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Cell Biology and Physiology of CLC Chloride Channels and Transporters

Tobias Stauber, Stefanie Weinert, Thomas J. Jentsch

10.1002/cphy.c110038

Source: Volume 2, Issue 3, July 2012

Published online: July 2012

Full Article on Wiley Online Library



Abstract

Proteins of the CLC gene family assemble to homo‐ or sometimes heterodimers and either function as Cl– channels or as Cl–/H+‐exchangers. CLC proteins are present in all phyla. Detailed structural information is available from crystal structures of bacterial and algal CLCs. Mammals express nine CLC genes, four of which encode Cl– channels and five 2Cl–/H+‐exchangers. Two accessory β‐subunits are known: (1) barttin and (2) Ostm1. ClC‐Ka and ClC‐Kb Cl– channels need barttin, whereas Ostm1 is required for the function of the lysosomal ClC‐7 2Cl–/H+‐exchanger. ClC‐1, ‐2, ‐Ka and ‐Kb Cl– channels reside in the plasma membrane and function in the control of electrical excitability of muscles or neurons, in extra‐ and intracellular ion homeostasis, and in transepithelial transport. The mainly endosomal/lysosomal Cl–/H+‐exchangers ClC‐3 to ClC‐7 may facilitate vesicular acidification by shunting currents of proton pumps and increase vesicular Cl– concentration. ClC‐3 is also present on synaptic vesicles, whereas ClC‐4 and ‐5 can reach the plasma membrane to some extent. ClC‐7/Ostm1 is coinserted with the vesicular H+‐ATPase into the acid‐secreting ruffled border membrane of osteoclasts. Mice or humans lacking ClC‐7 or Ostm1 display osteopetrosis and lysosomal storage disease. Disruption of the endosomal ClC‐5 Cl–/H+‐exchanger leads to proteinuria and Dent's disease. Mouse models in which ClC‐5 or ClC‐7 is converted to uncoupled Cl– conductors suggest an important role of vesicular Cl– accumulation in these pathologies. The important functions of CLC Cl– channels were also revealed by human diseases and mouse models, with phenotypes including myotonia, renal loss of salt and water, deafness, blindness, leukodystrophy, and male infertility. © 2012 American Physiological Society. Compr Physiol 2:1701‐1744, 2012.

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Figure 1. Figure 1.

The mammalian CLC family of chloride channels and transporters. The CLC family comprises nine members in mammals. This overview depicts their tissue expression (and of the appropriate β‐subunits (in red)), cellular function, and known human and mouse pathologies. The members of the first branch of the CLC family, ClC‐1, ‐2, ‐Ka, and ‐Kb, are plasma membrane Cl– channels. By contrast, the members of the second (ClC‐3, ‐4, and ‐5) and third (ClC‐6 and ‐7) subfamilies are Cl–/H+‐exchangers that localize predominantly to intracellular compartments of the endosomal/lysosomal pathway.
Figure 2. Figure 2.

Spongiform vacuolation in the white matter of ClC‐2 knockout (KO) mice. Semithin brain sections of the middle cerebellar peduncle of 2‐, 5‐, and 14‐month‐old wildtype (WT) and ClC‐2 KO mice show abundant vacuoles (asterisks) in the cerebellar white matter of 5‐ and 14‐month‐old KO but not WT animals. C, capillaries; scale bar, 20 μm. [Image adapted from, reference (35), with permission.]
Figure 3. Figure 3.

Roles of ClC‐K/barttin in transepithelial transport. (A) Schematic representation of renal NaCl reabsorption in the thick ascending limb (TAL) of Henle. Active transport by the basolateral Na,K‐ATPase drives NaCl uptake by the apical NKCC2 transporter. Cotransported K+ is recycled through the apical ROMK channel and Cl– leaves the cell through basolateral ClC‐Kb/barttin. (B) Schematic representation of K+ secretion in the stria vascularis of the cochlea. K+ is taken up by the basolateral NKCC1 transporter and the Na,K‐ATPase. While K+ exits through the apical KCNQ1/KCNE1 channel, Cl– is recycled by basolateral ClC‐Ka/barttin and ClC‐Kb/barttin channels.
Figure 4. Figure 4.

