Abstract.
Cl− channels are widely found anion pores that are regulated by a variety of signals and that play various roles. On the basis of molecular biologic findings, ligand-gated Cl− channels in synapses, cystic fibrosis transmembrane conductors (CFTRs) and ClC channel types have been established, followed by bestrophin and possibly by tweety, which encode Ca2+-activated Cl− channels. The ClC family has been shown to possess a variety of functions, including stabilization of membrane potential, excitation, cellvolume regulation, fluid transport, protein degradation in endosomal vesicles and possibly cell growth. The molecular structure of Cl− channel types varies from 1 to 12 transmembrane segments. By means of computer-based prediction, functional Cl− channels have been synthesized artificially, revealing that many possible ion pores are hidden in channel, transporter or unidentified hydrophobic membrane proteins. Thus, novel Cl−-conducting pores may be occasionally discovered, and evidence from molecular biologic studies will clarify their physiologic and pathophysiologic roles.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Cl− channels are expressed as selective anion pores, have been found in a variety of physiologic preparations and are ubiquitously distributed, including in oocytes. Cl− channels allow the passive diffusion of negatively charged ions along electrochemical gradients or the transport of positive charges, and they conduct other anions, such as HCO3 −, I−, SCN− and NO3 −. The physiological roles of Cl− channels in the plasma membrane and in the vesicular membrane of intracellular organelles were vigorously investigated in the 1980s. Measurement of Cl− current in various cells revealed that Cl− channel gates were regulated by membrane voltage, cell volume, extracellular ligands, intracellular ions (such as Ca2+, H+ or anions) or phosphorylation of residues by various protein kinases. Therefore, one method of classification of Cl− channels was based on their functions. To summarize the diversity of function, the following list may conducted on the basis of phenotype in vivo: (i) ligand-gated transmission in the post-synaptic membrane, (ii) stabilization of resting membrane potential in (skeletal) muscle, (iii) depolarization of smooth-muscle cells or possibly of retinal pigment epithelium, (iv) cell-volume regulation in various cells, (v) fluid transport in epithelia and (v) neutralization of H+ ions in lysosomal vesicles.
Identification of a molecule usually leads to a breakthrough in elucidating its function. γ-aminobutyric acid (GABA or glycine receptors are composed of heteromeric subunits forming a Cl− channel that plays roles in post-synaptic membranes, as mentioned above in (i). The most exciting topic in Cl− channel research was the cloning and functional expression of the first member of the ClC family, by Jentsch et al. [1]. Interestingly, mutations in the ClC family have revealed that the ClC family covers functional roles in diverse processes, as mentioned above in (ii) to (vi). Furthermore, the study of mice lacking some ClCs has taught us that morphogenesis is an additional role of Cl− channels in the normal development of organs [2]. These aspects of Cl− channels have been extensively discussed in a recent comprehensive review by Jentsch et al. [2]. In addition, Nilius [3–6] has extensively reviewed volume-sensitive and Ca2+-activated Cl− channels.
One method of classification of channel molecules is based on properties found by electrophysiologic analysis. Expression of ClC has revealed Cl− channels with conductance less than 10 pS. There are, however, three types of Cl− channels, as determined by their conductance in situ: small (<10 pS), middle (10–100 pS), and large (>100 pS [maxi-Cl−]) conductance channels. Some Cl− channels have different states of conductance, namely double- or multi-barreled type (table 1). Explanation of the relationship between variable magnitude of conductance and molecular structure is disclosed by extensive studies with mutations in ClC channels [7–9]. The molecular structure of Cl− channels is also diverse. The number of transmembrane segments (TMSs) of Cl− channels is variable; for example, CFTR [10] and ClC [2, 11] have 10 or 12 TMSs. The ClCA family, which has 5 TMSs, encodes middle-conductance, Ca2+-activated Cl− (CaC) channels [12, 13]. Newly found CaC channels include bestrophin (the vitelliform macular dystrophy [VMD] protein), which defines a new family of chloride channels [14] with 4 TMSs, and tweety, which encodes 5–6 TMSs [15]. The Isk channel encodes 1 TMS, forming a Cl− and K+ channel pore expressed in Xenopus oocytes [16].
Functional transporters or other channels occasionally allow anion conductance. An example is an amino acid, glutamate transporter expressed in Xenopus oocytes, in which a Cl− current has been observed after the binding of glutamate to the transporter [17]. Aquaporin-6 (AQP-6) encodes an Hg2+-sensitive water channel that is simultaneously expressed as a unique acid-dependent Cl− channel [18].
Crystal structural analysis performed on ClC channels has shown a unique homodimeric composition [19]. Interestingly, bacterial ClC shows a characteristics of Cl−/H+ transporters as well as Cl− channels [20]. Such a complex structure-function relation is found only in anion carrying protein, not in cation channels.
On the other hand, functional Cl− channel pores can be artificially designed and synthesized as 20–30 amino acids [21]. These peptides assemble to make a functional pore consisting of a cluster of 4 or 5 molecules. The proposed structure of the peptide shows an a-helix possessing hydrophilic alignment on one side and a hydrophobic cluster of amino acids on the other [21, 22]. As suggested by studies of peptide channels, construction of a non-selective anion or cation selective pore is easier than construction of a Na+ or K+ selective pore. Thus, the Cl− channel may be formed occasionally by unrelated clusters of TMSs that form a hydrophilic pore [23].
In this review, explanations of the molecular functions of Cl− channels are given, according to the above criteria, and consideration of the Cl− channel pore suggests the possibility of the future discovery of novel Cl− channels. Discussion of ligand-gated Cl− channels is omitted because of detailed reviews elsewhere [24, 25].
Diversity of Cl− channel function
Ligand-gated transmission in the post-synaptic membrane
There are a number of articles and reviews regarding the ligand-gated Cl− channel GABA and glycine receptor [26].
Stabilization of membrane potential
Excitation of the membrane is generally achieved by an influx of the cations Na+ and/or Ca2+. The subsequent recovery of membrane potential is driven by the efflux of K+ or by the influx of Cl−. The ClC-1 channel is a voltage-dependent type of channel that is a known example of a membrane potential stabilizer and is expressed in skeletal muscle. It has a single-channel conductance of 1 pS and contributes approximately 75% of the resting conductance of the muscle membrane, with a high open probability at negative membrane potential. The ClC-1 channel suppresses depolarizing inputs and stabilizes the membrane potentials [27–29]. Mutations in the ClC-1 channel cause either recessive or dominant congenital myotonia, in which the mutant ClC-1 channels act as dominant negative subunits of the ClC-1 channel pore [30]. Homodimers or heterodimers of mutants with wild-type subunits result in a dramatic shift in voltage dependency. A loss of membrane potential stabilization by the mutant leads to prolonged depolarization of excitable membrane, resulting in a myotonic phenotype. The ClC-2 channel is activated by hyperpolarization, acidic pH and swelling of the cell [31, 32]. Postsynaptic GABAA and glycine receptors are ligand-gated Cl− channels that yield hyperpolarizing or depolarizing currents, depending on the intracellular Cl− concentration [33]. Since the activity of ClC-2 determines intracellular Na+ and Cl− concentration [34], iCIC-2 is important in GABA and andglycine-induced hyperpolarization and depolarization. Activation of ClC-2 may help the ligandgated Cl− channels to yield hyperpolarizing currents. ClC-2 is thus assumed to play a role in membrane stabilization with hyperpolarizing transmission by GABA and glycine in neurons. The mapping of an epilepsy susceptibility locus close to the CLCN2 gene suggested that Clcn2-/ — mice might suffer from spontaneous seizures [35, 36]. However, the lack of ClC-2 in mice has resulted in retinal degeneration or male infertility without a lowered threshold of seizure by chemicals [37]. This is one example of the unexpected physiological role of Cl− channels revealed from work with knockout animals.
