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Slow diffusion of K⁺ in the T tubules of rat cardiomyocytes

¹Institute for Experimental Medical Research, Ullevaal University Hospital; ²Center for Heart Failure Research, University of Oslo; and ³Department of Cardiology, Ullevaal University Hospital, Oslo, Norway

Submitted 10 March 2006 ; accepted in final form 5 June 2006

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS ACKNOWLEDGMENTS REFERENCES

 
Cardiomyocyte contractility is regulated by the extracellular K⁺ concentration ([K⁺]_o). Potassium dynamics in the T tubules during the excitation-contraction cycle depends on the diffusion rate of K⁺, but this rate is not known. Detubulation of rat cardiomyocytes was induced by osmotic shock using formamide, which separated the surface membrane from the T tubules. Changes in current and membrane potential in voltage-clamped (–80 mV) and current-clamped control and detubulated cardiomyocytes were compared during rapid switches between 5.4 and 8.1 mM [K⁺]_o, and the results were simulated in a mathematical model. In the voltage-clamp experiments, the current changed significantly slower in control than in detubulated cardiomyocytes during the switch from 5.4 to 8.1 mM [K⁺]_o, as indicated by the times to achieve 25, 50, 90, and 95% of the new steady-state current [control (ms) t₂₅ = 98 ± 12, t₅₀ = 206 ± 20, t₉₀ = 570 ± 72, t₉₅ = 666 ± 92; detubulated t₂₅ = 61 ± 11, t₅₀ = 142 ± 17, t₉₀ = 352 ± 52, t₉₅ = 420 ± 69]. These time points were not significantly different either during the 8.1 to 5.4 mM [K⁺]_o switch or in current-clamped cardiomyocytes switching from 5.4 to 8.1 mM [K⁺]_o. Mathematical simulation of the difference current between control and detubulated cardiomyocytes gave a t-tubular diffusion rate for K⁺ of 85 µm²/s. We conclude that the diffusion of K⁺ in the T tubules is so slow that they constitute a functional compartment. This might play a key role in local regulation of the action potential, and thus in the regulation of cardiomyocyte contractility.

mathematical model; detubulation; rapid perfusion; electrophysiology

Although the diameters of the T tubules vary between 20 and 450 nm in cardiac myocytes, more than half have a diameter of 180–280 nm (18, 19). Despite the rather large diameter, the diffusion of Ca²⁺ in the T tubules has been reported to be slow compared with diffusion in bulk solution at the surface of the cardiomyocyte (1, 17). Moreover, Yao et al. (21) suggested that there is a restricted space on the surface of the cell membrane that limits the diffusion of cations. However, these studies did not investigate t-tubular diffusion rate for K⁺.

According to the Einstein diffusion relation (6), the diffusion of solutes depends on the temperature. However, an increase by 3°C would only increase the diffusion by 1%, which is not significant in our context. More importantly, the activity of several membrane proteins is substantially more affected by temperature variation. For example, the Na⁺-K⁺-ATPase has a temperature coefficient (Q10) of 2.1 (15). Therefore it is important to evaluate the diffusion rate of K⁺ in the T tubules at physiological and stable temperatures. For this reason we used a solution switcher system that keeps a constant temperature around the cell during the experiment.

In this study we determine the rate of diffusion of K⁺ in the T tubules by comparing the rate of change in holding current in voltage-clamped control and detubulated myocytes during a switch between two solutions with different K⁺ concentration. Shepherd and McDonough (17) observed that the depolarization of a cardiomyocyte due to a change in extracellular K⁺ occurred faster than the concentration change itself, owing to electrotonic spread of depolarization. Therefore we also measured the rate of change in membrane potential in current-clamped cardiomyocytes. A mathematical model was written in Matlab to simulate the experimental results.

	MATERIALS AND METHODS

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Animal and cardiomyocyte preparation.   Animals were cared for according to the Norwegian Animal Welfare Act, which conforms to the National Institutes of Health guidelines (NIH publication no. 85-23, revised 1996). Two animals were kept in each cage and housed in a temperature regulated room with a 12:12-h day-night cycling and had access to food and water ad libitum. Male Wistar rats (Møllegaard Breeding and Research Center, Skensved, Denmark) weighing 300 g were anesthetized with 68% N₂O, 29% O₂, and 2–3% isoflurane and ventilated on a respirator (Zoovent, Triumph Technical Services, Milton Keynes, UK). After a thoracotomy, heparin (170 IE in 0.2 ml, Leo, Buckinghamshire, UK) was administrated intravenously before excision of the heart. Single cardiomyocytes were isolated enzymatically as previously described (14).

