Approaches of Studying Lysosomal Ion Channels and Transporters: Comparison
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Please note this is a comparison between Version 1 by Armando Carpaneto and Version 2 by Lindsay Dong.

Lysosomes are acidic organelles, pH of about 4.6 [1], considered as the digestive system of the animal cell. They act as the major compartment for detoxification of both the outer and the inner content of the cell. In fact, lysosomes represent the key players in degradation, recycling, autophagy, cell death, cell proliferation, cell defence, immunity–autoimmunity processes and therefore in maintenance of cellular homeostasis.  A distinct set of channels and transporters regulates the ion fluxes across the lysosomal membrane. Malfunctioning of these transport proteins and the resulting ionic imbalance is involved in various human diseases, such as lysosomal storage disorders, cancer, as well as metabolic and neurodegenerative diseases. As a consequence, these proteins have stimulated strong interest for their suitability as possible drug targets.

  • lysosomes
  • ion channels
  • transporters

1. Introduction

1.1. Main Families of Lysosomal Channels and Transporters

The lysosomal ion channels and transporters known so far belong to a limited number of protein families (Table 1).
Table 1.
Lysosomal channels and transporters (related references inserted in the main text).
Channel/Transporter Transported Ion(s)
CLC-6 Cl−, H+
CLC-7
SLC38A7 Na+, aminoacids
SLC38A9
NHE3 Na+, H+
NHE5
NHE6
TPC1 Na+, Ca2+
TPC2
VGCCs Ca2+
TRPML1 Na+, Ca2+, Fe2+, Zn2+, cations
TRPML2
TRPML3
P2X4
BK K+
TMEM175
LRRC8 Cl−, organic anions
V-ATPase H+
CLN7 Cl−

1.2. Summary of the Experimental Methods to Investigate the Functional Properties of Lysosomal Ion Channels and Transporters

A summary of the methods, their respective advantages and disadvantages, together with the lysosomal channels/transporters to which they have been applied is presented in Table 2.

Table 2.
Summary of current methods for the functional characterization of endolysosomal channels and transporters.
Method Advantages Disadvantages Lysosomal Channels/Transporters
Incorporation into artificial membranes or liposomes Low level of background current noise Protein amount

Channel removed from native environment

Impurities		 TPC1, TPC2, TRPLM1
Solid-supported membrane-based electrophysiology Native environment

Automation

Suitable for screening		 No control of membrane potential

No control of luminal solution		 CLC-7, V-ATPase
Flux measurements on purified lysosomes Native environment

Large number of lysosomes tested		 No control of membrane potential

No control of luminal solution		 CLC-7
Patch-clamp electrophysiology on enlarged lysosomes Native environment

Direct

Robust		 Insufficient resolution to detect the activity of low turnover rate transporters

Interference by endogenous channels and transporters

Need of trained electrophysiologist		 TPCs, TRPML1, BK, LRRC8, TMEM175, LRRC8, CLN7
Targeting to the plasma membrane upon manipulation of sorting signals Well-characterized expression systems can be used

Easy to perform, even if sorting/retention signals are not known		 Different lipid environment may affect activity

Modification of protein by mutation or tag

Not applicable to all intracellular transmembrane proteins		 CLC-6, CLC-7, TPC2, GLUT8, LRRC8, CLN7
Nuclear membrane electrophysiology Easy access to cytoplasmic and luminal sides

Ligand conditions rigorously controlled

High temporal resolution

Simple protocol

High signal-to-noise ratio		 Not tolerant to high Vapp

Low quality and stability of giga-ohm seals

High background current

Difficult excised nuclear patches		 hTPC2
Patch-clamp electrophysiology on plant vacuoles Good knowledge of vacuolar endogenous channels

Large size

Ease of isolation

Low noise

Possibility of different patch configurations

Eukaryotic post-translational modifications		 Long protoplasting procedure

Need of trained electrophysiologist

Fused fluorescent protein could impair functionality

Differences with mammalian post translational modifications		 CLC-7, hTPC1, hTPC2
Patch-clamp electrophysiology on giant vacuoles from yeast cells Very high signal-to-noise ratio Need of preparation of giant cells hTPC2

