AJP - Heart Calcium Transients and Cell-Sarcomere
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Am J Physiol Heart Circ Physiol 290: H2155-H2162, 2006. First published January 6, 2006; doi:10.1152/ajpheart.00958.2005
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Cells expressing unique Na+/Ca2+ exchange (NCX1) splice variants exhibit different susceptibilities to Ca2+ overload

Cecilia Hurtado,1 Michele Prociuk,1 Thane G. Maddaford,1 Elena Dibrov,1 Nasrin Mesaeli,2 Larry V. Hryshko,3 and Grant N. Pierce1,4

1Division of Stroke and Vascular Disease and 3Institute of Cardiovascular Sciences, St. Boniface Hospital Research Centre, and 2Departments of Physiology and Biochemistry and Medical Genetics, Faculties of Medicine and 4Pharmacy, University of Manitoba, Winnipeg, Manitoba, Canada

Submitted 8 September 2005 ; accepted in final form 20 December 2005


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The Na+/Ca2+ exchanger (NCX) NCX1 exhibits tissue-specific alternative splicing. Such NCX splice variants as NCX1.1 and NCX1.3 are also differentially regulated by Na+ and Ca2+, although the physiological implications of these regulatory characteristics are unclear. On the basis of their distinct regulatory profiles, we hypothesized that cells expressing these different splice variants might exhibit unique responses to conditions promoting Ca2+ overload, such as during exposure to cardiac glycosides or simulated ischemia. NCX1.1 or NCX1.3 was expressed in human embryonic kidney (HEK)-293 cells or rat neonatal ventricular cardiomyocytes (NVC), and expression was confirmed by Western blotting and immunocytochemical analyses. HEK-293 cells lacked NCX1 protein before transfection. With use of adenoviral vectors, neonatal cardiomyocytes were induced to overexpress the NCX1.1 splice variant by nearly twofold, whereas the NCX1.3 isoform was expressed on the endogenous NCX1.1 background. Total expression was comparable for NCX1.1 and NCX1.3. Exposure of NVC to ouabain induced a significant increase in cellular Ca2+, an effect that was exaggerated in cells overexpressing NCX1.1, but not NCX1.3. The increase in intracellular Ca2+ was inhibited by 5 µM KB-R7943. Cardiomyocytes overexpressing NCX1.1 also exhibited a greater accumulation of intracellular Ca2+ in response to simulated ischemia than did cells expressing NCX1.3. Similar responses were observed in HEK-293 cells where NCX1.1 was expressed. We conclude that expression of the NCX1.3 splice variant protects against severe Ca2+ overload, whereas NCX1.1 promotes Ca2+ overload in response to cardiac glycosides and ischemic challenges. These results highlight the importance of ionic regulation in controlling NCX1 activity under conditions that promote Ca2+ overload.

NCX1.1; NCX1.3; ouabain; ischemia

THE SODIUM/CALCIUM EXCHANGER (NCX) NCX1 plays an important role in cardiac excitation-contraction coupling, serving as the primary mechanism for Ca2+ efflux (1). NCX1 has also been identified as having an important role in distinct types of cardiovascular pathophysiology, including ischemia-reperfusion injury, hypertension, and heart failure (14, 26, 27). Considerable effort has been focused on understanding the regulation of NCX1, and many investigators have focused on studying the structure-function relations of this protein at the molecular level (20). The NCX1 gene undergoes alternative splicing in a region corresponding to the COOH terminus of its large intracellular loop. Six cassette exons, A, B, C, D, E, and F, code for the alternatively spliced site (15). Exons A and B are mutually exclusive and are followed by various combinations of the remaining four cassette exons. Distinct splice variants of NCX1 are expressed in a tissue-specific manner (16, 22). In general, excitable tissues utilize NCX1 splice variants containing exon A, whereas exon B-containing splice variants are expressed in most other tissues. The only isoform found in the heart is NCX1.1, which expresses exons A, C, D, E, and F. Another isoform, NCX1.3, expresses exons B and D and is found in several nonexcitable tissues. It is the most abundant isoform in the kidney and is also expressed in vascular smooth muscle (31).

