|ISSN No. 1606-7754 Vol.9 No.3 December 2001|
Myofilament Ca2+ sensitivity in ventricular myocytes from streptozotocin-induced diabetic rat
FC Howarth, MA Qureshi
Department of Physiology, Faculty of Medicine and Health Sciences, United Arab Emirates University, Al Ain
Contractile dysfunction is a frequently reported complication of diabetic cardiomyopathy and many of the defects observed in the clinical setting have also been reported in experimentally-induced diabetes. We have investigated the relationship between intracellular Ca2+ concentration and cell length during the relaxation phase of contraction in ventricular myocytes from streptozotocin (STZ) – induced diabetic rats. Cell length and intracellular Ca2+ concentration were measured simultaneously in electrically stimulated (1 Hz) myocytes loaded with fura-2 and maintained at 35-36 C. The amplitude and time to peak shortening and Ca2+ transient were similar, however, the relaxation of contraction and the Ca2+ transient were significantly prolonged in myocytes from STZ-treated rats compared to controls. Myofilament Ca2+ sensitivity, which was assessed by plotting cell length against fura-2 fluorescence ratio during the relaxation phase of a contraction, was significantly increased in myocytes from STZ-treated (9.23 0.77 m/fura-2 fluorescence unit) compare to controls (4.84 0.78 m/fura-2 fluorescence unit). The slower time course of relaxation of contraction and Ca2+ transient may be explained by defective sarcoplasmic reticulum Ca2+ uptake and to a lesser extent mechanisms of plasma membrane Ca2+ efflux including Na+/Ca2+ exchange and Ca2+ATPase. In conclusion, the apparent increase in myofilament Ca2+ sensitivity may be attributed to slower cross-bridge cycling rate which in turn may be related to the alteration of expression of different myofilament myosin isoforms.
Key words: Diabetes, calcium transport, myofilament Ca2+ sensitivity, streptozotocin, ventricular myocytes
Contractile dysfunction can occur in the diabetic heart in the absence of major blood vessel disease1-4 and many of the contractile defects observed in the clinical setting have also been reported in experimentally-induced diabetes.1 The streptozotocin– induced diabetic rat is a widely used animal model for the study of diabetes.1 Administration of STZ (60 mg kg-1) to young adult rats causes selective pancreatic beta cell necrosis and as a consequence these animals are characteristically hyperglycaemic and hypoinsulinaemic.1,5-7 Reduced rates of shortening, re-lengthening and the extent of shortening have been variously reported in electrically stimulated myocytes from STZ–induced diabetic rats.5,8-10 The characteristics of contractile defects depend partly on the duration of STZ – treatment.5,7 We have investigated the relationship between intracellular Ca2+ concentration and cell length during the early and late stages of relaxation of contraction in ventricular myocytes from STZ – induced diabetic rats.
Materials & Methods
Induction of diabetes:
Diabetes was induced by a single injection of STZ administered to young male rats (230-270 g) according to previously described techniques5,7,10. Briefly, male Wistar rats (230-270 g) received a single intraperitoneal injection of STZ (60 mg kg-1; Sigma, S-0130).7 STZ was dissolved in a citrate buffer solution (0.1 M citric acid and 0.1 M sodium citrate, pH 4.5). Age-matched controls received an equivalent volume of citrate acid buffer solution alone. Control and diabetic rats were caged separately but housed under similar conditions. Both groups of animals were fed the same diet and water ad libitum until they were used 8 to 10 weeks after treatment. Blood glucose was measured with a glucose meter (One Touch II glucose meter, Lifescan Inc, USA) 3-5 days after administration of STZ and immediately prior to experiments to confirm diabetic state. Principles of laboratory animal care were followed throughout. Approval for this project was obtained from the Faculty of Medicine & Health Sciences Ethics Committee.
