Myofilament Calcium Sensitivity in Skinned Rat Cardiac Trabeculae
Role of Interfilament Spacing
The increase in myofilament Ca2+ responsiveness on an increase in sarcomere length (SL) is, in part, the cellular basis for Frank-Starling’s law of the heart. It has been suggested that a decrease in myofilament lattice spacing (LS) in response to an increase in SL underlies this phenomenon. This hypothesis is supported by previous studies in which reduced muscle width induced by osmotic compression was associated with an increase in Ca2+ sensitivity, mimicking those changes observed with an increase in SL. To evaluate this hypothesis, we directly measured LS by synchrotron x-ray diffraction as function of SL in skinned rat cardiac trabeculae bathed in 0% to 6% dextran solutions (MW 413 000). We found that EC50, [Ca2+] at which force is half-maximal, at SL between 1.95 and 2.25 μm did not vary in proportion to LS when 3% or 6% dextran solutions were applied. We also found that moderate compression (1% dextran) of skinned trabeculae at SL=2.02 μm reduced LS (LS=42.29±0.14 nm) to match that of uncompressed fibers at a long SL (SL=2.19 μm; LS=42.28±0.15 nm). Whereas increasing SL from 2.02 to 2.19 μm significantly increased Ca2+ sensitivity as indexed by the EC50 parameter (2.87±0.11 μmol/L to 2.52±0.12 μmol/L), similar reduction in myofilament lattice spacing achieved by compression with 1% dextran did not alter Ca2+ sensitivity (2.87±0.10 μmol/L) at the short SL. We conclude that alterations in myofilament lattice spacing may not be the mechanism that underlies the sarcomere length–induced alteration of calcium sensitivity in skinned myocardium.
The heart regulates ventricular output in response to changes in ventricular filling, a mechanism known as Frank-Starling’s law of the heart. This mechanism has two components1: a fast component caused by changes in myofilament calcium sensitivity,2,3⇓ and a slow component caused by a change in the calcium transient.4 In this study, we are concerned only with the fast component that is responsible for the beat-to-beat changes in ventricular output in response to ventricular filling.
It is not clear how the information concerning sarcomere length (SL) is transmitted to the myofilaments. Although thin filament proteins are not ruled out as potential regulators of length-dependent activation,5 a more unifying hypothesis suggests that the lateral spacing between actin and myosin influences crossbridge reactivity independent of the primary events involving Ca2+ binding and subsequent thin filament activation.6–8⇓⇓ As the interfilament spacing is reduced with increased SL,9–11⇓⇓ it is hypothesized that the probability of strong crossbridge formation is enhanced. Facilitated by the highly cooperative nature of myofilament activation, the sensitivity of the myofilaments to Ca2+ would thus be increased at longer SL. In support of this hypothesis, shrinkage of the myofilament lattice by osmotic compression using high molecular weight moieties has been shown to increase myofilament Ca2+ sensitivity at short SL, mimicking the impact of increased SL on myofilament lattice spacing and Ca2+ sensitivity.6–8,12–14⇓⇓⇓⇓⇓ The extent of shrinkage of the myofilament lattice by osmotic compression in those studies, however, was estimated from the reduction in muscle width. Muscle width in skinned preparations may not be strictly proportional to myofilament lattice spacing,15 an observation that we confirm in the present study in skinned isolated rat myocardium.
