Catalase Protects Cardiomyocyte Function in Models of Type 1 and Type 2 Diabetes

OVE26 diabetic mice (15) and catalase transgenic mice that overexpress catalase 60-fold in the heart were previously described (16). Catalase and OVE26 mice were maintained on the inbred background FVB. OVE26-positive mice were identified by the presence of small eyes caused by the cointegration of the GR19 gene, which is expressed in the eye. Catalase-transgenic mice were identified by PCR of tail DNA. Mice positive for the OVE26 diabetes gene and the catalase transgene were designated OVE26C. Age-matched OVE26, OVE26C, and FVB mice of both sexes were obtained at 18–22 weeks of age.

Ay type 2 diabetic mice on the KK background were obtained from Jackson Laboratories. Catalase transgenic mice were bred to Ay mice. Offspring positive for the Ay gene were recognized by agouti coat color. Agouti-positive catalase-positive mice were designated AyC. All agouti experiments were performed on the F1 generation. These mice were designated FK for their 50% FVB and 50% KK genetic background. All type 1 and type 2 mating pairs used nondiabetic females to prevent the influence of diabetic gestation on cardiac phenotype. Mice were maintained on a 12-h light/dark cycle and received food (Purina Laboratory Rodent Diet 5001) and water ad libitum. The U.S. Department of Agriculture-certified institutional animal care committee approved all animal procedures.

Northern blots were prepared and probed as previously described (17), using 10 μg of heart RNA from transgenic or control mice at 150 days of age. Oligonucleotide probes for mouse mRNAs for catalase and glyceraldehyde 3-phosphate dehydrogenase (GAPDH; GGAACATGTAGACCATGTA-GTTGAGGTCAATGAAG) were labeled with [γ-³²P]ATP and polynucleotide kinase. The strength of the hybridization signal was determined on a Molecular Dynamics phosphoimager. Signals for catalase were normalized to the signals for GAPDH.

Hearts were frozen in liquid nitrogen, pulverized, and homogenized in 50 mmol/l sodium phosphate buffer (pH 7) containing 1 mmol/l EDTA, 40 μmol/l butylated hydroxytoluene, and protease inhibitor cocktail. The protein estimation was carried out using a Pierce BCA kit. Samples equivalent to 10 μg proteins were separated on 4–20% Tris-glycine gradient gels and transferred to polyvinylidene difluoride membrane. The membranes were blocked in 5% milk and incubated overnight in 1:2,000 dilution goat anti-malondialdehyde (MDA) polyclonal antibody from Academy Biomedical. The bands were visualized using peroxidase-conjugated affinity-purified donkey anti-goat IgG and an enhanced chemiluminescence detection system (Amersham). Signals for MDA were normalized to the signals for β-actin obtained from the same blot.

Procedures were carried out as previously described (17). Fixation was carried out by vascular perfusion through the left ventricle. The perfusates included washouts at 4°C of 35 ml half-strength Karnovsky’s (18) fixative with 1% procaine, and 75 ml full-strength Karnovsky’s final fix. Thin sections were observed in an Hitachi 7500 transmission electron microscope at initial magnifications of 4,000–15,000 diameters.

To quantify the morphologic features of cardiac tissues in each of the animal groups, transmission electron micrographs of randomly selected areas of left-ventricular thin sections were printed at identical magnifications. Two observers who were unaware of their origin rated these prints. Observers rated the morphology on a scale of 1–4 relative to a figure exhibiting near-perfect cardiac morphology. Figures were rated for overall morphology, which included the presence of edematous areas, myelin figures, and overall organization; mitochondrial morphology, which included mitochondrial packing, density, and shape; and myofibril morphology, which examined linearity of the Z lines and the linearity, regularity, and continuity of myofibrils.

Single ventricular myocytes were isolated by enzymatic dissociation with Liberase Blendzyme 4 (Roche Diagnostics, Indianapolis, IN) from adult mice by modification of described procedures (4). Briefly, animals were injected with heparin (1,000 units/kg i.p.) and anesthetized with ketamine (150 mg/kg i.p.) and xylazine (22.5 mg/kg i.p.). Their hearts were rapidly removed and perfused (at 37°C) with oxygenated (5% CO₂/95% O₂) Krebs-Henseleit calcium-free buffer (in mmol: NaCl 135, KCl 4.0, MgCl₂ 1.0, NaH₂PO₄ 0.33, HEPES 10, glucose 10, and Butanedione 10, pH 7.4). Hearts were then perfused with the same Ca²⁺-free Krebs-Henseleit buffer with collagenase II (0.9 mg/ml) for 15–20 min until the heart became flaccid. After perfusion, the left ventricle was removed, minced, and resuspended with the same Ca²⁺-free Krebs-Henseleit buffer at room temperature. Extracellular Ca²⁺ was added incrementally back to 1.2 mmol/l. Isolated myocytes were maintained at room temperature in a serum-free medium consisting of medium 199 (Gibco) with Hanks’ salts containing 25 mmol HEPES for further mechanical and fluorescent studies. Mechanical properties remained relatively stable in myocytes for 8–10 h in the serum-free medium. Cell yield was 50–70%. There was no notable difference of yield between normal FVB, FK, and diabetic mice. Myocytes with obvious sarcolemmal blebs or spontaneous contractions were not used. Only rod-shaped myocytes with clear edges were used for recording of mechanical properties, intracellular Ca²⁺ transients, and ROS fluorescent measurement.