ClC‐K/barttin in the cochlea. (A) Model of potassium recycling in the cochlea of the inner ear. The endolymph in the cavity of the scala media displays a high K+ concentration of 140 mmol/L and a positive potential of +100 mV. Both the high [K+] and the potential are established by the stria vascularis (light blue). Both parameters are important for the depolarizing K+ current through mechanosensitive channels in the apical membrane of inner (red) and outer (green) hair cells. K+ leaves these sensory cells basolaterally into the perilymph, which displays the zero potential and low [K+] of normal extracellular space. While the perilymph is separated from the endolymph by tight junctions, potassium is transported back to the stria vascularis through a gap junction system. (B) The scheme represents a model of how potassium is secreted into the endolymph through the stria vascularis. In this multilayered epithelium, a layer of marginal cells, which are connected by tight junctions and apically face the endolymph, and a layer of basal cells, also connected by tight junctions, isolate an intrastrial space with a low K+ concentration. K+ enters this space through Kir4.1 from intermediate cells, which can receive K+ through a system of gap junctions, and it is taken up by marginal cells that secrete it into the endolymph. Cl– exits the marginal cells through basolateral ClC‐K/barttin channels. The presence of another, unidentified Cl– channel on the basolateral side is suggested by the normal endocochlear K+ concentration upon inner‐ear‐specific barttin deletion. [Models modified from, reference (343), with permission.]
Figure 5. Figure 5.

ClC‐K/barttin in the cochlea. (A) Model of potassium recycling in the cochlea of the inner ear. The endolymph in the cavity of the scala media displays a high K+ concentration of 140 mmol/L and a positive potential of +100 mV. Both the high [K+] and the potential are established by the stria vascularis (light blue). Both parameters are important for the depolarizing K+ current through mechanosensitive channels in the apical membrane of inner (red) and outer (green) hair cells. K+ leaves these sensory cells basolaterally into the perilymph, which displays the zero potential and low [K+] of normal extracellular space. While the perilymph is separated from the endolymph by tight junctions, potassium is transported back to the stria vascularis through a gap junction system. (B) The scheme represents a model of how potassium is secreted into the endolymph through the stria vascularis. In this multilayered epithelium, a layer of marginal cells, which are connected by tight junctions and apically face the endolymph, and a layer of basal cells, also connected by tight junctions, isolate an intrastrial space with a low K+ concentration. K+ enters this space through Kir4.1 from intermediate cells, which can receive K+ through a system of gap junctions, and it is taken up by marginal cells that secrete it into the endolymph. Cl– exits the marginal cells through basolateral ClC‐K/barttin channels. The presence of another, unidentified Cl– channel on the basolateral side is suggested by the normal endocochlear K+ concentration upon inner‐ear‐specific barttin deletion. [Models modified from, reference (343), with permission.]
Figure 6. Figure 6.

Localization of the intracellular CLC proteins to the endosomal/lysosomal pathway. The scheme illustrates the proposed subcellular localizations of the members of the second and third CLC subfamilies. While ClC‐5 localizes to early compartments of the endocytic pathway, ClC‐3 and ‐6 localize on late endosomes. The localization of ClC‐4 is less clear, and ClC‐7/Ostm1 is the only lysosomal CLC protein. The ATP‐consuming proton pump acidifies the compartments, from the extracellular pH 7.4 to pH 4.5 in lysosomes. At least in early endosomal compartments, the shunt current is provided by the CLC proteins. For all intracellular CLCs except ClC‐3, a Cl–/H+‐exchange activity has been shown. For ClC‐7, this activity has been shown to accumulate chloride in lysosomes.
Figure 7. Figure 7.

Neuronal degeneration in ClC‐3 KO mice. (A) Nissl‐stained frontal brain sections show severe neuronal cell loss that leads to a complete loss of the hippocampus in adult ClC‐3 KO mice. (B) Retinal sections of P28 WT and ClC‐3 KO mice reveal a complete degeneration of photoreceptors, with a loss of the photoreceptor outer segment (OS) as well as the nuclei that form the outer nuclear layer (ONL). RPE, retinal pigment epithelium; IS, photoreceptor inner segments; OPL, outer plexiform layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer. [Images adapted from, reference (402), with permission.]
Figure 8. Figure 8.

Basic electrophysiological properties of ClC‐4. (A) Two‐electrode voltage‐clamp traces of ClC‐4 expressed in Xenopus oocytes show strongly outwardly rectifying chloride currents which a rapidly activated at positive voltages. (B) Neutralizing the “gating glutamate” in ClC‐4 converts the Cl–/H+‐exchanger into a pure Cl– conductor which results in significant currents also in the negative voltage range. [Traces taken from, reference (119), with permission.]
Figure 9. Figure 9.

