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Am J Physiol Heart Circ Physiol 290: H1879-H1888, 2006. First published December 9, 2005; doi:10.1152/ajpheart.01045.2005
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Differential autocrine modulation of atrial and ventricular potassium currents and of oxidative stress in diabetic rats

Yakhin Shimoni, Don Hunt, Keyun Chen, Teresa Emmett, and Gary Kargacin

Department of Physiology and Biophysics, Health Sciences Centre, University of Calgary, Alberta, Canada

Submitted 4 October 2005 ; accepted in final form 2 December 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The autocrine modulation of cardiac K+ currents was compared in ventricular and atrial cells (V and A cells, respectively) from Type 1 diabetic rats. K+ currents were measured by using whole cell voltage clamp. ANG II was measured by ELISA and immunofluorescent labeling. Oxidative stress was assessed by immunofluorescent labeling with dihydroethidium, a measure of superoxide ions. In V cells, K+ currents are attenuated after activation of the renin-angiotensin system (RAS) and the resulting ANG II-mediated oxidative stress. In striking contrast, these currents are not attenuated in A cells. Inhibition of the angiotensin-converting enzyme (ACE) also has no effect, in contrast to current augmentation in V cells. ANG II levels are enhanced in V, but not in A, cells. However, the high basal ANG II levels in A cells suggest that in these cells, ANG II-mediated pathways are suppressed, rather than ANG II formation. Concordantly, superoxide ion levels are lower in diabetic A than in V cells. Several findings indicate that high atrial natriuretic peptide (ANP) levels in A cells inhibit RAS activation. In male diabetic V cells, in vitro ANP (300 nM–1 µM, >5 h) decreases oxidative stress and augments K+ currents, but not when excess ANG II is present. ANP has no effect on ventricular K+ currents when the RAS is not activated, as in control males, in diabetic males treated with ACE inhibitor and in diabetic females. In conclusion, the modulation of K+ currents and oxidative stress is significantly different in A and V cells in diabetic rat hearts. The evidence suggests that this is largely due to inhibition of RAS activation and/or action by ANP in A cells. These results may underlie chamber-specific arrhythmogenic mechanisms.

diabetes; cardiac potassium currents

DIABETES MELLITUS is a growing epidemic (59). Despite improved treatment, cardiovascular complications develop, becoming the leading cause of diabetes-related death (24, 59). Diabetic cardiomyopathy impairs mechanical function, (3, 38) and electrical abnormalities increase the propensity for some cardiac arrhythmias (1). Cardiac action potentials are prolonged in animal models of diabetes due to attenuated transient and sustained repolarizing K+ currents (39, 56). A reduction in current magnitude reflects downregulation of K+ channel expression, as determined in earlier work by us and others (33, 42, 43). Diminished outward currents presumably underlie the prolongation of the QT interval in the human electrocardiogram. Prolongation and dispersion of the QT interval, established predictors of arrhythmias and mortality (23), are common in diabetes (50).

In contrast to numerous studies on diabetic ventricles, few studies have addressed atrial pathology. This is of importance, because the most common arrhythmia (9) is atrial fibrillation (AF). AF leads to changes in several atrial currents (49, 58). A transient K+ current is attenuated, although not in all conditions (9). K+ currents controlling atrial repolarization are mainly similar to those in ventricles (21, 55), although differences in channel expression levels lead to differences in repolarization patterns (28).

Diabetes (in humans and in rat models) activates a cardiac renin-angiotensin system (RAS) (10, 11). This elevates cardiac levels of ANG II, which acts by autocrine, paracrine, or intracrine mechanisms (2). ANG II, through numerous signaling pathways, plays a major role in cardioprotective and maladaptive mechanisms (7, 52). Chronically elevated ANG II attenuates K+ currents (57) and enhances oxidative stress (25), an important feature of diabetes (13, 14). The involvement of ANG II in K+ current attenuation was suggested both by direct measurements of ANG II elevation (10, 11, 44) and by the fact that inhibition of either the angiotensin-converting enzyme (ACE) or blockade of ANG II receptors leads to augmentation of attenuated K+ currents (35, 39) and associated channel proteins (42, 43). Furthermore, relief of ANG II-mediated oxidative stress also augments these currents (41, 56). The autocrine regulation of K+ currents and enhanced ANG II levels and oxidative stress in diabetes is sex dependent (40, 41, 43, 44). ANG II levels are elevated and K+ currents are attenuated to a significantly greater extent in males. This is presumably due to inhibition of the RAS by estrogen (12).

