Integrin Activation in the Heart
A Link Between Electrical and Contractile Dysfunction?
Integrins mechanically link the cytoskeleton to the extracellular matrix in cardiac myocytes and are thereby involved in mechanotransduction. Integrins appear to be necessary for cardiac myocyte hypertrophy. To determine the effect of increased integrin ligation and signaling on adult cardiac function, a heart-specific truncated α5 integrin (gain of function) was conditionally expressed in mice. Four days later, we observed an 80% reduction in amplitude of the QRS complex, profound systolic dysfunction, decreased connexin43, loss of gap junctions, and abnormal intercalated discs. Surprisingly, isolated left ventricular myocytes contracted normally and exhibited normal Ca2+ transients. This suggested that cell/cell electrical and/or mechanical coupling was disrupted. To distinguish electrical from mechanical coupling deficits, we compared the papillary muscle force generated by electrically stimulated versus rapid cooling contractions in which intracellular Ca2+ is released without electrical depolarization. Both were decreased in the transgenic muscle. However, electrically stimulated contractions were more significantly reduced than rapid cooling contractures. This suggests a component of cell/cell electrical uncoupling. Optical mapping revealed a loss of the normal elliptical isochronal activation pattern implying a loss of preferential conduction through gap junctions. For the first time, we have shown that integrins can regulate both mechanical and electrical coupling in the adult heart, even in the absence of primary hemodynamic alterations. Furthermore, we demonstrated that unregulated integrin activation leads to both contractile dysfunction and arrhythmias.
The extracellular matrix in the heart is linked to the internal cytoskeleton and force generating proteins of myocytes by transmembrane proteins, including integrins. In cardiac myocytes, integrins are localized to the Z-discs and intercalated discs.1 In addition to providing a mechanical linkage, integrin/ligand binding results in activation of intracellular signaling cascades that influence diverse cellular processes.2
Cardiac hypertrophy develops in response to chronically increased wall stress. In general, the pathological consequences of long-standing pressure overload include increased extracellular matrix deposition, decreased myocardial contractility, and increased vulnerability to arrhythmias. The mechanism(s) that link and govern these seemingly diverse changes have remained elusive. Integrins are likely candidates for the molecules that transduce increased mechanical force into the intracellular biochemical signals that ultimately define the hypertrophic phenotype.3 Previous data showing that integrin expression increases during pressure overload in the heart indicate a potential role for integrins in the hypertrophic response.4 A variety of in vitro studies also support the concept that integrin signaling is involved in cardiac hypertrophy.5
We directly tested the hypothesis that enhanced integrin signaling in the adult heart, in the absence of altered loading conditions, contributes to the electrical and mechanical defects that are characteristic of pressure-overload hypertrophy. We used an α5 integrin with a truncated cytoplasmic domain, hereafter denoted α5-1. This mutant integrin is not subject to the usual affinity regulation by “inside-out” signaling and thus behaves as a classic gain of function mutant.6 In fact, α5-1-integrin expression increases both fibronectin matrix assembly and downstream tyrosine phosphorylation.6,7 We here establish that conditional expression of the α5-1 integrin in adult cardiomyocytes results in marked electrocardiographic and conduction abnormalities, contractile dysfunction and sudden death. We provide evidence that this phenotype results, at least in part, from loss of connexin43 (Cx43)-containing gap junctions, leading to cell-to-cell electrical and mechanical uncoupling.
Materials and Methods
The creation of double transgenic (Tg) mice, all molecular biology, histology, cell biology, optical acquisition, physiology, functional assays, and antibodies used are described in the online data supplement, available at http://circres.ahajournals.org.
Student’s t test (2-tailed, unpaired) compared doxycycline (Dox)-treated wild-type (WT) with Dox-treated α5-1 female mice. For studies using more than 1 time point or condition, ANOVA was performed with post hoc multiple comparisons procedures applied when appropriate. Probability of <0.05 was considered significant. Data presented as mean±SEM.