Proximal tubule endocytosis defect in ClC‐5 KO mice. Fluorescently labeled lactoglobulin (in blue) was injected into female Clcn5+/– mice. Due to random X‐chromosomal inactivation in females, proximal tubular cells (PTCs) either express (ClC‐5 stained in red) or lack ClC‐5 (indicated by arrows). Cells expressing ClC‐5 accumulated substantial amounts of lactoglobulin below the brush border while neighboring ClC‐5 KO cells had taken up much less protein indicating a reduced receptor‐mediated endocytosis in a cell‐autonomous manner. [Images modified from, reference (324), with permission.]
Figure 10. Figure 10.

Model for renal pathology caused by an impaired proximal tubular endocytosis as observed in ClC‐5 KO mice and in patients with Dent's disease. (A) Events in the early proximal tubule: After the small peptide PTH (parathyroid hormone) and various forms of vitamin D have passed the glomerular filter, loss of ClC‐5 results in an impaired megalin‐mediated endocytosis of PTH that would have been normally degraded in lysosomes. The uptake of 1,25(OH)2‐ and 25(OH)‐vitaminD3 together with their binding protein DBP, which requires binding to megalin, is also impaired (indicated by red minus symbols). Under normal conditions 1,25‐VitD reaches VitD receptors that regulate the transcription of nuclear target genes (not shown) and 25‐VitD is metabolized by mitochondrial enzymes: 1α‐hydroxylase (1α‐HYD) converts the precursor 25(OH)‐vitaminD3 to the active hormone 1,25(OH)2‐vitaminD3 (1,25‐VitD), whereas the 24‐hydroxylase (24‐HYD) inactivates 1,25‐VitD. (B) Events in the late proximal tubule: The defect in ClC‐5‐dependent endocytosis results in an increased luminal PTH concentration which leads to an enhanced stimulation of apical PTH receptors (indicated by green plus symbols). The enhanced stimulation of PTH receptors together with the decreased uptake of 1,25(OH)2‐vitaminD3 enhances the transcription of the mitochondrial enzyme 1α‐hydroxylase that converts the precursor 25(OH)‐vitaminD3 into the active metabolite 1,25(OH)2‐vitaminD3, while decreasing transcription of the catabolizing enzyme 24‐hydroxylase. However, 25(OH)‐vitaminD3 megalin‐dependent endocytosis to reach the enzyme is impaired, thus both 1,25(OH)2‐ and 25(OH)‐vitaminD3 are lost into the urine. There is a balance between the changed transcription of enzymes that cooperate to increase active vitamin D3, and the loss of the precursor and active form into the urine. Depending on the outcome (increased or decreased concentration of active vitamin D3 in serum), this leads to different clinical phenotypes. (C) Events in the distal tubule: In contrast to proximal tubules, 1,25(OH)2‐vitaminD3 enters distal tubular cells mainly by diffusion of the free hormone over the membrane and binds to the vitamin D3 receptor which activates the transcription of 1,25(OH)2‐vitaminD3 dependent genes after heterodimerization with the retinoid X receptor (RXR). As the concentration of 1,25(OH)2‐vitaminD3 is increased in the lumen of ClC‐5 KO kidneys, renal 1,25(OH)2‐vitaminD3 target genes are selectively affected. VDR, vitamin D receptor; RXR, retinoid X receptor; PTH‐R, parathyroid hormone receptor. [Model adapted from, reference (244), with permission.]
Figure 11. Figure 11.

Models for endosomal acidification. According to the classical model (A), Cl– channels provide the electrical shunt for the proton‐pumping V‐ATPase in endosomes. But in reality, the Cl–/H+‐exchanger ClC‐5 performs this role (B). Neutralizing its “gating glutamate” uncouples the Cl− conductance from H+ countertransport, which results in a pure Cl– conductor (C), going back to “classics” (A). [Model adapted from, reference (291), with permission.]
Figure 12. Figure 12.

Renal endosomal acidification of mice converting ClC‐5 into a chloride conductor. (A) Averaged traces of acridine orange fluorescence comparing ATP‐driven acidification of endosomal fractions from renal cortex of WT (green) and ClC‐5unc (red) (left panel) and WT and Clcn5– (black) mice (right panel). Acidification of WT and ClC‐5unc vesicles occurred with similar efficiency but was severely reduced with endosomes from mice in which ClC‐5 was lacking. (B) Fluorescently labeled dextran, a marker for fluid‐phase endocytosis (green), was injected into female Clcn5+/unc mice. Cells expressing ClC‐5unc accumulated much less dextran than neighboring cells expressing the 2Cl–/H+‐exchanger ClC‐5 (for details, see (291)). [Images taken from, reference (291), with permission.]
Figure 13. Figure 13.