Mechanical properties of ventricular myocytes were assessed using a video-based edge-detection system (IonOptix, Milton, MA) (4). The cells were field stimulated at a frequency of 1.0 Hz for 4 ms, using a pair of platinum wires that were placed on opposite sides of the dish chamber and connected to a MyoPacer Field Stimulator (IonOptix). The polarity of the stimulatory electrodes was reversed frequently to avoid possible buildup of electrolyte by-products. The myocytes being studied were displayed on the computer monitor using an IonOptix MyoCam camera, which rapidly scans the image area every 8.3 ms such that the amplitude and velocity of shortening/relengthening was recorded with good fidelity. Soft-edge software (IonOptix) was used to capture changes in cell length during shortening and relengthening. Cell shortening and relengthening were assessed using the following indexes: peak shortening, time to 90% peak shortening (TPS₉₀), time to 90% relengthening (TR₉₀), and maximal velocities of shortening and relengthening (±dL/dt).

A separate cohort of myocytes was loaded with Fura 2-AM (1.0 μmol/l) for 30 min, and fluorescence measurements were recorded with a dual-excitation fluorescence photomultiplier system (Ionoptix). Myocytes were placed in a dish chamber on the stage of an Olympus IX-70 inverted microscope and imaged through a Fluor 100× oil objective. Cells were exposed to light emitted by a 75-W lamp and passed through either a 360- or a 380-nm filter (bandwidths were ±15 nm), while being stimulated to contract at 1.0 Hz. Fluorescence emissions were detected between 480 and 520 nm by a photomultiplier tube after first illuminating the cells at 360 nm for 0.5 s and then at 380 nm for the duration of the recording protocol (333-Hz sampling rate). The 360-nm excitation scan was repeated at the end of the protocol, and qualitative changes in intracellular Ca²⁺ concentration ([Ca²⁺]_i) were inferred from the ratio. Intracellular Ca²⁺ transients were measured as changes in Fura 2 fluorescence intensity (FFI). ΔFFI was determined as the difference between the levels of Ca²⁺ in systolic and diastolic conditions (ΔFFI = peak FFI − baseline FFI). The time course of the fluorescence signal decay (τ: the duration where Ca²⁺ transient decays 67% from the peak level) was calculated to assess intracellular Ca²⁺ clearing rate.

The membrane-permeable probe 5-(6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate (CM-H₂DCFDA) enters cardiomyocytes and produces a fluorescent signal after intracellular oxidation by ROS such as hydrogen peroxide and hydroxyl radical (19). Intracellular oxidant stress was monitored by measuring changes in fluorescence resulting from intracellular probe oxidation. Isolated myocytes were loaded with 1 μmol/l CM-H₂DCFDA (Molecular Probes, Eugene, OR) for 30 min. The cardiomyocytes from each group were incubated at room temperature in M199 medium with normal glucose concentration (5.5 mmol/l) or high glucose (25 mmol/l). Myocytes were fixed in 4% paraformaldehyde after incubation and stored at 4°C for up to 16 h before imaging. Fluorescence intensity from individual cells was measured using an excitation wavelength of 485 nm and an emission wavelength of 530 nm. Cells were sampled at random in each preparation, using an Olympus IX70 inverted microscope equipped with a digital-cooled charge coupled device camera and ImagePro image analysis software (Media Cybernetics, Silver Spring, MD). Fluorescence was calibrated with InSpeck microspheres (Molecular Probes). Calibration curves were generated, and cell brightness was measured. The InSpeck microspheres were used as internal standards. After subtraction of background fluorescence, individual cell values were determined as the ratio of myocyte to standard bead fluorescence.

Data are expressed as the means ± SE. Statistical comparisons were performed by ANOVA and Bonferroni post hoc test among different groups. Significance was defined as P < 0.05.