Depolarization
In contrast to excitable membrane in which voltage-dependent cation channels play an important role in suppressive depolarization, in non-excitable cells, such as smooth muscle cells, the opening of Cl− channels drives the membrane potential VCl, which is approximately −20 mV and which is excitable depolarization. When excitatory ligand binds to smooth-muscle cells, a rise in intracellular Ca2+ occurs in response to the second messenger. The CaC channel has been recorded in a variety of electrophysiologic preparations and is ubiquitously distributed across all cell types [38], although not always fully evident. In isometric tension recordings, niflumic acid, a blocker of the CaC channel, inhibited spontaneous contractions. The CaC channel has received further attention in the study of secretory, epithelial, smooth-muscle and neuronal cell types, but the properties and phenotypic functions of the CaC channel are not as well characterized as those for Ca2+-activated K (CaK) channels [38–40].
CaC and CaK channels are frequently coexpressed and coactivated by elevation of Ca2+ concentration within the cell. Simultaneous activation of CaC and CaK channels may help stabilize plasma membrane potential. For example, vascular smooth-muscle cells possess both channels: one of the CaK channels is a largeconductance channel, called the maxi-K channel when encoded by Drosophila slow poke, but the molecular nature of the CaC channel is not known. Spontaneous transient inward currents (STICs) are associated with the simultaneous opening of a CaC channel after the periodic release of Ca2+ from intracellular stores. STICs are often co-incident with spontaneous transient outward currents carried by CaK channels [38]. In the absence of functional maxi-K channels when a defect in the b-subunit occurs, smooth-muscle cells tend to be constricted, resulting in high blood pressure [41]. Assuming that the remaining CaC function is exaggerated in the absence of the maxi-K channel, CaC would contribute to excitation of smooth muscle, leading to contractions [38, 41].
Molecular identification of CaC channels is difficult, since CaC channels are usually detected in cells used for expression of exogenous complementary DNA (cDNA). Thus, variable molecules have emerged at candidates of CaC channels. Proposed structures of various candidates of Cl− channels are shown in figure 1.
mClCA1 is isolated from airway epithelia expressing a CaC channel current. An extensive comparison has been made between the currents evoked by mCLCA1 expression and native CaC channel currents in various smooth-muscle cells [12, 13, 42, 43]. Dithiothreitol, a blocker of mCLCA1, had no effect on CaC channels in murine portal vein cells; again, channel kinetics and channel modulation differed. Therefore, most subsequent studies concluded that mCLCA1 alone does not comprise the CaC channel [44] but is a subunit that aids the expression of endogenous CaC.
Proteins encoded by these genes might be involved in tumorigenesis. hCLCA2 is critical for endothelial adhesion of cancer cells as a complex with integrin b4 [45–47]. This is a novel and interesting mechanism because several studies have suggested that Cl− channels play a role in cell growth and apoptosis, as described below.
Recently, molecular identification of (probably small conductance) CaC channels was achieved [14]. VMD (Best disease; MIM 153700) is an early onset, autosomal dominant disorder in which accumulation of lipofuscinlike material within and beneath the retinal pigment epithelia is associated with a progressive loss of central vision. The gene mutated in VMD, VMD2, was identified in 1998 [48, 49]. Bestrophin, a VMD gene product, is homologous to at least 3 other proteins within the human genome, 4 in the Drosophila genome and 24 in the Caenorhabditis elegans genome. An aberrant electrooculogram is noted when depolarization of retinal pigment epithelial cells is reduced, suggesting that bestrophin encodes a channel contributing to membrane depolarization. Induction of the light peak requires a ‘light-peak substance’ that is secreted by the neurosensory retina [50]. Transduction of the signal that induces the light peak requires signaling across the retinal pigment epithelial cell from a proposed receptor at the apical surface of the cells to activate one or more Cl− channels in the basolateral plasma membrane of the retinal pigment epithelial cell. Bestrophin is involved in the small- to middle-conductance Cl− channel in the basolateral membrane, contributing to depolarization of the membrane and leading to granule secretion. Bestrophin encodes 4 or 6 TMSs and induces a larger Cl− current in HEK cells after an increase in intracellular Ca2+ released from the caged compound activated by light. Human, Drosophila or nematode bestrophins (1 and 2) have different types of current-voltage relationships [14]. Furthermore, the mutant in VMD showed a dominant negative subunit and coexpression of mutant and wildtype heteromeric bestrophin, which provides less current than the wild-type homopolymer [14]. Although HEK cells still possess endogenous Cl− channels, these results strongly indicate that bestrophin encodes CaC channels. Bestrophin is bound to protein phosphatase 2, and this mechanism is considered to be functionally regulated by phosphorylation/dephosphorylation by this enzyme [51]. Exogenous bestrophin increases the magnitude of the Cl− channel current in HEK cells or in HeLa cells after exposure to hypotonic solution. Bestrophin may encode channels responsible for volume-sensitive Cl− channel current proposed to be <6 pS or may help to express these channels in the retinal pigment epithelial cells [52].
The functional role of CaC in vivo will be clarified in future analyses of these molecules.
Cell-volume regulation
When cells are exposed to hypotonic media, the cells gain volume by an influx of water, and an increase in intracellular Ca becomes evident [53]. This increase leads to loss of KCl via activation of Cl− and K+ channels, resulting in a loss of intracellular osmolarity as equilibrium with extracellular hypotonicity is reached. Subsequently, swollen cells recover normal volume (regulatory volume decrease). In early experiments, Ca2+-dependent channels were considered to play a central role in volume regulation, but studies have since identified other volumesensitive Cl− channels (VDCCs) or K+ channels that are not directly activated by cytosolic Ca2+. Direct coupling of membrane tension with channel opening is important, when stretch-activated Cl− or K+ channels, channels that are open by suction of patch pipettes, are found in the swollen cell. However, stretch-activated Cl− channels are not found in many cells. Swelling of the cell not only stretches the membrane surface but triggers unidentified, presumably cell-specific, signal transduction, resulting in the opening of VDCCs [53–55]. Candidates for signal transduction toward VDCCs include calmodulin [56], tyrosine kinase [57] and small molecular G proteins [58] (fig. 2), but study of the mechanisms await the molecular identification of the channel molecules.
Again, the molecular identification of VDCC has been difficult because of the absence of identification of the cell line of VDCCs (endogenous VDCCs in table 1). Various proteins are suspected to be molecular candidates for VDCCs. The three initial candidates, P-glycoprotein (P-gp) [59], pICln [60] and ClC-3, have since been discarded. On the basis of the well-known function of CFTR, hypothesized to be a bifunctional multiple-drug transporter, P-gp would adopt two mutually exclusive functional states, either as an adenosine triphosphate (ATP)-dependent drug transporter or a VDCC [61, 62]. However, this is unlikely because other laboratories could not repeat the original experiments on which this hypothesis was based.
The second protein that has been proposed for VDCCs is the ubiquitously expressed pICln [62]. The electrophysiologic and pharmacological properties of pICln-associated Cl− conductance in Xenopus oocytes differ from those of VDCC currents. pICln has a constitutive cytosolic and nuclear location. pICln interacts with splice factors, suggesting a role in gene expression and embryonic development [63, 64].
The ClC-3 protein [65], a member of the ClC family, was suggested recently as a molecular candidate for VDCCs. However, some characteristics of ClC-3-induced currents, such as single-channel kinetics, biophysical properties such as rectification, modulation by PKC and the large current amplitude under isotonic conditions, are at variance with those of VDCCs [66, 67]. In addition, ClC-3 is mainly located in endosomes, and no changes in VDCCs have been detected in cardiac muscles in mice lacking the ClC-3 gene [68]. Rather, disruption of the ClC-3 gene leads to morphologic abnormalities, suggesting that ClC-3 is important in maintaining pH and Cl− concentration within endosome vesicles, as discussed below. Recently, ClC-3 was again reported to be a fundamental molecular component of VDCCs in epithelial cells [69], in smooth muscle cells [70] and on the basis of antisense oligonucleotide experiments [69]. Some associated proteins related to recycling of the plasma membrane and endosomes have been isolated [71]. When the recycling is regulated by signal transduction induced by swelling, the complex of ClC-3 and an associated protein is a leading candidate in VDCCs [72].
