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The Cardiac Na+‐Ca2+ Exchanger: From Structure to Function

Michela Ottolia, Scott John, Adina Hazan, Joshua I. Goldhaber

10.1002/cphy.c200031

Source: Volume 12, Issue 1, January 2022

Published online: December 2021

Full Article on Wiley Online Library



Abstract

Ca2+ homeostasis is essential for cell function and survival. As such, the cytosolic Ca2+ concentration is tightly controlled by a wide number of specialized Ca2+ handling proteins. One among them is the Na+‐Ca2+ exchanger (NCX), a ubiquitous plasma membrane transporter that exploits the electrochemical gradient of Na+ to drive Ca2+ out of the cell, against its concentration gradient. In this critical role, this secondary transporter guides vital physiological processes such as Ca2+ homeostasis, muscle contraction, bone formation, and memory to name a few.

Herein, we review the progress made in recent years about the structure of the mammalian NCX and how it relates to function. Particular emphasis will be given to the mammalian cardiac isoform, NCX1.1, due to the extensive studies conducted on this protein. Given the degree of conservation among the eukaryotic exchangers, the information highlighted herein will provide a foundation for our understanding of this transporter family. We will discuss gene structure, alternative splicing, topology, regulatory mechanisms, and NCX's functional role on cardiac physiology. Throughout this article, we will attempt to highlight important milestones in the field and controversial topics where future studies are required. © 2021 American Physiological Society. Compr Physiol 12:1‐37, 2021.