2. Approaches Using Purified Proteins or Native Endolysosomal Membranes

2.1. Incorporation into Artificial Membranes or Liposomes

A planar bilayer is an artificial membrane formed across a small hole placed in a thin plastic partition that separates two aqueous compartments. In turn, lipid bilayers may be used to characterize the activity of lysosomal membrane proteins at the level of single molecular events by incorporating either liposomes containing purified proteins or native membranes vesicles derived from specific compartments/organelles.
Lipid bilayers have a very low level of background current noise so that it is possible to record single-channel currents. Accessibility of both sides of the bilayer and the possibility to clamp the membrane voltage allow studies of channel gating, ion conduction and selectivity, effect of ligands, etc.
An advantage of the bilayer approach is that it enables to examine the effect of the lipid environment on the channel, as bilayers may be formed by different types of lipids. Disadvantages include the requirement of a sufficient amount of protein, the fact that the channel is removed from its native environment and that there is no control of protein orientation within the membrane when purified protein is used. However, the main drawback, especially when native membrane vesicles are used, is the presence of impurities, whose activity may be erroneously attributed to the protein of interest.
Single channel events of immunopurified hTPC2 and hTPC1 reconstituted in artificial membranes were observed respectively by [8][9][10].
TRPML1 function was investigated in lipid bilayers using reconstitution of both endosomal vesicles derived from cells over-expressing TRPML1 and liposomes previously dialyzed with TRPML1 protein [11]. TRPML1 reconstituted in lipid bilayers showed spontaneous cation channel activity in the presence of asymmetric K+, voltage-dependent activation and multiple sub-conductance states.

2.2. Solid-Supported Membrane-Based Electrophysiology

A number of different electrogenic proteins (ion pumps, transporters and channels) have been tested using solid-supported membrane (SSM) electrophysiology [12][13][14][15]. More recently, this technique has been applied to intracellular transporters to overcome the inaccessibilty of endomembranes [16][17].
For such electrical measurements, the SSM is built on a planar gold electrode by depositing an alkane-thiol layer followed by a phospholipid monolayer on top of it. Proteoliposomes containing purified proteins or membrane vesicles and membrane fragments from native tissue containing the protein of interest are adsorbed on this SSM to form a capacitively coupled membrane system. The SSM behaves like a capacitor with the solution in the fluid compartment, and with the gold electrode being the capacitor plates.
Charge translocation, due to the protein’s transport activity, is initiated by providing a substrate or a ligand via rapid solution exchange. The transport-dependent transient currents correspond to the charging of the gold electrode sensor and the charging kinetics depends on the transport activity of the assayed protein. In general, the peak current value is used to analyse the stationary protein transport activity. Membrane fractions (from native tissue or transfected cells) enriched in plasma membrane or specific intracellular membranes can be obtained by sugar gradient fractionation, giving the possibility to study proteins expressed in intracellular organelles in their native environment.

2.3. Flux Measurements on Purified Lysosomes

Concentrative isotope uptake was previously used for measuring ion fluxes through ion channels in membrane vesicles [18]. Vesicle suspensions were incubated with the 22NaCl isotope and the amount of 22Na trapped within the vesicles was measured. This procedure allowed to measure a specific 22Na uptake and to identify the fraction of vesicles containing Na+ channels among a heterogeneous vesicle population [18] Concentrative uptake of 36Cl− due to a Cl− gradient was used to determine the conductance properties of Cl− channels extracted from Torpedo plasma membrane and reconstituted into liposomes [19]. The concentrative uptake method was employed to show that the Cl−/H+ antiporter CLC-7 is a major chloride permeation pathway in lysosomes [20]. Lysosomes isolated from rat liver by differential sedimentation were loaded with high concentrations of unlabelled chloride and then diluted into a buffer containing 36Cl−. The rapid uptake of 36Cl−, which was abolished by the external addition of valinomycin, suggested the presence of a specific electrogenic transport pathway for chloride. Additional experiments varying internal anions and in the presence of a pH gradient established the apparent permeability sequence and showed the coupling between Cl− and proton transport. Measurements performed with a Cl− gradient and monitoring the internal pH with the fluorescent dye 2′,7′-bis-(2-carboxyethyl)-5(6)-carboxyfluorescein (BCECF) showed that protons could move against the pH gradient as expected for a Cl−/H+ antiporter. Finally, the equilibrium potential for H+ flux, monitored by BCECF, was measured at a series of theoretical voltages set with K+/valinomycin. As a result of these experiments performed on isolated lysosomes, by using concentrative 36Cl− uptake combined with fluorescence measurements of proton fluxes, it he authors could establish that the lysosomal transport of Cl− and H+ is mediated by a Cl−/H+ antiporter, identified as CLC-7 [20]. This method allows to estimate fluxes in a large number of lysosomes under varying external conditions and maintaining the proteins in their native membrane. It can identify ion transport mechanisms across the lysosomal membrane, however its use to characterize in detail the functional activity of lysosomal membrane transporters seems difficult. It requires performing radioactivity measurements and it does not allow direct and precise control of the membrane potential preventing the study of the voltage dependence of the ion transport.