Ventricular myocyte isolation
Ventricular myocytes were isolated according to previously described techniques with minor modifications.7,11 Rats were killed by decapitation using a guillotine, hearts were rapidly removed and perfused by the Langendorff’s method with a physiological salt solution (isolation solution – see below) containing 0.75 mM Ca2+ at 37°C. Perfusion flow-rate was adjusted to 8 ml g heart-1 min-1 to allow for differences in heart weight between diabetic and control animals. When the preparation appeared stable, perfusion was switched to an isolation solution containing 0.1 mM EGTA for 4 min. The heart was then perfused with the isolation solution containing 0.05 mM Ca2+, 0.75 mg ml-1 collagenase (Worthington, LS004196) and 0.075 mg ml-1 protease (Sigma, P-5147). The enzyme solution was recirculated to give a total exposure time of 6 min. At the end of the enzyme perfusion, the heart was cut down and the ventricles dissected free and cut into small slices.
The ventricular tissue was shaken in 5 ml of enzyme solution containing 1% bovine serum albumin for 4 min at 37°C and then filtered through gauze (300 mm aperture). After addition of 5 ml isolation solution containing 0.75 mM Ca2+, the cell filtrate was centrifuged (400 rpm, 40 sec). The supernatant was removed and the cell pellet was resuspended in 10 ml of isolation solution containing 0.75 mM Ca2+. The shaking process was repeated a total of four times. Myocytes from shakes 2 and 3 were accumulated and stored at 4°C prior to use. Cell viability, defined by the rod-shape, was compared in ventricular myocytes from diabetic or control rat heart within 1 h after completion of the cell isolation. Cells were used during a period of 1-8 h after isolation.
Measurement of shortening and intracellular Ca2+ concentration
Myocytes were loaded with the fluorescent indicator fura-2 AM (Molecular Probes, USA, F-1221) as described previously.12 In brief, 6.25 ml of a 1.0 mM stock solution of fura-2 dissolved in dimethylsulphoxide (Sigma, D-5879) was added to 2.5 ml of cells to give a final fura-2 concentration of 2.5 mM. Myocytes were shaken gently for 10 min at room temperature and then centrifuged (200-500 rpm). The supernatant was then removed and replaced with 10 ml of fresh experimental salt solution. Myocytes were left for 30 min to ensure complete hydrolysis of the intracellular ester.
Shortening and intracellular Ca2+ concentration were measured simultaneously using video edge detection (Crystal Biotech USA, VED-114) and fluorescence photometry (Cairn Research, England) systems, respectively. Electrically stimulated (1 Hz) myocytes were superfused (3-5 ml min-1) with a physiological salt solution (experimental solution – see below) containing 1 mM Ca2+ at 35-36°C. The amplitude of shortening (expressed as a % of resting cell length), the time to peak (TPK) shortening and time to 50 % (T50), 70 % (T70) and 90 % (T90) relaxation were recorded for each contraction.
To measure intracellular Ca2+ concentration, myocytes were alternately illuminated by 340 nm and 380 nm light using a monochromator (Cairn Research, England), which changed the excitation light every 2 ms. The resultant fluorescent emission at 510 nm was recorded by a photomultiplier tube and the ratio of the emitted fluorescence at the two excitation wavelengths (340/380 ratio) was calculated to provide an index of intracellular Ca2+ concentration. The amplitude of the Ca2+ transient and TPK and T50 %, T70 % and T90 % relaxation of the Ca2+ transient were recorded for each contraction.
Spurgeon et al13 have argued that during the final relaxation phase of contraction, the myofilaments come into quasi-equilibrium with the intracellular Ca2+ concentration. We have plotted cell length against fura-2 fluorescence ratio during the relaxation phase of a contraction to obtain an index of Ca2+ sensitivity of the myofilaments.
Solutions: The composition of the basic isolation solution was (in mM) 130.0 NaCl, 5.4 KCl, 1.4 MgCl2, 0.4 NaH2PO4, 5 HEPES, 10 glucose, 20 taurine and 10 creatine set to pH 7.3 with NaOH. The experimental salt solution contained (in mM) 140 NaCl, 5 KCl, 1 MgCl2, 10 Glucose, 5 HEPES and 1 CaCl2 set to pH 7.4 with NaOH.