Very little is known about RAS activation and its potential modulation of K+ currents in atrial cells (A cells) under pathological conditions. A cells secrete a natriuretic peptide (ANP), which plays a key role in cardiovascular homeostasis (6, 19, 36, 53). Mice lacking ANP receptors develop hypertension and hypertrophy and exhibit ventricular arrhythmias and sudden death (18, 29). Cardiac-specific ANP receptor attenuation also compromises protection against adverse conditions such as pressure overload (30). Importantly, the protective role of ANP is linked to its ability to inhibit the RAS, thus countering detrimental effects of RAS activation (18, 30). Furthermore, the action of natriuretic peptides persists under diabetic conditions, when other protective mechanisms, such as the nitric oxide system, are compromised (53).

Based on earlier studies, we hypothesized that interaction of ANP and the RAS could result in different patterns of K+ current modulation in A cells and ventricular cells (V cells), as well as a potentially different degree of oxidative stress. Our objectives were 1) to establish whether atrial K+ currents are modified in diabetes, 2) to determine whether A cells exhibit RAS activation and elevated oxidative stress, 3) to investigate whether the RAS modulates atrial K+ currents, and 4) to investigate whether natriuretic peptides modulate K+ currents and oxidative stress under diabetic conditions and whether this relates to interaction with the RAS.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study conforms to the National Institutes of Health Guide for Care and Use of Laboratory Animals. All of the experiments were done according to guidelines established and approved by the Animal Care and Use Committee of the University of Calgary.

Animals. Age-matched control (untreated) and diabetic male Sprague-Dawley rats (250–300 g) were used. Type 1 diabetes was induced by intravenous injection of streptozotocin (100 mg/kg) 8–14 days before experiments. In one subset of experiments, diabetic female rats were also used.

Cell isolation was achieved by enzymatic dispersion. Rats were anesthetized by CO2 inhalation and euthanized by cervical dislocation. Hearts were perfused, after aortic cannulation, with a solution (at 37°C, bubbled with 100% O2) containing (in mM) 113 NaCl, 4.7 KCl, 1.2 MgSO4, 0.6 KH2PO4, 0.6 Na2HPO4, 12 NaHCO3, 12 KHCO3, 5.5 glucose, 10 HEPES, 30 taurine, and 10 2,3-butanedione monoxime (BDM). This was followed after 5–6 min by the same solution, also containing liberase blendzyme (0.02–0.25 mg/ml, Roche), trypsin (0.14 mg/ml), and 12.5 µM CaCl2. After 7–8 min, the free wall of the right ventricle or both atria were cut into pieces. After shaking was completed, tissue pieces were filtered and cell suspensions were collected and stored at room temperature in a solution containing no enzymes, 20 mM taurine, 5 mg/ml albumin, and 0.1 mM CaCl2.

Current recording. Cells were placed on a stage of an inverted microscope and perfused (at 21–22°C) with a solution containing (in mM) 150 NaCl, 5.4 KCl, 1 CaCl2, 1 MgCl2, 5 HEPES, and 5 glucose (brought to pH 7.4 with NaOH). L-type calcium current was blocked by 0.3 mM CdCl2. Currents were recorded by whole cell voltage clamp, using 500-ms pulses to membrane potentials ranging from –110 to +50 mV. Digitized currents (2 kHz) were normalized to cell size by dividing by cell capacitance. Recording pipettes (2–3 M{Omega} resistance) contained (in mM) 120 potassium aspartate; 30 KCl, 4 Na2ATP, 10 HEPES, 10 EGTA, 1 CaCl2, and 1 MgCl2 (pH 7.2 with KOH). Two currents were studied. The transient outward current (Ito), which determines early repolarization and the action potential plateau level, was measured (and is presented) both as the peak outward current and as the difference between peak current and the current at the end of 500-ms pulses.

The sustained outward current (Isus), measured at the end of 500-ms pulses, reflects a mixture of delayed rectifier currents (28) that determine late repolarization. Current densities were compared in the absence or presence of drugs. Results from different days were pooled. It should be noted that changes in current densities reflect changes in channel expression, which are measurable only after incubations of >5 h.

Superoxide production was detected by using dihydroethidium (DHE, Molecular Probes), a cell-permeable fluorescent dye that is oxidized by superoxide to ethidium bromide, a nucleic acid stain. Its fluorescent intensity indicates the relative level of superoxide production (14). Myocytes were suspended in a 800-µl solution containing (in mM) 120 NaCl, 5.4 KCl, 1.2 MgSO4, 1.2 Na H2PO4, 5.6 glucose, 20 NaHCO3, 10 BDM, and 5 taurine and 100 µM calcium and 0.2% fatty acid-free BSA. DHE (5 µM) was added to the cells, which were incubated in a light-protected incubator (30 min, 37°C). Suspensions were centrifuged (2 min, <1,000 g), and pellets were resuspended in 20–50 µl PBS. A small aliquot (10–15 µl) was fixed (with 90% glycerol) onto a slide.

