lechleiter, & Herman, 1999) and genetically encoded Ca2þ indicators (GECIs) (Demaurex, 2005; McCombs & Palmer, 2008). Fura-2 is one of the most widely used fluorescent dyes that permit ratiometric measurement of cytosolic Ca2þ. However, in cases where the channel is also present in the plasma membrane or other organelles (e.g., endoplasmic reticulum or mitochondria membranes), additional approaches are required to exclude the contribution of Ca2þ from other sources. GECIs provide a selective way to examine intracellular Ca2þ signaling because they can be restricted to desired intracellular compartments by fusing the construct to organelle-specific targeting motifs. For instance, fusing GCaMP3 to the N-terminus of TRPML1 allows the direct measurement of Ca2þ
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release through TRPML1 on lysosomal membranes (Shen et al., 2012). In addition to GCaMP3, 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 to study lysosomal Ca2þ channels activity at higher spatial and temporal resolutions.
1.2.3 Studying lysosomal channels in plasma membrane or in artificial membranes using patch clamping The patch-clamp technique allows high-resolution, low noise measurement of the ionic currents flowing through the cell membrane (Neher & Sakmann, 1976). It is known as the most powerful approach in the study of ion channels behaviors, for example, the ion selectivity, channel kinetics, and gating. Different configurations can be achieved to record the electrical activity of channels from a section of the cell membrane (known as patch) or the whole cell (Hamill, Marty, Neher, Sakmann, & Sigworth, 1981). For cell-attached mode, the patched membrane adheres tightly to the pipette, which maintains the intact membrane and intracellular environment. The whole-cell mode is achieved by rupturing the patch formed in the cell-attached mode through applying a quick suction or a pulse of voltage. It allows recording of the whole-cell current at an applied voltage (voltage clamp), or recording of the changes in the membrane potential where the current is kept constant (current clamp). The inside-out mode is achieved by pulling the pipette from the cellattached mode so that the cytosolic side of the membrane is exposed to bath solution. Withdrawing the pipette from whole-cell configuration establishes the outside-out mode, where the outside of the membrane is exposed to the bath solution. Because of intracellular localization and the relatively small size of vesicles, it was not feasible to directly measure the electrical activity of lysosomal channels in the past. Alternative approaches had to be employed. For example, by overexpressing or introducing some mutations, TRPML1 (Dong et al., 2008; Xu, Delling, Li, Dong, & Clapham, 2007), TPC2 (Brailoiu et al., 2010; Jha, Ahuja, Patel, Brailoiu, & Muallem, 2014; Wang et al., 2012), and ClCs (Jentsch, 2007; Stauber & Jentsch, 2013) can be redirected to the plasma membrane where they can be recorded 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 (Brailoiu et al., 2010; Pitt et al., 2010) have also studied in vitro by reconstituting the channel proteins into planar lipid bilayers. A drawback of this approach is that the proteins are studied in their nonnative membrane. Indeed, several of the channels appear to have quite different properties when recorded from lipid bilayers and when studied from the organelles, and a large controversy arises when these channels were studied in the nonnative membranes (Raychowdhury et al., 2004; Soyombo et al., 2006).
1.2.4 Study of lysosomal channels in lysosomes using lysosome patch clamping Although several ion channels have been shown to be localized in lysosomal membranes, the study of functions and properties of these lysosomal channels
2. Materials
has been hampered due to the inaccessibility of patch clamping. Recently, Xu et al. (2007) have overcome the difficulties and developed a novel technique called lysosome patch clamping, which provides a unique way to study the lysosomal channels in their native environments by directly recording isolated lysosomes (Dong et al., 2008, 2010; Saito, Hanson, & Schlesinger, 2007; Wang et al., 2012). The size of a lysosome is usually 2 h prior to performing patch-clamp recordings.
3.2 PIPETTES AND SOLUTIONS The pipettes (electrodes) commonly used for whole-lysosome recordings are similar to those for whole-cell recording except for a smaller size of the pipette tip. Pipettes are 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 under
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 Tyrode’s 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 [Ca2þ]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.
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 (Naþ/Ca2þ), moving from the lumen of lysosome to the cytosol.
