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. 2000 Dec 1;529(Pt 2):405–411. doi: 10.1111/j.1469-7793.2000.00405.x

Cytochalasin D reduces Ca2+ sensitivity and maximum tension via interactions with myofilaments in skinned rat cardiac myocytes

S C Calaghan *, E White *, S Bedut †, J-Y Le Guennec *
PMCID: PMC2270202  PMID: 11101650

Abstract

  1. The F-actin disrupter cytochalasin D depresses cardiac contractility, an effect previously ascribed to the interaction of cytochalasin D with cytoskeletal actin. We have investigated the possibility that this negative inotropic effect is due to the interaction of cytochalasin D with sarcomeric actin of the thin filament.

  2. Confocal images of Triton X-100-skinned myocytes incubated with a fluorescent conjugate of cytochalasin D revealed a longitudinally striated pattern of binding, consistent with a myofibrillar rather than cytoskeletal structure.

  3. Tension–pCa relationships were determined at sarcomere lengths (SLs) of 2.0 and 2.3 μm following 2 min incubation with 1 μm cytochalasin D. Cytochalasin D significantly reduced the pCa for half-maximal activation (pCa50) at both SLs. The shift in pCa50 was significantly greater at a SL of 2.3 μm compared with that at a SL of 2.0 μm. Cytochalasin D had no effect on the Hill co-efficient at either SL.

  4. Cytochalasin D significantly reduced the maximum tension at both SLs.

  5. We suggest that the length-dependent decrease in myofilament Ca2+ sensitivity in response to cytochalasin D is due to a decrease in the affinity of troponin C for Ca2+.

  6. Cytochalasin D has been used for many years as the agent of choice for disruption of cytoskeletal actin. However, we have demonstrated for the first time an interaction of cytochalasin D with sarcomeric actin of the thin filament, which can account for the effects of cytochalasin D on cardiac contractility.

Within the cardiac cell, actin is found both in the thin filament and in cytoskeletal microfilaments. Sarcomeric actin in the thin filament, through interaction with myosin, is responsible for contraction. Cytoskeletal actin has been linked with many processes including integration of intracellular space, determination of the shape of T-tubules and intracellular organelles, and modulation of membrane-bound ion channels and exchangers (Moses & Delcarpio, 1983; Messina & Lemanski, 1989; Bennett, 1990; Hilgemann, 1997). Cytochalasin D has been used as a tool to investigate specifically the role of cytoskeletal actin in a variety of cell types. Cytochalasin D binds to the barbed end of the actin filament at which net polymerisation occurs, thereby preventing addition of actin monomers, and may also bind to a subunit in the interior of an actin filament and ‘sever’ the filament in two (Brenner & Korn, 1979; Cooper, 1987). It has been thought that the relative stability of sarcomeric actin protects it from the effects of cytochalasin D.

In studies using single cardiac cells, isolated muscle and the intact perfused heart, exposure to cytochalasin D for between 2 min and 4 h has been shown to depress cardiac contractility (Maltsev & Undrovinas, 1997; Skobel & Kammermeier, 1997; Biermann et al. 1998; Howarth et al. 1998). This effect of cytochalasin D has been dissociated from changes in the L-type Ca2+ current, changes in action potential characteristics and a decrease in amplitude of the intracellular Ca2+ transient (Biermann et al. 1998; Howarth et al. 1998; Undrovinas & Maltsev, 1998). Cytochalasin D has been shown to slow both the rising phase and decay of Ca2+ transients in rat ventricular myocytes, and a modification of sarcoplasmic reticulum function by cytochalasin D via cytoskeletal actin has been suggested (Undrovinas & Maltsev, 1998).

However, although the negative inotropic effect of cytochalasin D has been interpreted in terms of a cytoskeletal action, Howarth et al. (1998) demonstrated a shift in the phase-plane relationship between contraction and Ca2+ in single cardiac myocytes following 2 min perfusion with cytochalasin D. This observation is consistent with a decrease in sensitivity of the myofilaments to Ca2+ and implies an interaction of cytochalasin D with sarcomeric actin. It is interesting to note, therefore, that the cyclic peptide phalloidin, which also interacts specifically with actin, but in contrast to cytochalasin D acts to stabilise the actin filament (Cooper, 1987), has been shown to increase myofilament Ca2+ sensitivity (Bukatina et al. 1995).

The aim of the present study was to characterise the acute effects of cytochalasin D on sarcomeric actin filaments. We report for the first time that the effects of cytochalasin D on contraction and the kinetics of the Ca2+ transient can be explained by an effect on the sarcomeric actin. Our data allow us to speculate as to the mechanism by which cytochalasin D exerts its effects on the thin filament.