2.4. Patch-Clamp Electrophysiology on Enlarged Lysosomes

Acidic lysosomal compartments in animal cells are small in size (diameter < 500 nm), and this property has strongly limited their use for patch-clamp studies and thus our knowledge about the transporter composition of the lysosome membrane. The importance of patching endolysosomal membranes relates to the fact that, in animal cells, a series of channels have been reported to be localized on both lysosomal and/or endosomal membranes and have been predicted to play roles in signalling events and endomembrane fusion [21][22][23][24][25][26]. A detailed protocol reporting the use of enlarged lysosomes for patch-clamp was published in 2017 [27], but the first patch-clamp recording on endosomal membranes was made possible thanks to the expression of a hydrolysis-deficient SKD1/VPS4B (E235Q) protein in HEK293 cells, which induced the formation of enlarged endosomes (3–6 µm in diameter) by blocking their transition to lysosomes, hence making them accessible to the patch-clamp approach [28]. This strategy allowed the characterization of an endosomal Ca2+ channel whose activity was affected by the luminal Cl− concentration [28].

3. Approaches Based on Alternative Targeting and Heterologous Expression

3.1. Targeting to the Plasma Membrane upon Manipulation of Sorting Signals

Most organelles are not easily amenable to classical uptake experiments or electrophysiological recordings, hampering the analysis of intracellular ion and solute transport processes. One exception to this is the large central vacuole of plant and yeast cells, which is directly and easily accessible after rupture of the plasma membrane. One possibility to circumvent the problem of membrane accessibility is to manipulate the subcellular localization of intracellular transport proteins by altering their targeting route, thus redirecting them to the vacuolar or plasma membranes.
In the secretory or endocytic pathway, coordinated vesicle trafficking delivers transport proteins to the correct destination membrane. Transport vesicles recruit their transmembrane protein cargo upon interaction of adaptor proteins with sorting or internalization signals within the cytoplasmic regions of the cargo proteins. Sorting or internalization signals consist of short, linear motifs, post-translational modifications or three-dimensional structural motifs [29][30][31]. Among the linear sequence motifs, dileucine-based motifs [DE]xxxL[LI] and tyrosine-based motifs YxxØ (where Ø represents a bulky, hydrophobic amino acid) are common. They are recognized by clathrin adaptor complexes (AP1 to AP5), Golgi-localized, γ-ear containing, Arf-binding proteins (GGAs) and stonins 1 and 2. GGAs select the dileucine-motif variant DxxLL and specifically control vesicle sorting at the trans-Golgi network.

3.2. Nuclear Membrane Electrophysiology

The nuclear membrane may represent a novel heterologous system to express endo-lysosomal channels and transporters, as it provides relatively easy access to both the cytoplasmic and luminal sides of the membrane, so that ionic and ligand conditions can be rigorously controlled [33][34]. The technique, whose advantages and disadvantages have been summarized in Table 2, has been recently used to characterize the hTPC2 channel [35]. In The researchtheir study, the authors generated a stable DT40TKO cell line expressing hTPC2 and lacking both functional InsP3R and RyR, two intracellular Ca2+ channels. Using the nuclear membrane patch-clamp technique, they detected a ~220 pS single-channel current activated by NAADP with K+ as the permeant ion.