Detubulation.   Single cardiomyocytes were detubulated using a hypo-osmotic shock as described by Kawai et al. (8). The detubulation procedure was evaluated on a Axiovert 100 LSM-510 confocal microscope (Zeiss). For this purpose, cardiomyocytes were gently spun down and resuspended in solution A containing (in mM) NaCl 140, HEPES 5, KCl 5.4, CaCl₂ 1, MgCl₂ 0.5, D-glucose 5.5, and NaH₂PO₄ 0.4 (pH 7.40) supplemented with 5 µM di-8-ANEPPS (Molecular Probes, Eugene, OR) for 10–20 min. Di-8-ANEPPS was kept as a stock solution (10 mM) in DMSO (D-5879, Sigma) plus 20% Pluronic acid (Pluronic F-127, P-6867, Molecular Probes). The cardiomyocytes were washed three times in solution A before imaging. The dye was excited with an argon laser at 488 nm, and emission was collected at 505–550 nm. Detubulation was also verified for each cardiomyocyte included in the analysis by comparing calculated cell volume to cell capacitance in both groups.

Rapid solution switcher.   The superfusion solutions were applied by a rapid switcher system (Fig. 1). Solutions flowed by gravity through disposable infusion sets with drop chamber and roller clamp to adjust flow. The tubes of the infusion sets were connected to silicone tubes with outer diameter (OD) 2.1 mm and inner diameter (ID) 1.15 mm that could be clamped by magnetic valves. The magnetic valves were controlled with a custom-made transistor-transistor logic-controller. The system allowed a rapid switch between five different solutions (but only two solutions were used in this study). The five tubes passed through a silicon tube (OD 7.5 mm, ID 6 mm) that countercurrent circulated prewarmed water to accurately control solution temperature. The perfusion tip was made of five stainless steel pipes (OD 1.3 mm, ID 0.6 mm) soldered together with silver alloy like a revolver and inserted into a pipe (OD 6 mm, ID 5 mm). The solution from each 1.3-mm pipe was collected in a mixing chamber at the tip of the 6-mm pipe and superfused on the cardiomyocyte. The perfusion system used in this study is quite similar to that of Levi et al. (11), but because of its size the device does not require much space and will fit even in small working spaces.

View larger version (23K): [in this window] [in a new window]   Fig. 1. The perfusion system. The solutions flowed by gravity from the solution reservoir through silicone tubes to the perfusion tip. The flow was controlled by a computer and a TTL-controller by clamping the tube with a magnetic valve. In the perfusion tip, prewarmed water was circulated countercurrent in silicon tubes (large arrows). Bottom right: schematic representation of the perfusion tip and the experimental chamber. Cardiomyocytes were plated on a glass coverslip that was placed in a holder made of polyoxymethylene and used as the floor of the experimental chamber. Suction was applied to the outlet of the experimental chamber to remove the solution during the experiment.

The time necessary to switch between two solutions was determined as previously described by Spitzer and Bridge (20) by measuring changes in liquid junction potential between a glass microelectrode (3–5 M) and the solutions flowing from the perfusion tip. The microelectrode was filled with 3 mM NaCl and was superfused with two solutions, X and Y, containing 147 mM Na-glucuronate with 3 mM NaCl and 100 mM Na-glucuronate with 50 mM NaCl, respectively.

Electrophysiological measurements.   The cardiomyocytes were patched by use of suction pipettes made from thin-walled standard borosilicate glass TW150F-4 capillary (World Precision Instruments, Sarasota, FL). The pipettes had a resistance of 2.3–4.0 M when filled with an internal solution containing (in mM) K-aspartate 120, KCl 25, MgCl₂ 0.5, NaCl 6, K₂-ATP 4, EGTA 0.06, HEPES 10 and D-glucose 10 (pH 7.20). The cardiomyocytes were superfused with a control solution containing (in mM) NaCl 140, HEPES 5, KCl 5.4, CaCl₂ 1.8, MgCl₂ 0.5, D-glucose 5.5, NaH₂PO₄ 0.4 (pH 7.40). During the test step, the cardiomyocytes were superfused with the control solution containing 8.1 mM KCl. Whole-cell voltage and current-clamp experiments were performed using an Axoclamp 200B amplifier (Axon Instruments, Union City, CA). The cardiomyocytes were voltage clamped at –80 mV to record the transmembrane current or current clamped to record the membrane potential during solution switch. Because of the relatively slow changes in membrane potential, the use of a conventional patch-clamp amplifier in these experiments was not a limitation. Voltage and current clamp protocols were written in pClamp 8.0 software (Axon Instruments). The recordings were digitized with a Digidata 1200B and analyzed with pClamp 9.0 software. The kinetics of the change in current or membrane potential induced by a rapid switch in [K⁺]_o was quantified as the time to reach 25, 50, 90, and 95% of the new steady-state current. Cell capacitance was calculated by integrating the capacitive currents elicited by hyperpolarizing steps (10 mV). Cell volume (v) was calculated using v = (lwd) (3), where l and w are the measured length and width. The cell depth (d) was assumed to be one-third of the cell width (3).