Accordingly, we set out to obtain a direct comparison between myofilament Ca2+ sensitivity and myofilament lattice spacing in skinned rat cardiac trabeculae under conditions of varying degrees of osmotic compression induced by dextran. Measurements of Ca2+ sensitivity were obtained at 3 SLs, and these measurements were correlated with direct measurement of myofilament lattice spacing obtained via synchrotron x-ray diffraction. We found that Ca2+ sensitivity did not vary in proportion to myofilament lattice spacing in osmotically compressed trabeculae. Furthermore, when the myofilament lattice was osmotically compressed to match the reduced lattice spacing as induced by increased SL, myofilament Ca2+ sensitivity was not affected. These findings suggest, therefore, that alterations in myofilament lattice spacing may not be the primary mechanism that underlies the sarcomere length–induced alteration of calcium sensitivity in skinned myocardium. A preliminary report of these studies has been published previously.16
Materials and Methods
Muscle Preparation and Experimental Protocol
Rats (LBNF-1 220 to 280 g) were anesthetized (sodium pentobartbital 50 mg/kg BW IP injection), and the hearts were rapidly excised and retrogradely perfused with a modified Krebs-Henseleit (K-H) solution with propranolol (5 μmol/L), to block nonspecific β-adrenergic activation, and carbamyl choline (10 μmol/L), to enhance phosphatase activity.17 Trabeculae were dissected from the right ventricular free wall and demembranated overnight with 1% Triton X-100 in standard relaxing solution (EGTA=10 mmol/L; ionic strength=180 mmol/L). All solutions contained the following (in mmol/L): 10 phosphocreatine, 100 N,N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid (BES), 0.1 leupeptin, 0.1 phenylmethylsulfonyl fluoride (PMSF), 1 dithiothreitol (DTT), and 4 U/mL creatine phosphokinase. The free Mg2+ and Mg-ATP concentration were calculated at 1 and 5 mmol/L, respectively. Relaxing and activating solutions were appropriately mixed to obtain a range of free [Ca2+] at 15°C. Trabeculae (n=22; 0% dextran) were 2 to 3 mm in length, 136±10 μm in width, and 104±10 μm in thickness. Ca2+-force relationships were determined at three sarcomere lengths (SL=1.95 μm, 2.10 μm, and 2.25 μm). Muscle length was adjusted to maintain constant SL throughout the contraction.18 SL was determined from the first order He-Ne laser light diffraction band. Steady-state force was recorded at each SL (Figure 1). Muscles that did not maintain 90% of maximal activation on completion of the experiment or that lost the laser diffraction band were discarded. In some experiments, 1%, 3%, or 6% (wt/volume) dextran (Sigma, 413 kDa) was added to the solutions (equilibration >30 minutes). X-ray diffraction experiments were performed as described previously.11
Each individual Ca2+-force relationship was fit to a modified Hill equation: Frel=[Ca2+]n/(EC50n+[Ca2+]n); Frel=relative force, EC50=[Ca2+] at which force is half-maximal, n=Hill coefficient. Separation of the equatorial 1,0 or 1,1 x-ray diffraction reflections was measured from detector images using the program FIT2D19 and converted to d10 lattice spacing using Bragg’s Law, which can then be converted to the interthick filament spacing by multiplying d10 by 2/√3. The differences among EC50, ΔEC50, LS, and ΔLS for each group at each SL were analyzed with a one-way ANOVA followed by a Student’s t test with a post hoc Bonferroni correction to assess differences among mean values. All data are shown as mean±SEM.
Myofilament Ca2+ Sensitivity
To assess the effect of sarcomere length on myofilament Ca2+ sensitivity, Ca2+-force relationships were determined at 3 SLs (1.95, 2.10, and 2.25 μm) in each of the groups studied (0%, 3%, and 6% dextran solution). Original recordings of force from a typical experiment are illustrated in Figure 1. Steady-state force was measured at each SL during each activating cycle; the zero force level was identified by instituting a quick ramp shortening in muscle length just before introduction of relaxing solution into the bath (active force was calculated as the difference between total force and relaxed force). Figure 1A shows a series of such quick length-release steps recorded at varying levels of activation (Ca2+ in μmol/L indicated on the right of each tracing) for trabeculae bathed with a 0%, 3%, or 6% dextran solution (SL=2.25 μm). A final maximal activation was administered to assess muscle preparation integrity. The relationship between active force development and Ca2+ concentration ([Ca2+]) was fit to the Hill equation (see Materials and Methods) at each dextran concentration as is illustrated in Figure 1B. Application of either 3% or 6% dextran induced a significant increase in myofilament Ca2+ sensitivity (Figure 1B and pooled data in Figure 2). The average parameters of the Hill fit obtained in each individual trabecula are summarized in the Table. Increasing SL from 1.95 to 2.10 μm and then to 2.25 μm caused a leftward shift of the Ca2+-force relationship (Figure 2) and a significant decrease in the EC50 parameter for each experimental group (Table). Addition of 3% dextran (wt/volume) significantly increased Ca2+ sensitivity at all 3 SLs from the control (0% dextran) group (Figure 2A). Further addition of dextran from 3% to 6% slightly increased Ca2+ sensitivity at all SLs, albeit not significantly (Figure 2B; Table). Neither changes in SL nor changes in dextran concentration had an appreciable affect on the steepness of the Ca2+-force relationship (Hill coefficient; Table).