Several large-conductance Cl− channels (maxi-Cl−) with single-channel conductance between 250 and 430 pS, behaving like VDCCs, have been described [73–77], including a Cl− channel with a conductance of 400 pS that is activated by osmotic cell swelling and that is sensitive to Gd3+. This channel is ATP conductive, with a PATP/PCl of 0.09, and is a candidate for the volume- and voltagedependent ATP-conductive pathway for swelling-induced ATP release (maxi-Cl− ATP) [77]. In contrast to the general role of anionic transport by Cl− channels, maxi-Cl− ATP can transmit a cationic excitatory signal to an adjacent cell. Swelling of the cell possessing maxi-Cl− ATP, by mechanical stress, releases ATP, which then activates the influx of Ca2+ into adjacent cells having a purinergic ATP receptor. This phenomenon is evident in the mechanism of renal tubuloglomerular feedback in macula densa cells [77, 78] (fig. 3). The higher the luminal Cl− concentration, the larger the volume of macula densa cells that have maxi-Cl− ATP in the basolateral membrane. Although macula densa cells are a continuous part of the water-impermeable loop of Henle, the luminal membrane of these cells has high water permeability, and the cells change volume easily. The basolateral maxi-Cl− ATP activated by an increase in volume releases ATP and Cl− toward adjacent smooth-muscle-like mesangial cells. Contraction of mesangial cells reduces renal blood flow, leading to a lower amount of luminal solute involving Cl− concentration. This feedback mechanism is suggested to be central in the renal response to salt overloading, and the identification of maxi-Cl− ATP molecules would be a clue to elucidating the mechanism of hypertension. A possibility of ATP conductivity of Cl− channels is suggested, but not evident, in brain and lung epithelial cells [2, 78].
We have reported that Drosophila tweety may encode a large-conductance CaC channel that has been detected on occasion in situ in endothelium, neurons or smoothmuscle cells [15]. These channels have five or six TMSs [15, 79]. There are three members of the family of tweety homologs (TTYH1-3) in mammals. When TTYH1-3 are expressed in Chinese hamster ovary (CHO) cells, a large current is evoked by addition of a Ca2+ ionophore or in 0.1 mM Ca2+ in a pipette fill, which is sensitive to 10–100 mM DIDS but resistant to NPPB or niflumic acid. A spliced variant of TTYH1 was activated in the expressed cells exposed to hypotonic solution. A single-channel recording of TTYH3 showed Cl conductance of 260 pS in a cytosolic Ca2+ concentration of >1 mM, while a 250-pS endogenous Cl− channel can also be found after prolonged depolarization in CHO cells. Mutants of TTYH3 reveal altered selectivity to anion, suggesting that TTYHs may encode large-conductance Cl− channels. The tweety locus is adjacent to the flightless locus and is regulated by the same promoter, but whether tweety is directly related to flying in the fly is not known [79]. The tweety gene is constantly expressed during development from the larva to the adult fly. There are two tweety families in Drosophila, and knockout of these genes has proved to be lethal. Thus, experimentation with tissueor time-specific knockout of these genes is essential to identification of their properties in vivo.
Another Cl− channel, the MID-1-related Cl− channel (MCLC), is a protein of 541 amino acids and four putative TMSs [80]. MID-1 encoded a yeast Saccharomyces cerevisiae membrane protein of two TMS that is expressed as a stretch-activated cation channel in mammalian CHO cells [81]. Using the MID-1 sequence as a probe, human MCLC has been cloned by BLAST search. MCLC is located in intracellular compartments, including the endoplasmic reticulum and the Golgi apparatus, and provides a 70-pS permeable Cl− channel in a planer lipid bilayer. MCLC is also considered to be a new class of VDCC expressed in intracellular compartments [80].
Fluid transport
The mechanism for contribution of Cl− channels to NaCl fluid transport across epithelial cells is well documented and is summarized elsewhere [2, 82, 83]. One of the most striking examples put forward is the case of the ClC-Kb channel. The renal thick ascending loop of Henle is an essential part of the osmotic concentration gradient in the interstitial space, since NaCl is transported from the lumen to the interstitium without movement of free water (fig. 3). Luminal Na+ entering the cell via Na/K/2Cl transport (NKCC) is reabsorbed by the basolateral Na+/K+ pump, whereas K+ is recycled back through apical K+ channels (ROMK and Kir1.1). The accumulated Cl− is transported via the ClC-Kb channel through the basolateral membrane by means of the electrochemical gradient. Finally, NaCl accumulates in the interstitium, and K+ remains in the lumen, where K+ creates positive lumen potential and drives other cations, such as Ca2+ and Mg2+, to the interstitial space through the paracellular pathway. Barttin is a binding protein of the ClC-Kb channel, with a two-membrane spanning domain that plays a role in the membrane expression of this channel. The channels ClC-Ka and ClC-Kb are constitutively open when co-expressed with their subunit Barttin [84]. Both channels are slightly outwardly rectifying, inhibited by extracellular acidosis and potentiated by an increase in extracellular calcium. Functional loss of NKCC, Kir1.1, ClC-Kb or Barttin is a cause of Bartter syndrome, in which nephrogenic diabetes insipides, hypokalemia, hypomagnesemia and hypocalcemia have been observed as variations in phenotype [82–84].
A similar schema can be illustrated for the cells of the Stria vascularis, which makes inner-ear fluid. The ClCKa and ClC-Kb channels are involved in the recycling of Cl− that accompanies the secretion of K+. A high concentration of K+ is necessary for hair cells to sense mechanical stimuli. Therefore, congenital deafness accompanied with diabetes insipidus is associated with genetic mutation in the ClC-Ka and ClC-Kb channels and in barttin protein [2, 4, 82, 83].
Another important Cl− channel is encoded by the cystic fibrosis transmembrane conductance regulator (CFTR) [85, 86]. Cystic fibrosis (CF), the most common lethal genetic disease among caucasians, is complicated by abnormal epithelial solute and fluid transport due to mutations in CFTR. CFTR belongs to the ATP-binding cassette (ABC) family of transporters, containing 12 predicted transmembrane helices and five cytoplasmic domains consisting of two nucleotide-binding domains (NBDs), a regulatory (R) domain containing numerous consensus phosphorylation sequences (fig. 1). The ABC superfamily regulates other processes in addition to its own role. For example, the sulfonylurea receptor, SUR, regulates K+ channels. Likewise, CFTR regulates Na channel (ENaC) and outward-rectified Cl− channel (ORCC) by excretion of ATP, although these are still controversial [87–89]. Work with truncation or mutation suggests that the first transmembrane domain (TMD-1) of CFTR, especially predicted a-helices 5 and 6, forms an essential part of the Cl− channel pore, whereas the first NBD domain is essential for its ability to regulate ORCCs [88]. All three functions, Cl− channels, ORCC and Na+ channel together contribute isotonic transport of fluid across the respiratory epithelia [89].