Figure 1. Figure 1. Modes of operation of the Na+‐Ca2+ exchanger. (A) In the forward mode (top) NCX transports one Ca2+ ion out of the cell in exchange for three Na+ ions entering. One net charge is moved across the membrane resulting in a measurable ionic current. This is the most physiologically relevant cycle. In the reverse mode (bottom), one Ca2+ ions enter the cell and three Na+ ions exit. The direction in which Ca2+ is moved is governed by both the ionic gradients of Na+ and Ca2+ and membrane potential 164. (B) The scheme shows NCX transport cycle. This electrogenic transporter transitions between the outward and inward‐facing states via intermediate occluding states, leading to the coupled but opposite movements of Na+ and Ca2+ ions across the plasma membrane 164.
Figure 2. Figure 2. Na+‐Ca2+ exchanger gene organization. Each panel depicts the chromosomal location, number of exons, size, mRNA example, and corresponding protein of the three human isoforms. Gene exon structures are derived from Ensembl. Exons are indicated as grey bars and include both coding and noncoding exons. Exons undergoing alternative splicing are shown in color and are named in accordance with Quednau and colleagues 220. 5′ and 3′ regions of the gene are not included so that the gene size given extends from the beginning of the first exon and ends at the 3′ end of the last exon. Of special note, NCX2 does not undergo alternative splicing and all exons encode for protein. NCX1 and NCX3 have one noncoding exon, both undergo alternative splicing. An example of mRNA profile of the indicated exchangers is shown under each corresponding gene. Graphs are to scale. The alternative spliced exons (in colors), which at the protein level result in changes in the calcium‐binding domain 2 (CBD2) 220, are depicted in blue.
Figure 3. Figure 3. Tissue distribution of the exchanger isoforms. The three exchanger isoforms and their corresponding splice variants are differentially distributed within tissues 133,136,143,220,221. Examples are given for heart, brain, kidney, and skeletal muscle. The tissue distribution has been assessed using reverse transcriptase‐polymerase chain reaction as described in Ref. 220. Numerous evidence indicates that the isoform NCX1.1 136,143,220 is the only exchanger expressed in cardiomyocytes, although a study has identified isoforms NCX1.3 and NCX1.4 133. The splice variants NCX1.6 (ACD) and NCX3.3 (BC) are absent from the table. These exchangers have been localized in the brain 136,220, but their cell specific expression is undetermined.
Figure 4. Figure 4. Progression of Na+‐Ca2+ exchanger topology. The cartoons show how the topology of the mammalian exchanger has evolved through the years since its cloning in 1990 185. (A) Initially, NCX1.1 was predicted to be organized into 12 transmembrane segments (TMS). The two highly conserved regions among the exchangers which are known as α‐repeats 245 are highlighted. These regions are essential for ion transport 183,184,195. (B) Subsequent studies 50,60,106,239 revealed that the first helix was a signal peptide not necessary for activity, restricting the mature protein to 11 TMSs with an extracellular N terminus. (C) Cysteine scanning mutagenesis and epitope tagging were then applied to investigate NCX1.1 architecture. Based on these studies the topology of NCX was reorganized into nine transmembrane segments with an intracellular C‐terminus and two reentrant loops between TMSs 2‐3 and 7‐8 35,112,114,186,187,212. (D) The crystal structure of the archaebacterial exchanger (NCX_Mj) 150 has revealed a protein of 10 TMSs, contrasting with the predicted 9 TMSs for the mammalian exchanger as shown in (C). (E) Topology of the mammalian NCX based on NCX_Mj organization. The new architecture does not include the previously modeled reentrant loops, which are now predicted to be a short extracellular loop between TMSs 2‐3, within the α1‐repeat, and a new transmembrane segment (TMS 8) in the α2‐repeat. Future studies should be conducted to confirm the absence of either or both of these two reentrant structures and to validate that the mammalian exchanger and the archaebacterial are had a similar organization. The position of the cysteine residues involved in the intramolecular disulfide bond 241 is also shown. Amino acids are numbered based on the protein lacking the signal sequence.
Figure 5. Figure 5. Inconsistencies between the topology of the Na+‐Ca2+ exchanger and the archaebacterial exchanger. (A) Cysteine scanning mutagenesis and epitope mapping studies have been instrumental in deciphering the topology of the cardiac exchanger 35,50,112,114,186,187,212. The figure shows the amino acids that were replaced with cysteine and found accessible to membrane‐impermeable sulfhydryl agents either from the cytoplasm (blue circles) or the external side of the membrane (orange circles). Based on these investigations, NCX was modeled to have an extracellular N‐terminus, and a cytoplasmic C‐terminus. Moreover, residues 120 and 125 within the α1‐repeat were found to be exposed to the intracellular environment as well as several residues within the α2‐repeat, constraining these regions into reentrant loops. This organization corresponds to the one shown in Figure 4C. (B) With the atomic structure of the archaebacterial exchanger at hand 150, NCX1.1 is now assumed to be organized in 10 transmembrane segments 228,265, as shown in Figure 4E. However, this new arrangement places several residues previously found accessible to the cytoplasmic application of sulfhydryl reagents (blue circles), now facing the extracellular environment, including the C‐terminus. Note that the new architecture does not include reentrant loops. More investigations are needed to fully elucidate the topology of the mammalian exchanger.
Figure 6. Figure 6. Sequence alignment of the α‐repeats. (A) The sequence alignment illustrates the conservation of the α repeat regions within the Na+‐Ca2+ exchanger family. This is consistent with a broad range of evidence underlying their essential role in ion transport 125,183,184,195. The α1‐repeat encompasses residues 96 to 150 of NCX1.1, while the α2‐repeat stretches from 797 to 849 (amino acid numbering of NCX1.1 without the signal sequence). These boundaries were selected as previously described in Ref. 245. The residues that coordinate Na+ and Ca2+ ions, as revealed by the atomic structure of the archaebacterial exchanger 150, are indicated with a red arrow. (B) In addition to the conservation between species, the α‐repeats show intramolecular homology. This forms the foundation for the idea that exchangers are a result of gene duplication.
Figure 7. Figure 7. Domains and residues relevant for NCX1.1 regulation. (A) Highlighted are the residues of the canine cardiac Na+‐Ca2+ exchanger found important for its regulation. For a detailed role of these regions in controlling NCX transport activity please see the main text. Briefly, the residues shown in pink constitute the XIP region, which is intimately involved with the Na+‐dependent inactivation 166. The two helix bundle domain (THB) follows in yellow and its structure is shown in (B) (PDB# 6BV7) 295. The role of this domain remains to be determined. The residues encompassing the two Ca2+ regulatory domains are shown in light blue (Ca2+‐binding domain 1, CBD1) and dark blue (Ca2+‐binding domain 2, CBD2), their atomic structure is shown in (C) and (D), respectively [CBD1 PDB# 2PDK 189 and CBD2 PDB# 2QVM 16]. At the distal end of the large cytoplasmic loop, there is a stretch of residues (in dark grey) forming an amphipathic helix, required for palmitoylation of cysteine at position 739 207. Single point mutations that affect Na+, Ca2+, and pH regulation are indicated.
Figure 8. Figure 8. The XIP region is essential for Na+‐dependent inactivation. (A) Sequence alignment of the XIP region (amino acids 219 to 238, note NCX_Mj has no XIP region). (B) Mutations within this region either accelerate or abrogate the time course of current decay due to raised intracellular Na+, so‐called “Na+‐dependent inactivation”. Modified, with permission, from Matsuoka S, et al., 1997 166.
Figure 9. Figure 9. Properties of the exchanger Ca2+‐binding domain 1 (CBD1). (A) Protein sequence alignment of the CBD1 region among human exchanger isoforms. Numbering is reported according to NCX1.1 sequence, excluding the signal peptide. Residues involved in ion coordination are highlighted with red arrows. (B) This domain coordinates 4 Ca2+ ions (PDB code 2DPK, Ca2+ ions are shown in yellow) 189 but only the ion coordinating sites Ca3 and Ca4 have been shown to be relevant for regulation of the cardiac exchanger 196. (C) The ionic current generated by the cardiac exchanger can be evaluated using the giant patch technique 100. Shown is the Ca2+ dependency of WT NCX1.1 ionic currents and the indicated mutants 196. Mutations E385A and D446A‐D447A disrupt the binding of Ca2+ from sites 3 and 4 respectively and decreases the sensitivity of NCX1.1 to cytoplasmic Ca2+ of approximately 10‐fold.