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3.3 LYSOSOME PATCH-CLAMP RECORDING Lysosome patch-clamp recordings are performed on manually isolated enlarged lysosomes as previously reported (Dong et al., 2008, 2010; Wang et al., 2012). All experiments are conducted at room temperature (w20 C).
3.3.1 Isolation of enlarged lysosomes Remove the glass coverslip that contains vacuolin 1-treated cells from the 24-well plate and place it in the perfusion chamber. Positively transfected cells are recognized by green fluorescence. Mount a pipette (electrode) to the electrode holder, and micromanipulate it to touch the cell containing enlarged lysosomes to be patched. The patch pipette is pressed against the cell and quickly pulled away to slice the cell membrane. Enlarged lysosomes are allowed to release into the recording chamber by pushing the top of the cell with the same pipette (Figure 3).
3.3.2 Whole-lysosome patch clamping After an enlarged lysosome is released into the bath, a new pipette is mounted. To prevent backflow of the bath solution into the pipette and to prevent the pipette from getting plugged with debris, a slight positive pressure is applied to the pipette before the pipette is dipped into the bath solution. Manipulate the pipette until its tip is just above the isolated enlarged lysosomes without touching it. Set the holding potential 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 enlarged lysosomes, and then release the positive pressure. Watch for a reduction of the test pulse-induced current, and apply a slight negative pressure to obtain a tight (giga ohm) seal between the pipette and the lysosome membrane. There are several ways to control the positive or negative pressure at the tip of the pipette. The method we commonly use is to apply pressure or suction by mouth from the end of the tube connected to the pipette. Notably, the tube connected to the pipette holder must be firmly anchored to the head stage so as to minimize the vibration while applying pressure or suction. When a tight seal is formed, a current transient is normally observed. Pipette capacitance compensation is performed to reduce the transient. In order to achieve a whole-lysosome configuration, a quick suction by mouth or a brief voltage pulse is applied. The successful break-in is verified by the reappearance of capacitance transients (sharp capacitance spike with fast decay kinetics) in response to the 5-mV test pulse (Figure 4(A)). Care must be taken to ensure that the lysosome does not enter lysosome cytoplasmic-side-out patch configuration, which, unfortunately, happens quite often. During the experiment, this can be monitored as a loss of capacitance transients and a reduction in current noise. However, one should bear in mind that the fluid level in the perfusion chamber can also affect the capacitance transients. Because of the ubiquitous expression of TRPML1, alternatively, the detection of endogenous TRPML1 current induced by PI(3,5)P2 or ML-SA1 (a commonly used 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
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).
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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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 is accompanied by depolarization (Vlumen becomes more negative) of the lysosome recorded in the current clamp mode. (See color plate)
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 ¼ 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.
3.3.3 Other patch configurations In addition to whole-lysosome mode, other patch configurations are also available for lysosome patch-clamp recording. The lysosome-attached mode is obtained
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.
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 mucolipin 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 gainof-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 Csþ-based solution (147 mM Cs-methanesulfonate) was used as the pipette solution for both configurations. (See color plate) Adopted from Dong et al. (2008).
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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 Naþ, Kþ, 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, Xu’s 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., Alzheimer’s 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).
REFERENCES Accardi, A., & Miller, C. (2004). Secondary active transport mediated by a prokaryotic homologue of ClC Cl channels. Nature, 427, 803e807. Bach, G. (2005). Mucolipin 1: endocytosis and cation channelda review. Pflu¨gers Archiv: European Journal of Physiology, 451, 313e317. Bassi, M. T., Manzoni, M., Monti, E., Pizzo, M. T., Ballabio, A., & Borsani, G. (2000). Cloning of the gene encoding a novel integral membrane protein, mucolipidindand identification of the two major founder mutations causing mucolipidosis type IV. American Journal of Human Genetics, 67, 1110e1120. Bertl, A., Blumwald, E., Coronado, R., Eisenberg, R., Findlay, G., Gradmann, D., et al. (1992). Electrical measurements on endomembranes. Science (New York, NY), 258, 873e874. Brailoiu, E., Churamani, D., Cai, X., Schrlau, M. G., Brailoiu, G. C., Gao, X., et al. (2009). Essential requirement for two-pore channel 1 in NAADP-mediated calcium signaling. The Journal of Cell Biology, 186, 201e209. Brailoiu, E., Rahman, T., Churamani, D., Prole, D. L., Brailoiu, G. C., Hooper, R., et al. (2010). An NAADP-gated two-pore channel targeted to the plasma membrane uncouples triggering from amplifying Ca2þ signals. The Journal of Biological Chemistry, 285, 38511e38516.