Oxidatively stressed cells respond by induction of protective enzymes. In OVE26 hearts expression of catalase mRNA was increased by 40% compared with control hearts (Fig. 1). This was less induction than that reported for streptozotocin-induced (20,21) and alloxan-induced (22) diabetic models, but it was similar to results obtained with hearts from the BB rat, an autoimmune model of type 1 diabetes, which exhibited a 52% increase in catalase (23). The moderate increase in expression of catalase suggests that the heart is responding to diabetes-induced oxidative stress.

Transmission electron microscopy was carried out to determine whether the catalase transgene could reduce the damage to morphology in left ventricular tissue produced by diabetes (Fig. 2). Typical morphology of FVB, OVE26, and OVE26C hearts is shown in Figs. 2A–C. Consistent with our previous observations (10), heart tissue from diabetic OVE26 animals showed disorganization of normal architecture, myelin figures, damaged mitochondria, and disrupted myofibrils. To verify our impression that overexpression of catalase was effective in reducing morphologic damage, semiquantitative ratings were made of random electron micrographs by observers blind to the origin of the picture. These figures were judged for overall morphology, mitochondrial structure, and myofibrillar substructure, as described in research design and methods. The results of this analysis (Fig. 2D) confirmed that catalase overexpression provided significant protection from diabetes-induced damage on all parameters of cardiac morphology.

Observation at higher magnification revealed focal areas in OVE26 hearts with severely damaged mitochondria. This is most apparent when comparing high-magnification pictures of typical FVB mitochondria (Fig. 2E) and severely damaged OVE26 mitochondria (Fig. 2F). Similarly damaged mitochondria were never seen in control hearts and almost never seen in OVE26C hearts. In some micrographs (Fig. 2G), relatively healthy-looking myofibers were juxtaposed to severely damaged mitochondria. This suggests that mitochondria were especially sensitive to diabetes-induced damage.

Diabetes significantly impaired contractility in isolated cardiomyocytes (Fig. 3). OVE26 diabetes produced at least a 30% reduction in percent shortening, rate of shortening, and rate of relengthening. Figure 3 shows that catalase overexpression in OVE26C mice completely prevented this effect: contractility in FVB and OVE26C myocytes was indistinguishable.

As shown in Fig. 4, recovery of the Ca²⁺ transient occurred more rapidly in control FVB cardiomyocytes than in OVE26 myocytes. The effect of the catalase transgene on the Ca²⁺ transient was small. Calcium recovery times in OVE26 and OVE26C myocytes were not significantly different, and both were significantly slower than that in FVB cardiomyocytes.

MDA is a by-product of lipid peroxides and thus an indicator for oxidative damage. Protein bands of ∼35, ∼50, and ∼80 kDa gave increased signals with antibody to MDA in OVE26 hearts (Fig. 5). This effect was at least partially reversed by overexpression of catalase. The 50-kDa band, which was the most prominent MDA adduct, was quantified relative to β-actin staining and found to be increased by 2.5-fold in OVE26 hearts (P < 0.02) compared with FVB control hearts. The intensity of the 50-kDa band was not significantly different in FVB and OVE26C hearts.

Figure 6 shows ROS production measured with CM-H₂DCFDA. Normal extracellular glucose produced similar ROS intensities among the cardiomyocytes from all three groups. Treatment of myocytes with high glucose (25 mmol/l) for 60 min did not affect the ROS level in control cardiomyocytes, but it caused a significant increase in the OVE26 cardiomyocytes. However, this effect of high glucose on ROS production was prevented by overexpression of catalase.

Mitochondrial electron transport is an important source of superoxide anion. Inhibition of electron transport with rotenone or thenoyltrifluoroacetone eliminated the excess ROS production of OVE26 myocytes in high glucose (Fig. 7).

Catalase transgenic mice were bred to Ay diabetic mice to test whether catalase overexpression would be protective in this model of type 2 diabetes. Mice were analyzed at 130–145 days of age. F1 offspring that were Ay positive had significantly higher blood glucose levels than their Ay-negative siblings (266 ± 31 vs. 155 ± 13, P < 0.05). Ay-positive mice were also significantly more obese than their Ay-negative siblings (48 ± 0.9 vs. 36 + 2.5, P < 0.01). Ay diabetes significantly impaired cardiomyocyte contractility (Fig. 8), as indicated by reduced peak shortening and lower rates of shortening and relengthening. Overexpression of catalase provided almost complete protection against the reduction in contractility.

Our results show that catalase protects against diabetic cardiomyopathy. Nontransgenic diabetic hearts express 40% more mRNA from their endogenous catalase gene than control hearts. By increasing catalase activity 60-fold with a cardiac-specific transgene, we were able to significantly reduce the primary characteristics of diabetic cardiomyopathy. The transgene prevented morphologic damage to mitochondria and myofibrils. Impaired cardiomyocyte contractility was eliminated, and increased ROS production by high glucose was significantly reduced. Catalase was found to be effective in models of type 1 and type 2 diabetes.