Minimal mathematical model for ATP‐dependent vesicle acidification. Mathematical modeling for the ATP‐dependent acidification of a minimal vesicle containing just an H+ pump, an H+ leak, and either no Cl– conductance (−/−; black traces), a 2Cl–/H+‐exchanger (+/+; green traces) or a Cl– channel (unc/unc; red traces). For details, see (448). (A) Simulations predict a more efficient vesicular acidification with the exchanger despite H+ efflux due to Cl–/H+ antiport compared to the acidification predicted with a Cl– channel. There is no acidification without any Cl– conductance. (B) Upon acidification an inside‐positive potential is predicted with a Cl– channel whereas the lumen becomes negative with a Cl–/H+‐exchanger. (C) The calculated intravesicular Cl– concentration is higher with a Cl–/H+‐exchanger than with a Cl– channel. The initial conditions were: pHi = 7.2; pHo = 7.2; [cation+]i = 140 mmol/L; [cation+]o = 140 mmol/L; [Cl–]i = 50 mmol/L; [Cl–]o = 50 mmol/L; U = 0 mV. [Graphs adapted from, reference (448), with permission.]
Figure 14. Figure 14.

Neuronal storage material in ClC‐6 and ClC‐7 KO mice. Electron micrography shows the accumulation of electron‐dense deposits (arrows) in ClC‐6‐deficient hippocampal neurons and in cortical neurons of ClC‐7 KO mice. The localization of storage material is restricted to the initial axon segment in ClC‐6‐deficient neurons, whereas it is scattered through the entire soma of ClC‐7 KO neurons. nuc, nucleus; scale bars, 2 μm. [Images modified from, references (327) (left panel) and (183) (right panel) , with permission.]
Figure 15. Figure 15.

Subcellular localization of ClC‐7 and Ostm1 in mouse fibroblasts. Top panel: Costaining for ClC‐7 (green) and the lysosomal marker lamp‐1 (red) revealed localization of ClC‐7 on lysosomal compartments in mouse fibroblasts. Lower panel: Immunofluorescence of ClC‐7 (green) and its β‐subunit Ostm1 (red) showed colocalization of the two subunits. [Images modified from, reference (448), with permission.]
Figure 16. Figure 16.

Bone phenotype of ClC‐7 mouse models. Microcomputed tomography images of tibiae show increasing severity of osteopetrosis from Clcn7unc/unc to Clcn7–/– mice, which even lack the bone marrow cavity. [Images adapted from, reference (448), with permission.]
Figure 17. Figure 17.

Enlarged late endosomal/lysosomal compartments in ClC‐7‐deficient PTCs. Kidney sections from mice with a chimeric ClC‐7 deletion in renal proximal tubules were immunostained for ClC‐7 (green), the late endosomal/lysosomal protein lamp‐1 (red) and the proximal tubule marker villin (blue). Lamp‐1‐positive vesicles are drastically enlarged in ClC‐7‐deficient cells (KO) compared to their appearance in ClC‐7‐expressing cells (WT). [Images modified from, reference (445), with permission.]
Figure 18. Figure 18.

Lysosomal transport characterization. carbonyl cyanide 3‐chlorophenylhydrazone (CCCP)