Temperature measurements.   Temperature stability was measured with a J-type thermocouple made of constantan and iron (Teck Instruments AS, Tranby, Norway). It was placed in the chamber 200 µm from the perfusion tip, where a cardiomyocyte would be during an experiment, and connected to a reference thermocouple placed in a temperature regulated bath (37.0 ± 0.1°C). The voltage differences between the measuring and the reference thermocouple were amplified by a custom-made direct-current amplifier and recorded digitally through the Digidata 1200B.

Mathematical model.   The T tubule was considered a cylinder with length L, divided into N compartments (Fig. 2). The length of each compartment was L = L/N. In each compartment, the change in [K⁺] (C/t) was affected by the diffusion to or from adjacent compartments and was determined by

where J_N is the rate of K⁺ influx into compartment N, D is the diffusion constant in the T tubule, A_N is the cross-sectional area of the T tubule, C_N is the [K⁺] in compartment N (C₀ is the initial change in [K⁺] or stimulus), V_N is the volume of compartment N, and t is time. To mimic the initial change in [K⁺] at the mouth of the T tubules (C₀), we used the junction potential recordings scaled to concentration values. The length of the T tubule was set to L = 10 µm; the radius of the T tubule was residual because it was divided by V_N but was set numerically to 1 in the model. Thus the model was not dependent on T tubule radius.

View larger version (19K): [in this window] [in a new window]   Fig. 2. Simulation of K+ diffusion in the T tubules. A: graphic representation of the T tubule used for simulations. The T tubule was considered a cylinder with length L = 10 µm, consisting of N = 50 compartments. The diffusion of K+ from one compartment to the next is represented by J. B: expected changes in holding currents at various values of diffusion rate (D).

Statistics.   Values are expressed as means ± SE. Comparisons between means were made by Student's t-test (homoscedastic, two-tailed distribution). Differences between sample means were considered significant when P < 0.05.

	RESULTS

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Rapid solution switcher.

In this study we evaluate the diffusion rate of solutes in the T tubules by measuring changes in the transmembrane current-membrane potential in whole cell voltage/current clamp during rapid switch between two solutions. To limit stretch activation of currents and avoid cardiomyocyte displacement during the experiment, it was important that the flow of solution was equal in both channels. In these experiments, the flow was set to 1 drop/s in the drop chamber. Ten drops passed during 10.2 ± 0.3 and 10.0 ± 0.4 s in the two channels, giving a flow of 2.67 ± 0.07 and 2.75 ± 0.12 ml/min, respectively, both n = 10 [not significant (NS)]. When switching between solutions, there was a delay from the switch command by the computer until the new solution reached the cardiomyocyte. This delay corresponds to the time to activate the magnetic valves and the time to exchange the solution in the mixing chamber of the perfusion tip. To determine this delay, the measured liquid junction potentials from 26 sweeps from two separate experiments were averaged. We observed transient changes in the initial phase of the junction potential recording that probably were due to turbulent flow in the pipette tip. These transient changes were not observed in the current recordings. Thus the change in junction potential started 311 ms (switch from solution X to Y) and 326 ms (Y X) after activation of the magnetic valves and changed by 19 mV with a time constant -63 ms. Starting from these time points, we calculated the time for the junction potential to reach 25, 50, 90, and 95% of the maximal response. During switch from solution X to solution Y, t₂₅ = 50 ms, t₅₀ = 72 ms, t₉₀ = 143 ms, and t₉₅ = 169 ms. When switching from solution Y to solution X, t₂₅ = 46 ms, t₅₀ = 70 ms, t₉₀ = 160 ms, and t₉₅ = 218 ms. Thus the time course of the solution switch was similar in the two perfusion lines.