DHE fluorescence was quantified by using an Olympus IX70 inverted epifluorescence microscope and a SPOT RT-cooled CCD camera. All images (1,600 x 1,200 pixels) were collected with the same camera settings. DHE fluorescence was analyzed with the use of two methods (41). First, a lower pixel intensity threshold was determined (using SPOT software) for images of individual nuclei, so that only light from the nucleus remained in the image. The mean fluorescence intensity of pixels above threshold for each nucleus was determined by using software designed for this purpose. In the second method, a boundary was drawn around each nucleus and the mean fluorescence intensity of pixels within the nuclear boundary (region of interest) was determined from an intensity histogram by using Photoshop software. Both methods of analysis gave similar results.

Cellular ANG II content was measured by ELISA, using a commercial kit (Peninsula). Standard curves were constructed, and optical densities of samples (in triplicates) were read from these curves (all values falling within the calibration curve). ANG II levels were normalized for protein content, measured in the same samples (using a bicinchoninic acid protein assay kit from Pierce). Values were similar to those reported by others (10).

ANG II immunofluorescence. ANG II is mainly localized in subcellular granules (10, 11). In our experiments, atria and ventricles were embedded in optimum cutting temperature compound (Electron Microscopy Sciences), frozen in 100% ETOH on dry ice for 30 min, and stored at –80°C. Frozen tissue was cut into 8-µm sections, using a cryostat (–20°C). Sections were mounted on coated slides (VWR) and stored at –80°C. For ANG II labeling, sections were air dried (room temperature) and fixed (1% formaldehyde). After being washed (3 times) with PBS, the tissue was permeabilized by using 1% Triton- X (10 min, room temperature). Nonspecific binding was blocked by using 2% BSA + 5% normal goat serum. A rabbit anti-ANG II antibody (Peninsula) was added (1:200 dilution in PBS + 2% BSA) and left overnight in a humid chamber. After tissues were washed in PBS, secondary antibody (anti-rabbit IgG conjugated to Alexa 488; Molecular Probes) was added (1:100 in PBS, 2% BSA). After 1 h in a humid chamber, the slides were washed in PBS. Finally, a mounting medium was added (Biomedia), and the slides were covered and stored at 4°C. As a control, ANG II (0.06 µg) was added to 1 µl of antibody. The preabsorption of ANG II antibody yielded unstained slides. These verified antibody specificity and were used to determine background fluorescence.

Quantification of ANG II was done as with DHE, using atrial and ventricular sections labeled with anti-ANG II. As noted above, background was determined from sections exposed to preabsorbed primary antibody, followed by secondary antibody. Multiple background images were collected for each preparation, and the background value used for the analysis was taken as the mean of the background light intensities determined from these images. All sections were labeled at the same time under the same conditions, and images were collected using the same camera settings. Background fluorescence was subtracted, and the number of pixels in each image with light intensities above background was determined. This value was used as one measure of ANG II levels. A second measure was obtained by computing the total above-background light intensity. This was done by summing the intensities of all of the pixels in the image that had values above background. Similar results were obtained from both methods of analysis. Several control and diabetic hearts were used, and a large number of images (>30) were collected and analyzed for each tissue type. As can be seen in one of the images shown in RESULTS (see Fig. 5A), some tissue sections did not occupy the entire field in an image. Inclusion of these images in the analysis could bias the results because there were fewer cells in the field. To avoid this possibility, the percentage of image area occupied by cardiac tissue was determined for each image and used to correct the fluorescence intensity values for that image.


Figure 5
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  		Fig. 5. Immunofluorescence images of atrial tissue showing ANG II in control (A) and diabetic (B) rats. C: summary of analysis of 34 sections from 3 control and 38 sections from 3 diabetic rats. Fluorescence intensities in sections from diabetic animals were normalized to fluorescence of sections from controls, prepared at the same time. Analysis was done after subtraction of background fluorescence and correction for image area occupied by tissue in each image as described in METHODS and Fig. 4 legend. Fluorescence intensities in images from diabetic sections were not significantly different from controls (P > 0.9). Bar in B = 20 µm.