‐induced alkalinization assay of lysosomes that were preloaded with the ratiometric pH indicator Oregon Green‐dextran. The protononophore rapidly dissipated lysosomal pH in Clcn7+/+ (green), Clcn7unc/unc (red), and Clcn7–/– (black) fibroblasts, demonstrating the presence of countercurrents. Lysosomes of Clcn7unc/unc mice reached a more alkaline pH than Clcn7–/– lysosomes, suggesting that the uncoupled transporter (ClC‐7unc) mediates a Cl– conductance. However, Clcn7unc/unc lysosomes reached a less alkaline steady‐state pH than WT lysosomes. Since the steady‐state pH is determined by voltage, which depends on the Cl– diffusion potential, the difference in lysosomal pH suggests higher lysosomal chloride concentration in WT than in Clcn7unc/unc lysosomes. Alkalinization of Clcn7–/– lysosomes by protonophore treatment argues for lysosomal conductances in addition to ClC‐7 most likely by a so far unknown cation conductance. Mathematical modeling of such an experiment (on the lower right) confirms the plausibility of the conclusions (for details, see (448)). The simulated vesicle contains either a Cl–/H+‐exchange activity, or a Cl– conductance or none of these, corresponding to the three genotypes (same color code as in the experimental data). In addition, the vesicle contains a cation conductance and a large proton conductance simulating CCCP. The initial conditions were: pHi = 4.75; pHo = 6.4; [cation+]i = 100 mmol/L; [cation+]o = 10 mmol/L; [Cl–]i = 120 or 80 mmol/L as indicated; [Cl–]o = 20 mmol/L. With the same initial [Cl–], the luminal pH of vesicles containing a Cl–/H+‐exchange activity or a Cl– conductance reaches the same value. Lower initial luminal [Cl–] in vesicles with a Cl– conductance may explain the different final pH values. [Graphs taken from, reference (448), with permission.]


Figure 1.

The mammalian CLC family of chloride channels and transporters. The CLC family comprises nine members in mammals. This overview depicts their tissue expression (and of the appropriate β‐subunits (in red)), cellular function, and known human and mouse pathologies. The members of the first branch of the CLC family, ClC‐1, ‐2, ‐Ka, and ‐Kb, are plasma membrane Cl– channels. By contrast, the members of the second (ClC‐3, ‐4, and ‐5) and third (ClC‐6 and ‐7) subfamilies are Cl–/H+‐exchangers that localize predominantly to intracellular compartments of the endosomal/lysosomal pathway.



Figure 2.

Spongiform vacuolation in the white matter of ClC‐2 knockout (KO) mice. Semithin brain sections of the middle cerebellar peduncle of 2‐, 5‐, and 14‐month‐old wildtype (WT) and ClC‐2 KO mice show abundant vacuoles (asterisks) in the cerebellar white matter of 5‐ and 14‐month‐old KO but not WT animals. C, capillaries; scale bar, 20 μm. [Image adapted from, reference (35), with permission.]



Figure 3.

Roles of ClC‐K/barttin in transepithelial transport. (A) Schematic representation of renal NaCl reabsorption in the thick ascending limb (TAL) of Henle. Active transport by the basolateral Na,K‐ATPase drives NaCl uptake by the apical NKCC2 transporter. Cotransported K+ is recycled through the apical ROMK channel and Cl– leaves the cell through basolateral ClC‐Kb/barttin. (B) Schematic representation of K+ secretion in the stria vascularis of the cochlea. K+ is taken up by the basolateral NKCC1 transporter and the Na,K‐ATPase. While K+ exits through the apical KCNQ1/KCNE1 channel, Cl– is recycled by basolateral ClC‐Ka/barttin and ClC‐Kb/barttin channels.



Figure 4.

ClC‐K/barttin in the cochlea. (A) Model of potassium recycling in the cochlea of the inner ear. The endolymph in the cavity of the scala media displays a high K+ concentration of 140 mmol/L and a positive potential of +100 mV. Both the high [K+] and the potential are established by the stria vascularis (light blue). Both parameters are important for the depolarizing K+ current through mechanosensitive channels in the apical membrane of inner (red) and outer (green) hair cells. K+ leaves these sensory cells basolaterally into the perilymph, which displays the zero potential and low [K+] of normal extracellular space. While the perilymph is separated from the endolymph by tight junctions, potassium is transported back to the stria vascularis through a gap junction system. (B) The scheme represents a model of how potassium is secreted into the endolymph through the stria vascularis. In this multilayered epithelium, a layer of marginal cells, which are connected by tight junctions and apically face the endolymph, and a layer of basal cells, also connected by tight junctions, isolate an intrastrial space with a low K+ concentration. K+ enters this space through Kir4.1 from intermediate cells, which can receive K+ through a system of gap junctions, and it is taken up by marginal cells that secrete it into the endolymph. Cl– exits the marginal cells through basolateral ClC‐K/barttin channels. The presence of another, unidentified Cl– channel on the basolateral side is suggested by the normal endocochlear K+ concentration upon inner‐ear‐specific barttin deletion. [Models modified from, reference (343), with permission.]



Figure 5.