Temperature stability was measured during a switch in flow between the two lines. Figure 3 shows a typical temperature recording. At a flow of 2.7 ± 0.1 ml/min (n = 20), the mean temperature in 22 experiments was 36.9 ± 0.2°C during flow from line 1 and 36.9 ± 0.2°C during flow from line 2 (NS).

View larger version (6K): [in this window] [in a new window]   Fig. 3. Temperature stability of the perfusion system. Temperature was monitored 200 µm from the tip of the perfusion outlet during rapid changes of solutions indicated by the arrows. Temperature remained constant throughout the whole experiment.

To verify the success of detubulation, cardiomyocytes were stained with di-8-ANEPPS and scanned on the confocal microscope (Fig. 4). Formamide treated cardiomyocytes showed no staining of the T tubules. All control myocytes showed regular staining of the T tubules. In accordance with previous findings (8), there was no significant difference in cell volume upon detubulation (23.6 ± 1.8 pl in control vs. 20.4 ± 2.2 pl in detubulated cardiomyocytes). However, cell capacitance was lowered by 30% (199 ± 9 pF in control and 140 ± 14 pF in detubulated cardiomyocytes, P < 0.01). Cell capacitance was plotted as a function of cell volume (Fig. 4C), and this showed that detubulated cardiomyocytes had lower capacitance than control cardiomyocytes with the same volume, distinctly separating the two groups.

View larger version (43K): [in this window] [in a new window]   Fig. 4. Effect of detubulation on cell capacitance and cell volume. Representative images of control (A) and detubulated cardiomyocytes (B) stained with di-8-ANEPPS and scanned with a confocal microscope at comparable z-heights. The lack of staining of the T tubules in B demonstrates that the cardiomyocytes were detubulated before experiments. C: cell capacitance vs. calculated volume for all control () and detubulated () cardiomyocytes studied. A linear regression was calculated for control (solid line, = 116, = 3.5) and detubulated (dotted line, = 50, = 4.4) cardiomyocytes. Mean values and SE are represented by square symbols with error bars.

To determine the diffusion kinetics in the T tubules, we followed the change in current induced by a rapid switch of [K⁺]_o in voltage-clamped control and detubulated cardiomyocytes. The delay due to change of solution in the perfusion tip was not significantly different in the control and the detubulated group (Table 1). However, the times to achieve steady-state holding current in elevated [K⁺]_o (t₂₅, t₅₀, and t₉₀) were all significantly shorter in the detubulated cardiomyocytes (Table 1). Figure 5A shows the mean curves of the current recorded during switch from 5.4 to 8.1 mM K⁺ in control and detubulated cardiomyocytes, with the baseline (–0.55 ± 0.04 nA in control and –0.47 ± 0.05 nA in detubulated cardiomyocytes, NS) adjusted to 0. The maximum change in current was 31% smaller in the detubulated cardiomyocytes (0.91 ± 0.07 nA in control and 0.63 ± 0.10 nA in detubulated, P < 0.05). Moreover, the densities of the changes in current during switch in [K⁺]_o were not significantly different in control and detubulated cardiomyocytes (4.72 ± 0.45 pA/pF and 4.87 ± 0.96 pA/pF, respectively). The difference between the two mean curves shown in Fig. 5A represents the change in current due to the change of [K⁺] in the T tubules (Fig. 5C). To visualize the difference in kinetics between the two groups, the maximum change in current was normalized to 1. The time to achieve maximal response was shorter in the detubulated group than in the control group (Fig. 5B and Table 1). Also shown in Fig. 5B is the time course of the solution switch (recorded junction potential).

View larger version (24K): [in this window] [in a new window]   Fig. 5. Change in holding current and membrane potential during change in extracellular [K+]. A–D: switch from 5.4 to 8.1 mM K+. E–H: 8.1 to 5.4 mM K+. A and E: basal currents were adjusted to 0 and the times of start of the current change were set equal. Curves show mean currents from control cardiomyocytes (black, A: n = 9, E: n = 7) and detubulated cardiomyocytes (light gray, A: n = 7, E: n = 5). B and F: the mean currents from control and detubulated cardiomyocytes were normalized so that the steady-state current equals 1. The dotted line represents the time course of the solution switch around the cell. C and G: the difference between the current recordings from control and detubulated cardiomyocytes (A and E) is traced in dark gray. The simulated data are showed in the dotted line. D and H: the recorded membrane potentials were adjusted to 0 and the time of start of the change in potential were set equal. Curves show mean potential from 10 control cardiomyocytes (black) and 6 detubulated cardiomyocytes (light gray).