Creation of Mice With Cardiac-Specific, Inducible Expression of α5-1 Integrin
Twenty-one potential transresponder (TR) founder mice were born. Polymerase chain reaction identified 5 that carried the α5-1 transgene. All 5 transmitted the α5-1 transgene when mated to codon-optimized, tetracycline-regulated, transactivator (TA) mice (Line JAM8585).8 The double-Tg (TR/TA) offspring, hereafter referred to as α5-1 mice, were treated with Dox to induce α5-1 expression. Immunohistochemical analysis identified 2 founders that uniformly expressed the α5-1 integrin in all cardiac myocytes. Both lines, JAM2954 and JAM 2938, had similar phenotypes. JAM 2954 was used in all subsequent experiments.
Induction of α5-1 Integrin Leads to Sudden Death
Double-Tg α5-1 mice at 6 to 12 weeks of age had normal body size, appearance, and survival. Before administration of Dox, α5-1-integrin expression was undetectable (Figure 1A). Two to 6 hours after Dox administration, α5-1-integrin protein was detected in Tg hearts and maximal expression was reached within 72 to 96 hours (Figure 1A). Immunofluorescence microscopy of isolated myocytes demonstrated the punctuate membrane localization of α5-1 integrin on cardiac myocytes (Figure 1B). This pattern of localization was consistent with activated integrin clustering. The endogenous α5 integrin had a similar pattern with additional staining representative of inactive integrin localization (Figure 1B). Expression of α5-1 integrin was uniform throughout the myocardium (Figure 1C). α5-1 Integrin expression was not detected in any other tissues of the Tg mice, or in any tissues of control mice. After 96 hours of Dox, some α5-1 mice exhibited labored breathing and sudden death. Within 2 weeks, most α5-1, but no WT, mice had died. Administration of Dox to WT mice had no discernable effects. Postmortem examination of the α5-1-integrin mice that died revealed no pericardial or pleural effusions or ascites. A small increase in heart weight/ body weight and heart weight/tibia length was observed in Tg mice after 4 days (Table). Isolated cardiomyocytes demonstrated no significant change in surface area (Table). Immunofluorescence microscopy revealed a substantial increase in fibronectin matrix deposition in α5-1 hearts (Figure 1D, Tg, versus 1E, WT). Most subsequent analyses were conducted 4 days after Dox administration unless stated otherwise. Preliminary studies suggested more severe left ventricular (LV) dysfunction in female versus male mice (data not shown). Therefore, to avoid sex-specific effects, only female mice were analyzed.
Electrocardiographic and Hemodynamic Abnormalities
Surface ECGs revealed that heart rate was modestly slowed in α5-1 versus WT mice (Table). α5-1 Versus WT mice had mild prolongation of the PR interval (40.0±1.4 versus 33.1±1.4 ms; P=0.013) and QRS duration (16.7±1.7 versus 12.9±0.05 ms; P=0.03). The most striking electrocardiographic feature was the marked decrease in the amplitude of the P waves (0.13±0.016 versus 0.22±0.014 mV; P=0.0007) and QRS complexes (0.24±0.04 versus 1.12±0.05 mV; P<0.0001) in α5-1 versus WT mice (Figure 2A). One time, we recorded a spontaneous tachyarrhythmia in an α5-1 mouse (Figure 2B). Echocardiography performed at 4 days showed a marked reduction in global LV systolic function, no change in LV wall thickness, but increased LV internal dimensions in diastole and particularly in systole (Table). LV fractional shortening and cardiac output were markedly reduced, and LV catheterization revealed lower LV systolic pressure, reduced +dP/dt and −dP/dt, and increased LV end-diastolic pressure (Table). No changes were seen in LV function after 2 days of Dox treatment (data not shown). There were no significant echocardiographic or hemodynamic differences among Dox-treated WT mice, untreated WT, or untreated double-Tg mice (data not shown).