Intracellular Na+ and Ca2+, besides being the substrates for the exchanger, are also its allosteric modulators: Na+ acts at the intracellular surface of the protein to produce Na+-dependent inactivation (9, 10), and Ca2+ can alleviate this inactivation (10). High concentrations of intracellular Na+ increase the degree of inactivation (presumably by increasing the population of inactive exchangers). In the absence of intracellular Ca2+, the exchanger enters an inactive state. Ca2+ binding to a high-affinity regulatory site within the cytoplasmic loop of the exchanger removes this inactivation (17, 19). Consequently, elevations in cytosolic Ca2+ increase NCX1 activity.

NCX1 regulation has been extensively studied using the giant patch-clamp technique (8) in membrane patches from ventricular cells (7) and from Xenopus laevis oocytes expressing different NCX isoforms (4, 5). This technique has been used to demonstrate that NCX1 splice variants function differently with respect to their ionic regulatory properties. Specifically, inactivation of the reverse-transport mode (i.e., Ca2+ entry) is significantly more pronounced in the NCX1.3 splice variant (5, 12). Furthermore, an increase in intracellular Ca2+ reduces Na+-dependent inactivation in NCX1.1, but not NCX1.3. There is no clear understanding about the physiological significance of these distinct ionic regulatory patterns or the basis for splice variant diversity.

In this study, we sought to gain insight into the potential role of ionic regulatory differences under conditions that promoted cellular Ca2+ overload. Because increasing intracellular Ca2+ concentration ([Ca2+]i) decreases Na+-dependent inactivation in NCX1.1, but not NCX1.3, we hypothesized that by promoting such conditions (i.e., high intracellular Na+ and Ca2+), we might be able to discern differences in cellular Ca2+ handling. Adenoviral-induced overexpression of the NCX1.1 or NCX1.3 splice variant in neonatal ventricular cardiomyocytes (NVC) and stable transfections of human embryonic kidney (HEK)-293 cells were therefore used to determine whether functional differences were apparent in cells exposed to simulated ischemia or in response to ouabain treatment.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cells

HEK-293 cells. HEK-293 cells (Quantum) were grown in DMEM-5% FBS. HEK-293 cells expressing the NCX1 isoforms were generated by cotransfection of the pShuttle-CMV plasmid carrying the canine cDNA of NCX1.1 or NCX1.3 and the pcDNA3.1-Hygro plasmid using Lipofectamine (Invitrogen) and subsequent selection with hygromycin (200 µg/ml). Clones that expressed similar levels of NCX1.1 and NCX1.3 protein were chosen for the study.

NVC. NVC were isolated from the ventricles of 0- to 24-h-old Sprague-Dawley rats as described by Doble et al. (3). NVC were plated in F-10-DMEM, 10% FBS, and 10% horse serum at a density of 0.75 x 106 cells per 35-mm collagen-coated culture dish. On the following day, the cells were rinsed once with DMEM and maintained in DMEM containing 0.5% FBS, 20 nmol/l selenium, 10 µg/ml insulin, 10 µg/ml transferrin, 2 mg/ml BSA, and 20 µg/ml ascorbic acid, and adenovirus was added. The cells were studied 44–52 h after infection.

Adenoviral Vectors

Canine NCX1.1 or NCX1.3 cDNA was cloned into the pShuttle-CMV plasmid (Quantum) for generation of recombinant adenovirus vectors. Viral amplification was performed after single-plaque purification. NVC were infected using 1.5 viruses/cell. An adenovirus that codes for beta-galactosidase (Ad-beta-Gal; Quantum) was used as control.

Western Blot

Samples were lysed with RIPA buffer (50 mM Tris, 150 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 1 mM EDTA, and 1 mM EGTA, pH 7.5, with 1 mM PMSF, 1 mM benzamidine, and a protease inhibitor cocktail). After 50 µg of total protein in Laemmli sample buffer were loaded per gel lane, 4–16% Tris·HCl-acrylamide gradient gel (Bio-Rad) was used for SDS-PAGE. Proteins were transferred onto nitrocellulose membrane in a wet transfer apparatus overnight at 4°C at 35 V. The primary antibodies were anti-NCX (R3F1 monoclonal, 1:1,000 dilution; Swant), antiactin A2066 (1:2,000 dilution; Sigma), and anti-Na+-K+-ATPase {alpha}1-subunit (1:300 dilution; Developmental Studies Hybridoma Bank). The membrane was incubated with the appropriate horseradish peroxidase-conjugated secondary antibody, and signal was developed using West Pico chemiluminescence substrate (Pierce). Signal was collected in a Bio-Rad detection system and quantified by densitometry analysis using Quantity One software (Bio-Rad).