Neutralization of H+ ions in lysosomal vesicles
The lysosome is an intracellular vesicular organelle that plays a role in protein degradation and has a low interior pH. To maintain this low pH, the vesicle possesses H+-ATPase-accumulating H+ ions. Accumulation of H+ ions is back-leaked by the electrochemical gradient until cationic deflection due to charge accumulation is diminished. To neutralize the voltage deflection and maintain pH, an acid-activated Cl− channel is required in order to accumulate Cl− in the interior of the vesicle. ClC-5, located in renal proximal tubules, is a well-known example of a Cl− channel that plays a role in protein degradation [2, 90–94]. A functional defect in this gene causes progressive renal failure with nephrocalcinosis. A defect in this channel apparently does not show abnormal metabolism of Cl−, water or Na+ ions. Thus, loss of Cl− permeability per se does not induce progressive loss of nephron function. Megalin, which is closely related to metabolism of 1,25(OH)2D3, is degraded in the lysosomes of renal proximal tubules [94]. Similarly, several proteins, such as parathyroid hormone (PTH) and micromolecular proteins are not degraded in the proximal tubules. Lack of degradation of megalin and PTH in mice lacking ClC-5 is believed to lead to a high luminal concentration of these calcitropic proteins, resulting in microalbuminuria, hyperphosphaturia and hypercalciuria [93]. Hypercalciuria, however, has not been observed in ClC-5 knockout mice used in different studies [94].
A defect in the Cl− channel in lysosomes (ClC-3) may further cause morphologic disorders. ClC-3 knockout mice are viable but smaller. They survive >1 year, but show severe degeneration of the CA1 region of the hippocampus and the retina, resulting in a complete loss of photoreceptors. Electrophysiological analysis of hippocampal slices from juvenile ClC-3 knockout mice revealed no major functional abnormalities, except for a slight increase in the amplitude of miniature excitatory postsynaptic currents [68]. Another model lacking ClC-3 revealed significant kyphosis, in addition to brain anomalies, and no major change in swelling-activated Cl− permeability [95]. Therefore, ClC-3 is an endosomal Cl− channel that is related to the degradation of proteins and that plays a role in morphogenesis in expressed tissue.
A defect in ClC-7 in the endosomes of osteoclasts reveals a striking phenotype, osteopetrosis, in which bone remodeling, reabsorption and calcification are not processed inherently. Osteoclasts generate and secrete H+, by means of H+-ATPase, into the surface of bone in order to digest mineral crystals. Ruffled border, a closed space on the reabsorption surface where secreted H+ accumulates, has a pH of 4.0–6.0. Osteoclasts lacking ClC-7 are unable to maintain a low pH. The mechanism is the same as that for maintenance of low pH in lysosomes [96].
AQP-6 is a member of water channel but is activated by a low pH, to express Cl− channel in renal collecting segments. AQP-6 is located in endosomes, suggesting that the functional role of this channel is similar to the above ClCs, although mice lacking this gene have not yet been studied [11, 97].
Do Cl− channels play a role in tumorigenesis?
Cl− channels are thought to contribute to cell growth. Glioma Cl− channels are upregulated in glioma cells. Some Cl− channel blockers induce suppression of gene p21 [98]. Similarly, blocking of VDCC-related Cl− channels prevents apoptosis of various cells [99]. As described above, the hClCA2 family may be related to tumorigenesis. hClCA1 is also related to carcinoma cells [100]. These data may suggest a novel role for Cl− channels in biology, but evidence at the level of a single gene is needed.
Blockers of Cl− channels
A number of reagents block Cl− channels. Most of these reagents, however, are not specific for one class of Cl− channels. Disulfonic stilbenes, for example, are widely used blockers of any Cl− channel but only some anion transporters. Niflumic acid is employed to block endogenous CaC current in Xenopus oocytes. But niflumic acid is also used to inhibit cation current [101]. Table 2 summarizes widely used reagents for blocking most Cl− channels, where IC50 (mean inhibitory concentration) less than 100 mM is described. Classic anion channel blockers inhibit CaC and VDCC at micromolar concentrations. In general, ClC is resistant to blockade with classical blockers. ClC-1 alone can be inhibited by 9-AC, DPC and niflumic acid in the micromolar range. ClC-2 is inhibited by these reagents at millimolar concentrations but by Zn2+ ion at micromolar concentrations. CFTR is an ATP-binding cassette family, blocked by sulfonylureas such as glibenclamide at over 100 mM. DIDS inhibits CFTR but only from inside the cell. Thiazolidinone derivatives are potent inhibitors of CFTR at less than 10 mM. [102] In a search for more specific reagents, chlorotoxin was found and has been advertised as a selective Cl− channel blocker toxin [103]. However, it may be ineffective [104]. Direct CFTR activator have been characterized. These include the xanthines and the flavonoids, of which the isoflavonoid genistein is the most potent activator. Interestingly, phenylglycine (2-[(2-1H-indol-3-yl-acetyl)-methylamino]-N-(4-isopropylphenyl)-2-phenylacetamide) or sulfonamide [6-(ethylphenylsulfamoyl)-4-oxo-1,4-dihydroquinoline-3-carboxylic acid cycloheptylamide] prolongs the single channel open probability of DF508CFTR (most frequent mutant in CF patients). These reagents may be useful in treating of CF patients [105].
Unique reagents are being developed to discriminate between CaC channels and VDCCs. The antimalarial drug mefloquine and the antidepressant fluoxetine (Prozac), considered to be selective 5-hydroxytryptamine reuptake inhibitors, block CaC channels and have IC50 values of approximately 5 mM [106]. VDCCs are inhibited specifically by antioestrogens such as clomiphen, nafoxidine and tamoxifen, and by antimalarials such as mefloquine. A large-conductance, volume-dependent Cl− channel is blocked by Gd3+ [77]. Macloride antibiotics, erythromycin and clarithromycin may inhibit Cl current in some cells [107].
Structure and permeation of Cl− channels
There are several Cl− channel structures, suggested by their hydrophobic analysis (fig. 1). Mechanisms of Cl− channel permeation have been successfully studied by means of X-ray analysis of ClC channel crystals. This sophisticated method has revealed a complex structure of ClC channels and a binding motif of Cl− ions [19, 108]. The structure revealed a complex fold of 18 α-helices per subunit with at least two Cl− ions bound in the center of each protopore [109, 110]. These studies have shown that a lone Cl− ion bound to the center of the ClC pore is pushed out by a second ion that enters the pore and takes its place. The pushing of an ion by another is favorable for energy barriers that reduce the largest free-energy gap. This center portion is the narrowest part of the pore, formed by residues Tyr-445 and Ser-107 [19] and stabilizes Cl− ions from the water shell, which prevents the passage of uncharged solutes through this gate. A critical glutamic acid residue was identified whose side chain seems to occupy a third Cl− ion binding site in the closed state and that moves away to allow Cl− binding [111, 112]. ClC has double-barreled pores, the pathways of which are determined by mutation analysis [7–9]. Compared with the structure of the K+ channel with six TMSs, the structure of ClC is far more complex and not predicted by hydrophobic analysis. Interestingly, crystallography shows prokaryotic ClC to be a H+/Cl− transporter rather than an anion channel [20]. Furthermore, recent data support this finding, so that ClC0, 1, 2 and Ka/b function as ion channels, and ClC3-7 as Cl−/H+ exchangers [113].
The crystal structures of other Cl− channels are unavailable, including that for CFTR. Extensive studies with mutants suggested that the TMS6 and NBD domains are important in the permeation pathway [114]. However, the conducting pore of CFTR [115], glycine receptor [116] or even a cationic channel can be functionally and structurally reproduced in artificial membranes, by means of synthetic peptides. The peptide is designed to make a cylinder with an uneven distribution of hydrophobic and hydrophilic alignment (fig. 4). At least 18–20 amino acids are required in order to penetrate the lipid bilayers. The sequence requires positively charged amino acids in the entrance of the cylinder. Four or five oligomers combine to form a conducting pore. To enhance the formation of oligomers, tethering of the peptide by an artificial chain (lysine) and ring (β-turn structure) effectively reproduces the anion-conducting pore in the artificial membrane [20–22]. The computer based structure of TMS2 of glycine receptor (the sequence of amino acids from the second transmembrane of the glycine receptor) suggested that Cl− ions without a water shell are bound to the center of the pore [21, 22]. Using this technique, homo- and heteropolymerization of TMS2 and TMS6 of CFTR have been probed to form anion-conducting pores resembling the single-channel conductance of CFTR. However, TMS1, −3, −4 and −5 fail to form ion-conducting pores [115]. The artificial peptide Cl− channel may be used to ameliorate poor secretion of fluid in lung epithelia of patients with cystic fibrosis [115]. Interestingly, under the rule of amino acid sequences, many unexpected peptides are potential Cl− channels. In other words, many possible Cl− channel pores are hidden in various TMSs, as indicated by calculation of cylinder conformation with uneven hydrophobicity. Several molecules have been thought to be Cl− channels; they be subunits of ubiquitously expressed Cl− channels or may be due to pore formation by unexpected TMS [21, 22, 116, 117]. Cl− channels are classified by their physiologic roles. Studies of the family of CIC channels have revealed a majority of their functions. Additional evidence of novel Cl− pores by various molecules with different TMSs has been growing. Functional evidence that is based on the study of molecular expression of defective molecules in vitro or in vivo requires a molecule-based classification of the diversity of Cl− channels in the future.