Decreased Force Production in α5-1 Myocardium
To determine the mechanisms of the profound systolic dysfunction following α5-1-integrin induction, we assessed in vitro LV papillary muscle function. Maximal isometric force production and relaxation were measured under conditions of comparable preload. Papillary muscles from α5-1 hearts (n=8) showed a severe reduction in maximal developed tension compared with WT muscles (n=7; Figure 2C). Similar reductions were seen at both slow and fast stimulation frequencies (data not shown) and at low and high extracellular [Ca2+] (Figure 2D). The force reduction was not purely a result of tissue damage as histological sections of papillary muscles from a separate group of Dox-treated α5-1 mice showed only modest areas of replacement fibrosis (supplement Figure IA through IC). Isolated left atrial preparations from α5-1 mice demonstrated similar reductions in developed force, indicating that the α5-1 integrin also provoked deleterious effects in the atria (supplemental Figure II).
Preserved Contractility, Ca2+ Transients, and L-type Ca2+ Currents in Isolated Myocytes
To further define the nature of the contractile impairment in α5-1-integrin hearts, we assessed sarcomeric shortening and Ca2+ transients in isolated cardiac myocytes (WT, n=19; Tg, n=23). The yield of Ca2+-tolerant, striated, rod-shaped, quiescent myocytes was similar in WT and Tg mice. Surprisingly, we found that both sarcomere shortening and Ca2+ transients were of normal amplitude in the α5-1-integrin myocytes compared with WT myocytes (2.77±0.56% versus 2.56±0.43%, P=NS; and 2.70±0.21 versus 2.38±0.12 F/F0, P=NS, respectively) (Figure 2E). Furthermore, peak L-type Ca2+ current density was not different in LV myocytes from Dox-treated WT (n=8) and Tg (n=7) mice (10.3±1.0 versus 12.7±1.0 pA/pF; P=NS). Therefore, the prominent ventricular and papillary muscle dysfunction was apparently not caused by impaired function of individual myocytes. This suggested that myocyte death, impaired force transduction in muscle, or impaired electrical activation of the myocardium may explain the dysfunction seen in the whole-heart and multicellular muscle preparations.
Increased Cardiac Membrane Permeability and Fibrosis
To test the hypothesis that reduced cardiac function was attributable to membrane permeability and myocyte death, cardiac myocytes, in vivo, were examined for the ability to exclude Evans Blue dye. Fluorescent microscopic examination revealed scattered Evans Blue uptake into individual cardiac myocytes of the α5-1 integrin-expressing mice (Figure 3A), but not in Dox-treated WT mice. Specifically, less than 1% of cells were permeable to Evans Blue dye after 2 days, ≈9% after 3 days, and ≈33% after 4 days. Masson’s trichrome staining revealed minimal fibrosis in α5-1 hearts after 4 days of Dox (data not shown). However, increased cardiac fibrosis was observed in α5-1 mice allowed to recover for 14 days (Figure 3B).
Alterations in Cardiac Gene Expression Induced by α5-1 Integrin
To examine the molecular responses to α5-1-integrin expression in the heart, we performed both microarray (NIA 15K mouse clone set, chip A) and more focused GEarray (SuperArray Inc, Bethesda, Md) analysis of mRNA expression. To maximize our ability to detect early downstream changes directly linked to signaling via the α5-1 integrin, rather than general compensatory changes caused by ventricular dysfunction, analysis was performed 48 hours after α5-1 expression was initiated. Microarray analysis confirmed the rapid and robust induction of α5-integrin mRNA (310-fold). Other major increases in mRNA included the heat shock proteins HSP70/grp78 (9.6×) and HSP90/gp96 (8×), binding protein (BiP) (10.5×), and calreticulin (8.5×). No changes were observed in the sarcoplasmic reticulum calcium ATPase (SERCA2) or the sarcolemmal sodium/potassium ATPase. Analysis with GEarray Q series, focused DNA microarrays (SuperArray Bioscience Corporation) demonstrated no change in transcripts involved in apoptosis such as bax, bcl-2, inducible nitric oxide synthase, FasL, and p53 (data not shown). Selective protein immunoblot analysis determined that increases in mRNA were accompanied by proportional changes in protein expression. For example, the 8.5-fold increase in calreticulin mRNA was associated with an 8-fold increase in calreticulin protein (Figure 3C). Similarly, as a control for a nonupregulated mRNA, SERCA2a protein expression was unchanged (Figure 3D) and expression of the sodium/calcium exchanger protein remained unchanged out to 6 days after Dox administration (Figure 3E).