Biotinylation of Surface Membrane Proteins

Biotinylation was performed on NVC adherent to the culture plate or HEK-293 cells in suspension. Biotin reagent (Z-Link-Sulfo-NHS-Biotin, Pierce Biotechnology) was added to the cells in a final concentration of 3 mM. After 30 min of incubation at 4°C, the cells were washed with 100 mM glycine in PBS and lysed in RIPA buffer containing protease inhibitors. The lysate was stored at –80°C for future analysis.

NCX Immunoprecipitation

Goat antibiotin (Pierce) or anti-NCX R3F1 (Swant) monoclonal antibody was used to precipitate NCX protein from the biotinylated samples. Briefly, antibody was added to precleared cell lysate containing 100–200 µg of the total protein. After incubation with rotation for 16 h at 4°C, the protein-antibody complex was precipitated with immobilized protein G (Calbiochem), washed, extracted with sample buffer, and used for SDS-PAGE and Western blots. Antibiotin horseradish peroxidase-conjugated antibody (Sigma-Aldrich) was used to recognize biotinylated NCX precipitated with the R3F1 antibody, and R3F1 antibody was used to recognize NCX precipitated with the antibiotin antibody.

Immunocytochemistry

Cells on coverslips were fixed for 12 min in 4% paraformaldehyde in PBS, permeabilized with 0.1% Triton X-100, and blocked with 2% milk-0.1% Triton X-100 in PBS. R3F1 primary antibody to NCX (1:150 dilutions) was followed by Alexa-conjugated goat anti-mouse secondary antibody (1:700 dilutions). Nuclei were stained with Hoechst 33342. Cells were mounted on glass slides using FluorSave (Calbiochem). All cells were imaged on a Zeiss fluorescent microscope.

Measurement of NCX Activity

NCX activity in the cardiomyocytes was measured as intracellular Na+-dependent 45Ca2+ uptake following the protocol described by Vemuri et al. (29). The cells were plated on 35-mm petri dishes, washed twice with an Na+ loading solution (140 mM NaCl, 2 mM MgCl2, and 10 mM MOPS, pH 7.4), and incubated at 4°C for 10 min with the same Na+ loading solution with the addition of 5 mM ouabain. The solution was then replaced by 2 ml of uptake medium containing 25 µM CaCl2, 0.3 µCi/ml 45Ca2+, 10 mM MOPS (pH 7.4), 5 mM ouabain, and either 140 mM KCl or 140 mM NaCl at room temperature. After 2 min, the cells were rapidly washed three times with 3 ml of an ice-cold solution containing 140 mM KCl and 1 mM EGTA, pH 7.4, to stop the uptake reaction. The cells were then collected with 500 µl of 0.5 N NaOH. A 20-µl aliquot of the lysate was used for measurement of protein concentration, and 100 µl were used for radioactive counting in a Beckman scintillation counter. Measurements were performed in duplicate. NCX activity was calculated from the difference between the uptake in the KCl solution and the uptake in the NaCl solution (blank) and was expressed as 45Ca2+ counts per milligram of protein per 2 min of uptake reaction. In HEK-293 cells, NCX activity was measured as Ca2+ entry after equimolar replacement of extracellular Na+ with extracellular Li+ in fura 2-loaded cells as described below with a spectrofluorometer used to measure NCX activity.

Intracellular Ca2+ Measurements

NVC were incubated with 2 µM fura 2-AM and 0.04% pluronic acid in HEPES buffer at room temperature for 15 min and then deesterified for 30 min. HEK-293 cells were incubated with 2 µM fura 2-AM and 0.04% pluronic acid in HEPES buffer at 37°C for 30 min and then deesterified for 30 min. Spectrofluorometric measurements were performed on cells attached to a coverslip and then placed on the stage of a microscope or in a cuvette. [Ca2+]i was expressed as the 340-to-380-nm ratio of fura-2 fluorescence.