Effect of a rapid change of [K⁺]_o on the membrane potential.

Nonuniform voltage clamp of the t-tubular membrane could give inaccurate measurements of currents from the T tubules, e.g., the currents coming from the surface membrane would be more efficiently monitored than currents emerging from the T tubules. Thus, to verify our observations in the voltage-clamp experiments, we performed current-clamp experiments during switch of [K⁺]_o. Switching between 5.4 and 8.1 mM K⁺ induced a change in the membrane potential of 9.9 ± 0.3 mV in control and 9.2 ± 0.5 mV in detubulated cardiomyocytes (NS). This is close to the theoretically expected change in membrane potential based on the Nernst equation. The resting membrane potential was –69 ± 1 mV in the control cardiomyocytes and –66 ± 2 mV in detubulated cardiomyocytes (P < 0.05). The kinetics of the change in membrane potential upon [K⁺]_o switch (t₂₅, t₉₀, and t₉₅) were not significantly different from those in the voltage-clamp experiments (Table 2). During exposure to 8.1 mM K⁺, the cardiomyocytes became unstable. Therefore the recorded change in potential during switch from 8.1 to 5.4 mM was not analyzed.

Simulations were only performed on the voltage-clamp data. The simulated data converged to the experimental data after 26 iterations. This resulted in a D of 85 µm²/s. At this diffusion rate, a monoexponential curve fit (tolerance R² > 0.99) of the simulated data gave = 471 ms in the step from 5.4 to 8.1 mM and = 478 ms in the step from 8.1 to 5.4 mM.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS ACKNOWLEDGMENTS REFERENCES

 
In the present study we examined the diffusion rate of K⁺ in T tubules of rat cardiomyocytes. Changes in current or membrane potential over time in voltage- or current-clamped control and detubulated cardiomyocytes were compared, and the difference was taken as the t-tubular component. The diffusion rate for K⁺ in the T tubules was modeled and was found to be slow, 85 µm²/s, which is about ninefold slower than in water (13).

The current and membrane potential changed with a time constant several orders of magnitude slower than the change in solution. Thus the change of solution around the cell was sufficiently fast for the present experiments. In control cardiomyocytes, the slow change of current and membrane potential can be explained by the presence of T tubules where the diffusion of K⁺ is limited. However, in the detubulated cardiomyocytes, the change should originate only from altered K⁺ currents across the surface membrane. Thus the change in current should occur immediately after the change of [K⁺]_o at the surface. However, as seen in Fig. 5B, the time course of the solution switch was faster than the actual change in transmembrane current. This has several possible explanations: 1) The detubulated cardiomyocytes were not completely detubulated. In some of the cardiomyocytes we observed remaining T tubules near the surface membrane. However, T tubules were not observed deep in the cardiomyocytes. 2) The diffusion could be restricted in the caveolae of the surface membrane. 3) The cardiomyocytes adhered to laminin coated coverslips during the experiments. Thus they were probably not equally accessible on all sides, which could restrict the change of solution. 4) The diffusion of solutes could be restricted near the surface membrane. Yao et al. (21), e.g., proposed that negatively charged polysaccharides, glycocalyx, reduce diffusion rates of ions. However, it was not the aim of this paper to determine the nature of such restrictive diffusion space. Furthermore, it is likely that restrictive diffusion due to caveolae, laminin adherence, or inherent surface membrane properties would be similar in detubulated and control cardiomyocytes. Thus the difference in current changes observed between control and detubulated cardiomyocytes in our experiments should originate only from restricted diffusion of K⁺ in the T tubules.