Although application of dextran caused a general increase in myofilament Ca2+ sensitivity at any SL, the length-dependent shift of the Ca2+-force relationship appeared not to be altered by dextran treatment. We quantified this by computing ΔEC50, the difference between EC50 at the short and long sarcomere length in each muscle (SL=1.95 and 2.25 μm, respectively). Indeed, ΔEC50 (in μmol/L) was unaffected by dextran treatment (Table and Figure 4B). Previous studies have employed pCa50 (−log[EC50]) to index calcium sensitivity.6–8,12–14⇓⇓⇓⇓⇓ Employing this method also revealed no statistically significant difference in length-dependent activation (Table). Ca2+-saturated maximum force decreased 20%, on average, with a reduction in SL from 2.25 to 1.95 μm in each experimental group, consistent with previous reports.3
Application of dextran may affect maximum force development.8,20,21⇓⇓ Accordingly, we directly determined the effect of dextran application on maximum force in a separate group of trabeculae at SL=2.20 μm (n=6). We found that consecutive application of 1%, 3%, and 6% dextran increased maximal force development, on average, by 13.3±4.0%, 12.2±4.6%, and 11.9±5.2%, respectively, compared with that in trabeculae in the absence of dextran (P<0.01). This observation is consistent with previous reports.7,8,21⇓⇓
Osmotic Compression and Myofilament Lattice Spacing
We simultaneously measured myofilament lattice spacing (LS) and SL in skinned cardiac trabeculae (n=8 to 10 for each group studied) using synchrotron x-ray diffraction.11 Figure 3 illustrates typical x-ray diffraction patterns obtained in trabeculae at SL=2.1 μm in relaxing solutions containing 0%, 3%, or 6% dextran. There was an evident outward displacement of the reflections with increasing dextran concentration in the solution, indicative of a reduction in myofilament lattice spacing. Figure 4A shows that LS was an inverse function of SL at each dextran concentration (n=8 to 10). Furthermore, LS was reduced at every SL in trabeculae immersed in a 3% dextran solution compared with control trabeculae, and it was further reduced in trabeculae that were bathed in a 6% dextran solution. At SL=2.10 μm, LS was significantly reduced (P<0.001) with a 3% dextran solution (37.9±0.19 nm) from control (42.6±0.20 nm); adding 6% dextran to the relaxing solution further reduced LS (34.6±0.12 nm) significantly from both the control and the 3% dextran group at this SL.
The impact of changes in SL on myofilaments lattice spacing, as reflected by the slope of the relationships displayed in Figure 4A, was markedly reduced at 3% and 6% dextran. That is, LS became a less sensitive function of SL with increasing amounts of dextran in the solution. To quantify this observation, we defined the decrease in LS when SL was increased from 1.95 to 2.25 μm as ΔLS in each muscle. The average ΔLS, as well as the ΔEC50 parameters are shown in Figure 4B. ΔLS was greatest in the control trabeculae (1.67±0.16 nm), was markedly reduced in trabeculae bathed in the 3% dextran solution (0.70±0.04 nm), and reduced even further in trabeculae bathed in a 6% dextran solution (0.47±0.04 nm). Each of these values was significantly different from the other (P<0.01). Thus, length-dependent activation, as indexed by either the ΔEC50 or the ΔpCa50 parameter, was unaffected by dextran compression, whereas changes in myofilament lattice spacing (ΔLS) induced by changes in SL were significantly reduced on compression by 3% or 6% dextran. LS remained inversely proportional to SL under all levels of dextran compression. Therefore, these results indicate that the impact of interfilament spacing on myofilament Ca2+ sensitivity is not constant but rather varies with the overall compression state of the cardiac sarcomere.