Calreticulin Upregulation May Contribute to the α5-1 Cardiac Phenotype
Previous work has shown that constitutive overexpression of calreticulin in neonatal cardiac myocytes leads to cardiomyopathy, heart block, and arrhythmias that are similar to those we previously found in mice with constitutive expression of α5-1.9,10 The electrophysiological abnormalities in the calreticulin-overexpressing mice were attributed to a decrease in Cx43 expression. Therefore, we looked for downregulation of Cx43 in our mice. Immunoblot analysis revealed a progressive reduction in Cx43 levels between days 2 and 4 after α5-1 induction (Figure 3F). This decrease in Cx43 protein was associated with reduced abundance of Cx43 mRNA, suggesting transcriptional regulation of this protein. Immunostaining of cardiac tissues revealed a substantial decrease in the number of Cx43-positive gap junctions (Figure 4A). Mean data from WT (n=6) and Tg (n=5) hearts showed ≈50% decrease in the density of Cx43 clusters (366±39 versus 706±43 clusters/unit myocyte area; P=0.0002) and the average size of the Cx43 clusters (0.50±0.02 versus 0.97±0.06 μm2; P<0.0001) in Tg versus WT hearts. A similar decrease in density and size of Cx43 clusters was observed in the left atria of Tg mice (supplemental Figure III). Transmission electron microscopy 3 days post-α5-1 induction revealed prominent architectural abnormalities of most intercalated discs (Figure 4B and 4C). In contrast to the changes in the intercalated discs, myocytes with intact membranes had normally organized sarcomeric structure (Figure 4B). However, many permeable myocytes had lost their normal sarcomeric structure and organization (data not shown).
To further investigate the relationship among Cx43 expression, electrocardiographic abnormalities, and cardiac function, we compared the number of Cx43 positive foci with QRS amplitude and fractional shortening in a group of mice with a range of quantitative changes in each of these parameters. To obtain this phenotypic spectrum, a group of mice (WT, n=6; Tg n=5) were treated with Dox for 4 days, and then mice were followed for 14 days after discontinuation of Dox. ECGs and echocardiograms were obtained at multiple time points and hearts were harvested at the end of the 14-day withdrawal period. We found that QRS amplitude and LV fractional shortening both declined in parallel during Dox treatment, reached a nadir at day 5 to 7 post-Dox, and then increased slightly thereafter (Figure 5A and 5B). There was a strong correlation between QRS amplitude and fractional shortening in individual mice (Figure 5C). Moreover, there was also a significant correlation between the number of Cx43-containing gap junctions and the QRS amplitude on the final ECG (Figure 5D). This suggests that a reduced number of gap junctions could contribute to the ECG abnormalities. To determine whether the alterations in Cx43 were specific to this Tg model, or whether such changes were a more general feature of cardiac disease, we assessed Cx43-containing gap junction density in a widely used pressure-overload model of murine cardiac hypertrophy (supplemental Figure IV). We found that the density of gap junctions per unit tissue area was decreased by 24% in aortic banded mice (591±20* versus 776±79 clusters/unit tissue area). However, the average size of each gap junction was not significantly decreased (0.76±0.05 versus 1.0±0.13 μm2; see the online data supplement for additional details).
Optical Mapping Reveals Impaired Electrical Propagation
Optical mapping was performed 3 days after α5-1 induction; Tg hearts exhibited greater total activation times (9.5±3.7 ms) versus WT hearts (6.7±1.8 ms; P=0.01). This finding implies ≈30% slower propagation in the α5-1 hearts. WT hearts showed a typical elliptical shape of the isochronal activation maps, with faster conduction along the direction of the fibers (Figure 6A). Hearts from α5-1 mice showed a more circular shape of the isochrones, implying diminished anisotropy of electrical propagation (Figure 6A). The anisotropic ratio (ie, the longitudinal versus the transverse axes of the 1-ms isochrone) was greater in WT than α5-1 hearts (1.9±0.4 versus 1.3±0.2; P=0.01). Action potential duration at 30%, 50%, and 70% recovery from the peak amplitude were not statistically different in WT and α5-1 hearts (6.7±1.9 versus 7.3±2.9, 12.5±3.1 versus 12.6±4.9, and 23.5±3.7 versus 24.2±7.9 ms, respectively). Action potential increase times were similar for WT (2.1±0.7) and Tg (2.2±0.9 ms) hearts.