Figure 10. Figure 10. Properties of the exchanger Ca2+‐binding domain 2 (CBD2). (A) Protein sequence alignment of CBD2 regions among human exchanger isoforms. The protein regions generated by alternative splicing (see Figure 2) are shown. Key residues for Ca2+ coordination are indicated. The lysine at position 585 does not directly coordinate Ca2+ ions but maintains CBD2 structural integrity by coordinating Asp 552 and Glu 648 in the absence of Ca2+ 16,93. Residue 683 (Glu) corresponds to Glu 648 in isoforms expressing exons AD and to Glu 647 in exon BD expressing exchangers. Numbering is reported according to NCX1.1 sequence, excluding the signal peptide. (B) The alternative splicing within CBD2 affects Ca2+ coordination. NMR and X‐ray structures show that isoforms expressing exon A allows coordination of two Ca2+ ions (PDB code 2QVM) 16,93,95, while those expressing exon B (PDB code 2KLT) 93 are unable to coordinate Ca2+. The presence of a positive charge at position 578 and a cysteine at position 585 in exon B expressing CBD2s prevents Ca2+ coordination 16,93,95. (C) Example of ionic currents recorded from exchangers expressing either exon A or B is shown. While the steady state current of the brain isoform NCX1.4 (AD exon) augmented in the presence of 10 μM cytoplasmic Ca2+, the kidney NCX1.3 (BD exon) steady state ionic current was insensitive to changes in intracellular Ca2+ concentration. The binding of Ca2+ to CBD2 enables exon A expressing isoforms to overcome the Na+‐dependent inactivation with elevation in intracellular Ca2+ 49. In contrast, exon B expressing exchangers lack this important regulatory feature. Modified, with permission, from Dunn J, et al., 2002 49.
Figure 11. Figure 11. The ion coordinating residues are conserved between the archaebacterial and mammalian exchangers. (A) The atomic structure of the archaebacterial exchanger revealed 12 amino acids within the α‐repeats (TMS 2 yellow, TMS 3 green, TMS 7 pink, and TMS 8 cyan) that coordinate Na+ and Ca2+ ions (PDB code 3V5U) 150. (B) The residues involved in ion coordination in NCX_Mj are shown. In parenthesis are reported the homologous residues in the mammalian NCX1.1 (numbered without taking into account the signal peptide). The three sodium coordinating sites are identified as SExt, SMid, and SInt. The Ca2+ ion binding site is referred to as SCa 150. New evidence indicates that SMid coordinates water instead of Na+ 151,162.
Figure 12. Figure 12. Residues relevant for NCX transport activity. The location of residues whose replacement perturbed NCX1.1 transport activity is shown. Amino acid numbering is based on the sequence of the canine NCX1.1 exchanger, excluding the signal peptide. The indicated residues were mutated one at a time and the activity of the resulting exchanger was analyzed either electrophysiologically or as Na‐dependent 45Ca2+ uptake 47,112,114,125,183,184,195. Mutations within the α‐repeats drastically affect transport activity. This is consistent with these regions harboring the residues involved in ion coordination, as revealed by the crystal structure of NCX_Mj 150 (see Figure 11). The amino acids coordinating the transported ions (marked with a red circle) are well conserved between NCX_Mj and the mammalian exchanger (see Figure 11) and their replacement impairs activity (grey circle).
Figure 13. Figure 13. Molecular mechanisms of ion transport. (A) Superimposed structures of NCX_Mj obtained in the presence of high Na+ (100 mM, gold, PDB code 5HXE) or low Na+ (10 mM, blue, PDB code 5HWY), Ca2+ ions were absent 151. Binding of Na+ (red spheres) elicits conformational changes throughout the protein. The movements of the external half of TMS 7 govern the transition from outward facing state to the occluded configuration 151. In the absence of Na+, TMS 7 is a contiguous “straight” helix. Upon binding of Na+ to SExt, TMS 7 bends (forming two helices) resulting in its movement away from TMS 6. This allows TMSs 1 and 6 to slide perpendicular to the membrane and transition the protein through occluded state (not shown) to the inward state (not shown). (B) Cysteines scanning mutagenesis allowed the mapping of the residues of the canine cardiac exchanger (NCX1.1) accessible from the cytoplasmic side. Shown are the homologous NCX_Mj residues mapped onto the inward model 125,150. Cysteine introduced at positions highlighted in cyan were accessible to cytoplasmically applied membrane‐impermeable sulfhydryl agent independently of NCX1.1 conformational state of. Residues labeled in orange were accessible to intracellular sulfhydryl agents only when the exchanger was constrained to the inward state. Residues labeled in grey were unaffected by sulfhydryl agents as assessed by ionic current inhibition. (C) Surface representation of NCX_Mj viewed from the cytoplasm. The inward‐facing model predicts two separate cytoplasmic pathways for ions to reach their binding sites 150 and cysteine scanning mutagenesis conducted in the mammalian exchanger support this hypothesis 125. Residues are color coded as described in (B). (D) NCX_Mj residues corresponding to those of NCX1.1 tested for reactivity to sulfhydryl reagents are highlighted and indicated in the table. (B–D) Modified, with permission, from John SA, et al., 2013 125, National Academy of Sciences.
Figure 14. Figure 14. pH regulation of NCX1.1. (A) Cytoplasmic acidification inhibits NCX activity; conversely, alkaline pH enhances NCX ionic currents and renders NCX insensitive to both Ca2+ and Na+ (not shown) 45,46,124. Outward exchanger currents were recorded from oocytes expressing the canine cardiac exchanger NCX1.1 using the giant patch technique. (B) Detailed electrophysiological studies show that mutations that disrupt binding of Ca2+ either to CBD1 (Δ446–499) or CBD2 (E516L) do not decrease NCX pH sensitivity 124. This result suggests that displacement of Ca2+ from the regulatory sites is not essential to confer pH sensitivity to the full‐length exchanger. This contrasts with experiments conducted with the isolated Ca2+‐binding domains (CBD1 and CBD2) showing that protons can displace Ca2+ from their binding sites 22. (C) Replacement of histidine 125 and 165 with alanine drastically decreases the sensitivity of the cardiac exchanger to cytosolic protons and in the case of position 165 also affected the cooperativity of the binding. Measurements were conducted on the canine cardiac exchanger expressed in Xenopus laevis oocytes as detailed in 124. (A,C) Modified, with permission, from John S, et al., 2018 124, © 2018, Rockfeller University Press.
Figure 15. Figure 15. NCX1.1 distribution in cardiac cells. The cartoon summarizes the distribution of NCX within adult cardiomyocytes and its coupling to other proteins essential for excitation‐contraction coupling. (A) The exchanger has been localized in proximity to voltage‐dependent Na+ (Nav,). The influx of Na+ via voltage‐dependent Na+ channels drives NCX to function in reverse mode and therefore prime the dyadic cleft with Ca2+ 140,271. (B) Changes in NCX1.1 expression (overexpression or knockout) regulate the extent of the voltage‐dependent Ca2+ channels (LTCCs) Ca2+‐dependent inactivation process, suggesting that these two proteins are in close proximity within the dyadic cleft. (C) NCX forms a macromolecular complex with the Na+/K+ ATPase (NKA), inositol triphosphate receptors (IP3R) and ankyrin (ANKB) 172. Genetic disorders due to defective expression of ankyrin result in disruption of this macromolecular complex leading to cardiac arrhythmia 173. (D) NCX appears to be in tight connection with the transient receptor potential channel (TRPC3) channels. The current model predicts that the influx of Na+ via TRPC3 tunes NCX transport activity by changing the Na+ levels in its proximity. If elevated enough, Na+ may reverse NCX augmenting intracellular Ca2+ levels 53,54.
Figure 16. Figure 16. NCX1 genetically modified mouse lines. (A) The cartoon shows a timeline of the mouse lines generated with altered expression levels of NCX. (B,C) Overexpression or ablation of NCX affects the cardiac action potential duration. The effects on the action potential waveforms have been connected to changes in K+ currents density 219, although both Ca2+ channels and NCX may contribute to this phenotype. Additional studies are needed to fully elucidate the impact of NCX on both the contractile and electrical properties of the heart. (B,C) Modified, with permission, from Bögeholz N, et al., 2016 20, © 2016, Elsevier.