Simulated Ischemia-Reperfusion

Cells were perfused with a control HEPES buffer containing 140 mM NaCl, 6 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2, 6 mM HEPES (pH 7.4), and 10 mM dextrose, bubbled with 100% O2. To simulate ischemia, we used a modification of the protocol described in detail elsewhere (13). After control perfusion, the perfusate was switched to an ischemia-mimetic solution containing 130 mM NaCl, 8 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2, 6 mM HEPES (pH 6.0), 2.5 mM sodium cyanide, and 10 mM sodium lactate. If present, KB-R7943 was included in the ischemic buffer and during the first 10 min of reperfusion.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Figure 1 depicts immunostaining for NCX1 in control, NCX1.1, and NCX1.3 stably transfected HEK-293 cells and adenovirus-transfected NVC. The R3F1 monoclonal antibody was used to detect NCX1.1 and NCX1.3. NCX1 is absent in control HEK-293 cells. In the other groups, NCX1 localized to the membrane of cells. More intense NCX1 staining was observed in NVC infected with adenovirus for NCX1.1 (Ad-NCX1.1) or NCX1.3 (Ad-NCX1.3) transgenes than in control cardiomyocytes. In cells expressing the transgene, NCX1 was also observed within the cells, particularly surrounding the nuclei. Most likely, this represents protein within the endo/sarcoplasmic reticulum that was not completely processed.


Figure 1
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  		Fig. 1. Immunostaining of Na+/Ca2+ exchanger (NCX) as a function of expression of different NCX isoforms. Human embryonic kidney (HEK)-293 cells and neonatal ventricular cardiomyocytes (NVC) expressing NCX1.1 and NCX1.3 are shown. Images of NCX (green) and nuclear staining with Hoechst 33342 (blue) are superimposed. Exposure conditions were identical for the 3 images of each cell type. Insets: higher magnification.

 
Western blot analysis (Fig. 2) showed the characteristic 120- and 160-kDa bands corresponding to NCX1 in control NVC and transfected cells. The calculated size of the protein is 120 kDa. Under nonreducing or limited reducing conditions, the exchanger can also be observed as a 160-kDa band. The change in size is due to the formation of intramolecular disulfide bridges between cysteine residues (24). No bands were detected in control HEK-293 cells (Fig. 2A). In transfected HEK-293 cells, NCX1.3 was slightly smaller than NCX1.1, because NCX1.3 codes for 36 fewer amino acids than NCX1.1. NCX1 expression in HEK-293 cells transfected with the NCX1.3 splice variant was 155 ± 35% of that observed for cells expressing NCX1.1, which was considered to be 100% (P > 0.05, n = 4). NCX transgene expression was similar between Ad-NCX1.1- and Ad-NCX1.3-transfected cells (Fig. 2B). NCX1.3 expression was 102 ± 15% (P > 0.05, n = 3) compared with NCX1.1 expression, which was considered to be 100%. We did not compare relative amounts with control cells, because the transgene is of canine origin and the antibody has a different species affinity (21). Qualitatively similar results were obtained in three independent experiments. In NVC infected with a control adenovirus expressing beta-Gal, NCX expression levels were similar to those in uninfected cells. Also adenoviral transfection did not alter expression of the Na+ pump, and actin was used as a loading control (Fig. 2B). For comparative purposes, equal amounts of total protein from the different experimental groups are shown in Fig. 2C.


Figure 2
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  		Fig. 2. Western blot analysis of NCX expression. A and B: NCX content in samples from HEK-293 cells and NVC, respectively. C: comparison of NCX expression in different experimental groups. D: NCX localization at plasmalemma membrane in NVC. Membrane proteins were biotinylated and immunoprecipitated. NCX was analyzed by SDS-PAGE and detected using the R3F1 antibody.