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Calcraft, P. J., Ruas, M., Pan, Z., Cheng, X., Arredouani, A., Hao, X., et al. (2009). NAADP mobilizes calcium from acidic organelles through two-pore channels. Nature, 459, 596e600. Cang, C., Bekele, B., & Ren, D. (2014). The voltage-gated sodium channel TPC1 confers endolysosomal excitability. Nature Chemical Biology, 10, 463e469. Cang, C., Zhou, Y., Navarro, B., Seo, Y. J., Aranda, K., Shi, L., et al. (2013). mTOR regulates lysosomal ATP-sensitive two-pore Na(þ) channels to adapt to metabolic state. Cell, 152, 778e790. Cheng, X., Shen, D., Samie, M., & Xu, H. (2010). Mucolipins: Intracellular TRPML1-3 channels. FEBS Letters, 584, 2013e2021. Chen, T. W., Wardill, T. J., Sun, Y., Pulver, S. R., Renninger, S. L., Baohan, A., et al. (2013). Ultrasensitive fluorescent proteins for imaging neuronal activity. Nature, 499, 295e300. Christensen, K. A., Myers, J. T., & Swanson, J. A. (2002). pH-dependent regulation of lysosomal calcium in macrophages. Journal of Cell Science, 115, 599e607. Coen, K., Flannagan, R. S., Baron, S., Carraro-Lacroix, L. R., Wang, D., Vermeire, W., et al. (2012). Lysosomal calcium homeostasis defects, not proton pump defects, cause endolysosomal dysfunction in PSEN-deficient cells. The Journal of Cell Biology, 198, 23e35. Demaurex, N. (2005). Calcium measurements in organelles with Ca2þ-sensitive fluorescent proteins. Cell Calcium, 38, 213e222. Dong, X. P., Cheng, X., Mills, E., Delling, M., Wang, F., Kurz, T., et al. (2008). The type IV mucolipidosis-associated protein TRPML1 is an endolysosomal iron release channel. Nature, 455, 992e996. Dong, X. P., Shen, D., Wang, X., Dawson, T., Li, X., Zhang, Q., et al. (2010). PI(3,5)P(2) controls membrane trafficking by direct activation of mucolipin Ca(2þ) release channels in the endolysosome. Nature Communications, 1, 38. Graves, A. R., Curran, P. K., Smith, C. L., & Mindell, J. A. (2008). The Cl/Hþ antiporter ClC-7 is the primary chloride permeation pathway in lysosomes. Nature, 453, 788e792. Grynkiewicz, G., Poenie, M., & Tsien, R. Y. (1985). A new generation of Ca2þ indicators with greatly improved fluorescence properties. The Journal of Biological Chemistry, 260, 3440e3450. Hamill, O. P., Marty, A., Neher, E., Sakmann, B., & Sigworth, F. J. (1981). Improved patchclamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflu¨gers Archiv: European Journal of Physiology, 391, 85e100. Hay, J. C. (2007). Calcium: a fundamental regulator of intracellular membrane fusion? EMBO Reports, 8, 236e240. Huang, P., Zou, Y., Zhong, X. Z., Cao, Q., Zhao, K., Zhu, M. X., et al. (2014). P2X4 forms functional ATP-activated cation channels on lysosomal membranes regulated by luminal pH. The Journal of Biological Chemistry, 289, 17658e17667. Huynh, C., & Andrews, N. W. (2005). The small chemical vacuolin-1 alters the morphology of lysosomes without inhibiting Ca2þ-regulated exocytosis. EMBO Reports, 6, 843e847. Ibraheem, A., & Campbell, R. E. (2010). Designs and applications of fluorescent proteinbased biosensors. Current Opinion in Chemical Biology, 14, 30e36. Jentsch, T. J. (2007). Chloride and the endosomalelysosomal pathway: emerging roles of CLC chloride transporters. The Journal of Physiology, 578, 633e640. Jha, A., Ahuja, M., Patel, S., Brailoiu, E., & Muallem, S. (2014). Convergent regulation of the lysosomal two-pore channel-2 by Mg(2)(þ), NAADP, PI(3,5)P(2) and multiple protein kinases. The EMBO Journal, 33, 501e511.