Impaired Electrical Coupling Contributes to Reduced Contractility
Finally, to confirm the relationship between reduced Cx43 expression and reduced contractility, we compared electrically stimulated (ES) contractions of papillary muscles to rapid cooling contractures (RCCs) of the same muscles (WT, n=6; Tg, n=5). The contractions (twitches) induced by field stimulation rely on the swift transmission of electrical impulses between myocytes to produce coordinated muscle contraction. In contrast, RCC induces nearly complete release of Ca2+ from the sarcoplasmic reticulum independent of electrical coupling (Figure 6B). Therefore, RCC should produce a maximal contraction, even in the absence of effective electrical coupling between myocytes. Thus, an increased size of RCC relative to that of twitch would suggest an impairment of electrical coupling. Indeed, we found that α5-1 papillary muscles had reduced amplitude of both ES contractions and RCCs. However, the ES twitches were reduced to a greater degree, resulting in an increased ratio of RCC/ES amplitude in the α5-1 versus WT hearts (Figure 6C).
Integrin engagement with extracellular matrix ligands is required for cardiac myocyte hypertrophy in response to adrenergic stimulation.5 In addition, the abundance of different integrin subunits expressed by myocytes changes during cardiac hypertrophy.4,11 Although it seems likely that integrin signaling is involved in the development of cardiac hypertrophy, there are many gaps in our knowledge regarding the function of integrins in the adult myocardium.3 In a previous study, we constitutively overexpressed WT α5 integrin or the truncated α5-1 integrin in the mouse heart.9 Mice overexpressing the native integrin had no demonstrable cardiac phenotype. In contrast, expression of the α5-1-integrin subunit lacking the critical regulatory intracellular cytoplasmic domain resulted in uniform neonatal lethality. Now, we have used conditional transgenesis to determine the effects of expressing the same α5-1 integrin de novo in cardiac myocytes of adult mice. We found that α5-1-integrin expression rapidly resulted in electrophysiological abnormalities and profound ventricular contractile dysfunction. Thus, as illustrated in Figure 7, unregulated integrin activation may contribute to several components of pathological hypertrophy in adult myocardium.
One potential explanation for the observed phenotype of the mice in the present study is cardiac myocyte death induced by expression of the altered integrin. Using Evans Blue dye, we found progressively increased numbers of myocytes with sarcolemmal permeability beginning around day 3 after α5-1-integrin induction. Additionally, animals allowed to recover from α5-1 expression had increased cardiac fibrosis, further suggesting myocyte injury or death. Although this level of myocyte injury or death could cause severe global hypokinesis or the dramatic changes in surface ECG complexes, we believe this is unlikely, because the reductions in ECG amplitude preceded the marked increases in cell permeability by at least 1 to 2 days. Moreover, there are many animal models of cardiomyopathy and many human diseases that include variable degrees of myocardial cell death without significant reductions in QRS amplitude. Danik et al observed markedly reduced QRS amplitudes in mice with reduced Cx43 even in the absence of obvious myocardial injury or dysfunction.12 These pieces of evidence suggest that factors other than myocyte dropout are contributing to the severe contractile and electrical abnormalities in the α5-1 hearts.
The mechanism(s) of myocyte injury in the α5-1 mice are unclear. However, overexpressing the native α5 integrin did not cause a cardiac phenotype.9 Interestingly, the mutant α5-1 protein was not directly toxic to cardiac myocytes as high-level expression of α5-1 integrin in a mosaic pattern (approximately half of the myocytes), rather than in all myocytes, had no apparent pathophysiological consequences.9 In other words, cardiac myocytes expressing the α5-1 integrin were protected when adjacent cells did not express the mutant integrin. This suggests the possibility that cell/cell interactions mediate the effects of the unregulated integrin. The presence of myocytes not expressing the α5-1 integrin may allow the heart to maintain normal cardiac function because the unaffected cells could support the propagation of electrical, mechanical, or biochemical signals to the affected cells.