 
The level of NCX protein expressed at the sarcolemmal membrane (where it is functional) will determine the level of activity of the expressed protein. To verify that similar amounts of NCX1.1 and NCX1.3 were directed to the cell membrane in transfected cells, all proteins that were exposed at the extracellular surface of intact cells were biotinylated and immunoprecipitated. NCX1 was then analyzed by SDS-PAGE. Representative experiments are shown in Fig. 2D. Similar results were obtained with the reciprocal experiment (NCX immunoprecipitation and detection of biotinylated protein). The level of biotinylated NCX1.3 was slightly higher than that of biotinylated NCX1.1 in both cell types: the ratio of NCX1.3 to NCX1.1 was 1.52 ± 0.24 for HEK-293 cells (n = 5) and 1.09 ± 0.20 for NVC (n = 5).

NCX1 activity was determined in fura 2-loaded HEK-293 cells by measuring the increase in intracellular Ca2+ when extracellular Na+ was replaced by Li+. This intervention generates an outwardly directed Na+ gradient, leading to an NCX1-mediated increase in intracellular Ca2+ (reverse-transport mode). Replacing extracellular Na+ with Li+ had no effect on intracellular Ca2+ in control HEK-293 cells (Fig. 3A), because HEK-293 cells do not express NCX. However, a rapid increase in intracellular Ca2+ was observed in cells expressing the NCX1.1 or NCX1.3 splice variants. Pooled data are shown in Fig. 3B. The increase in intracellular Ca2+ was significantly larger in cells expressing NCX1.1 than in those expressing NCX1.3 on the basis of the change in fura 2 fluorescence ratio (4.05 ± 0.08 and 1.60 ± 0.24, respectively, P < 0.05, n = 4–5). Because NCX1 expression levels at the membrane were slightly greater for NCX1.3 than for NCX1.1 (Fig. 2D), these data indicate that, under our experimental conditions, reverse-mode activity was less for NCX1.3 than for NCX1.1. Because this particular assay measures only reverse-mode transport, the observed differences are likely to reflect the intrinsic regulatory characteristics of each splice variant.


Figure 3
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  		Fig. 3. NCX activity. A: representative recordings of fura 2 fluorescence in HEK-293 cells during removal and reapplication of extracellular Na+ (open arrow). Time breaks indicate 20-s interruption in data collection, while the solution was changed. B: mean data from 4–5 experiments in A. C: NCX activity in NVC measured as Na+ gradient-dependent 45Ca2+ uptake. Data are expressed as percentage of NCX activity in control noninfected cardiomyocytes. Values are means ± SE (n = 4–8). *P < 0.05 vs. control. #P < 0.05 vs. NCX1.1.

 
Differences in the rate of recovery of intracellular Ca2+ on reintroduction of extracellular Na+ (Fig. 3A) were observed between NCX1.1- and NCX1.3-expressing HEK-293 cells. Because of technical limitations, we could not collect data during the first ~20 s after solution exchange. In NCX1.1-expressing cells, intracellular Ca2+ was completely restored to control levels when we restarted acquisition and was even rising (a positive slope was observed). In contrast, in NCX1.3-expressing cells at the same time of recording, intracellular Ca2+ was still decreasing toward the pretreatment level (negative slope), indicating that inhibition of exchange activity was slowly being removed. The rates of change of intracellular Ca2+ during reapplication of extracellular Na+ (fura 2 ratio/s) were calculated to be 0.005 ± 0.0007 for NCX1.1 and –0.006 ± 0.0017 for NCX1.3 (P < 0.05).

Figure 3C illustrates the results obtained in control and transfected neonatal cardiomyocytes, where NCX activity was measured as the Na+ gradient-dependent 45Ca2+ uptake. In the Ad-NCX1.1 and Ad-NCX1.3 groups, NCX activity increased 77 ± 0.15% and 15 ± 0.01% over that observed in nontransfected control cells (P < 0.05 vs. control). Ad-beta-Gal infection up to 50 multiplicity of infection units did not affect NCX activity (102 ± 1% of activity in noninfected cells, P > 0.05 vs. control, n = 4). Thus, in NVC and transfected HEK-293 cells, less reverse-mode NCX activity is shown by the NCX1.3 than the NCX1.1 splice variant.