Figure 1. Modes of operation of the Na+‐Ca2+ exchanger. (A) In the forward mode (top) NCX transports one Ca2+ ion out of the cell in exchange for three Na+ ions entering. One net charge is moved across the membrane resulting in a measurable ionic current. This is the most physiologically relevant cycle. In the reverse mode (bottom), one Ca2+ ions enter the cell and three Na+ ions exit. The direction in which Ca2+ is moved is governed by both the ionic gradients of Na+ and Ca2+ and membrane potential 164. (B) The scheme shows NCX transport cycle. This electrogenic transporter transitions between the outward and inward‐facing states via intermediate occluding states, leading to the coupled but opposite movements of Na+ and Ca2+ ions across the plasma membrane 164.


Figure 2. Na+‐Ca2+ exchanger gene organization. Each panel depicts the chromosomal location, number of exons, size, mRNA example, and corresponding protein of the three human isoforms. Gene exon structures are derived from Ensembl. Exons are indicated as grey bars and include both coding and noncoding exons. Exons undergoing alternative splicing are shown in color and are named in accordance with Quednau and colleagues 220. 5′ and 3′ regions of the gene are not included so that the gene size given extends from the beginning of the first exon and ends at the 3′ end of the last exon. Of special note, NCX2 does not undergo alternative splicing and all exons encode for protein. NCX1 and NCX3 have one noncoding exon, both undergo alternative splicing. An example of mRNA profile of the indicated exchangers is shown under each corresponding gene. Graphs are to scale. The alternative spliced exons (in colors), which at the protein level result in changes in the calcium‐binding domain 2 (CBD2) 220, are depicted in blue.