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Ramsey, I. S., Delling, M., & Clapham, D. E. (2006). An introduction to TRP channels. Annual Review of Physiology, 68, 619e647. Raychowdhury, M. K., Gonzalez-Perrett, S., Montalbetti, N., Timpanaro, G. A., Chasan, B., Goldmann, W. H., et al. (2004). Molecular pathophysiology of mucolipidosis type IV: pH dysregulation of the mucolipin-1 cation channel. Human Molecular Genetics, 13, 617e627. Saito, M., Hanson, P. I., & Schlesinger, P. (2007). Luminal chloride-dependent activation of endosome calcium channels: patch clamp study of enlarged endosomes. The Journal of Biological Chemistry, 282, 27327e27333. Samie, M., Wang, X., Zhang, X., Goschka, A., Li, X., Cheng, X., et al. (2013). A TRP channel in the lysosome regulates large particle phagocytosis via focal exocytosis. Developmental Cell, 26, 511e524. Shaner, N. C., Steinbach, P. A., & Tsien, R. Y. (2005). A guide to choosing fluorescent proteins. Nature Methods, 2, 905e909. Shen, D., Wang, X., Li, X., Zhang, X., Yao, Z., Dibble, S., et al. (2012). Lipid storage disorders block lysosomal trafficking by inhibiting a TRP channel and lysosomal calcium release. Nature Communications, 3, 731. Shen, D., Wang, X., & Xu, H. (2011). Pairing phosphoinositides with calcium ions in endolysosomal dynamics: phosphoinositides control the direction and specificity of membrane trafficking by regulating the activity of calcium channels in the endolysosomes. BioEssays: News and Reviews in Molecular, Cellular and Developmental Biology, 33, 448e457. Song, Y., Dayalu, R., Matthews, S. A., & Scharenberg, A. M. (2006). TRPML cation channels regulate the specialized lysosomal compartment of vertebrate B-lymphocytes. European Journal of Cell Biology, 85, 1253e1264. Soyombo, A. A., Tjon-Kon-Sang, S., Rbaibi, Y., Bashllari, E., Bisceglia, J., Muallem, S., et al. (2006). TRP-ML1 regulates lysosomal pH and acidic lysosomal lipid hydrolytic activity. The Journal of Biological Chemistry, 281, 7294e7301. Stauber, T., & Jentsch, T. J. (2013). Chloride in vesicular trafficking and function. Annual Review of Physiology, 75, 453e477. Sumoza-Toledo, A., Lange, I., Cortado, H., Bhagat, H., Mori, Y., Fleig, A., et al. (2011). Dendritic cell maturation and chemotaxis is regulated by TRPM2-mediated lysosomal Ca2þ release. FASEB Journal: Official Publication of the Federation of American Societies for Experimental Biology, 25, 3529e3542. Sun, M., Goldin, E., Stahl, S., Falardeau, J. L., Kennedy, J. C., Acierno, J. S., Jr., et al. (2000). Mucolipidosis type IV is caused by mutations in a gene encoding a novel transient receptor potential channel. Human Molecular Genetics, 9, 2471e2478. Takahashi, A., Camacho, P., Lechleiter, J. D., & Herman, B. (1999). Measurement of intracellular calcium. Physiological Reviews, 79, 1089e1125. Venkatachalam, K., Hofmann, T., & Montell, C. (2006). Lysosomal localization of TRPML3 depends on TRPML2 and the mucolipidosis-associated protein TRPML1. The Journal of Biological Chemistry, 281, 17517e17527. Wang, X., Zhang, X., Dong, X. P., Samie, M., Li, X., Cheng, X., et al. (2012). TPC proteins are phosphoinositide- activated sodium-selective ion channels in endosomes and lysosomes. Cell, 151, 372e383. Weinert, S., Jabs, S., Supanchart, C., Schweizer, M., Gimber, N., Richter, M., et al. (2010). Lysosomal pathology and osteopetrosis upon loss of Hþ-driven lysosomal Cl accumulation. Science (New York, NY), 328, 1401e1403.