METHODS

Preparation of skinned cardiac myocytes

Experimental procedures conformed to national guidelines. Single ventricular myocytes were isolated enzymatically according to the method of Pucéat et al. (1990). Briefly, male Wistar rats (200–250 g) were killed by cervical dislocation and their hearts were removed quickly and mounted on a Langendorff apparatus. Hearts were perfused retrogradely with calcium-free physiological saline solution (PSS; see below) at 37°C. When the coronary circulation had cleared of blood, PSS supplemented with 1.3–1.4 mg ml−1 collagenase (Worthington type 4), 0.1 mm CaCl2 and 0.15 mm EGTA was recirculated for 30–40 min. The left ventricle was gently dissociated by agitation in PSS. Following isolation, myocytes were suspended in PSS containing 0.3 mm Ca2+ and 0.25% bovine serum albumin (BSA), pH adjusted to 7.6 with NaOH.

Cells were rapidly skinned by incubation in relaxing solution (pCa 9; see below) containing 0.3% (v/v) Triton X-100 at 4°C. The cell suspension was gently agitated at room temperature for 6 min. Cells were then rinsed twice in relaxing solution and maintained at 4°C for up to 10 h (Cazorla et al. 1999).

Fluorescence confocal microscopy

Skinned myocytes were incubated with 1 μm cytochalasin D- BODIPY FL conjugate (Molecular Probes Europe) in relaxing solution and imaged by confocal laser microscopy (Leica True Confocal Scanner SP). An argon laser provided excitation light at 488 nm and emissions were collected between 500 and 600 nm. Sections of ≈2 μm thickness were taken longitudinally through the cell. Because of the time taken to prepare cells for confocal microscopy, images were taken between 5 and 15 min following incubation with fluorescent cytochalasin D.

Mechanical recording and sarcomere length measurement

The experimental apparatus and cell attachment procedure were adapted from those described previously (Pucéat et al. 1990; Cazorla et al. 1999). Experiments were carried out in Petri dishes pre-treated with PSS containing 0.25% BSA to prevent adherence of the cells to the bottom of the chamber. Myocytes were observed through an inverted microscope (Nikon Diaphot 300, France). Each end of a skinned myocyte was attached to a thin glass rod (length 20–30 mm; diameter 1 mm), which was drawn to a fine point. Rods were attached with optical glue (NOA 60, Norland Products Inc., North Brunswick, NJ, USA) which was polymerised by UV illumination for 3 min to give a strong attachment with low compliance. To allow the recording of tension, one rod was attached to a piezoresistive strain gauge (Model AE 801, SensoNor a.s., Horten, Norway) and connected to an amplifier to yield a sensitivity of 55 mV μN−1 with a noise level below 0.15 μN. Compliance of the strain gauge was 0.165 μm μN−1 and unloaded resonance frequency was 500 Hz. Tension was recorded using a Grass chart recorder (Model 7400). The cell was positioned at the tip of a conical microcapillary that received the outlet of 10 microcapillaries connected to 5 ml syringes (flow rate of 200 μl min−1). Experiments were performed at room temperature (25°C). Cells were studied at a sarcomere length of either 2.0 (± 0.1) or 2.3 (± 0.1) μm (set length ± predicted inhomogeneity based on Fourier analysis; see below). Cells were stretched by means of a hydraulic micromanipulator (MO-103, Charenton, Narishige) and sarcomere length was determined on-line using a fast-Fourier transformation of a video image of the cell using sections of cell between the force transducer (Gannier et al. 1993). Sarcomere length was monitored on-line throughout all experiments at a rate of 50 Hz. Cells were exposed to the full activating solution (pCa 4.5) to test the quality of the attachment and recovery to baseline. A reduction in maximum tension may occur following repeated exposure to pCa 4.5, and therefore the number of tension-pCa curves recorded in each cell was kept to the minimum required to perform the experiment. Figure 1 illustrates a representative cell attached at both ends to glass rods and held at a sarcomere length of 2.3 μm.

Figure 1. Photomicrograph of a Triton X-100-skinned rat cardiac myocyte.

Figure 1

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The myocyte is attached to two glass rods by optical glue and held at a sarcomere length of 2.3 μm in relaxing solution (pCa 9). The image shows the points of attachment of the cell to the glass rods, which are associated with the white opaque areas slightly behind the tips of the glass rods. Throughout the course of the experiment, sarcomere length was monitored on-line using a fast-Fourier transformation of a video image of the cell in the region between the tips of the glass rods. Off-line measurement of 20 randomly selected sarcomeres gives an estimate of sarcomere length for this cell of 2.28 ± 0.029 μm. The scale bar represents 12 μm. For all experiments, cells were studied at a sarcomere length of either 2.0 μm or 2.3 μm.