Figure 3. Tissue distribution of the exchanger isoforms. The three exchanger isoforms and their corresponding splice variants are differentially distributed within tissues 133,136,143,220,221. Examples are given for heart, brain, kidney, and skeletal muscle. The tissue distribution has been assessed using reverse transcriptase‐polymerase chain reaction as described in Ref. 220. Numerous evidence indicates that the isoform NCX1.1 136,143,220 is the only exchanger expressed in cardiomyocytes, although a study has identified isoforms NCX1.3 and NCX1.4 133. The splice variants NCX1.6 (ACD) and NCX3.3 (BC) are absent from the table. These exchangers have been localized in the brain 136,220, but their cell specific expression is undetermined.


Figure 4. Progression of Na+‐Ca2+ exchanger topology. The cartoons show how the topology of the mammalian exchanger has evolved through the years since its cloning in 1990 185. (A) Initially, NCX1.1 was predicted to be organized into 12 transmembrane segments (TMS). The two highly conserved regions among the exchangers which are known as α‐repeats 245 are highlighted. These regions are essential for ion transport 183,184,195. (B) Subsequent studies 50,60,106,239 revealed that the first helix was a signal peptide not necessary for activity, restricting the mature protein to 11 TMSs with an extracellular N terminus. (C) Cysteine scanning mutagenesis and epitope tagging were then applied to investigate NCX1.1 architecture. Based on these studies the topology of NCX was reorganized into nine transmembrane segments with an intracellular C‐terminus and two reentrant loops between TMSs 2‐3 and 7‐8 35,112,114,186,187,212. (D) The crystal structure of the archaebacterial exchanger (NCX_Mj) 150 has revealed a protein of 10 TMSs, contrasting with the predicted 9 TMSs for the mammalian exchanger as shown in (C). (E) Topology of the mammalian NCX based on NCX_Mj organization. The new architecture does not include the previously modeled reentrant loops, which are now predicted to be a short extracellular loop between TMSs 2‐3, within the α1‐repeat, and a new transmembrane segment (TMS 8) in the α2‐repeat. Future studies should be conducted to confirm the absence of either or both of these two reentrant structures and to validate that the mammalian exchanger and the archaebacterial are had a similar organization. The position of the cysteine residues involved in the intramolecular disulfide bond 241 is also shown. Amino acids are numbered based on the protein lacking the signal sequence.