ClC‐K/barttin in the cochlea. (A) Model of potassium recycling in the cochlea of the inner ear. The endolymph in the cavity of the scala media displays a high K+ concentration of 140 mmol/L and a positive potential of +100 mV. Both the high [K+] and the potential are established by the stria vascularis (light blue). Both parameters are important for the depolarizing K+ current through mechanosensitive channels in the apical membrane of inner (red) and outer (green) hair cells. K+ leaves these sensory cells basolaterally into the perilymph, which displays the zero potential and low [K+] of normal extracellular space. While the perilymph is separated from the endolymph by tight junctions, potassium is transported back to the stria vascularis through a gap junction system. (B) The scheme represents a model of how potassium is secreted into the endolymph through the stria vascularis. In this multilayered epithelium, a layer of marginal cells, which are connected by tight junctions and apically face the endolymph, and a layer of basal cells, also connected by tight junctions, isolate an intrastrial space with a low K+ concentration. K+ enters this space through Kir4.1 from intermediate cells, which can receive K+ through a system of gap junctions, and it is taken up by marginal cells that secrete it into the endolymph. Cl– exits the marginal cells through basolateral ClC‐K/barttin channels. The presence of another, unidentified Cl– channel on the basolateral side is suggested by the normal endocochlear K+ concentration upon inner‐ear‐specific barttin deletion. [Models modified from, reference (343), with permission.]



Figure 6.

Localization of the intracellular CLC proteins to the endosomal/lysosomal pathway. The scheme illustrates the proposed subcellular localizations of the members of the second and third CLC subfamilies. While ClC‐5 localizes to early compartments of the endocytic pathway, ClC‐3 and ‐6 localize on late endosomes. The localization of ClC‐4 is less clear, and ClC‐7/Ostm1 is the only lysosomal CLC protein. The ATP‐consuming proton pump acidifies the compartments, from the extracellular pH 7.4 to pH 4.5 in lysosomes. At least in early endosomal compartments, the shunt current is provided by the CLC proteins. For all intracellular CLCs except ClC‐3, a Cl–/H+‐exchange activity has been shown. For ClC‐7, this activity has been shown to accumulate chloride in lysosomes.



Figure 7.

Neuronal degeneration in ClC‐3 KO mice. (A) Nissl‐stained frontal brain sections show severe neuronal cell loss that leads to a complete loss of the hippocampus in adult ClC‐3 KO mice. (B) Retinal sections of P28 WT and ClC‐3 KO mice reveal a complete degeneration of photoreceptors, with a loss of the photoreceptor outer segment (OS) as well as the nuclei that form the outer nuclear layer (ONL). RPE, retinal pigment epithelium; IS, photoreceptor inner segments; OPL, outer plexiform layer; INL, inner nuclear layer; IPL, inner plexiform layer; GCL, ganglion cell layer. [Images adapted from, reference (402), with permission.]



Figure 8.

Basic electrophysiological properties of ClC‐4. (A) Two‐electrode voltage‐clamp traces of ClC‐4 expressed in Xenopus oocytes show strongly outwardly rectifying chloride currents which a rapidly activated at positive voltages. (B) Neutralizing the “gating glutamate” in ClC‐4 converts the Cl–/H+‐exchanger into a pure Cl– conductor which results in significant currents also in the negative voltage range. [Traces taken from, reference (119), with permission.]



Figure 9.

Proximal tubule endocytosis defect in ClC‐5 KO mice. Fluorescently labeled lactoglobulin (in blue) was injected into female Clcn5+/– mice. Due to random X‐chromosomal inactivation in females, proximal tubular cells (PTCs) either express (ClC‐5 stained in red) or lack ClC‐5 (indicated by arrows). Cells expressing ClC‐5 accumulated substantial amounts of lactoglobulin below the brush border while neighboring ClC‐5 KO cells had taken up much less protein indicating a reduced receptor‐mediated endocytosis in a cell‐autonomous manner. [Images modified from, reference (324), with permission.]



Figure 10.