Figure 4 shows the changes in intracellular Ca2+ in control and transduced cardiomyocytes in response to 10 min of treatment with 1 mM ouabain, which should produce near-complete inhibition of the Na+ pump, leading to a progressive rise in intracellular Na+ and Ca2+. Ouabain gradually increased resting Ca2+ in control and Ad-NCX1.3 cardiomyocytes. A much larger change was observed in Ad-NCX1.1-transfected cardiomyocytes. The increase in fura 2 fluorescence ratio relative to the pretreatment value was 0.63 ± 0.08 in control cells, 0.95 ± 0.12 in Ad-NCX1.3, and 1.98 ± 0.12 in Ad-NCX1.1 (P < 0.05 vs. control, P < 0.05 vs. Ad-NCX1.3) at the end of the 10-min ouabain treatment. Although the value for NCX1.3 at the 10-min time point was not statistically different from control, there was a tendency for larger changes in resting Ca2+ concentration throughout the entire treatment. For all earlier time points, there were statistical differences between all three groups during treatment. No differences were observed between noninfected controls and Ad-beta-Gal-infected cells in response to ouabain treatment (Fig. 4B). The effects of KB-R7943, a preferential blocker of the reverse mode of NCX (Ca2+ influx) on the different groups of cardiomyocytes, are shown in Fig. 4C. When 5 µM KB-R7943 was present in the buffer solution 10 min before and during ouabain application, the ouabain-induced increase in intracellular Ca2+ was significantly attenuated in all three groups.


Figure 4
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  		Fig. 4. Effect of 10 min of treatment with 1 mM ouabain on resting Ca2+ as a function of NCX expression shown as fura 2 fluorescence ratio. A: noninfected NVC (control, {blacksquare}) and NVC infected with Ad-NCX1.1 (bullet) or Ad-NCX1.3 ({blacktriangleup}). Values are means ± SE (n = 11–17). *P < 0.05 vs. control. #P < 0.05 vs. NCX1.1. B: noninfected NVC and NVC infected with Ad-beta-galactosidase (Ad-beta-Gal). C: without (solid line) and with (dashed line) 5 µM KB-R7943. Values are means ± SE (n = 3–17). §P < 0.05 vs. without KB-R7943.

 
The transduced cardiomyocytes were further challenged with an ischemia-reperfusion protocol to evaluate the behavior of different splice variants. Cells typically responded to the 90-min ischemic insult with an increase in resting intracellular Ca2+ followed by a transient rise in intracellular Ca2+ during reperfusion and then a slow return to baseline levels (Fig. 5). The increase in Ca2+ was higher in cells expressing NCX1.3 than in control cells, and these changes were even more pronounced in cells overexpressing NCX1.1.


Figure 5
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  		Fig. 5. Effect of simulated ischemia and reperfusion on resting Ca2+ levels as a function of expression of different NCX isoforms in NVC shown as fura 2 fluorescence. Values are means ± SE (n = 23–28). *P < 0.05 vs. control. #P < 0.05 vs. Ad-NCX1.1.

 
Figure 6 shows the changes in intracellular Ca2+ during simulated ischemia and reperfusion in HEK-293 cells. During ischemia, intracellular Ca2+ rose in all three groups. Intracellular Ca2+ was significantly higher during the entire period of ischemia in NCX1.1-expressing than in control cells. There was a tendency for intracellular Ca2+ to be higher during ischemia in the NCX1.3-expressing HEK-293 cells than in control cells, but the values did not reach statistical significance. During the last 40 min of ischemia, intracellular Ca2+ was significantly higher in NCX1.1- than in NCX1.3-expressing HEK-293 cells. On reperfusion, intracellular Ca2+ was significantly higher in NCX1.1- and NCX1.3-expressing cells than in control cells. The difference in resting intracellular Ca2+ between the end of reperfusion and before treatment was not significant for control cells (0.99 ± 0.03 before vs. 1.032 ± 0.019 after, P > 0.05, n = 15) but was significantly different for NCX1.1 (0.98 ± 0.025 before vs. 1.10 ± 0.016 after, P < 0.05, n = 13) and NCX1.3 (1.022 ± 0.016 before vs. 1.14 ± 0.04 after, P < 0.05, n = 12). Similar to our ouabain treatment results (Fig. 4), these increases in [Ca2+]i were blocked in all groups when cells were pretreated with 5 µM KB-R7943 (Fig. 6B).