Figure 5. Inconsistencies between the topology of the Na+‐Ca2+ exchanger and the archaebacterial exchanger. (A) Cysteine scanning mutagenesis and epitope mapping studies have been instrumental in deciphering the topology of the cardiac exchanger 35,50,112,114,186,187,212. The figure shows the amino acids that were replaced with cysteine and found accessible to membrane‐impermeable sulfhydryl agents either from the cytoplasm (blue circles) or the external side of the membrane (orange circles). Based on these investigations, NCX was modeled to have an extracellular N‐terminus, and a cytoplasmic C‐terminus. Moreover, residues 120 and 125 within the α1‐repeat were found to be exposed to the intracellular environment as well as several residues within the α2‐repeat, constraining these regions into reentrant loops. This organization corresponds to the one shown in Figure 4C. (B) With the atomic structure of the archaebacterial exchanger at hand 150, NCX1.1 is now assumed to be organized in 10 transmembrane segments 228,265, as shown in Figure 4E. However, this new arrangement places several residues previously found accessible to the cytoplasmic application of sulfhydryl reagents (blue circles), now facing the extracellular environment, including the C‐terminus. Note that the new architecture does not include reentrant loops. More investigations are needed to fully elucidate the topology of the mammalian exchanger.


Figure 6. Sequence alignment of the α‐repeats. (A) The sequence alignment illustrates the conservation of the α repeat regions within the Na+‐Ca2+ exchanger family. This is consistent with a broad range of evidence underlying their essential role in ion transport 125,183,184,195. The α1‐repeat encompasses residues 96 to 150 of NCX1.1, while the α2‐repeat stretches from 797 to 849 (amino acid numbering of NCX1.1 without the signal sequence). These boundaries were selected as previously described in Ref. 245. The residues that coordinate Na+ and Ca2+ ions, as revealed by the atomic structure of the archaebacterial exchanger 150, are indicated with a red arrow. (B) In addition to the conservation between species, the α‐repeats show intramolecular homology. This forms the foundation for the idea that exchangers are a result of gene duplication.


Figure 7. Domains and residues relevant for NCX1.1 regulation. (A) Highlighted are the residues of the canine cardiac Na+‐Ca2+ exchanger found important for its regulation. For a detailed role of these regions in controlling NCX transport activity please see the main text. Briefly, the residues shown in pink constitute the XIP region, which is intimately involved with the Na+‐dependent inactivation 166. The two helix bundle domain (THB) follows in yellow and its structure is shown in (B) (PDB# 6BV7) 295. The role of this domain remains to be determined. The residues encompassing the two Ca2+ regulatory domains are shown in light blue (Ca2+‐binding domain 1, CBD1) and dark blue (Ca2+‐binding domain 2, CBD2), their atomic structure is shown in (C) and (D), respectively [CBD1 PDB# 2PDK 189 and CBD2 PDB# 2QVM 16]. At the distal end of the large cytoplasmic loop, there is a stretch of residues (in dark grey) forming an amphipathic helix, required for palmitoylation of cysteine at position 739 207. Single point mutations that affect Na+, Ca2+, and pH regulation are indicated.


Figure 8. The XIP region is essential for Na+‐dependent inactivation. (A) Sequence alignment of the XIP region (amino acids 219 to 238, note NCX_Mj has no XIP region). (B) Mutations within this region either accelerate or abrogate the time course of current decay due to raised intracellular Na+, so‐called “Na+‐dependent inactivation”. Modified, with permission, from Matsuoka S, et al., 1997 166.


Figure 9. Properties of the exchanger Ca2+‐binding domain 1 (CBD1). (A) Protein sequence alignment of the CBD1 region among human exchanger isoforms. Numbering is reported according to NCX1.1 sequence, excluding the signal peptide. Residues involved in ion coordination are highlighted with red arrows. (B) This domain coordinates 4 Ca2+ ions (PDB code 2DPK, Ca2+ ions are shown in yellow) 189 but only the ion coordinating sites Ca3 and Ca4 have been shown to be relevant for regulation of the cardiac exchanger 196. (C) The ionic current generated by the cardiac exchanger can be evaluated using the giant patch technique 100. Shown is the Ca2+ dependency of WT NCX1.1 ionic currents and the indicated mutants 196. Mutations E385A and D446A‐D447A disrupt the binding of Ca2+ from sites 3 and 4 respectively and decreases the sensitivity of NCX1.1 to cytoplasmic Ca2+ of approximately 10‐fold.