References
Jentsch T. J., Steinmeyer K. and Schwarz G. (1990) Primary structure of Torpedo marmorata chloride channel isolated by expression cloning in Xenopus oocytes. Nature 348: 510–514
Jentsch T. J., Stein V., Weinreich F. and Zdebik A. A. (2002) Molecular structure and physiological function of chloride channels. Physiol. Rev. 82: 503–568
Nilius B. (2001) Chloride channels go cell cycling. J. Physiol. 532: 581
Nilius B. and Droogmans G. (2003) Amazing chloride channels: an overview. Acta Physiol. Scand. 177: 119–147
Nilius B. and Droogmans G. (2002) Calcium activated chloride channels in vascular endothelial cells. In: Ca2+-activated Cl channels, pp. 327–344, Fuller K. (ed.), Academic Press, New York
Nilius B., Eggermont J., Voets T. and Droogmans G. (1996) Volume-activated Cl channels. Gen. Pharmacol. 27: 1131–1140
Fahlke C., Rhodes T. H., Desai R. R. and George A. L. Jr. (1998) Pore stoichiometry of a voltage-gated chloride channel. Nature 394: 687 690
Lin C. W. and Chen T. Y. (2000) Cysteine modification of a putative pore residue in ClC-0: implication for the pore stoichiometry of ClC chloride channels. J. Gen. Physiol. 116: 535–546
Chen T. Y. (2005) Structure and function of ClC channels. Annu. Rev. Physiol. 67: 809–839
Schwiebert E. M., Egan M. E., Hwang T. H., Fulmer S. B., Allen S. S., Cutting G. R. et al. (1995) CFTR regulates outwardly rectifying chloride channels through an autocrine mechanism involving ATP. Cell 81: 1063–1073
Jentsch T. J. (2002) Chloride channels are different. Nature 415: 276–277
Gandhi R., Elble R. C., Gruber A. D., Schreur K. D., Ji H. L., Fuller C. M. et al. (1998) Molecular and functional characterization of a calcium-sensitive chloride channel from mouse lung. J. Biol. Chem. 273: 32096–32101
Gruber A. D., Elble R. C., Ji H. L., Schreur K. D., Fuller C. M. and Pauli B. U. (1998) Genomic cloning, molecular characterization and functional analysis of human CLCA1, the first human member of the family of Ca2+-activated Cl channel proteins. Genomics 54: 200–214
Sun H., Tsunenari T., Yau K. W. and Nathans J. (2000) The vitelliform macular dystrophy protein defines a new family of chloride channels. Proc. Natl. Acad. Sci. USA 99: 4008–4013
Suzuki M. and Mizuno A. (2004) A novel Cl channel related to Drosophila flightless locus. J. Biol. Chem 279: 22461–22468
Attali B., Guillemare E., Lesage F., Honore E., Romey G, Lazdunski M. et al. (1993) The protein IsK is a dual activator of K+ and Cl− channels. Nature 365: 850–852
Wadiche J. I. and Kavanaugh M. P. (1998) Macroscopic and microscopic properties of a cloned glutamate transporter/chloride channel. J. Neurosci. 18: 7650–7661
Yasui M., Hazama A., Kwon T. H., Nielsen S., Guggino W. B. and Agre P. (1999) Rapid gating and anion permeability of an intracellular aquaporin. Nature 402: 184–187
Dutzler R., Campbell E. B., Cadene M., Chait B. T. and MacKinnon R. (2002) X-ray structure of a ClC chloride channel at 3.0 Å reveals the molecular basis of anion selectivity. Nature 415: 287–294
Accardi A. and Miller C. (2004) Secondary active transport mediated by a prokaryotic homologue of ClC Cl− channels. Nature 427, 803–807
Grove A., Iwamoto T., Montal M. S., Tomich J. M. and Montal M. (1992) Synthetic peptides and proteins as models for poreforming structure of channel proteins. Methods Enzymol. 207: 510–525
Montal M. O., Iwamoto T., Tomich J. M. and Montal M. (1993) Design, synthesis and functional characterization of a pentameric channel protein that mimics the presumed pore structure of the nicotinic cholinergic receptor. FEBS Lett. 320: 261–266
Montal O. M., Buhler L. K., Iwamoto T., Tomich J. M. and Montal M. (1993) Synthetic peptides and four-helix bundle proteins as model systems for the pore-forming structure of channel proteins. I. Transmembrane segment M2 of the nicotinic cholinergic receptor channel is a key pore-lining structure. J. Biol. Chem. 268: 14601–14607
Conley E. (1996) Cl GABAA. In: Ion Channel Factsbook vol. 1, pp. 293–365, Conley E. and Brammer W. (eds.), Academic Press, London
Farrant M. and Cull-Candy S. (1993) GABA receptors, granule cells and genes. Nature 361: 302–303
Conley E. (1996) Cl glycine. In: Ion Channel Factsbook, vol. 1, pp. 367–399, Academic Press, London
Pusch M., Ludewig U., Rehfeldt A. and Jentsch T. J. (1995) Gating of the voltage-dependent chloride channel ClC-0 by the permeant anion. Nature 373: 527–531
Jentsch T. J. and Gunther W.(1997) Chloride channels: an emerging molecular picture. Bioessays 119:117-126
Rychkov G. Y., Pusch M., Astill D. S., Roberts M. L., Jentsch T. J. and Bretag A. H. (1996) Concentration and pH dependence of skeletal muscle chloride channel ClC-1. J. Physiol. 497: 423–435
Kubisch C., Schmidt-Rose T., Fontaine B., Bretag A. H. and Jentsch T. J. (1998) ClC-1 chloride channel mutations in myotonia congenita: variable penetrance of mutations shifting the voltage dependence. Hum. Mol. Genet. 7: 1753–1760
Thiemann A., Grunder S., Pusch M. and Jentsch T. J. (1992) A chloride channel widely expressed in epithelial and non-epithelial cells. Nature. 356: 57–60
Grunder S., Thiemann A., Pusch M. and Jentsch T. J. (1992) Regions involved in the opening of CIC-2 chloride channel by voltage and cell volume. Nature 360: 759–762
Stanley K. The role of an inwardly rectifying chloride conductance in postsynaptic inhibition. (1994) J. Neurophysiol. 72: 273–284
Dinudom A., Young J. A. and Cook D. I. (1993) Na and Cl conductances are controlled by cytosolic Cl− concentration in the intralobular duct cells of mouse mandibular glands. J. Membr. Biol. 135: 289–295
Smith R. L., Clayton G. H., Wilcox C. L., Escudero K. W. and Staley K. J. (1995) Differential expression of an inwardly rectifying chloride conductance in rat brain neurons: a potential mechanism for cell-specific modulation of postsynaptic inhibition. J. Neurosci. 15: 4057–4067
Saley K., Smith R., Schaack J., Wilcox C. and Jentsch T. J. (1996) Alteration of GABAA receptor function following gene transfer of the CLC-2 chloride channel. Neuron 17: 543–551
Bösl M. R., Stein V., Hübner C., Zdebik A. A., Jordt S. E., Mukhophadhyay A. K. et al. (2001) Male germ cells and photoreceptors, both depending on close cell-cell interactions, degenerate upon ClC-2 Cl-channel disruption. EMBO J. 20: 1289–1299
Conley E. C. and Brammar W. J. (1996) CaCC. In: The Ion Channel Facts Book, vol. 2, pp. 524–571, Academic Press, London
Baron A., Pacaud P., Loirand G., Mironneau C. and Mironneau J. (1991) Pharmacological block of Ca2+-activated Cl− current in rat vascular smooth muscle cells in short-term primary culture. Pflugers Arch. 419: 553–558