Model for renal pathology caused by an impaired proximal tubular endocytosis as observed in ClC‐5 KO mice and in patients with Dent's disease. (A) Events in the early proximal tubule: After the small peptide PTH (parathyroid hormone) and various forms of vitamin D have passed the glomerular filter, loss of ClC‐5 results in an impaired megalin‐mediated endocytosis of PTH that would have been normally degraded in lysosomes. The uptake of 1,25(OH)2‐ and 25(OH)‐vitaminD3 together with their binding protein DBP, which requires binding to megalin, is also impaired (indicated by red minus symbols). Under normal conditions 1,25‐VitD reaches VitD receptors that regulate the transcription of nuclear target genes (not shown) and 25‐VitD is metabolized by mitochondrial enzymes: 1α‐hydroxylase (1α‐HYD) converts the precursor 25(OH)‐vitaminD3 to the active hormone 1,25(OH)2‐vitaminD3 (1,25‐VitD), whereas the 24‐hydroxylase (24‐HYD) inactivates 1,25‐VitD. (B) Events in the late proximal tubule: The defect in ClC‐5‐dependent endocytosis results in an increased luminal PTH concentration which leads to an enhanced stimulation of apical PTH receptors (indicated by green plus symbols). The enhanced stimulation of PTH receptors together with the decreased uptake of 1,25(OH)2‐vitaminD3 enhances the transcription of the mitochondrial enzyme 1α‐hydroxylase that converts the precursor 25(OH)‐vitaminD3 into the active metabolite 1,25(OH)2‐vitaminD3, while decreasing transcription of the catabolizing enzyme 24‐hydroxylase. However, 25(OH)‐vitaminD3 megalin‐dependent endocytosis to reach the enzyme is impaired, thus both 1,25(OH)2‐ and 25(OH)‐vitaminD3 are lost into the urine. There is a balance between the changed transcription of enzymes that cooperate to increase active vitamin D3, and the loss of the precursor and active form into the urine. Depending on the outcome (increased or decreased concentration of active vitamin D3 in serum), this leads to different clinical phenotypes. (C) Events in the distal tubule: In contrast to proximal tubules, 1,25(OH)2‐vitaminD3 enters distal tubular cells mainly by diffusion of the free hormone over the membrane and binds to the vitamin D3 receptor which activates the transcription of 1,25(OH)2‐vitaminD3 dependent genes after heterodimerization with the retinoid X receptor (RXR). As the concentration of 1,25(OH)2‐vitaminD3 is increased in the lumen of ClC‐5 KO kidneys, renal 1,25(OH)2‐vitaminD3 target genes are selectively affected. VDR, vitamin D receptor; RXR, retinoid X receptor; PTH‐R, parathyroid hormone receptor. [Model adapted from, reference (244), with permission.]



Figure 11.

Models for endosomal acidification. According to the classical model (A), Cl– channels provide the electrical shunt for the proton‐pumping V‐ATPase in endosomes. But in reality, the Cl–/H+‐exchanger ClC‐5 performs this role (B). Neutralizing its “gating glutamate” uncouples the Cl− conductance from H+ countertransport, which results in a pure Cl– conductor (C), going back to “classics” (A). [Model adapted from, reference (291), with permission.]



Figure 12.

Renal endosomal acidification of mice converting ClC‐5 into a chloride conductor. (A) Averaged traces of acridine orange fluorescence comparing ATP‐driven acidification of endosomal fractions from renal cortex of WT (green) and ClC‐5unc (red) (left panel) and WT and Clcn5– (black) mice (right panel). Acidification of WT and ClC‐5unc vesicles occurred with similar efficiency but was severely reduced with endosomes from mice in which ClC‐5 was lacking. (B) Fluorescently labeled dextran, a marker for fluid‐phase endocytosis (green), was injected into female Clcn5+/unc mice. Cells expressing ClC‐5unc accumulated much less dextran than neighboring cells expressing the 2Cl–/H+‐exchanger ClC‐5 (for details, see (291)). [Images taken from, reference (291), with permission.]



Figure 13.

Minimal mathematical model for ATP‐dependent vesicle acidification. Mathematical modeling for the ATP‐dependent acidification of a minimal vesicle containing just an H+ pump, an H+ leak, and either no Cl– conductance (−/−; black traces), a 2Cl–/H+‐exchanger (+/+; green traces) or a Cl– channel (unc/unc; red traces). For details, see (448). (A) Simulations predict a more efficient vesicular acidification with the exchanger despite H+ efflux due to Cl–/H+ antiport compared to the acidification predicted with a Cl– channel. There is no acidification without any Cl– conductance. (B) Upon acidification an inside‐positive potential is predicted with a Cl– channel whereas the lumen becomes negative with a Cl–/H+‐exchanger. (C) The calculated intravesicular Cl– concentration is higher with a Cl–/H+‐exchanger than with a Cl– channel. The initial conditions were: pHi = 7.2; pHo = 7.2; [cation+]i = 140 mmol/L; [cation+]o = 140 mmol/L; [Cl–]i = 50 mmol/L; [Cl–]o = 50 mmol/L; U = 0 mV. [Graphs adapted from, reference (448), with permission.]