Figure 6
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  		Fig. 6. Changes in intracellular Ca2+ concentration during simulated ischemia and reperfusion in HEK-293 cells as a function of NCX expression shown as fura 2 fluorescence. A: control nontransfected HEK-293 cells and NCX1.1- and NCX1.3-expressing cells. B: without (top line) and with (bottom line) 5 µM KB-R7943. Values are means ± SE (n = 7–11). *P < 0.05 vs. control. #P < 0.05 vs. NCX1.1. §P < 0.05 vs. without KB-R7943.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Overview

In this study, we expressed two distinct NCX splice variants, NCX1.1 and NCX1.3, in HEK-293 cells and NVC. We then used experimental conditions designed to promote intracellular Na+ and Ca2+ overload to examine changes in intracellular Ca2+ levels for these different cell types. Specifically, cells were exposed to high-dose ouabain or simulated ischemia-reperfusion. The overall goal was to determine whether cellular Ca2+ handling would be altered by expression of unique exchanger splice variants. We demonstrate that intracellular Ca2+ handling differs considerably depending on which splice variant is present, implying that ionic regulatory differences between NCX1.1 and NCX1.3 play an important role in regulating exchange activity during intracellular Na+ and Ca2+ overload.

Ionic Regulation of NCX

Ionic regulatory mechanisms can profoundly influence NCX activity. Both transport substrates, Na+ and Ca2+, allosterically modify the activity of this transporter. Na+-dependent inactivation describes the process whereby intracellular Na+ promotes exchanger inactivation, analogous to the gating of ion channels (11). The XIP region on the intracellular loop of the cardiac exchanger NCX1.1 has been identified as serving a prominent role in mediating Na+-dependent inactivation (18). This process occurs with an intracellular Na+ concentration required for 50% of peak activity of 15–25 mM (5). Therefore, at physiological levels of intracellular Na+, inactivation through this mechanism is unlikely to occur. Intracellular Na+ can be dramatically elevated under pathophysiological conditions (e.g., ouabain treatment or ischemia-reperfusion injury), such that Na+-dependent inactivation could prominently alter the population of active exchangers. However, there is virtually no information on whether this change in the population of active exchangers occurs.

Ca2+-dependent regulation of the exchanger describes the influence of cytosolic Ca2+ on NCX activity. Ca2+ binds to a high-affinity, regulatory site on the cytoplasmic loop of the exchanger and promotes exchange activity (10). In the absence of cytoplasmic Ca2+, the exchanger enters an inactive state, even if the electrochemical gradients strongly favor exchange activity. The Ca2+ concentration required for 50% of peak activity is ~0.2–0.3 µM, which is intermediate between diastolic and systolic Ca2+ levels in cardiomyocytes. This process has been demonstrated to influence exchange activity in a variety of intact cell types (6, 7, 23, 30) and has been thoroughly characterized using the giant excised patch-clamp technique (19, 28).

Differences in Ionic Regulation Between NCX1.1 and NCX1.3

Substantial interactions have been demonstrated between the Na+- and Ca2+-dependent regulatory processes when examined by giant excised patch-clamp studies. For example, all mutations within the XIP region of the exchange that modify Na+-dependent inactivation also produce alterations in the Ca2+ regulatory phenotype (18). Similarly, mutations within the high-affinity, regulatory Ca2+ binding site lead to alterations in Na+-dependent inactivation (19). In the cardiac exchanger NCX1.1, elevated cytosolic Ca2+ can completely alleviate Na+-dependent inactivation (19). Thus, under conditions of elevated Na+ and Ca2+, it might be anticipated that substantial NCX activity would still occur, inasmuch as the effects of regulatory Ca2+ would be predicted to eliminate any inactivation mediated by increases in intracellular Na+. In contrast, NCX1.3 lacks this interaction, such that prominent Na+-dependent inactivation occurs irrespective of intracellular Ca2+ levels (5). Therefore, it seems reasonable to assume that, in cells expressing this splice variant, Na+-dependent inactivation would still occur under conditions of elevated intracellular Na+ and Ca2+. Our experimental results provide substantial support for these anticipated responses.