The protection produced by catalase indicates that ROS are an essential intermediate in diabetic cardiomyopathy. This hypothesis was strengthened by the finding that several protein bands in diabetic hearts contained more MDA, a by-product of lipid peroxidation, than nondiabetic hearts. Catalase reversed the increase in MDA staining for the most prominent of these bands, confirming that catalase reduces diabetes-induced oxidative stress. Identification of the specific MDA-modified proteins may indicate which cardiac compartments are most stressed by diabetes and which proteins contribute to impaired myocyte function. In our previous (10,11) studies, we demonstrated that a transgene for the heavy metal-binding antioxidant protein MT was protective against diabetic cardiomyopathy. Unlike MT, which has several functions, catalase has only one major action in the cell: detoxification of hydrogen peroxide. Compared with MT, catalase was just as effective, except that it did not restore the normal rate of calcium reuptake. The additional protection by MT could be attributable to nonantioxidant actions of MT, or it could be a result of its broader range of antioxidant actions than catalase. MT can scavenge hydroxyl radical, superoxide, and peroxynitrite, whereas catalase is largely limited to hydrogen peroxide. One or more of the ROS that catalase cannot detoxify may therefore be involved in impaired calcium reuptake. On the other hand, these results show clearly that it is not essential to protect against all ROS in order to provide significant protection from diabetic cardiomyopathy. This should be of advantage in devising effective antioxidant therapies.

Our ROS measurement studies and morphology results implicate mitochondria as a major source of oxidative stress in OVE26 myocytes. Mitochondria are an obvious candidate because they are the major source of superoxide radical (24), they produce more ROS in a diabetic environment (9,25), and mitochondrial damage is commonly associated with cardiomyopathy (26) and diabetes (27). When diabetic myocytes were incubated in high glucose, they generated excess ROS compared with normal cardiomyocytes. This could be prevented by treatment with rotenone or thenoyltrifluoroacetone, inhibitors of electron transport from cytochromes I and II, respectively. If mitochondria are the major source of ROS, they should be exposed to the most oxidative stress and may suffer the most damage. Consistent with this, we observed in electron micrographs of OVE26 ventricles that mitochondria were the most obviously disrupted cellular component. If this damage impairs downstream mitochondrial electron transport complexes III (28) and IV (29), then the half-life of ubisemiquinone will be prolonged and ROS production increased. It is possible that diabetes produces initial damage to mitochondria, which increases ROS production, resulting in progressively greater damage to cardiomyocytes. If this scenario is correct, then catalase limits the damage diabetes inflicts on cardiomyocytes by providing protection from initial damage to mitochondria and/or protecting against enhanced mitochondrial ROS production.

The catalase transgene also protected cardiomyocytes of the agouti mouse. Diabetic pathology is very different in agouti and OVE26 models. Agouti mice are insulin resistant, obese, and moderately hyperglycemic. OVE26 mice are lean and severely hyperglycemic. OVE26 hearts are not insulin resistant, as judged by insulin-dependent phosphorylation of Akt (G.Y., N.S.M., R.V.D., S.X., M.X., E.C.C., P.N.E., unpublished results). Despite the differences, agouti diabetic cardiomyocytes showed deficits in contractility similar to OVE26 myocytes: percent contractility, rate of contractility, and rate of relaxation were all impaired. These results indicate that the insulin-resistant moderately hyperglycemic diabetes of agouti mice produces similar contractile impairment as insulinopenic, severely hyperglycemic diabetes of OVE26 mice. However, our preliminary results (not shown) suggest that calcium reuptake is either not impaired or much less impaired in agouti myocytes than in OVE26 myocytes. Catalase restored contractility to near-normal levels in agouti myocytes, as it did in OVE26 myocytes. The ability of a specific antioxidant protein to protect cardiomyocytes strongly implicates ROS as a mechanism of damage in type 2 diabetes. That both type 1 and 2 diabetes should act through ROS mechanisms is reasonable. Although there are many hypothesized mechanisms of increased ROS production in diabetes, all are initiated by hyperglycemia. Both the agouti and OVE26 models demonstrated hyperglycemia, albeit to a different extent. Because insulin resistance alone has been shown to impair cardiomyocyte function (30), it is likely that this added to the detrimental effects of the mild hyperglycemia in the agouti mouse. The fact that catalase was effective in agouti cardiomyocytes provides one of the first indications that ROS are an essential cause of diabetic cardiomyopathy in type 2 diabetes

This work was supported by grants from the National Institutes of Health (HL62892, HL6677, and RR17702), the Commonwealth of Kentucky Research Challenge Trust Fund, and the University of Louisville Center for Genetics and Molecular Medicine.

We thank Donna Laturnus for technical assistance with electron microscopy.