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Xu, H., Delling, M., Li, L., Dong, X., & Clapham, D. E. (2007). Activating mutation in a mucolipin transient receptor potential channel leads to melanocyte loss in varitintwaddler mice. Proceedings of the National Academy of Sciences of the United States of America, 104, 18321e18326. Zeevi, D. A., Frumkin, A., Offen-Glasner, V., Kogot-Levin, A., & Bach, G. (2009). A potentially dynamic lysosomal role for the endogenous TRPML proteins. The Journal of Pathology, 219, 153e162. Zhang, J., Campbell, R. E., Ting, A. Y., & Tsien, R. Y. (2002). Creating new fluorescent probes for cell biology. Nature Reviews Molecular Cell Biology, 3, 906e918. Zhang, F., Jin, S., Yi, F., & Li, P. L. (2009). TRP-ML1 functions as a lysosomal NAADP-sensitive Ca2þ release channel in coronary arterial myocytes. Journal of Cellular and Molecular Medicine, 13, 3174e3185. Zhang, F., & Li, P. L. (2007). Reconstitution and characterization of a nicotinic acid adenine dinucleotide phosphate (NAADP)-sensitive Ca2þ release channel from liver lysosomes of rats. The Journal of Biological Chemistry, 282, 25259e25269. Zhao, Y., Araki, S., Wu, J., Teramoto, T., Chang, Y. F., Nakano, M., et al. (2011). An expanded palette of genetically encoded Ca(2)(þ) indicators. Science (New York, NY), 333, 1888e1891.
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Xi Z. Zhong, Xian-Ping Dong1 Department of Physiology and Biophysics, Dalhousie University, Halifax, Nova Scotia, Canada 1
Corresponding author: E-mail: [email protected]
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 Ca2þ 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
Methods in Cell Biology, Volume 126, ISSN 0091-679X, http://dx.doi.org/10.1016/bs.mcb.2014.10.022 © 2015 Elsevier Inc. All rights reserved.
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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).
1.1 LYSOSOME ION CHANNELS An important feature of the lysosome is an acidic luminal pH (pH w4e5) that ensures lysosomal hydrolases to function properly. The acidic luminal pH is established by the vacuolar type Hþ-ATPase, a well-studied Hþ transporter present on lysosomal membranes (Lloyd-Evans & Platt, 2011; Luzio et al., 2000; Luzio, Pryor, et al., 2007; Mindell, 2012). Although Hþ transport has been the most extensively studied ion movement across lysosomal membranes, recent studies have also indicated that lysosomal membranes are permeable to many other ions, including Naþ, Kþ, and Cl (Cang et al., 2013; Cang, Bekele, & Ren, 2014). Advances in modern cell biology and physiological techniques, together with classical genetic and biochemical approaches, have allowed us to identify a plethora of ion transport proteins in lysosomal membranes (Figure 1), including transient receptor potential mucolipin 1 (TRPML1) (Cheng, Shen, Samie, & Xu, 2010; Dong et al., 2008,
1. Lysosome
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/Hþ exchanger); Hþ-ATPase, proton-pump ATPase.
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/Hþ 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 Ca2þ and Fe2þ permeabilities (Bach, 2005). Impaired TRPML1-mediated Ca2þ/Fe2þ 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 Ca2þrelease 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 Ca2þ 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, Xu’s group at the University of Michigan and others have suggested that TPC1 and TPC2 are in fact highly Naþ-selective channels with very limited Ca2þ permeability (Cang et al., 2013, 2014; Wang et al., 2012). ClCs: ClCs Cl channels (Cl/Hþ 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 Hþ-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 Hþ, rather than ion channels (Accardi & Miller, 2004; Graves et al., 2008; Jentsch, 2007; Weinert et al., 2010).
1.2 METHODS FOR STUDYING LYSOSOMAL ION CHANNELS 1.2.1 Methods to study lysosomal channel localization One step of characterizing the lysosomal channels is to identify their intracellular localizations. Fluorescent proteins fused to the target proteins provide a useful tool to virtualize protein localization in live cells. A number of commonly used fluorescent proteins 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 of GFP fused-TRPML1 revealed that TRPML1 is specifically localized in late endosomes and lysosomes in a variety of cells (Dong et al., 2008). Because overexpression might cause an artificial accumulation of the proteins in cellular compartments,
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.