Data analysis and statistics
Signal Averager v 6.37 (Cambridge Electronic Design, England) was used to acquire and analyse the experimental data. Results are presented as mean ± SEM from (n) cells. Statistical comparisons were made using the independent samples t-test. Significance values of less than 0.05 were considered significant.
General characteristics of STZ-treated and control rats: Diabetes was confirmed in STZ-treated rats by a 4.5-fold elevation of blood glucose and a significant (p < 0.05) reduction of non-fasting plasma insulin (Table 1). Body and heart weights were significantly (p < 0.01) lower in diabetic rats compared to age-matched controls.
Characteristics of ventricular myocytes from STZ-treated and control rats: The percentage of viable myocytes (defined by the rod shape of the cell) isolated from STZ-treated rats was lower than that from the controls despite compensating for the differences in the size of hearts by adjusting perfusion flow rates during cell isolation (Table 2). Using light microscopy, there was no obvious visual difference between rod shaped myocytes from STZ-treated compared to control rats.
Table 1: Characteristics of experimental animals
|Body weight (g)||291.20 ± 11.09 (5)||199.67 ± 7.67 (6)**|
|Heart weight (g)||1.24 ± 0.04 (5)||1.00 ± 0.00 (6) **|
|Blood glucose (mg dl-1)||84.00 ± 4.37 (5)||321.83 ± 25.84 (6)**|
|Plasma insulin (ng ml-1)||1.98 ± 0.35 (9)||0.82 ± 0.23 (9)*|
|Data are mean ± SEM, (n animals), ** P<0.01, * P<0.05.|
Table 2: Characteristics of ventricular myocytes
(% rod shaped cells)
|68.80 ± 2.27 (5)||37.50 ± 2.09 (6)**|
|Resting cell length (mm)||96.86 ± 2.40 (28)||101.47 ± 2.27 (33)|
|Peak amplitude shortening|
(% resting cell length)
|6.61 ± 0.66 (17)||5.74 ± 0.99 (20)|
|Time to peak shortening (ms)||119.65 ± 4.49 (17)||126.05 ± 4.27 (20)|
(Fluorescence ratio units)
|1.36 ± 0.06 (18)||1.27 ± 0.04 (20)|
|Ca2+ transient amplitude|
(Fluorescence ratio units)
|0.34 ± 0.03 (18)||0.32 ± 0.03 (20)|
|Time to peak Ca2+ transient (ms)||69.42 ±4.74 (18)||67.50 ± 3.35 (20)|
|Data are mean ± SEM, (n cells), ** P < 0.01.|
Resting cell length and fura-2 ratio were not significantly (p > 0.05) altered by STZ-treatment (Table 2).
Effects of STZ-induced diabetes on myocyte contraction and the Ca2+ transient: Averaged fast time base simultaneous recordings of cell length and fura-2 fluorescence ratio in electrically stimulated (1 Hz) myocytes from control and STZ-treated rats are shown in figures 1 (a) and (b), respectively. The peak amplitude of shortening and Ca2+ transient and the TPK shortening and Ca2+ transient were not significantly (P > 0.05) altered by STZ-treatment (Table 2). The time course of relaxation of contraction and the Ca2+ transient in electrically stimulated (1 Hz) myocytes from STZ-treated and control rats is shown in Figure 2. The T70 % and T90 % relaxation of contraction (a) and the T50 %, T70 % and T90 % of relaxation of the Ca2+ transient (b) were significantly (p < 0.05) prolonged in myocytes from STZ-treated compared to control rats.
The relationship between the magnitude of cell length and the corresponding fura-2 fluorescence ratio in myocytes from control and STZ-treated rats are shown in figures 1 (c) and (d), respectively. For a given contraction, the data proceed in an anticlockwise direction and the early and late phases of relaxation, which are indicated with arrows, were fitted with regression lines. The slope of the regression during the early phase of relaxation was not significantly (P > 0.05) different in STZ (50.93 ± 8.95 mm/fura-2 fluorescence unit, n =9) compared to control (44.26 ± 5.28 mm/fura-2 fluorescence unit, n=10) myocytes. However, during the late phase of relaxation the slope of regression was significantly (P < 0.01) increased in STZ (9.23 ± 0.77 mm/fura-2 fluorescence unit, n=9) compared to control (4.84 ± 0.78 mm/fura-2 fluorescence unit, n=10) suggesting that myofilament Ca2+ sensitivity is increased in myocytes from STZ-treated rats.