Ca2+ Sensitivity With Moderate Amounts of Osmotic Compression
At the outset of the present study, we choose dextran concentrations consistent with previous reports that used either 2.5%7 or between 0% to 5%8,14,21⇓⇓ dextran concentrations. Indeed, the Ca2+ sensitivity of control trabeculae at the long SL was approximately matched by application of 3% dextran at the short SL (see Table). We found, however, that under our conditions, both 3% and 6% dextran reduced myofilament lattice spacing beyond the physiological effect of an increase in SL in control, uncompressed skinned trabeculae. That is, application of 3% dextran compressed the myofilament lattice beyond that observed at even the largest SL under control conditions (Figure 4). Therefore, to specifically test the notion that lattice spacing is the primary determinant for length-dependent myofilament Ca2+ sensitivity in skinned myocardium, we performed an additional set of experiments. The results are summarized in Figure 5. Skinned trabeculae were treated with 1% dextran to osmotically compress the myofilament lattice at a short SL so as to precisely match that of control, uncompressed trabeculae at a longer SL. Treatment of skinned cardiac trabeculae at a SL=2.02 μm with 1% dextran reduced LS to 42.29±0.14 nm, a value that was equivalent to LS at SL=2.19 μm in the untreated trabeculae (42.28±0.15 nm). Note that the LS at SL=2.02 μm with 0% dextran was 43.02±0.21 nm. Average Ca2+ sensitivities, as indexed by the EC50 parameter, under these conditions are shown in the bar graph inset of Figure 5. Increasing SL from 2.02 μm to 2.19 μm resulted in a significant decrease in EC50, consistent with the results shown in Figure 2 and Table. However, application of 1% dextran at SL=2.02 μm did not significantly affect Ca2+ sensitivity, despite the reduction in myofilament lattice spacing (whether indexed by EC50 or pCa50).
The notion that the interfilament spacing constitutes the molecular length sensor is a prominent feature of current theories regarding the mechanism that underlies Frank-Starling’s law of the heart.6–8,12–14,22⇓⇓⇓⇓⇓⇓ The most compelling evidence in support of this theory comes from studies in which shrinkage of the myofilament lattice in skinned muscle preparations by osmotic compression is accompanied by increased Ca2+ responsiveness of the contractile apparatus.6–8,12,13⇓⇓⇓⇓
The impact of sarcomere length (SL) on myofilament Ca2+ sensitivity, as indexed by either by the ΔEC50 or the ΔpCa50 parameter (Table), was unaffected by application of dextran, whereas the slope of the relationship between myofilament lattice spacing and sarcomere length (ΔLS) was considerably reduced with increasing dextran concentration in the bathing solutions (Figure 4). This reduction in the length dependence of myofilament lattice spacing would be expected to lead to a reduction of the length dependence of myofilament activation. Osmotic compression in this study, however, did not affect the length dependence of the myofilament Ca2+ sensitivity in rat trabeculae, consistent with a recent report on rat ventricular myocytes.23 Other studies have reported a reduced length dependency on compression of the myofilament lattice by dextran.8,12⇓ In those studies, Ca2+ sensitivity was indexed by pCa50. It should be noted that the relation between pCa and [Ca2+] is nonlinear; therefore, the choice of parameter to index myofilament Ca2+ sensitivity could potentially affect data interpretation. Because reaction rates are usually proportional to the free concentration of a substrate, we believe that the use of the EC50 parameter to index myofilament Ca2+ sensitivity is more appropriate than pCa50. However, this factor alone cannot explain the different findings because neither the ΔEC50 nor the ΔpCa50 parameter was significantly affected by osmotic compression in the present study. The precise reason for the discrepancies between previous studies8,12⇓ and our current study is unclear; a species specific difference in the effect of dextran on length-dependent activation may underlie these results. Clearly, this issue requires further study.