1.2.2 Methods to study lysosomal Ca2þ channels Ca2þ plays an indispensable role in a variety of intracellular processes. To accomplish their functions, lysosomes also frequently fuse with the plasma membrane and other cellular membranes such as endosomes, autophagosomes, and phagosomes. As with the synaptic vesicle fusion with the plasma membrane, lysosome membrane fusion with other membranes is also Ca2þ-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 the lysosome itself (and/or other organelles) is the main Ca2þ source for membrane fusion processes (Morgan et al., 2011; Pryor et al., 2000). Indeed, lysosomes are emerging as important intracellular Ca2þ stores with luminal [Ca2þ] of approximately 0.5 mM (Christensen, Myers, & Swanson, 2002). Abnormal lysosomal Ca2þ hemostasis is associated with numerous lysosomal storage diseases (LloydEvans et al., 2010; Luzio, Pryor, et al., 2007). In the study of lysosomal Ca2þ-permeable channels, Ca2þ imaging provides a direct way to evaluate channel-mediated Ca2þ release/uptake. Two distinct types of Ca2þ sensors are available: small molecular fluorescent Ca2þ indicator dyes (Grynkiewicz, Poenie, & Tsien, 1985; Takahashi, Camacho, Lechleiter, & Herman, 1999) and genetically encoded Ca2þ indicators (GECIs) (Demaurex, 2005; McCombs & Palmer, 2008). Fura-2 is one of the most widely used fluorescent dyes that permit ratiometric measurement of cytosolic Ca2þ. However, in cases where the channel is also present in the plasma membrane or other organelles (e.g., endoplasmic reticulum or mitochondria membranes), additional approaches are required to exclude the contribution of Ca2þ from other sources. GECIs provide a selective way to examine intracellular Ca2þ signaling because they can be restricted to desired intracellular compartments by fusing the construct to organelle-specific targeting motifs. For instance, fusing GCaMP3 to the N-terminus of TRPML1 allows the direct measurement of Ca2þ
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release through TRPML1 on lysosomal membranes (Shen et al., 2012). In addition to GCaMP3, 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 to study lysosomal Ca2þ channels activity at higher spatial and temporal resolutions.
1.2.3 Studying lysosomal channels in plasma membrane or in artificial membranes using patch clamping The patch-clamp technique allows high-resolution, low noise measurement of the ionic currents flowing through the cell membrane (Neher & Sakmann, 1976). It is known as the most powerful approach in the study of ion channels behaviors, for example, the ion selectivity, channel kinetics, and gating. Different configurations can be achieved to record the electrical activity of channels from a section of the cell membrane (known as patch) or the whole cell (Hamill, Marty, Neher, Sakmann, & Sigworth, 1981). For cell-attached mode, the patched membrane adheres tightly to the pipette, which maintains the intact membrane and intracellular environment. The whole-cell mode is achieved by rupturing the patch formed in the cell-attached mode through applying a quick suction or a pulse of voltage. It allows recording of the whole-cell current at an applied voltage (voltage clamp), or recording of the changes in the membrane potential where the current is kept constant (current clamp). The inside-out mode is achieved by pulling the pipette from the cellattached mode so that the cytosolic side of the membrane is exposed to bath solution. Withdrawing the pipette from whole-cell configuration establishes the outside-out mode, where the outside of the membrane is exposed to the bath solution. Because of intracellular localization and the relatively small size of vesicles, it was not feasible to directly measure the electrical activity of lysosomal channels in the past. Alternative approaches had to be employed. For example, by overexpressing or introducing some mutations, TRPML1 (Dong et al., 2008; Xu, Delling, Li, Dong, & Clapham, 2007), TPC2 (Brailoiu et al., 2010; Jha, Ahuja, Patel, Brailoiu, & Muallem, 2014; Wang et al., 2012), and ClCs (Jentsch, 2007; Stauber & Jentsch, 2013) can be redirected to the plasma membrane where they can be recorded 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 (Brailoiu et al., 2010; Pitt et al., 2010) have also studied in vitro by reconstituting the channel proteins into planar lipid bilayers. A drawback of this approach is that the proteins are studied in their nonnative membrane. Indeed, several of the channels appear to have quite different properties when recorded from lipid bilayers and when studied from the organelles, and a large controversy arises when these channels were studied in the nonnative membranes (Raychowdhury et al., 2004; Soyombo et al., 2006).