 
To test the validity of the general method of analysis used to compare DHE fluorescence or anti-ANG II immunofluorescence in different cell or tissue samples, fluorescent microspheres were suspended in solution and different dilutions of the spheres were pipetted onto microscope slides. Images of the suspensions in randomly chosen fields on the slides were then collected, and the fluorescence in the images was analyzed by determining both the total number of pixels above background in the images and the total fluorescence intensity of all the pixels that were above background. Figure 1 summarizes the results of this analysis for six experiments. In five of the experiments, the microspheres were diluted in water and backgrounds that eliminated out-of-focus light were subtracted from the images. In the sixth experiment, the spheres were diluted in water and then added to suspensions of unlabeled isolated cells that provide a background of autofluorescence that was then subtracted from the images. Results from the two types of experiments are combined in Fig. 1. The results shown indicate that, although there is a slight deviation from linearity for the least concentrated samples, the method of analysis was able to predict relative changes in concentration quite well. The deviation of the results from the expected relative fluorescence at the higher dilutions is likely due to the greater effect of variance in the background light level on the measurements made when the number of beads in the fields was low.


Figure 1
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  		Fig. 1. Analysis of images of diluted fluorescent microspheres. Spheres were diluted as described in METHODS. In each experiment, 6–9 image fields were randomly collected for each dilution, and fluorescence from image fields was expressed relative to mean fluorescence from the most concentrated sample. Two measures of fluorescence were used. A: total number of pixels in an image with light intensities above a threshold intensity (determined as described in METHODS). B: total intensity of all pixels above threshold intensity. Results are from 6 independent experiments. Solid lines in A and B indicate relative fluorescence expected from known dilutions of samples. *P < 0.05, significant differences from fluorescence of starting dilution.

 
Drugs. Quinapril-HCl was prepared as stock solution in 0.5% ethanol and diluted to the final concentration (1 µM for in vitro experiments and 6 mg/l for in vivo experiments). Final ethanol concentrations were <0.005%, and the pH of the solutions was 7.4.

Statistics. Results are given as means ± SE. Mean values were compared by using t-test or ANOVA, with the Student-Newman-Keuls multiple comparisons test. P values of <0.05 were considered significant.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
The first set of experiments compared peak and sustained outward currents in A cells from control and diabetic rats. In contrast to V cells (39), there was no significant current attenuation in diabetic A cells. Figure 2A shows current traces in atrial myocytes from control (Fig. 2A, left) and diabetic (Fig. 2A, right) rats. Figure 2B shows current-voltage relationships for Ito (Fig. 2B, left) and Isus (Fig. 2B, right) in control (n = 14) and diabetic (n = 27) cells (3 rats from each group).


Figure 2
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  		Fig. 2. Outward K+ currents in atrial cells. A: current traces obtained in response to pulses given from –80 mV to potentials ranging from –10 to +50 mV in atrial cells (A cells) from a control (left) and a diabetic (right) rat (same protocol used in subsequent figures). STZ, strepozotocin. B: mean current densities as a function of membrane potential [transient (Ito; left) and sustained (Isus; right) outward current] in cells from control (bullet) and diabetic ({blacksquare}) rats. C: difference currents (Idiff = Ito – Isus) are also not significantly different.

 
In addition to lack of current attenuation in A cells, an even more striking finding was a difference in current modulation. Diabetes leads to elevated ANG II levels in V cells, which contribute to current attenuation (39, 44). Incubation of V cells with the ACE inhibitor quinapril (>5 h) augments both Ito and Isus (39). In contrast, quinapril had no effect on either Ito or Isus in A cells. This is shown in Fig. 3A, which illustrates mean Ito (Fig. 3A, left) and Isus (Fig. 3A, middle) densities (at +50 mV) in the absence (36 cells, 5 rats) or presence (26 cells, 5 rats) of quinapril (1 µM, >5 h). For comparison, Fig. 3B shows the effects of this protocol in V cells.


Figure 3
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  		Fig. 3. Effects of the angiotensin-converting enzyme (ACE) inhibitor quinapril. A: in A cells, quinapril (hatched bars) has no significant effect (1 µM, 5–9 h) on Ito (left), Isus (middle), or Idiff (right). B: in ventricular cells (V cells), quinapril (same dose and duration) significantly augments Ito, Isus, and Idiff (*P < 0.005).

 
These results suggest that the RAS is less (or not) activated in diabetic A cells or that the effects of ANG II are suppressed. Our laboratory (44) previously measured ANG II levels in V cells, using ELISA. However, this was not feasible with A cells, because enzymatic dispersion yields considerably fewer cells. We therefore used immunofluorescent ANG II labeling of ventricular and atrial sections (as in Ref. 11). Concordant with our earlier results and with Ref. 11, we confirmed that ANG II labeling is significantly increased (see METHODS) in diabetic V cells, compared with control (4 rats, >30 sections in each group). This is shown in Fig. 4, which also shows the lack of labeling when the ANG II antibody was preabsorbed with ANG II (Fig. 4C).