Saleh S. N. and Greenwood I. A. (2005) Activation of chloride currents in murine portal vein smooth muscle cells by membrane depolarization involves intracellular calcium release. Am. J. Physiol. Cell. Physiol. 288: C122–131
Brenner R., Perez G. J., Bonev A. D., Eckman D. M., Kosek J. C., Wiler S. W. et al. (2000) Vasoregulation by the b subunit of the calcium-activated potassium channel. Nature 407: 870–876
Evans S. R., Thoreson W. B. and Beck C. L. (2004) Molecular and functional analyses of two new calcium-activated chloride channel family members from mouse eye and intestine. J. Biol. Chem. 279: 41792–41800
Ritzka M., Stanke F., Jansen S., Gruber A. D,. Pusch L., Woelfl S. et al. (2004) The CLCA gene locus as a modulator of the gastrointestinal basic defect in cystic fibrosis. Hum. Genet. 115: 483–491
Britton F. C., Ohya S., Horowitz B. and Greenwood I. A. (2002) Comparison of the properties of CLCA1 generated currents and I[Cl(Ca)] in murine portal vein smooth muscle cells. J Physiol. 539: 107–117
Gruber, A. D. and Pauli B. U. (1999) Tumorigenicity of human breast cancer is associated with loss of the Ca2+ activated chloride channel CLCA2. Cancer Res. 59: 5488–5491
Pauli B. U., Abdel-Ghany M., Cheng H. C., Gruber A. D., Archibald H. A. and Elble, R. C. (2000) Molecular characteristics and functional diversity of CLCA family members. Clin. Exp. Pharmacol. Physiol. 27: 901–905
Ghany A., Cheng H. C., Elble R. C. and Pauli B. U. (2001) The breast cancer beta(4) integrin and endothelial human CLCA2 mediate lung metastasis. J. Biol. Chem. 276: 25438–25446
Petrukhin K., Koisti M. J., Bakall B., Li W., Xie G., Marknell T. et al. (1998) Identification of the gene responsible for Best macular dystrophy. Nat. Genet. 19: 241–224
Marquardt A., Stohr H., Passmore L. A., Kramer F., Rivera A. and Weber B. H. (1998) Mutations in a novel gene, VMD2, encoding a protein of unknown properties cause juvenile-onset vitelliform macular dystrophy (Best’s disease). Hum. Mol. Genet. 7: 1517–1525
Gallemore R. P., Hughes B. A. and Miller S. S. (1998) Lightinduced responses of the retinal pigment epithelium. In: The Retinal Pigment Epithelium, Marmor M. F. and Wolfensberger T. J. (eds.), pp. 175–198, Oxford University Press, New York
Marmorstein L. Y., McLaughlin P. J., Brett Stanton J. B., Yan L., Crabb J. W. and Marmorstein A. D. (2002) Bestrophin interacts physically and functionally with protein phosphatase 2A. J. Biol. Chem. 277: 30591–30597
Fischmeister R. and Hartzell H. C. (2005) Volume-sensitivity of the bestrophin family of chloride channels. J. Physiol. 562: 477–491
Nilius B., Voets T., Eggermont J. and Droogmans G. (1999) VRAC: a multifunctional volume regulated anion channel in vascular endothelium. In: Chloride Channels, Kozlowski R. (ed.), pp. 47–63. Isis Medical Media Limited, Oxford
Okada Y. (1997) Volume expansion-sensing outward-rectifier Cl channel: fresh start to the molecular identity and volume sensor. Am. J. Physiol. 273: C755–C789
Clapham D. E. (1998) The list of potential volume-sensitive chloride currents continues to swell (and shrink). J. Physiol. Lond. 111: 623–624
Huang P., Liu J., Di A. K., Robinson N. C., Musch M. W., Kaetzel M. A. et al. (2001) Regulation of human CLC-3 channels by multifunctional Ca2+/calmodulin-dependent protein kinase. J. Biol. Chem. 276: 20093–20100
Lepple-Wienhues A., Szabo I., Laun T., Kaba N. K., Gulbins E. and Lang F. (1998) The tyrosine kinase p56(lck) mediates activation of swelling-induced chloride channels in lymphocytes. J. Cell. Biol. 141: 281–286
Nilius B., Voets T., Prenen J. Barth H., Aktories K., Kaibuchi K. et al.(1999) Role of RhoA and Rho kinase in the activation of volume-regulated anion channels in bovine endothelial cells. J. Physiol. 516: 67–74
Valverde M. A., Diaz M., Sepulveda F. V., Gill D. R., Hyde S. C. and Higgins C. F. (1992) Volume-regulated chloride channels associated with the human multidrug-resistance P-glycoprotein. Nature 355: 830–833
Paulmichl M., Li Y., Wickman K., Ackerman M., Peralta E. and Clapham D. (1992) New mammalian chloride channel identified by expression cloning. Nature 356: 238–241
Valverde M. A., Diaz M., Sepulveda F. V., Gill D. R., Hyde S. C. and Higgins C. F. (1992) Volume-regulated chloride channels associated with the human multidrug-resistance P-glycoprotein. Nature 355: 830–833
Valverde M. A., Bond T. D., Hardy S. P. Taylor J. C., Higgins C. F., Altamirano J. et al. (1996) The multidrug resistance P-glycoprotein modulates cell regulatory volume decrease. EMBO J. 15: 4460–4468
Krapivinsky G., Pu W., Wickman K., Krapivinsky L. and Clapham D. E. (1998) pICln binds to a mammalian homolog of a yeast protein involved in regulation of cell morphology. J. Biol. Chem. 273: 10811–10814
Pu W. T., Wickman K. and Clapham D. E. (2000) ICln is essential for cellular and early embryonic viability. J. Biol. Chem. 275: 12363–12366
Kawasaki M., Fukuma T., Yamauchi K., Sakamoto H., Marumo F. and Sasaki S. (1999) Identification of an acid-activated Cl channel from human skeletal muscles. Am. J. Physiol. Cell. Physiol. 277: C948–C954
Li X., Shimada K., Showalter L. A. and Weinman S. A. (2000) Biophysical properties of ClC-3 differentiate it from swelling-activated chloride channels in Chinese hamster ovary-K1 cells. J. Biol. Chem. 275: 35994–35998
Weylandt K. H., Valverde M. A., Nobles M., Raguz S., Amey J. S., Diaz M. et al. (2001) Human ClC-3 is not the swelling-activated chloride channel involved in cell volume regulation. J. Biol. Chem. 276: 17461–17467
Stobrawa S. M., Breiderhoff T., Takamori S., Engel D., Schweizer M., Zdebik A. A. et al. (2001) Disruption of CIC-3, a chloride channel expressed on synaptic vesicles, leads to a loss of the hippocampus. Neuron 29: 185–196
Vessey J. P., Shi C., Jollimore C. A., Stevens K. T., Coca-Prados M., Barnes S. et al. (2004) Hyposmotic activation of ICl, swell in rabbit nonpigmented ciliary epithelial cells involves increased ClC-3 trafficking to the plasma membrane. Biochem. Cell Biol. 82: 708–718
Zhou J. G., Ren J. L., Qiu Q. Y., He H. and Guan Y. Y. (2005) Regulation of intracellular Cl− concentration through volumeregulated ClC-3 chloride channels in A10 vascular smooth muscle cells. J. Biol. Chem. 280: 7301–7308
Salazar G., Love R., Styers M. L., Werner E., Peden A., Rodriguez S. et al. (2004) AP-3-dependent mechanisms control the targeting of a chloride channel (ClC-3) in neuronal and nonneuronal cells. J. Biol. Chem. 279: 25430–25439