1.2.4 Study of lysosomal channels in lysosomes using lysosome patch clamping Although several ion channels have been shown to be localized in lysosomal membranes, the study of functions and properties of these lysosomal channels
2. Materials
has been hampered due to the inaccessibility of patch clamping. Recently, Xu et al. (2007) have overcome the difficulties and developed a novel technique called lysosome patch clamping, which provides a unique way to study the lysosomal channels in their native environments by directly recording isolated lysosomes (Dong et al., 2008, 2010; Saito, Hanson, & Schlesinger, 2007; Wang et al., 2012). The size of a lysosome is usually 2 h prior to performing patch-clamp recordings.
3.2 PIPETTES AND SOLUTIONS The pipettes (electrodes) commonly used for whole-lysosome recordings are similar to those for whole-cell recording except for a smaller size of the pipette tip. Pipettes are 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 under
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 Tyrode’s 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 [Ca2þ]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.
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 (Naþ/Ca2þ), moving from the lumen of lysosome to the cytosol.
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3.3 LYSOSOME PATCH-CLAMP RECORDING Lysosome patch-clamp recordings are performed on manually isolated enlarged lysosomes as previously reported (Dong et al., 2008, 2010; Wang et al., 2012). All experiments are conducted at room temperature (w20 C).
3.3.1 Isolation of enlarged lysosomes Remove the glass coverslip that contains vacuolin 1-treated cells from the 24-well plate and place it in the perfusion chamber. Positively transfected cells are recognized by green fluorescence. Mount a pipette (electrode) to the electrode holder, and micromanipulate it to touch the cell containing enlarged lysosomes to be patched. The patch pipette is pressed against the cell and quickly pulled away to slice the cell membrane. Enlarged lysosomes are allowed to release into the recording chamber by pushing the top of the cell with the same pipette (Figure 3).
3.3.2 Whole-lysosome patch clamping After an enlarged lysosome is released into the bath, a new pipette is mounted. To prevent backflow of the bath solution into the pipette and to prevent the pipette from getting plugged with debris, a slight positive pressure is applied to the pipette before the pipette is dipped into the bath solution. Manipulate the pipette until its tip is just above the isolated enlarged lysosomes without touching it. Set the holding potential 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 enlarged lysosomes, and then release the positive pressure. Watch for a reduction of the test pulse-induced current, and apply a slight negative pressure to obtain a tight (giga ohm) seal between the pipette and the lysosome membrane. There are several ways to control the positive or negative pressure at the tip of the pipette. The method we commonly use is to apply pressure or suction by mouth from the end of the tube connected to the pipette. Notably, the tube connected to the pipette holder must be firmly anchored to the head stage so as to minimize the vibration while applying pressure or suction. When a tight seal is formed, a current transient is normally observed. Pipette capacitance compensation is performed to reduce the transient. In order to achieve a whole-lysosome configuration, a quick suction by mouth or a brief voltage pulse is applied. The successful break-in is verified by the reappearance of capacitance transients (sharp capacitance spike with fast decay kinetics) in response to the 5-mV test pulse (Figure 4(A)). Care must be taken to ensure that the lysosome does not enter lysosome cytoplasmic-side-out patch configuration, which, unfortunately, happens quite often. During the experiment, this can be monitored as a loss of capacitance transients and a reduction in current noise. However, one should bear in mind that the fluid level in the perfusion chamber can also affect the capacitance transients. Because of the ubiquitous expression of TRPML1, alternatively, the detection of endogenous TRPML1 current induced by PI(3,5)P2 or ML-SA1 (a commonly used 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
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).
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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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 is accompanied by depolarization (Vlumen becomes more negative) of the lysosome recorded in the current clamp mode. (See color plate)
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 ¼ 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.
3.3.3 Other patch configurations In addition to whole-lysosome mode, other patch configurations are also available for lysosome patch-clamp recording. The lysosome-attached mode is obtained
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.
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 mucolipin 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 gainof-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 Csþ-based solution (147 mM Cs-methanesulfonate) was used as the pipette solution for both configurations. (See color plate) Adopted from Dong et al. (2008).
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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 Naþ, Kþ, 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, Xu’s 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., Alzheimer’s 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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