Figure 4
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  		Fig. 4. Differential interference contrast (A) and immunofluorescence (B) images of same tissue section showing distribution of anti-ANG II labeling in rat ventricular myocytes. Immunofluorescence images of ventricular tissue sections comparing anti-ANG II labeling in control (C) and diabetic rat (D) heart. E: image of ventricular section after preabsorption of anti-ANG II with ANG II. F: summary results from analysis of 39 sections from 3 control and 40 sections from 3 diabetic animals. Results show fluorescence intensity in images of sections from diabetic and nondiabetic rat hearts plotted as percentage of fluorescence intensity in images of sections from nondiabetic animals. Fluorescence intensities in tissue sections were determined after background fluorescence was subtracted (determined from sections, as in E). For this experiment, 13 background images were collected and intensity of the background light was determined for each image. Mean light intensity in background images was used for the analysis (background fluorescence varied by ~5% in 13 images). Images were also corrected for fractional area occupied by tissue as discussed in METHODS. Fluorescence intensities in images from diabetic sections were significantly different from control (P ≤ 10–4). Similar results were obtained when total number of pixels with intensities above background was determined for images. From latter analysis, number of above-background pixels in tissue sections from diabetic animals was 623 ± 122% of number of above-background pixels in sections from nondiabetic animals (P ≤ 10–4). Bars in B, D, and E = 20 µm.

 
Based on the electrophysiological results with A cells (above), we hypothesized that no differences in ANG II would be found between control and diabetic A cells. The results with atrial sections confirmed this, although, surprisingly, ANG II labeling was high in both control and diabetic A cells, as shown in Fig. 5.

These results highlight a major difference between diabetic V and A cells in that diabetes elevates ANG II levels only in ventricles. If current attenuation depends on increased levels of ANG II, the absence of ANG II changes presumably underlies the lack of attenuation of Ito and Isus magnitude in diabetic A cells. Furthermore, high ANG II levels in control A cells may underlie the smaller baseline currents, compared with V cells. In V cells, ACE inhibition presumably suppresses ANG II levels within 5 h, because K+ currents are augmented after incubation with quinapril for 5–9 h. It is possible that with high baseline ANG II levels in A cells, ACE inhibition might not reduce ANG II levels sufficiently to enable augmentation of K+ currents (Fig. 3).

We subsequently tested whether atrial-ventricular differences result from a suppression of ANG II effects in A cells, rather than from differences in ANG II levels per se.

ANG II induces oxidative stress, mainly through the activation of NADPH oxidase and the generation of superoxide ions (25). These can be measured by using fluorescent DHE. DHE interacts with superoxide, binding to nuclear DNA proportionately to superoxide levels (14). Superoxide ion levels were compared in A and V cells, as a measure of ANG II-induced oxidative stress. DHE fluorescence was significantly higher in V cells from diabetic (n = 3) compared with control (n = 3) rats. This confirms the presence of oxidative stress, a prominent feature of diabetes (13, 14). Figure 6, A–C, shows V cells from a control (Fig. 6A) and a diabetic (Fig. 6B) rat, as well as mean fluorescence intensity (Fig. 6C) in one paired comparison. We subsequently isolated V and A cells from diabetic rats and compared DHE fluorescence in cells from the same hearts. In five out of seven rats, A cells had significantly (P < 0.005) or very significantly (P < 0.0001) lower DHE fluorescence than V cells, indicating lower atrial oxidative stress. Figure 6, D–F, shows sample cells and the mean fluorescence intensity from one of these comparisons.


Figure 6
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  		Fig. 6. Ventricular myocytes labeled with dihydroethidium (DHE), a marker of superoxide ions. Control (A) and diabetic (B) cells are shown. C: mean fluorescence intensities, measured in 18 individual nuclei from control cells and 19 nuclei from diabetic cells, indicate a significant increase in diabetic conditions (**P ≤ 0.001). AU, arbitrary units. DHE labeling in V (D) and A (E) cells were obtained from the same diabetic rat. F: mean fluorescence in 40 individual ventricular and 41 individual atrial nuclei. Atrial nuclei show significantly (***P < 0.005) lower DHE labeling, indicating lower superoxide ion levels and lower oxidative stress. Bars in B and E = 20 µm.

 
These experiments show that A cells are partially protected from the development of oxidative stress that occurs in V cells. This may underlie the absence of K+ current attenuation in these cells. The lack of sensitivity of atrial K+ currents to quinapril and a lower oxidative stress suggest that the RAS is either less activated in diabetic A cells and/or that its effector pathways are inhibited.