As in the current work, the dramatic reduction in QRS amplitude was also seen when the α5-1 integrin was expressed neonatally.9 This electrocardiographic phenotype resembled a phenotype attributable to reduced Cx43 expression in the mouse heart.12 Concomitant with decreased Cx43, that study demonstrated a slowing of conduction velocity, decreased QRS amplitude, arrhythmia inducibility, and sudden death. Indeed, we found similar sharp reductions in Cx43 in the α5-1 integrin-expressing hearts. Specifically, we observed a decrease in both density and size of Cx43 junctions that correlated to the decrease in fractional shortening and QRS amplitude. Additionally, optical mapping studies in our α5-1 hearts revealed loss of the normal anisotropic pattern of electrical activation (preferential conduction in the longitudinal fiber direction).
To determine whether the decreased Cx43 expression is common to other forms of cardiac pathology, we compared our α5-1 model with a widely studied model of compensated cardiac hypertrophy induced by 4 weeks of aortic banding. We found that sustained pressure-overload hypertrophy was also associated with a reduction in the density of Cx43-containing gap junctions (see online data supplement). The magnitude of the reduction was less severe in the banded mice than in the α5-1 mice. Interestingly, we saw a reduction in the size of Cx43 gap junctions in α5-1 mice but not after banding. Despite finding some differences in the 2 models we studied, our data and the bulk of published evidence suggest that both abundance and localization of Cx43 are consistently altered in experimental and human heart failure.13,14 Whether enhanced integrin signaling is the common mechanism remains to be determined. However, our data imply that integrin activation may be an initiating factor.
In contrast to the Cx43 knockout mice studied by Danik et al,12 we found reduced P wave amplitudes in the α5-1 mice. In addition, we found reduced atrial Cx43 density and impaired left atrial force development (see online data supplement). The preservation of p wave amplitude in the Cx43 knockout mice finding is likely explained by the fact that Cx43 is expressed at lower levels in the atria than the ventricle. The loss of Cx43 in the atria might be compensated for by increased Cx40 expression.15 The diminution of p wave amplitude in our mice suggests that enhanced integrin signaling affects the expression and/or localization of additional proteins in electrical and mechanical junctions (ie, Cx40) of atria. Additionally, the decrease in Cx43-containing gap junction size in the α5-1 mice indicates that there is a loss of cell/cell junctional integrity in the atria similar to that seen at the intercalated discs of the ventricle (see online data supplement).
One marked advantage of conditional transgenesis is the ability to study early changes in gene expression in response to α5-1-integrin induction rather than compensatory changes that occur in response to heart failure. In this study, we identified a remarkable increase in calreticulin mRNA and protein expression 2 days after transgene induction. This was of interest for several reasons. First, although calreticulin has a wide range of proposed functions, it is thought to have a role in mediating integrin signaling.16 Calreticulin binds to the GFFKR domain of the α-cytoplasmic domain of α5 integrin,17 the same domain deleted in α5-1 integrin. Calreticulin binding to this domain is postulated to activate the integrin, resulting in phosphorylation of calcium channels in some cell types.18,19 Second, calreticulin is essential to integrin-mediated adhesion.19 Furthermore, overexpression of calreticulin in neonatal hearts resulted in severe conduction defects and perinatal lethality, a phenotype resembling what we found with constitutive expression of the α5-1 integrin.10 Collectively, these data suggest that calreticulin may link integrin activation to the loss of Cx43 and subsequent abnormalities in conduction and contractility.