Our data, using 3% and 6% dextran suggest that, at a minimum, the interfilament spacing theory must be amended to include a variable impact of myofilament lattice spacing on myofilament Ca2+ sensitivity, depending on the overall extent of compression of the myofilament lattice. The increase in Ca2+ sensitivity on the decrease in myofilament lattice spacing in the 3% dextran solution is consistent with the interfilament spacing hypothesis (Figures 2 and 4⇑; Table). However, the further reduction in myofilament lattice spacing in 6% dextran without a further increase in Ca2+ sensitivity cannot readily be explained by this theory. The effect of dextran treatment on myofilament Ca2+ sensitivity has previously been shown to be limited to approximately 5% dextran wt/volume, with higher concentrations being without an effect on Ca2+ sensitivity.20,24⇓ Moreover, when dextran concentrations exceeded 10%, myofilament Ca2+ sensitivity may even be reduced.8 Hence, those data and our present data are consistent with the notion that the relation between interfilament spacing and myofilament Ca2+ sensitivity is highly nonlinear. This concept is illustrated in Figure 6 where Ca2+ sensitivity is plotted as function of myofilament lattice spacing. It is apparent that the effect of dextran treatment on Ca2+ sensitivity was limited to a narrow range of dextran concentrations, 1% to 3% under our conditions, whereas the myofilament lattice continued to shrink with increasing concentration of dextran in the solution. To differentiate whether the changes in Ca2+ sensitivity with dextran compression were the specific result of changes in lattice spacing, rather than some other, coincidental effect of osmotic compression, we performed a more critical test using 1% dextran. If interfilament spacing were the major determinant of crossbridge reactivity, then the Ca2+ sensitivity of moderately compressed trabeculae at the short SL should be similar to that of uncompressed trabeculae at the long SL given an equivalent lattice spacing. The data in Figures 5 and 6⇓ show that this was not the case, again regardless whether pCa50 or EC50 was used to index Ca2+ sensitivity. Hence, our data are not consistent with the notion that changes in myofilament lattice spacing per se upon changes in SL within the physiological range are responsible for the increase in myofilament Ca2+ sensitivity in skinned cardiac muscle.
This conclusion differs from previous studies that provided support for the interfilament spacing hypothesis.7,8,21⇓⇓ In those studies, however, changes in fiber width on either dextran compression or changes in SL were used to infer changes in myofilament lattice spacing. We found that the effect of different amounts of osmotic compression on muscle width is not equivalent to the effect on lattice spacing, as has been observed previously in skeletal muscle,15 albeit not uniformly.22 This is illustrated in Figure 7, where muscle width (as measured by microscopy) is plotted as a function of myofilament lattice spacing (as measured by x-ray diffraction) under conditions of varied SL and dextran concentration. The solid line (line of identity) shows the relationship that would be expected if muscle width were directly proportional to myofilament lattice spacing. These data show that application of 3% or 6% dextran reduced muscle width to a greater extent than myofilament lattice spacing. More importantly, SL change had a greater impact on muscle width than on myofilament lattice spacing. The deviation from proportionality increased with increases in dextran concentration. The mechanism that underlies these observations is unclear, but may result from a differential effect of dextran compression on muscle diameter and muscle width, such that compression caused the muscle to become more circular in cross-sectional shape. Alternatively, there may be a specific effect of dextran on the myofilament lattice alone, whereas changes in SL appear to affect all structures of the skinned trabecula, that is, both myofilament lattice spacing and extra-sarcomeric proteins. Nevertheless, under our conditions, to achieve a muscle width at the short SL equal to that at the long SL, approximately 2% to 3% dextran in the bathing solutions would have been required. Considering the nonlinear effect of dextran treatment on Ca2+ sensitivity (Figure 6), it is also apparent that 2% to 3% dextran treatment of skinned trabeculae would, in fact, be expected to significantly increase Ca2+ sensitivity, as indeed has been observed by the Moss group.7 By employing direct measurement of myofilament lattice spacing in skinned isolated cardiac trabeculae using x-ray diffraction, the confounding factor of using fiber width to estimate myofilament lattice spacing was eliminated in the present study.