Earlier work suggested that the RAS can be inhibited by the ANP. We hypothesized that if the more abundant ANP in A cells (6, 19, 53) inhibits effects of the RAS, then the addition of exogenous ANP may reverse some of the effects of activated RAS in V cells. We therefore compared the effects of ANP on ventricular K+ currents in control and diabetic cells. Earlier reports suggested dose-dependent actions of ANP, with maximal effects at 1 µM (54). We used ANP at concentrations ranging from 100 nM to 1 µM.

In V cells from diabetic (male) rats, in which the RAS activity and ANG II levels are elevated (10, 44), 300 nM ANP (5–9 h) significantly augmented both Ito and Isus, as shown in Fig. 7. Ito densities (at +50 mV) were 13.3 ± 1.1 (32 cells, 3 rats) and 18.4 ± 1.25 (33 cells, 3 rats) pA/pF (P < 0.004) in the absence or presence of ANP, respectively. The corresponding values for Isus were 4.1 ± 0.1 and 5.1 ± 0.2 pA/pF (P < 0.00001). An addition of 1 µM ANP produced similarly significant augmentation of both currents (5 rats, 51 untreated cells and 42 treated cells). In a small group of cells, 100 nM ANP significantly augmented Isus. Ito was also augmented but not significantly in the smaller sample of cells.


Figure 7
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  		Fig. 7. Effects of atrial natriuretic peptide (ANP) on diabetic V cells. A: current traces were obtained in absence (left) or presence (right) of ANP (300 nM, 6 h) in cells from diabetic males. B: mean densities (at +50 mV) of Ito (left) and Isus (middle) in absence (open bars) or after 300 nM ANP (hatched bars), showing significant augmentation of both currents (*P < 0.005 and **P < 0.0005) in these cells. C: ANP (1 µM) did not significantly alter mean Ito and Isus densities (at +50 mV) in cells from diabetic females.

 
In V cells from control rats, in which the RAS is not activated, even the higher concentration of ANP was without effect. The mean densities (at +50 mV) of Ito in the absence and presence of ANP (1 µM, 5–9 h) were 24.7 ± 2.1 (21 cells, 3 rats) and 25.1 ± 1.0 (19 cells, 3 rats) pA/pF, respectively (P > 0.05). The corresponding values for Isus were 7.3 ± 0.3 and 7.5 ± 0.6 pA/pF (P > 0.05) (results not shown).

ANP also had no effects in V cells from diabetic females, in which ANG II levels are not augmented (44), indicating that RAS preactivation is required for ANP action. This result is also shown in Fig. 7.

In diabetic A cells, the RAS is apparently not activated above control (Fig. 5), and/or effects of ANG II may be blocked. Concordantly, ANP was without effect. Mean Ito densities (at +50 mV) in the absence and presence of ANP (1 µM, 5–9 h) were 12.4 ± 1.3 (27 cells, 5 rats) and 13.3 ± 1.4 (24 cells, 5 rats) pA/pF, respectively (P > 0.05). The corresponding values for Isus were 5.9 ± 0.3 and 6.2 ± 0.4 pA/pF (P > 0.05) (not shown).

ANP has been shown to protect against oxidative damage (31). We thus investigated whether ANP inhibits induction of oxidative stress by ANG II. Cells were prepared from three diabetic rats and divided into two groups. One was incubated in ANP (1 µM for 5h), and the other was left untreated. Subsequent labeling of cells from both groups showed that in all three pairs, ANP caused a significant attenuation in DHE fluorescence, indicating reduced levels of superoxide ions. One example is shown in Fig. 8.


Figure 8
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  		Fig. 8. ANP inhibits ANG II-induced oxidative stress. DHE-labeled diabetic V cells in absence (A) or presence (B) of ANP (1 µM, 5 h). C: mean fluorescence intensity of 57 individual nuclei from untreated cells and 36 nuclei from ANP-treated cells. Mean fluorescence was significantly (*P < 10–4) reduced by ANP, indicating reduced superoxide levels. Similar results were obtained in two additional experiments. Bar in B = 10 µm.

 
Our results suggest that preactivation of the RAS is required for augmentation of Ito and Isus by ANP, which acts by inhibiting this system. ANP effects were tested under conditions in which the RAS is either not activated or is directly inhibited. First, male diabetic rats were given the ACE inhibitor quinapril in vivo for 3 wk (6 mg/l in the drinking water) before induction of diabetes. ANG II content in V cells from diabetic rats, without (n = 4) or after quinapril treatment (n = 4), was 47.1 ± 9.2 and 3.6 ± 0.5 pg/mg protein, respectively (P < 0.005), as shown in Fig. 9A.