Functional Differences in Intracellular Ca2+ Handling

In our study, we expressed NCX1.1 and NCX1.3 in HEK-293 cells and neonatal rat ventricular myocytes. In general, expression was slightly greater for NCX1.3 in both cell types. We used two separate means of elevating intracellular Na+ and Ca2+ levels: treatment with 1 mM ouabain and a simulated ischemia-reperfusion paradigm. It has been clearly established that the NCX plays a prominent role in mediating intracellular Ca2+ changes for both of these interventions (25, 27). Our results consistently revealed that overexpression of the cardiac exchanger NCX1.1 resulted in a greater increase in intracellular Ca2+ in response to these treatments. Similar conclusions arose from studies evaluating the consequences of overexpressing NCX1.1 in transgenic mice. In these studies, cardiac-specific NCX1.1 overexpression increased the injury associated with ischemia-reperfusion (2). In contrast, substantially smaller increases in Ca2+ resulted from forced expression of NCX1.3 than NCX1.1; however, expression levels were typically higher for NCX1.3 than for NCX1.1. Intuitively, this implicates the prominent role of reverse-mode NCX in mediating ouabain- and ischemia-reperfusion-induced Ca2+ elevations. That is, although both of these interventions would progressively reduce forward-mode exchange and augment reverse-mode exchange, it is difficult to imagine how increasing exchanger expression could reduce forward-mode exchange.

Our results highlight the intriguing possibility that ionic regulatory differences between two NCX1 splice variants may contribute prominently to NCX function under pathophysiological conditions. Overall, it seems very unlikely that differences in expression levels could account for the observed differences. That is, if expression levels alone were responsible for modulating the extent of intracellular Ca2+ elevation by reverse-mode exchange, we would have predicted more pronounced intracellular Ca2+ elevation in cells expressing NCX1.3 because its expression levels were generally higher. In fact, the opposite was observed. One reasonable possibility is that ionic regulatory mechanisms lead to a more pronounced inactivation of NCX1.3 under conditions of elevated intracellular Na+ and Ca2+. This prediction follows from the observation that intracellular Ca2+ does not alleviate Na+-dependent inactivation for this splice variant. It is important to recognize that other possibilities could explain the difference in Ca2+ overload as a function of isoform type. For example, we cannot rule out the possibility that the isoforms may have different stoichiometry and kinetics for Ca2+ transport (26a), altered phosphorylation profiles that may affect function (7a, 13a, 23a), different sensitivities to intracellular pH, differential responses to membrane depolarization (23a, 26a), or subtle differences in membrane localization or insertion that may alter how these proteins induce Ca2+ movements into the cell. However, the same responses to the two isoforms were observed in two different cell types, suggesting that differences in membrane localization or insertion are not likely to be responsible.

In conclusion, our results are most compatible with the idea that ionic regulation of NCX can prominently modify the behavior of this transporter in its native cellular environment. This was demonstrated by exposing cells to conditions that elevated intracellular Na+ and Ca2+ levels. The NCX1.3 isoform is not present in cardiac tissue in vivo. Its placement in cardiomyocytes in the present proposal did allow us to directly compare the isoforms under identical cellular conditions. It remains to be determined whether these results can be directly extrapolated to native tissues in vivo that express NCX1.3, such as the kidney and vascular tissue. However, it is tempting to speculate that the properties of NCX1.3 might confer some resistance to ischemia-reperfusion injury in these tissues because of its unique pattern of ionic regulation. These findings may be relevant to future gene therapy and/or comparative physiology and pharmacology between smooth and cardiac muscles. The differences in function between these two isoforms may also explain the different roles of the exchanger in the two tissues during control and disease conditions.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
This work was supported by a grant from the Heart and Stroke Foundations of Manitoba and New Brunswick. C. Hurtado was a Trainee of the Heart and Stroke Foundation of Canada, N. Mesaeli was a New Investigator of the Heart and Stroke Foundation of Canada, and L. V. Hryshko holds a Canada Research Chair.


   FOOTNOTES
 

Address for reprint requests and other correspondence: G. N. Pierce, Div. of Stroke & Vascular Disease, St. Boniface Hospital Research Centre, 351 Tache Ave., Winnipeg, MB, Canada R2H 2A6

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

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