Hermoso M., Satterwhite C. M. andrade Y., Hidalgo J., Wilson S. M., Horowitz B. et al. (2002) ClC-3 is a fundamental molecular component of volume-sensitive outwardly rectifying Cl channel and volume regulation in HeLa cells and Xenopus laevis oocytes. J. Biol. Chem. 205: 132–200
Mitchell C. H., Wang L. and Jacob T. J. C. (1997) A large-conductance chloride channel in pigmented ciliary epithelial cells activated by GTP gamma S. J. Membr. Biol. 158: 167–175
Diaz M., Bahamonde M. I., Lock H., Munoz F. J., Hardy S. P., Posas F. et al. (2001) Okadaic acid-sensitive activation of Maxi Cl channels by triphenylethylene antioestrogens in C1300 mouse neuroblastoma cells. J. Physiol. 536: 79–88
Kokubun S., Saigusa A. and Tamura T. (1991) Blockade of Cl channels by organic and inorganic blockers in vascular smooth muscle cells. Pflugers Arch. 418: 204–213
Sabirov R. Z., Dutta A. K. and Okada Y. (2001) Volume-dependent ATP-conductive large-conductance anion channel as a pathway for swelling-induced ATP release. J. Gen. Physiol. 118: 251–266
Bell P. D., Lapointe J. Y., Sabirov R., Hayashi S., Peti-Peterdi J. Manabe K. et al. (2003) Macula densa cell signaling involves ATP release through a maxi anion channel. Proc. Natl. Acad. Sci. USA 100: 4322–4327
Bell P. D., Lapointe J. Y. and Peti-Peterdi J. (2003) Macula densa cell signaling. Annu. Rev. Physiol. 65: 481–500
Campbell H. D., Kamei M., Claudianos C., Woollatt E., Sutherland G. R., Suzuki Y. et al. (2000) Human and mouse homologues of the Drosophila melanogaster tweety (tty) gene: a novel gene family encoding predicted transmembrane proteins. Genomics 68: 89–92
Nagasawa M., Kanzaki M., Iino Y., Morishita Y. and Kojima I. (2001) Identification of a novel chloride channel expressed in the endoplasmic reticulum, Golgi apparatus, and nucleus. J. Biol. Chem. 276: 20413–20418
Kanzaki M., Nagasawa M., Kojima I., Sato C., Naruse K., Sokabe M. et al. (1999) Molecular identification of a eukaryotic, stretch-activated nonselective cation channel. Science 285: 882–886
Uchida S. and Sasaki S. (2005) Function of chloride channels in the kidney. Annu. Rev. Physiol. 67: 759–778
Jentsch T. J. (2005) Chloride transport in the kidney: lessons from human disease and knockout mice. J. Am. Soc. Nephrol. 16: 1549–1561
Estevez R., Boettger T., Stein V., Birkenhager R., Otto E., Hildebrandt F. et al. (2001) Barttin is a Cl− channel beta-subunit crucial for renal Cl− reabsorption and inner ear K+ secretion. Nature 414: 558–561
Rommens J. M., Iannuzzi M. C., Kerem B., Drumm M. L., Melmer G., Dean M. et al. (1989) Identification of the cystic fibrosis gene:chromosome walking and jumping. Science 245: 1059–1065
Riordan J. R., Rommens J. M., Kerem B., Alon N., Rozmahel R., Grzelczak Z. et al. (1989) Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science 245: 1066–1072
Berger H. A., Anderson M. P., Gregoryt R. J., Thompson S., Howard P. W., Maurer R. A. et al. (1991) Identification and regulation of the cystic fibrosis transmembrane conductance regulator-generated chloride channel. J. Clin. Invest. 88: 1422–1431
Hallows K. R., Raghuram V., Kemp B. E., Schwiebert E. M., Morales M. M., Lopes A. G. et al. (1998) Chloride channel and chloride conductance regulator domains of CFTR, the cystic fibrosis transmembrane conductance regulator. Proc. Natl. Acad. Sci. USA 95: 2674–2679
Schwiebert E. M., Benos D. J., Egan M. E., Stutts M. J. and Guggino W. B. (1999) CFTR is a conductance regulator as well as a chloride channel. Physiol. Rev. 79: S145–S166
Piwon N., Gunther W., Schwake M., Bosl M. R. and Jentsch T. J. (2000) ClC-5 Cl-channel disruption impairs endocytosis in a mouse model for Dent’s disease. Nature 408: 369–373
Lloyd S. E., Pearce S. H., Fisher S. E., Steinmeyer K., Schwappach B., Scheinman S. J. et al. (1996) A common molecular basis for three inherited kidney stone diseases. Nature 379: 445–449
GuÈnther W., LuÈchow A., Cluzeaud F., Vandewalle A. and Jentsch T. J. (1998) ClC-5, the chloride channel mutated in Dent’s disease, colocalizes with the proton pump in endocytotically active kidney cells. Proc. Natl Acad. Sci. USA 95: 8075–8080
Nykjaer A., Dragun D., Walther D., Vorum H., Jacobsen C., Herz J. et al. (1999) An endocytic pathway essential for renal uptake and activation of the steroid 25-(OH) vitamin D3. Cell 96: 507–515
Christensen E. I., Devuyst O., Dom G., Nielsen R., Van der Smissen P., Verroust P. et al. (2003) Loss of chloride channel ClC-5 impairs endocytosis by defective trafficking of megalin and cubilin in kidney proximal tubules. Proc. Natl. Acad. Sci. USA. 100: 8472–8477
Yoshikawa M., Uchida S., Ezaki J., Rai T., Hayama A., Kobayashi K. et al. (2002) CLC-3 deficiency leads to phenotypes similar to human neuronal ceroid lipofuscinosis. Genes Cells 7: 597–605
Kasper D., Planells-Cases R., Fuhrmann J. C., Scheel O., Zeitz O., Ruether K. et al. (2005) Loss of the chloride channel ClC-7 leads to lysosomal storage disease and neurodegeneration. EMBO J. 24: 1079–1091
Yasui M., Kwon T. H., Knepper M. A., Nielsen S. and Agre P. (1999) Aquaporin-6: an intracellular vesicle water channel protein in renal epithelia. Proc. Natl. Acad. Sci. USA. 96: 5808–5813
Jiang B., Hattori N., Liu B., Nakayama Y., Kitagawa K. and Inagaki C. (2004) Suppression of cell proliferation with induction of p21 by Cl(-) channel blockers in human leukemic cells. Eur J Pharmacol. 488: 27–34
Shimizu T., Numata T. and Okada Y. (2004) A role of reactive oxygen species in apoptotic activation of volume-sensitive Cl(-) channel. Proc. Natl. Acad. Sci. USA. 101: 6770–6773
Loewen M. E., Bekar L. K., Walz W., Forsyth G. W. and Gabriel S. E. (2004) pCLCA1 lacks inherent chloride channel activity in an epithelial colon carcinoma cell line. Am. J. Physiol. Gastrointest. Liver Physiol. 287: G33–G41
Gogelein H., Dahlem D., Englert H. C. and Lang H. J. (1990) Flufenamic acid, mefenamic acid and niflumic acid inhibit single nonselective cation channels in the rat exocrine pancreas. FEBS Lett. 268: 79–82
Ma T., Thiagarajah J. R., Yang H., Sonawane N. D., Folli C., Galietta L. J. V. et al. (2002) Thiazolidinone CFTR inhibitor identified by high-throughput screening blocks cholera toxininduced intestinal fluid secretion J. Clin. Invest. 110: 1651–1658
Fuller M. D., Zhang Z. R., Cui G., Kubanek J. and McCarty N. A. (2004) Inhibition of CFTR channels by a peptide toxin of scorpion venom. Am. J. Physiol. Cell. Physiol. 287: C1328–C1341