Figure 9
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  		Fig. 9. Effects of quinapril treatment. A: ANG II content (measured by ELISA) in diabetic male V cells in absence (open bar) or presence (hatched bar) of quinapril (6 mg/l for 3 wk) in drinking water before STZ injection (continued until cell isolation). After quinapril treatment, ANP no longer augments outward currents. B: current traces in absence (left) or presence (right) of ANP (1 µM, 9 h). C: mean Ito (left), Isus (middle), and Idiff (right) densities (at +50 mV) in absence or presence of ANP in cells from quinapril-treated rats.

 
In V cells from these quinapril-treated diabetic rats, Ito and Isus are still lower than in the control, despite the reduction in ANG II levels (see DISCUSSION). However, in these cells, ANP (1 µM) no longer augments Ito and Isus, as shown in Fig. 9, B and C. Mean Ito (at +50 mV) was 14.8 ± 1.0 (37 cells) and 15.2 ± 0.8 (39 cells) pA/pF in the absence or presence of ANP, respectively (P > 0.05). Corresponding Isus values were 4.6 ± 0.2 and 4.9 ± 0.2 pA/pF (P > 0.05).

Finally, V cells from three diabetic male rats were exposed to ANP in the presence of excess (300 nM) ANG II. As shown in Fig. 10, the augmentation of Ito and Isus by ANP was significantly reduced, suggesting that excess ANG II counters the inhibitory effect of ANP on the RAS.


Figure 10
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  		Fig. 10. ANG II suppresses ANP effects. A: currents obtained in male diabetic V cells (3 rats) with no ANP (left, n = 18), after ANP (1 µM, 6.5 h, n = 15), and with ANP and 300 nM ANG II (n = 15), added 30 min before ANP (8 h of ANP). B: mean Ito (left), Isus (middle), and Idiff (right) densities (at +50 mV) show significant (*P < 0.05) augmentation of both currents by ANP, as well as significant suppression of this effect by ANG II.

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
In summary, the present work provides several novel results. The major finding is that cardiac autocrine mechanisms, modulating K+ currents and oxidative stress, are strikingly different in V and A cells. In contrast to the situation in V cells, atrial K+ currents are not attenuated after the onset of Type 1 diabetes (Fig. 1). In addition, atrial K+ currents are insensitive to the ACE inhibitor quinapril and to ANP, in contrast to V cells (Figs. 2 and 6). A further novel and important finding is that oxidative stress is lower in A cells than in V cells from the same diabetic hearts (Fig. 5). ANP may be a major contributor to the lower oxidative stress in A cells, as suggested by the reversal of augmented oxidative stress in diabetic V cells by ANP (Fig. 7). Finally, diabetic conditions augment ANG II in V, but not A, cells (Figs. 3 and 4).

Interpretation. Our hypothesis was that the activation of the RAS and ensuing ANG-mediated effects would be modified by ANP. Basal levels of ANP are higher in A cells (6) and presumably prevent activation of the RAS (Fig. 4) that occurs in V cells (Fig. 3) after onset of diabetic conditions (11, 44). Lack of RAS activation presumably underlies the lack of K+ current attenuation in A cells. However, ANG II levels are unexpectedly high in control A cells (Fig. 4), suggesting that ANP also inhibits ANG II signaling.

It is not yet known how ANP inhibits the RAS. ANP may inhibit both formation and/or secretion of ANG II, as well as its subsequent actions (19). ANP can inhibit p38MAPK and NF-{kappa}B (45), which mediate some ANG II actions (5).

ANG II signaling is highly complex (7) and involves multiple pathways. The interaction with ANP may occur at the level of cGMP. cGMP, which is the major second messenger that is elevated by ANP (6, 19, 53), is increasingly recognized as a modulator of gene expression (19, 32) and could therefore be a positive modulator of some K+ channels. ANG II inhibits cGMP signaling (17), which could be one of the mechanisms leading to current attenuation. The presence of ANP may counteract this action of ANG II, as well as ANG II-mediated oxidative stress.

Several lines of evidence support this. ANP augments K+ currents and suppresses oxidative stress in V cells from diabetic males (Figs. 6 and 7), but not if the RAS is inhibited before the onset of diabetes by quinapril (Fig. 8) or if the RAS is not activated, as in control males or in diabetic females (44). Furthermore, excess ANG II suppresses current augmentation by ANP (Fig. 10), suggesting a reciprocal inhibition between ANG II and ANP (17). This complexity is supported by reports showing inhibition of some ANP actions by ANG II (34).