The STZ rat model of diabetes is well characterized and widely used for the study of diabetic cardiomyopathy.1 As reported in several previous studies rats developed hyperglycaemia within 3 days after injection of STZ and at 8-10 weeks the STZ-treated rats showed significantly less body weight gain, increased plasma glucose and reduced non-fasting plasma insulin concentration in comparison with age-matched controls.5,7,14 The cell viability, defined by the rod-shape, was significantly lower in myocytes from STZ-treated rats. It has been suggested that hearts from STZ-treated rats may be more sensitive to collagenase digestion than controls.14 Attempts to equalise viability between STZ-treated and control preparations by adjusting isolation solution flow rates to allow for differences in heart weight did not significantly affect the outcome.
Consistent with several previous studies the resting cell length5-8 and fura-2 fluorescence ratio6,14 were not significantly altered by STZ-treatment. Reductions15 and increases16 of resting Ca2+ have also been previously reported in STZ myocytes.
Consistent with some previous studies the peak amplitude of shortening and the Ca2+ transient were not significantly altered in myocytes from STZ-treated compared to control rats.6,14 Increases7 or decreases8,9 in the amplitude of shortening and decreases10,15 in the amplitude of the Ca2+ transient have also previously been reported in myocytes after STZ-treatment.
The TPK shortening and the Ca2+ transient were not altered in myocytes from STZ-treated compared to control rats. Prolongation of TPK shortening in myocytes after STZ-treatment has also been reported in previous studies.5,7,8,9
These inconsistences in the amplitude and time course of shortening and Ca2+ transient may partly be attributable to differences in treatment protocols, isolation or experimental methodologies. Previous studies in our laboratory have shown that the kinetics of shortening alters as treatment time is increased.7
The T70 % and T90 % relaxation of contraction and T50 %, T70 % and T90 % relaxation of the Ca2+ transient were significantly prolonged in myocytes from STZ-treated compared to control rats. Prolonged relaxation of contraction5,8,9 and decay of the Ca2+ transient have been previously reported.5,15 Mechanisms underlying the prolonged time course of relaxation of contraction may include depressed SR Ca2+ ATPase17-20 or altered Na+/Ca2+ exchange activity.18, 21
During the final phase of relaxation of contraction, the myofilaments come into quasi-equilibrium with the intracellular Ca2+ concentration13 and by plotting cell length against fura-2 fluorescence ratio during the relaxation phase of a contraction, it is possible to obtain an index of Ca2+ sensitivity of the myofilaments.13 The slope of the regression during the late (but not the early) phase of relaxation of contraction was significantly increased in myocytes from STZ-treated rats suggesting that myofilament sensitivity to Ca2+ is increased after STZ-treatment. However, in the absence of any alteration to peak amplitude of shortening or Ca2+ transient it is suggested that the apparent increase in myofilament Ca2+ sensitivity may be attributed to a reduction of cross-bridge cycling rate which is a feature that has been previously reported in papillary muscle.22 The reduction in cross-bridge cycling may in turn be related to the alteration of myosin isoform from V1 to V323 a feature that has been frequently reported in STZ-induced diabetes.24,25
The slower time course of relaxation of contraction and Ca2+ transient may be explained by defective sarcoplasmic reticulum Ca2+ uptake and to a lesser extent mechanisms of plasma membrane Ca2+ efflux including Na+/Ca2+ exchange and Ca2+ ATPase. The apparent increase in myofilament Ca2+ sensitivity may be attributed to slower cross-bridge cycling rate which in turn may be related to the alteration of expression of different myofilament myosin isoforms.
This study was supported by a grant from the Faculty of Medicine & Health Sciences, United Arab Emirates University, Al Ain, U.A.E., and an award from H.H. Sheikh Hamdan Bin Rashid Al Maktoum Awards for Medical Sciences.