Figure 14.

Neuronal storage material in ClC‐6 and ClC‐7 KO mice. Electron micrography shows the accumulation of electron‐dense deposits (arrows) in ClC‐6‐deficient hippocampal neurons and in cortical neurons of ClC‐7 KO mice. The localization of storage material is restricted to the initial axon segment in ClC‐6‐deficient neurons, whereas it is scattered through the entire soma of ClC‐7 KO neurons. nuc, nucleus; scale bars, 2 μm. [Images modified from, references (327) (left panel) and (183) (right panel) , with permission.]



Figure 15.

Subcellular localization of ClC‐7 and Ostm1 in mouse fibroblasts. Top panel: Costaining for ClC‐7 (green) and the lysosomal marker lamp‐1 (red) revealed localization of ClC‐7 on lysosomal compartments in mouse fibroblasts. Lower panel: Immunofluorescence of ClC‐7 (green) and its β‐subunit Ostm1 (red) showed colocalization of the two subunits. [Images modified from, reference (448), with permission.]



Figure 16.

Bone phenotype of ClC‐7 mouse models. Microcomputed tomography images of tibiae show increasing severity of osteopetrosis from Clcn7unc/unc to Clcn7–/– mice, which even lack the bone marrow cavity. [Images adapted from, reference (448), with permission.]



Figure 17.

Enlarged late endosomal/lysosomal compartments in ClC‐7‐deficient PTCs. Kidney sections from mice with a chimeric ClC‐7 deletion in renal proximal tubules were immunostained for ClC‐7 (green), the late endosomal/lysosomal protein lamp‐1 (red) and the proximal tubule marker villin (blue). Lamp‐1‐positive vesicles are drastically enlarged in ClC‐7‐deficient cells (KO) compared to their appearance in ClC‐7‐expressing cells (WT). [Images modified from, reference (445), with permission.]



Figure 18.

Lysosomal transport characterization. carbonyl cyanide 3‐chlorophenylhydrazone (CCCP)

‐induced alkalinization assay of lysosomes that were preloaded with the ratiometric pH indicator Oregon Green‐dextran. The protononophore rapidly dissipated lysosomal pH in Clcn7+/+ (green), Clcn7unc/unc (red), and Clcn7–/– (black) fibroblasts, demonstrating the presence of countercurrents. Lysosomes of Clcn7unc/unc mice reached a more alkaline pH than Clcn7–/– lysosomes, suggesting that the uncoupled transporter (ClC‐7unc) mediates a Cl– conductance. However, Clcn7unc/unc lysosomes reached a less alkaline steady‐state pH than WT lysosomes. Since the steady‐state pH is determined by voltage, which depends on the Cl– diffusion potential, the difference in lysosomal pH suggests higher lysosomal chloride concentration in WT than in Clcn7unc/unc lysosomes. Alkalinization of Clcn7–/– lysosomes by protonophore treatment argues for lysosomal conductances in addition to ClC‐7 most likely by a so far unknown cation conductance. Mathematical modeling of such an experiment (on the lower right) confirms the plausibility of the conclusions (for details, see (448)). The simulated vesicle contains either a Cl–/H+‐exchange activity, or a Cl– conductance or none of these, corresponding to the three genotypes (same color code as in the experimental data). In addition, the vesicle contains a cation conductance and a large proton conductance simulating CCCP. The initial conditions were: pHi = 4.75; pHo = 6.4; [cation+]i = 100 mmol/L; [cation+]o = 10 mmol/L; [Cl–]i = 120 or 80 mmol/L as indicated; [Cl–]o = 20 mmol/L. With the same initial [Cl–], the luminal pH of vesicles containing a Cl–/H+‐exchange activity or a Cl– conductance reaches the same value. Lower initial luminal [Cl–] in vesicles with a Cl– conductance may explain the different final pH values. [Graphs taken from, reference (448), with permission.]

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Article Title: Cell Biology and Physiology of CLC Chloride Channels and Transporters
 

How to Cite

Tobias Stauber, Stefanie Weinert, Thomas J. Jentsch. Cell Biology and Physiology of CLC Chloride Channels and Transporters. Compr Physiol 2012, 2: 1701-1744. doi: 10.1002/cphy.c110038