Maertens C., Wei L., Tytgat J., Droogmans G. and Nilius B. (2000) Chlorotoxin does not inhibit volume-regulated, calcium-activated and cyclic AMP-activated chloride channels. Br. J. Pharmacol. 129: 791–801
Pedemonte N., Sonawane N. D., Taddei A., Hu J., Zegarra-Moran O,. Suen Y. F. et al. (2005) Phenylglycine and sulfonamide correctors of defective DF508 and G551D cystic fibrosis transmembrane conductance regulator chloride-channel gating. Mol. Pharmacol. 67: 1797–1807
Maertens C., Wei L., Droogmans G. and Nilius B. (2000) Inhibition of volume-regulated and calcium-activated chloride channels by the antimalarial mefloquine. J. Pharmacol. Exp. Ther. 295: 29–36
Nakaya Y., Kuzu N., Nakamura Y. and Sone S. (1998) Suppressive action of erythromycin on cultured tracheal epithelial cell chloride channel activated by interferon-gamma. Jpn. J. Antibiot. 51: 152–154
Corry B., O’Mara M. and Chung S. H. (2004) Conduction mechanisms of chloride ions in ClC-type channels. Biophys. J. 86: 846–60
Estevez R., Pusch M., Ferrer-Costa C., Orozco M. and Jentsch T. J. (2004) Functional and structural conservation of CBS domains from CLC chloride channels. J. Physiol. 557: 363–378
Chen T. Y. (2005) Structure and function of ClC channels. Annu. Rev. Physiol. 67: 809–839
Babini E. and Pusch M. (2004) A two-holed story: structural secrets about ClC proteins become unraveled? Physiology. 19: 293 299
Estevez R., Pusch M., Ferrer-Costa C., Orozco M. and Jentsch T. J. (2004) Functional and structural conservation of CBS domains from CLC chloride channels. J. Physiol. 557: 363 378
Scheel O., Zdebik A. A., Lourdel S. and Jentsch T. J. (2005) Voltage-dependent electrogenic chloride/proton exchange by endosomal CLC proteins. Nature 436: 424–427
Riordan J. R. (2005) Assembly of functional CFTR chloride channels. Annu. Rev. Physiol. 67: 701–718
Montal O. M., Reddy G. L., Iwamoto T., Tomich J. M. and Montal M. (1994) Identification of an ion channel-forming motif in the primary structure of CFTR, the cystic fibrosis chloride channel. Proc. Natl. Acad. Sci. USA. 91: 1495–1499
Broughman J. R., Shank L. P., Takeguchi W., Schultz B. D., Iwamoto T., Mitchell K. E. et al. (2002) Distinct structural elements that direct solution aggregation and membrane assembly in the channel-forming peptide M2GlyR. Biochemistry 41: 7350–7358
Gao L., Broughman J. R., Iwamoto T., Tomich J. M., Venglarik C. J. and Forman H. J. (2001) Synthetic chloride channel restores glutathione secretion in cystic fibrosis airway epithelia. Am. J. Physiol. Lung Cell. Mol. Physiol. 281: L24–L30
Bormann J., Hamill O. P. and Sakmann B. (1987) Mechanism of anion permeation through channels gated by glycine and gamma-aminobutyric acid in mouse cultured spinal neurones. J. Physiol. 385: 243–286
Berger H. A., Anderson M. P., Gregory R. J., Thompson S., Howard P. W. Maurer R. A. et al. (1991) Identification and regulation of the cystic fibrosis transmembrane conductance regulator-generated chloride channel. J. Clin. Invest. 88: 1422–1431
Edwards J. C., Tulk B. and Schlesinger P. H. (1998) Functional expression of p64, an intracellular chloride channel protein. J. Membr. Biol. 163: 119–127
Li X., Shimada, K., Lori A., Showalter L. A. and Weinman S. A. (2000) Biophysical properties of ClC-3 differentiate it from swelling activated chloride channels in Chinese hamster ovary-K1 cells. J. Biol. Chem. 275: 35994–35998
Zhu G., Zhang Y., Xu H. and Jiang C.. (1998) Identification of endogenous outward currents in the human embryonic kidney (HEK 293) cell line. J. Neurosci. Methods 81: 73–83
Baron A., Pacaud P., Loirand G., Mironneau C. and Mironneau J. (1991) Pharmacological block of Ca(2+)-activated Cl− current in rat vascular smooth muscle cells in short-term primary culture. Pflugers Arch. 419: 553–558
McDonough S., Davidson N., Lester H. A. and Mccarty N. A. (1994) Novel pore-lining residues in CFTR that govern permeation and openchannel block. Neuron 13: 623–634
Schmieder S., Lindenthal S., Banderali U. and Ehrenfeld J. (1998) Characterization of the putative chloride channel xClC-5 expressed in Xenopus laevis oocytes and comparison with endogenous chloride currents. J. Physiol. 511: 379–393
Janssen L. J. and Sims S. M. (1994) Spontaneous transient inward currents and rhythmicity in canine and guinea-pig tracheal smooth muscle cells. Pflugers Arch. 427: 473–480
Aromataris E. C., Astill D. S., Rychkov G. Y., Bryant S. H., Bretag A. H. and Roberts M. L. (1999) Modulation of the gating of CIC-1 by S-(−)-2-(4-chlorophenoxy) propionic acid. Br. J. Pharmacol. 126: 1375–1382
Picollo A., Liantonio A., Didonna M. P., Elia L., Camerino D. C. and Pusch M. (2004) Molecular determinants of differential pore blocking of kidney CLC-K chloride channels. EMBO J. 5: 584 589
Julien M., Verrier B., Cerutti M., Chappe V., Gole M., Devauchelle G. et al. (1999) Cystic fibrosis transmembrane conductance regulator (CFTR) confers glibenclamide sensitivity to outwardly rectifying chloride channel (ORCC) in Hi-5 insect cells. J. Membr. Biol. 168: 229–239
Bachmann A., Russ U. and Quast U. (1999) Potent inhibition of the CFTR chloride channel by suramin. Naunyn-Schmiedebergs Arch. Pharmacol. 360: 473–476
Valverde M. A., Mintenig G. M. and Sepuldeva F. V. (1993) Differential effects of tamoxifen and I− on three distinguishable chloride currents activated in T84 intestinal cells. Pflugers Arch. 425: 552–554
Maertens C., Droogmans G., Chakraborty P. and Nilius B. (2001) Inhibition of volume-regulated anion channels in cultured endothelial cells by the anti-oestrogens clomiphene and nafoxidine. Br. J. Pharmacol. 132: 135–142
Hogg R. C., Wang Q. and Large W. A. (1993) Time course of spontaneous calcium-activated chloride currents in smooth muscle cells from the rabbit portal vein. J. Physiol. 464: 15–31
Author information
Authors and Affiliations
Corresponding author
Additional information
Received 28 July 2005; received after revision 25 August 2005; accepted 21 September 2005
Rights and permissions
Open Access This is an open access article distributed under the terms of the Creative Commons Attribution Noncommercial License ( https://creativecommons.org/licenses/by-nc/2.0 ), which permits any noncommercial use, distribution, and reproduction in any medium, provided the original author(s) and source are credited.
About this article
Cite this article
Suzuki, M., Morita, T. & Iwamoto, T. Diversity of Cl− Channels. Cell. Mol. Life Sci. 63, 12–24 (2006). https://doi.org/10.1007/s00018-005-5336-4
Published:
Issue Date:
DOI: https://doi.org/10.1007/s00018-005-5336-4