Although calreticulin is a Ca2+-binding protein, we found no evidence of reduced cytoplasmic Ca2+ cycling, or of changes in other key Ca2+ cycling proteins (SERCA2a, sodium/calcium exchanger). In keeping with these observations, the amplitude of contractions and Ca2+ transients was unchanged in myocytes from the α5-1 hearts. Thus, we do not think that impaired myocyte excitation/contraction coupling was a major factor in the contractile dysfunction seen in our mice. Our findings are somewhat reminiscent of those seen in some other Tg mice with cardiomyopathy.20,21 In those studies, mice were reported to have severe cardiomyopathy, but normal or enhanced Ca2+ transients in isolated myocytes. Hence, impaired force transmission and electrical conduction between or within myocytes may also contribute to whole-heart dysfunction.
Additional mechanisms may contribute to loss of Cx43 containing gap junctions. Mice with cardiac targeted activation of c-Jun N-terminal kinase (JNK) developed cardiomyopathy and sudden death.22 These mice had reduced Cx43 expression and loss of gap junctions. The latter changes were proposed to account for the contractile dysfunction. In support of a potential link between integrin activation and JNK expression, it was shown that mechanical strain caused a conformational change in αvβ3 integrin, resulting in integrin activation mediated by phosphatidylinositol 3-kinase. Subsequent engagement with extracellular ligands resulted in JNK activation.23 Thus, our model may mimic the effects of increased mechanical strain because the α5-1-integrin signals as if it were continuously activated. Yamada et al showed that there are distinct pathways that regulate Cx43 expression and mechanical junction proteins.24 Importantly, integrins are a common component of both pathways. In keeping with their hypothesis, our findings imply that both electrical and mechanical coupling between myocytes may be altered in the setting of enhanced integrin signaling.
To compare the effects of reduced myocardial contractility (ie, caused by myocyte death) versus loss of electrical coupling, we used an isolated papillary muscle model to measure the contractile force resulting from electrical stimulation or rapid cooling. Electrical stimulation requires intact electrical coupling between cells. In contrast, rapid cooling directly releases intracellular Ca2+ stores to stimulate contraction and does not require electrical depolarization or coupling. The amplitude of electrically stimulated contractions was reduced more than the amplitude of rapid cooling contractions in α5-1 hearts. This finding supports a role for electrical uncoupling in the contractile dysfunction.
Independent of the precise mechanism through which integrin regulation affects gap junctions, our results support the growing evidence that inherited or acquired connexinopathies may cause heart failure and arrhythmias.13,14,25–27 Efficient cardiac contraction requires tightly coordinated contraction of all myocytes that form the complex 3D structure of the organ. This is normally achieved via a specialized conducting system and a large number of low resistance gap junctions between adjacent myocytes. It is well known that modest alterations in electrical activation in the intact heart, as occurs with left bundle branch block, reduce overall cardiac function. Even at the individual myocyte level, synchronized release of Ca2+ from a large number of sites is crucial to maintain normal cellular function.28 Others have found that reduced electrical coupling between myocytes robs the heart of contractile efficiency.21,26 Therefore, it seems likely that myocyte loss and decreased cell-to-cell connections (gap junctions and intercalated discs) contributed to the contractile dysfunction in the α5-1 hearts by electrically and mechanically uncoupling the myocytes.
Taken together, our findings suggest a broader role than has previously been appreciated for integrin regulation in the development of cardiomyocyte injury and global cardiac dysfunction. Specifically, we believe there is a novel link between integrin regulation, connexin expression, electrical activation, and contractility in the heart.
We thank Chris Hunter, Iva Neveux, Ying Yang, Jennifer Hall, Judy Bradley, and the Mayo Clinic Scottsdale Transgenic Facility for technical assistance.
Sources of Funding
Funding provided by NIH grants R01-HL070511 and P50-HL52338 and a grant from the Department of Veterans Affairs.
Original received May 3, 2006; revision received October 30, 2006; accepted November 1, 2006.
Gullberg D, Velling T, Lohikangas L, Tiger CF. Integrins during muscle development and muscular dystrophies. Front Biosci. 1998; 3: 1039–1050.