Figure 10. Properties of the exchanger Ca2+‐binding domain 2 (CBD2). (A) Protein sequence alignment of CBD2 regions among human exchanger isoforms. The protein regions generated by alternative splicing (see Figure 2) are shown. Key residues for Ca2+ coordination are indicated. The lysine at position 585 does not directly coordinate Ca2+ ions but maintains CBD2 structural integrity by coordinating Asp 552 and Glu 648 in the absence of Ca2+ 16,93. Residue 683 (Glu) corresponds to Glu 648 in isoforms expressing exons AD and to Glu 647 in exon BD expressing exchangers. Numbering is reported according to NCX1.1 sequence, excluding the signal peptide. (B) The alternative splicing within CBD2 affects Ca2+ coordination. NMR and X‐ray structures show that isoforms expressing exon A allows coordination of two Ca2+ ions (PDB code 2QVM) 16,93,95, while those expressing exon B (PDB code 2KLT) 93 are unable to coordinate Ca2+. The presence of a positive charge at position 578 and a cysteine at position 585 in exon B expressing CBD2s prevents Ca2+ coordination 16,93,95. (C) Example of ionic currents recorded from exchangers expressing either exon A or B is shown. While the steady state current of the brain isoform NCX1.4 (AD exon) augmented in the presence of 10 μM cytoplasmic Ca2+, the kidney NCX1.3 (BD exon) steady state ionic current was insensitive to changes in intracellular Ca2+ concentration. The binding of Ca2+ to CBD2 enables exon A expressing isoforms to overcome the Na+‐dependent inactivation with elevation in intracellular Ca2+ 49. In contrast, exon B expressing exchangers lack this important regulatory feature. Modified, with permission, from Dunn J, et al., 2002 49.


Figure 11. The ion coordinating residues are conserved between the archaebacterial and mammalian exchangers. (A) The atomic structure of the archaebacterial exchanger revealed 12 amino acids within the α‐repeats (TMS 2 yellow, TMS 3 green, TMS 7 pink, and TMS 8 cyan) that coordinate Na+ and Ca2+ ions (PDB code 3V5U) 150. (B) The residues involved in ion coordination in NCX_Mj are shown. In parenthesis are reported the homologous residues in the mammalian NCX1.1 (numbered without taking into account the signal peptide). The three sodium coordinating sites are identified as SExt, SMid, and SInt. The Ca2+ ion binding site is referred to as SCa 150. New evidence indicates that SMid coordinates water instead of Na+ 151,162.


Figure 12. Residues relevant for NCX transport activity. The location of residues whose replacement perturbed NCX1.1 transport activity is shown. Amino acid numbering is based on the sequence of the canine NCX1.1 exchanger, excluding the signal peptide. The indicated residues were mutated one at a time and the activity of the resulting exchanger was analyzed either electrophysiologically or as Na‐dependent 45Ca2+ uptake 47,112,114,125,183,184,195. Mutations within the α‐repeats drastically affect transport activity. This is consistent with these regions harboring the residues involved in ion coordination, as revealed by the crystal structure of NCX_Mj 150 (see Figure 11). The amino acids coordinating the transported ions (marked with a red circle) are well conserved between NCX_Mj and the mammalian exchanger (see Figure 11) and their replacement impairs activity (grey circle).