3.3. Patch-Clamp Electrophysiology on Plant Vacuoles

3.3.1. Sorting Routes and Signals to the Tonoplast and the Lysosomal Membrane

While performing more functions than the animal lysosome, i.e., storage of ions/metabolites and regulation of the turgor of the plant cell, the central lytic vacuole can be considered the lysosomal counterpart. This compartment (up to 40 μm in diameter) may occupy more than 80% of the cellular volume [36][37] in many cell types. Vacuoles are highly suitable for patch-clamp studies [38][39][40][41], because of simplicity of isolation and large size. For these reasons, many types of endogenous ion channels and transporters have been identified and characterized in great detail [36][42]. Besides macroscopic currents recorded in the whole-vacuole configuration [43][44], single channel events could be detected in excised vacuolar membrane patches, both in the cytosolic-side-out [45] and vacuolar-side-out configurations [46], Moreover, fluorescent indicator dyes have also been employed together with the patch-clamp technique [47][48][49][50][51]. The choice of ionic conditions and/or the use of appropriate Arabidopsis knock-out lines, allow to significantly reduce the density of the background currents, despite the presence of different types of endogenous ion transport systems.
From a topographic point of view, vacuoles and lysosomes occupy the same position within the secretory pathway: In general terms, they are one of the two endpoints of secretory traffic, the other being the cell surface. Like other secretory proteins, proteins of the tonoplast and the LM start their life in the ER [52]. The best characterized pathways to their final destinations pass through the Golgi apparatus and the trans-Golgi network (TGN). From there, a direct route proceeds via multivesicular bodies (MVB, also termed late endosomes) to the plant tonoplast or the LM, whereas an indirect route first leads to the plasma membrane and then to the final destination via endocytosis and MVB [53][54][55].
Membrane proteins are sorted along the endomembrane system through interactions between sorting motifs in their cytosolic domains and components of the different coat complexes that allow membrane traffic from one compartment to another. As Tintroduced in Section 3.1, two classes of motifs present in the cytosolic head or tail of membrane proteins have been clearly identified as lysosomal and tonoplast sorting signals: Dileucine-based [D/E]XXXL[L/I] or DXXLL, and tyrosine-based YXXØ, where X can be any amino acid and Ø represents a bulky hydrophobic residue [53][54]. These motifs interact with components of the AP1-4 or GGA adaptor complexes necessary to recruit clathrin or other membrane coats that promote the formation of coated vesicles, either from the TGN, the plasma membrane or between early and late endosomes (notice that, in plant cells, the TGN also play the role of animal early endosomes). Dileucine-based sorting signals for the tonoplast are recognized by AP1 [56], AP3 [57] or AP4 [58]. AP5 is involved in maintaining lysosome integrity [59], but its extact role has to be elucidate both in mammals and plants.
It should also be underlined that most membrane proteins also present ER-exit motifs that allow efficient initiation of traffic from this compartment, independently of their final destination. These diacidic (D/E-X-D/E), dihydrophobic or diaromatic (FF, YY, LL or FY) motifs interact with components of the COPII complex that initiates traffic from the ER (Barlowe, 2005; Marti et al., 2010). Finally, the ER quality control machinery usually prevents misfolded or unassembled newly synthesized polypeptides from trafficking [54]. Consistently with all these requirements for traffic and sorting, deletions or domain substitutions that destroy the tonoplast sorting signals often lead to mislocalization to the plasma membrane [60][61][62][57], whereas those that affect the ER-exit motifs or general folding result in ER retention [61][62][63]. The Golgi/TGN-mediated routes to the tonoplast and lysosomal membrane seem to use conserved mechanisms and signals [64]. This occurs despite the marked architectural differences in key compartments of the secretory pathway, perhaps most strikingly the Golgi apparatus: It is a single perinuclear membrane complex controlled by microtubules in most animal cells as opposed to a system composed of hundreds of apparently independent Golgi units moving along actin filaments all over any plant cell analyzed. These features rise as yet unresolved questions about the conservation of ER-to-Golgi and Golgi-to-endosome traffic mechanisms [65][66] and may also be related to the origin of the Golgi-bypassing routes to the tonoplast mentioned above. It has also been determined that potential N-glycosylation sequons are much less frequent in tonoplast proteins than in those of the plant plasma membrane, and that Golgi-modified Asn-linked oligosaccharide chains, abundantly present in plant plasma membrane proteins, are not detactable in tonoplast proteins of the same cells [67]. The major lysosomal membrane proteins are instead extensively N-glycosylated with oligosaccharide chains modified by Golgi enzymes. This “glycocalyx” is believed to protect the luminal loops of membrane proteins from degradation by lysosomal proteases [68]. In silico analysis of proteomes suggests that protection from vacuolar proteases has instead evolved by limiting the length of luminal domains of tonoplast proteins [67]. It is finally evident that a single large vacuole that occupies most of the cellular space in many fully expanded plant cells is quite different from the myriad of small lysosomes, at least in terms of surface/volume ratio.

3.3.2. The Plant Vacuole as a Heterologous Expression System of Lysosomal Channels and Transporters

In line with similarity discussed above in trafficking and targeting between tonoplast and lysosomal membrane proteins, mutants plants from Arabidopsis thaliana lacking specific endogenous vacuolar channels or transporters can be used for the expression of the respective homologous animal lysosomal proteins (interestingly, oocytes of Xenopus laevis are the system of choice for ion channels and transporters localized in the plasma membrane [69][70][71][72][73]). Plants of Arabidopsis can be grown on soil in a growth chamber under controlled light and temperature conditions. The cDNA of the animal intracellular channel or transporter is cloned into a suitable plant expression vector conveying high protein expression. Fusion with a fluorescent marker may be helpful to verify the expression and localization of the protein. By using a well-established protocol [74], protoplasts can be transiently transformed; see for a schematic overview. The efficiency of transformation can be estimated by GFP fluorescence of the protoplasts. After one to four days, vacuoles can be easily released from transformed protoplasts for subsequent patch-clamp experiments.