Ross RS, Pham C, Shai SY, Goldhaber JI, Fenczik C, Glembotski CC, Ginsberg MH, Loftus JC. Beta1 integrins participate in the hypertrophic response of rat ventricular myocytes. Circ Res. 1998; 82: 1160–1172.
Bauer JS, Varner J, Schreiner C, Kornberg L, Nicholas R, Juliano RL. Functional role of the cytoplasmic domain of the integrin alpha 5 subunit. J Cell Biol. 1993; 122: 209–221.
Wu C, Bauer JS, Juliano RL, McDonald JA. The alpha 5 beta 1 integrin fibronectin receptor, but not the alpha 5 cytoplasmic domain, functions in an early and essential step in fibronectin matrix assembly. J Biol Chem. 1993; 268: 21883–21888.
Valencik ML, McDonald JA. Cardiac expression of a gain-of-function alpha(5)-integrin results in perinatal lethality. Am J Physiol Heart Circ Physiol. 2001; 280: H361–H367.
Terracio L, Rubin K, Gullberg D, Balog E, Carver W, Jyring R, Borg TK. Expression of collagen binding integrins during cardiac development and hypertrophy. Circ Res. 1991; 68: 734–744.
Danik SB, Liu F, Zhang J, Suk HJ, Morley GE, Fishman GI, Gutstein DE. Modulation of cardiac gap junction expression and arrhythmic susceptibility. Circ Res. 2004; 95: 1035–1041.
Saffitz JE, Kleber AG. Effects of mechanical forces and mediators of hypertrophy on remodeling of gap junctions in the heart. Circ Res. 2004; 94: 585–591.
Dedhar S, Gray V. Isolation of a novel integrin receptor mediating Arg-Gly-Asp-directed cell adhesion to fibronectin and type I collagen from human neuroblastoma cells. Association of a novel beta 1-related subunit with alpha v. J Cell Biol. 1990; 110: 2185–2193.
Kwon MS, Park CS, Choi K, Ahnn J, Kim JI, Eom SH, Kaufman SJ, Song WK. Calreticulin couples calcium release and calcium influx in integrin-mediated calcium signaling. Mol Biol Cell. 2000; 11: 1433–1443.
Su Z, Yao A, Zubair I, Sugishita K, Ritter M, Li F, Hunter JJ, Chien KR, Barry WH. Effects of deletion of muscle LIM protein on myocyte function. Am J Physiol Heart Circ Physiol. 2001; 280: H2665–H2673.
Chu G, Carr AN, Young KB, Lester JW, Yatani A, Sanbe A, Colbert MC, Schwartz SM, Frank KF, Lampe PD, Robbins J, Molkentin JD, Kranias EG. Enhanced myocyte contractility and Ca2+ handling in a calcineurin transgenic model of heart failure. Cardiovasc Res. 2002; 54: 105–116.
Petrich BG, Gong X, Lerner DL, Wang X, Brown JH, Saffitz JE, Wang Y. c-Jun N-terminal kinase activation mediates downregulation of connexin43 in cardiomyocytes. Circ Res. 2002; 91: 640–647.
Katsumi A, Naoe T, Matsushita T, Kaibuchi K, Schwartz MA. Integrin activation and matrix binding mediate cellular responses to mechanical stretch. J Biol Chem. 2005; 280: 16546–16549.
Yamada K, Green KG, Samarel AM, Saffitz JE. Distinct pathways regulate expression of cardiac electrical and mechanical junction proteins in response to stretch. Circ Res. 2005; 97: 346–353.
Gutstein DE, Morley GE, Vaidya D, Liu F, Chen FL, Stuhlmann H, Fishman GI. Heterogeneous expression of Gap junction channels in the heart leads to conduction defects and ventricular dysfunction. Circulation. 2001; 104: 1194–1199.
Ai X, Pogwizd SM. Connexin 43 downregulation and dephosphorylation in nonischemic heart failure is associated with enhanced colocalized protein phosphatase type 2A. Circ Res. 2005; 96: 54–63.
Litwin SE, Zhang D, Bridge JH. Dyssynchronous Ca(2+) sparks in myocytes from infarcted hearts. Circ Res. 2000; 87: 1040–1047.