Why does Ca2+ sensitivity increase with application of dextran (Figures 1, 2, and 6⇑⇑; Table)? It may be argued that osmotic compression increases the number of attached crossbridges consistent with the increase in maximum force following dextran treatment. Yet, despite a significant increase in maximum force development on application of 1% dextran, there was no effect on myofilament Ca2+ sensitivity. Thus, osmotic compression at a short SL is not functionally equivalent to lengthening a sarcomere. Our data suggests that the effect of dextran treatment on Ca2+ sensitivity may be independent of changes in myofilament lattice spacing. Studies of skeletal muscle25 suggest that there is a collapse of myosin heads against the myosin backbone over a range of osmotic pressures corresponding to about 1% to 3% dextran, ie, a structural effect of osmotic compression that is independent of lattice spacing. This could lead to the observed lack of change in Ca2+ sensitivity on increasing the dextran concentration above 3% because a decrease in crossbridge radius might be expected to counteract the reduction in lattice spacing. In principle, evidence for such a change in myosin structure could be obtained by analysis of the x-ray equatorial reflection intensities.25 Whereas the minimal x-ray exposure times used to enhance sample lifetime in the present studies precluded such analysis, preliminary estimates of the 1,1 to 1,0 first order reflection intensity ratios in our study were consistent with previous findings in skeletal muscle.25 Whether or not this mechanism underlies the increase in myofilament Ca2+ sensitivity on application of 3 to 6% dextran must await studies optimized for this purpose. Clearly, the possibility of structural changes in addition to the lattice spacing be must taken into account when interpreting any study in which dextran is employed to compress the myofilament lattice.
We have previously shown a similar dependence of interfilament spacing on sarcomere length in intact relaxed rat myocardium as compared with that in relaxed skinned trabeculae, albeit at a reduced interfilament spacing.11 Our study does not directly address whether such a change in interfilament spacing underlies the length-dependent change in myofilament Ca2+ sensitivity in intact myocardium.26 It has been observed that the effect of SL on Ca2+ sensitivity appears to be less in intact myocardium compared with skinned myocardium,26 despite the similar impact of SL changes on interfilament spacing.11 Hence, there appear to be differences between intact and skinned cardiac muscle, such that our results on skinned myocardium may not be applicable to intact heart. Nevertheless, if altered myofilament lattice spacing on a change in SL in skinned myocardium is not responsible for changes in Ca2+ sensitivity, as suggested here, what mechanism could account for length-dependent activation? It has been suggested that changes in Ca2+ sensitivity with length involve modulation of cooperative crossbridge binding to the thin filament.27 Although sarcomere geometry and filament overlap may contribute to the availability of additional crossbridges along the ascending limb of the length-tension relationship,28 it alone cannot account for the magnitude of the increase in force development.3,7,29⇓⇓ It has been suggested that thin filament isoform proteins regulate the response to changes in length by some as of yet undetermined mechanism.5 Among alternative, plausible mechanisms that may be considered are those involving other sarcomeric proteins, such as the giant sarcomeric protein titin, which spans the entire sarcomere from Z-disk to Z-disk with interaction sites on both actin, myosin, and thin filament regulatory proteins.30–33⇓⇓⇓
In conclusion, we found that myofilament Ca2+ sensitivity did not vary in proportion to myofilament lattice spacing in osmotically compressed trabeculae. Furthermore, when the myofilament lattice was osmotically compressed so as to match the reduced lattice spacing as induced by increased sarcomere length, myofilament Ca2+ sensitivity was not significantly affected. These findings suggest that alterations in myofilament lattice spacing may not be the fundamental mechanism that underlies the sarcomere length–induced alteration of calcium sensitivity in skinned myocardium.
This work was supported, in part, by the Cardiovascular Sciences Program Training Grant T32 07692 (J.P.K.), a national Grant-in-Aid from the American Heart Association 9950459N (T.C.I.), and NIH grants HL52322 and HL62426 (P.P.d.T.). Use of the Advanced Photon Source was supported by the US Department of Energy, Basic Energy Sciences, Office of Energy Research, under Contract No. W-31-109-ENG-38. BioCAT is a US NIH supported Research Center (RR08630). We thank Darold Perry, Dr Robert Fischetti, and Dr Karen Bischoff for assistance with the x-ray diffraction studies.
Original received December 27, 2000; resubmission received October 22, 2001; revised resubmission received November 7, 2001; accepted November 7, 2001.
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