Several studies have assessed the diffusion rate of ions on the extracellular face of cardiomyocytes. Shepherd and McDonough (17) found a diffusion rate for Ca²⁺ in the T tubules of 95 µm²/s by comparing Ca²⁺ currents in ventricular cardiomyocytes from guinea pigs with atrial cardiomyocytes from rabbits (no T tubules). This diffusion rate is similar to that for K⁺ in the present study. Blatter and Niggli (1) found that a wavelike Ca²⁺ gradient traveled along the T tubules of guinea pig ventricular cardiomyocytes at a velocity of 3.4-16.3 µm/s. Assuming that the cardiomyocytes are 20 µm thick, the maximal diffusion velocity proposed by Blatter and Niggli would result in a complete exchange of solutes in the T tubules after 600 ms, which is similar to our results. Yao et al. (21) showed that the diffusion rate varies between species, but this could not be accounted for by differences in T tubule diameter or relative membrane area within the T tubules. A possible explanation could be that the cardiomyocyte membranes in different species are not equally affected by cell isolation procedures (e.g., collagenase exposure, Ref. 5). To avoid species- and cell-type-dependent differences, our study assessed the diffusion rate of K⁺ in the T tubules by comparing control and detubulated rat ventricular cardiomyocytes.

Both transmembrane potential and current depend on [K]_o and are largely linked to the properties of the inward rectifier (I_K1). During a change in [K⁺]_o, both the driving force for K⁺ and the conductance of I_K1 will change (16). Shepherd and McDonough (17) observed that the change in current was significantly slower than the change in membrane potential during a switch from 5.4 to 10.8 mM K⁺. The authors suggested that this discrepancy was due to electrotonic spread of depolarization, meaning that the depolarization of the cardiomyocyte due to a change in [K]_o proceeded much faster than the concentration change itself. In the present study, however, current- and voltage-clamp experiments showed similar results with regard to kinetics. This suggests that membrane potential and current are equally and directly dependent on [K]_o. Moreover, because both capacitance and maximum change in current were lowered by 30% in detubulated cardiomyocytes, the total density of active K⁺ channels was the same in the T tubules as on the surface membrane. This was also shown by Komukai et al. (9).

Detubulation of cardiomyocytes could alter Na⁺-K⁺-ATPase properties. Because the Na⁺-K⁺-ATPase is dependent on [K⁺]_o, this could complicate our interpretation that current and voltage changes were solely due to changes in K⁺ diffusion. However, this is unlikely for the following reasons: 1) Changes in [K⁺]_o in the 5–8 mM range will not greatly affect the pump rate (7). 2) Both current and voltage recordings had similar kinetics. 3) A change in Na⁺-K⁺-ATPase current would be negligible compared with the current recorded.

The slow diffusion of K⁺ in the T tubules indicates that they constitute functional compartments where K⁺, and probably other ions, can accumulate or deplete. Slow diffusion can be due to the tortuous structure of the T tubules and their varying diameter throughout the cell (18). Moreover, we speculate that gel properties in the lumen of T tubules as well as ion binding sites within the tubular membranes and glycocalyx can cause slow diffusion (17). Thus a "fuzzy space" (10) could also be present on the extracellular face of the cell membrane, in particular within the T tubules. The T tubules serve as a pathway for rapid transmission of the action potential to all parts of the cell, but exchange of solutes with the intercellular fluid is probably quite slow. Hence local control of the intratubular fluid composition will be quite important for maintaining excitability throughout the t-tubular structure. In skeletal muscle it has been speculated that intratubular K⁺ accumulation can be an important cause of fatigue during high-intensity exercise (16). Analogously, slow or dyssynchronous activation of T tubules in the heart may cause a slow rise of the Ca²⁺ transient (12). Thus both maintained t-tubular structure and tight control of the intratubular fluid composition are important for normal activation of the myocardium. It is possible that altered t-tubular function can be important for development of heart disease, for instance heart failure.

In conclusion, this study demonstrates that the diffusion rate for K⁺ in the T tubules of rat cardiomyocytes is slow, 85 µm²/s. This means that K⁺ ions, and probably other ions, can accumulate or become depleted in distinct areas in the T tubules during the contraction cycle. This compartmentalized distribution of ions probably plays an important role in the spread of the action potential, and possibly also for contractile properties of the cardiomyocytes.

	GRANTS

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We are grateful to the Norwegian Council for Cardiovascular Diseases, Anders Jahre's Fund for Promotion of Science and The Research Council of Norway for financial support.

	ACKNOWLEDGMENTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS ACKNOWLEDGMENTS REFERENCES

 
We thank Roy Trondsen for mechanical workshop expertise.

	FOOTNOTES

 

Address for reprint requests and other correspondence: F. Swift, Institute for Experimental Medical Research, Ullevaal Univ. Hospital, Kirkeveien 166, N-0407 Oslo, Norway (e-mail: fredrik.swift{at}medisin.uio.no)

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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