3.4. Patch-Clamp Electrophysiology on Giant Vacuoles from Yeast Cells

Escherichia coli and budding yeast (Saccharomyces cerevisiae) are frequently used to heterologously express plasma membrane proteins as complement assay to analyze their transport activity, similarly to other commonly used heterologous expression systems, such as Xenopus laevis oocytes and mammalian cell cultures. Budding yeast, a unicellular eukaryote, has a vacuole similar to plant cells. Its cell size is too small for whole-cell or whole- vacuolar patch-clamp measurements. Nevertheless, in 1990, vacuoles prepared from a tetraploid yeast strain, which are larger than haploid yeast cells, were employed and the vacuolar cation channel YVC1 was characterized [75]. As an alternative method for patch-clamp experiments on microorganisms, Yabe and co-workers developed a method to generate “giant” E. coli protoplasts (spheroplasts), as large as 5 to 10 μm in diameter, by digesting the cell wall by enzymatic treatment and addition of a peptidoglycan synthase inhibitor in the following incubation (named Spheroplast Incubation or SI method) [76]. Giant E. coli was used for patch-clamp recordings of H+ pump activity of respiratory chain F0 F1-ATPase [77]. Furthermore, the SI method was modified for yeast giant cell preparation using a 1,3-β-d-glucan synthase inhibitor. Enlarged yeast contained a huge central vacuole which apparently occupied more than 80% of the cellular volume. Using this system, the activity of yeast V-type H+-ATPase was evaluated directly from the vacuolar membrane [78]. Interestingly, the elaborate methods for inactivation of individual genes encoding endogenous channels in yeast enable the vacuole membrane to convert into a suitable expression platform with low background activity, providing a high signal-to-noise ratio for precise characterization of ion channels. In fact, various plant vacuolar proteins have been characterized using a biochemical approach with yeast mutant vacuoles [79][80][81]. This includes the detailed characterization of mung bean (Vigna radiata) proton-pumping pyrophosphatase (H+-PPase) [82] and the vacuole-localized K+ channel NtTPK1 from tobacco (Nicotiana tabacum cv. SR1) [83][84].

4. Outlook on Novel Techniques Complementing Direct Functional Studies

4.1. Cryo-Electron Microscopy

Cryo-electron microscopy (cryo-EM) has the ability to provide 3D structural information of biological molecules and assemblies by imaging non-crystalline specimens (single particles). Latest advances in detector technology and software algorithms have allowed the determination of biomolecular structures at near-atomic resolution [85][86].

4.2. Molecular Dynamics Simulations

X-ray crystallography and more recently cryo-EM provide us with an ever increasing number of atomic-resolution structures of membrane channels and transporters, allowing molecular dynamics (MD) simulations to study the mechanisms underlying their behavior [87][88][89]. Moreover, the increasing computational power permits simulations reaching tens or hundreds of microseconds, which are time scales approaching those of electrophysiological measurements.

4.3. Genome Editing

Genome editing (also called gene editing) is a group of technologies that give scientists the ability to change an organism’s DNA. These technologies allow genetic material to be added, removed or altered at particular locations in the genome. Several approaches to genome editing have been developed. A recent one, known as CRISPR, has generated a lot of excitement in the scientific community because it is faster, cheaper, more accurate and more efficient than other existing genome editing methods [90].

4.4. Nanoscopy

Stimulated emission depletion (STED) microscopy is one of the techniques that make up super-resolution microscopy. It creates super-resolution images by the selective deactivation of fluorophores, minimizing the area of illumination at the focal point and thus enhancing the achievable resolution for a given system [91] bypassing the diffraction limit of light microscopy to increase resolution. This technique was used to reveal the close physical relationship between clusters of ryanodine receptors (RyRs) in the terminal cisternae of the sarcoplasmic reticulum (SR) and TPC2 channels on the lysosomal membrane. It has been proposed that TPC2-RyR clusters act as “trigger zones” in which TPCs are stimulated to create highly localized elementary Ca2+ signals that subsequently lead to the opening of the RyR in the SR/ER membrane, resulting in global signals via Ca2+-induced Ca2+ release [92].
This entry is adapted from 10.3390/cells11060921

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