Figure 13. Molecular mechanisms of ion transport. (A) Superimposed structures of NCX_Mj obtained in the presence of high Na+ (100 mM, gold, PDB code 5HXE) or low Na+ (10 mM, blue, PDB code 5HWY), Ca2+ ions were absent 151. Binding of Na+ (red spheres) elicits conformational changes throughout the protein. The movements of the external half of TMS 7 govern the transition from outward facing state to the occluded configuration 151. In the absence of Na+, TMS 7 is a contiguous “straight” helix. Upon binding of Na+ to SExt, TMS 7 bends (forming two helices) resulting in its movement away from TMS 6. This allows TMSs 1 and 6 to slide perpendicular to the membrane and transition the protein through occluded state (not shown) to the inward state (not shown). (B) Cysteines scanning mutagenesis allowed the mapping of the residues of the canine cardiac exchanger (NCX1.1) accessible from the cytoplasmic side. Shown are the homologous NCX_Mj residues mapped onto the inward model 125,150. Cysteine introduced at positions highlighted in cyan were accessible to cytoplasmically applied membrane‐impermeable sulfhydryl agent independently of NCX1.1 conformational state of. Residues labeled in orange were accessible to intracellular sulfhydryl agents only when the exchanger was constrained to the inward state. Residues labeled in grey were unaffected by sulfhydryl agents as assessed by ionic current inhibition. (C) Surface representation of NCX_Mj viewed from the cytoplasm. The inward‐facing model predicts two separate cytoplasmic pathways for ions to reach their binding sites 150 and cysteine scanning mutagenesis conducted in the mammalian exchanger support this hypothesis 125. Residues are color coded as described in (B). (D) NCX_Mj residues corresponding to those of NCX1.1 tested for reactivity to sulfhydryl reagents are highlighted and indicated in the table. (B–D) Modified, with permission, from John SA, et al., 2013 125, National Academy of Sciences.


Figure 14. pH regulation of NCX1.1. (A) Cytoplasmic acidification inhibits NCX activity; conversely, alkaline pH enhances NCX ionic currents and renders NCX insensitive to both Ca2+ and Na+ (not shown) 45,46,124. Outward exchanger currents were recorded from oocytes expressing the canine cardiac exchanger NCX1.1 using the giant patch technique. (B) Detailed electrophysiological studies show that mutations that disrupt binding of Ca2+ either to CBD1 (Δ446–499) or CBD2 (E516L) do not decrease NCX pH sensitivity 124. This result suggests that displacement of Ca2+ from the regulatory sites is not essential to confer pH sensitivity to the full‐length exchanger. This contrasts with experiments conducted with the isolated Ca2+‐binding domains (CBD1 and CBD2) showing that protons can displace Ca2+ from their binding sites 22. (C) Replacement of histidine 125 and 165 with alanine drastically decreases the sensitivity of the cardiac exchanger to cytosolic protons and in the case of position 165 also affected the cooperativity of the binding. Measurements were conducted on the canine cardiac exchanger expressed in Xenopus laevis oocytes as detailed in 124. (A,C) Modified, with permission, from John S, et al., 2018 124, © 2018, Rockfeller University Press.


Figure 15. NCX1.1 distribution in cardiac cells. The cartoon summarizes the distribution of NCX within adult cardiomyocytes and its coupling to other proteins essential for excitation‐contraction coupling. (A) The exchanger has been localized in proximity to voltage‐dependent Na+ (Nav,). The influx of Na+ via voltage‐dependent Na+ channels drives NCX to function in reverse mode and therefore prime the dyadic cleft with Ca2+ 140,271. (B) Changes in NCX1.1 expression (overexpression or knockout) regulate the extent of the voltage‐dependent Ca2+ channels (LTCCs) Ca2+‐dependent inactivation process, suggesting that these two proteins are in close proximity within the dyadic cleft. (C) NCX forms a macromolecular complex with the Na+/K+ ATPase (NKA), inositol triphosphate receptors (IP3R) and ankyrin (ANKB) 172. Genetic disorders due to defective expression of ankyrin result in disruption of this macromolecular complex leading to cardiac arrhythmia 173. (D) NCX appears to be in tight connection with the transient receptor potential channel (TRPC3) channels. The current model predicts that the influx of Na+ via TRPC3 tunes NCX transport activity by changing the Na+ levels in its proximity. If elevated enough, Na+ may reverse NCX augmenting intracellular Ca2+ levels 53,54.


Figure 16. NCX1 genetically modified mouse lines. (A) The cartoon shows a timeline of the mouse lines generated with altered expression levels of NCX. (B,C) Overexpression or ablation of NCX affects the cardiac action potential duration. The effects on the action potential waveforms have been connected to changes in K+ currents density 219, although both Ca2+ channels and NCX may contribute to this phenotype. Additional studies are needed to fully elucidate the impact of NCX on both the contractile and electrical properties of the heart. (B,C) Modified, with permission, from Bögeholz N, et al., 2016 20, © 2016, Elsevier.
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Article Title: The Cardiac Na+‐Ca2+ Exchanger: From Structure to Function
 

How to Cite

Michela Ottolia, Scott John, Adina Hazan, Joshua I. Goldhaber. The Cardiac Na+‐Ca2+ Exchanger: From Structure to Function. Compr Physiol 2021, 12: 2681-2717. doi: 10.1002/cphy.c200031