Further work is required to elucidate the signaling pathways involved, to establish which channel isoforms are altered in V and A cells, and to determine whether there is a selective sensitivity to ANP of these isoforms.

Concordant with our study, other pathologies also exhibit similar atrial-ventricular differences. For example, in hyperthyroid dogs, ANG II receptor densities increase in ventricles but not in atria (37). In a rat model of cirrhosis, ventricular but not atrial K+ currents are attenuated, although mechanisms underlying this difference are unknown (51). Several cardiac pathologies lead to elevation in natriuretic peptides in both ventricles and atria (6). However, transcriptional control of peptide expression in atria and ventricles is different, because this expression is suppressed by ACE inhibition in ventricles but not atria (6). This suggests a different linkage between the RAS and ANP pathways in the two cell types. This is supported by our findings (Fig. 4) that diabetes leads to increased ANG II levels in V but not A cells.

In contrast to diabetic conditions, atrial K+ currents are modulated under other pathophysiological conditions, including metabolic stress, heart failure, or AF (49). Oxidative stress during AF is associated with augmented superoxide production (8), as well as current attenuation (48, 49), although oxidative stress may occur after abnormal (mechanical) function, rather than being a causative agent. Interestingly, ventricular tachypacing and AF are also associated with augmented RAS activity (15), with elevated ANG II levels, presumably underlying oxidative stress (8). ACE inhibitors and ANG II receptor blockers are reportedly of benefit for AF (16, 45), although this may be limited to subsets of patients (16). In paced canine hearts, ANG II levels increase more in atria than in ventricles (15). This suggests that different pathologies activate different mechanisms and/or that significant species differences are present.

The complexity of regulatory mechanisms may also involve temporal changes in the involvement of the RAS. Shinagawa et al. (45) found short- but not long-term protective effects of ACE inhibition in AF. This may be linked to similar time-limited effects of ANP (47).

In diabetes, plasma ANP levels are elevated (22), but the expression of a closely related (brain natriuretic) peptide is chamber dependent, increasing only in atria (4). Interestingly, regulation of ANP actions may be defective in diabetes (27), although other reports indicate the persistence of protective action (53). Our results suggest that, at least in diabetes, high natriuretic peptide levels in A cells prevent further increase of ANG II and limit oxidative stress, presumably by inhibiting RAS activation. This results in unaltered atrial K+ currents that are insensitive to ACE inhibition.

Limitations. ANP was not directly measured in this study. However, numerous reports suggest that ANP is found predominantly in A cells (6). Ventricular ANP can increase in some pathological conditions (6). Our results suggest that if this occurs in diabetic conditions, the magnitude is small and insufficient to prevent activation of the RAS and increases in ANG II (Fig. 3 and Refs. 10, 11, and 44). Furthermore, exogenous ANP augments attenuated ventricular (Fig. 6) but not atrial currents. The differences in oxidative stress (Fig. 5), largely ANG II dependent (13, 14, 25), also suggest lower ANP levels in V cells. Non-ANG II-related causes of oxidative stress (13) may explain why atrial and ventricular differences were not found in all cases.

Another limitation of this study lies in the fact that right and left atria were pooled. Several studies suggest differences between right and left atria, in terms of oxidative stress (8) or in some ionic currents (20). However, the transient outward current and most delayed rectifier currents were not different in the right and left (canine) atria (20). It is, however, possible that currents in right and left diabetic rat atria may have shown differences that we could not detect in these experiments.

Finally, we cannot at present distinguish between ANG II that is located in subcellular organelles and ANG II that has been released. These two pools will obviously have different functional significance.

Significance of study. Our results suggest possible differences in the generation of arrhythmias in ventricles and atria. The results also highlight functional aspects of the interaction between the RAS and natriuretic peptides, which are increasingly recognized as major participants in adaptive and maladaptive processes involved in cardiac cellular homeostasis.


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 ABSTRACT
 METHODS
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This study was supported by grants from the Canadian Institutes of Health Research (to Y. Shimoni and G. Kargacin) and from the Heart and Stroke Foundation of Alberta (to Y. Shimoni and G. Kargacin)


   FOOTNOTES
 

Address for reprint requests and other correspondence: Y. Shimoni, Health Sciences Centre, 3330 Hospital Dr. NW, Calgary, AB, Canada T2N 4N1 (e-mail: shimoni{at}ucalgary.ca)

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.


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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
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A. Nygren, M. L. Olson, K. Y. Chen, T. Emmett, G. Kargacin, and Y. Shimoni
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