Effect of cytochalasin D on tension-pCa relationship

Initial experiments were conducted to select a range of pCa solutions which would enable determination of the tension-pCa relationship at each sarcomere length. To investigate the effect of cytochalasin D on tension-pCa, cells were perfused with relaxing solution containing cytochalasin D (1 or 40 μm) for 2 min, and the pCa curve repeated as before but with cytochalasin D present in all solutions. Cells were then washed with control relaxing solution for 6 min and the pCa curve determined again in the absence of cytochalasin D. As sarcomere length was measured on-line throughout the course of the experiment, we were able to ascertain that the sarcomere length was stable during measurement of tension-pCa relationships and during application of cytochalasin D. Furthermore, we saw no effect of cytochalasin D on passive tension at either sarcomere length. Therefore we are able to rule out the possibility that any effects of cytochalasin D on the skinned myocyte that we observed were due to a change in the compliance of the attachment points.

Solutions

The PSS used during the cell isolation contained (mm): NaCl 117; KCl 5.7; NaHCO3 4.4; KH2PO4 1.5; MgCl2 1.7; Hepes 21; glucose 11; taurine 20 (pH 7.15). For determination of the tension-pCa relationship, the relaxing solution contained (mm): phosphocreatine 12; KCl 78.4; Na+ 30.6; imidazole 30; EGTA 10; dithiothreitol 0.3; pMg2+ 2.5; pMgATP 2.5; pCa 9.0. The full activating solution (pCa 4.5) contained 79.4 mm KCl and pCa was 4.5. The pH of both relaxing and activating solutions was adjusted to 7.1 with either acetic acid or KOH. Solutions with pCa between 9 and 4.5 were made by mixing proportions of the relaxing and full activating solutions. Stock solutions of cytochalasin D (at 1 and 40 mm) were made up in dimethylsulphoxide (DMSO) and stored at −20°C. The final concentration of DMSO in the cytochalasin-containing solutions was 0.1%; control solutions also contained 0.1% DMSO. A stock solution of cytochalasin D-BODIPY FL conjugate (10 mm) was made up in DMSO and stored at −20°C. All solutions containing cytochalasin D were protected from the light.

Data analysis

Tension-pCa relationships were fitted according to the Hill equation using Origin (Microcal Software Inc., version 3.5): T/Tmax (relative tension) =[Ca2+]nH/(KnH+[Ca2+]nH), where nH is the Hill coefficient and pCa50 (pCa for half-maximal activation) = -log10K. In some experiments, the value of Tmax was fixed in the Hill equation and ×2 used to establish the best fit between experimental and theoretical values. All data are presented as means ±s.e.m. for n observations. Where appropriate Student’s paired or unpaired t test was used to test for statistical significance.

RESULTS

Binding of fluorescent cytochalasin D within skinned myocytes

Figure 2 shows sections taken through the sub-sarcolemmal region (Fig. 2A) and the centre (Fig. 2B) of a representative cell incubated with the fluorescent conjugate of cytochalasin D for 15 min. In the section taken through the centre of the cell, fluorescent staining was evident in clear longitudinal stripes, consistent with the structure of the myofibrils in cardiac myocytes. Bright punctate labelling was also present randomly spaced along the stripes. Staining was weaker in the sub-sarcolemmal region of the cell. There was no evidence of sub-sarcolemmal staining consistent with the organisation of actin microfilaments.

Figure 2. Binding of fluorescent cytochalasin D within skinned myocytes.

Figure 2

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Fluorescent confocal micrographs taken through the sub-sarcolemmal region (A) and the centre (B) of a Triton X-100-skinned rat ventricular myocyte incubated with cytochalasin D-BODIPY FL conjugate (1 μm) for 15 min. Scale bar represents 20 μm. The clear longitudinal stripes evident in B are consistent with myofibrillar labelling and the interaction of fluorescent cytochalasin D with sarcomeric actin of the thin filament. The bright punctate labelling randomly spaced along the stripes is not consistent with microfilament structure in the adult myocyte and may be due to accumulation of high concentrations of fluorescent cytochalasin D.

Effect of cytochalasin D on tension-pCa relationship

Figure 3 illustrates the protocol used to determine the effect of the actin-disrupting drug cytochalasin D on the tension-pCa relationship in skinned rat cardiac myocytes. In this example, following 2 min exposure to 1 μm cytochalasin D, the tension produced by each pCa was reduced, and the maximum tension produced at a saturating pCa of 4.5 was slightly depressed. Furthermore, there was a shift in the sensitivity to Ca2+ so that the pCa50 was below 5.5 following perfusion with 1 μm cytochalasin D, whereas in the control cell, pCa50 lay between 5.75 and 5.625. With 40 μm cytochalasin D, which was the concentration of cytochalasin D used to reduce shortening by 80% in intact myocytes in the recent study of Howarth et al. (1998), the only pCa to produce an increase in tension was pCa 4.5. The maximal tension produced at pCa 4.5 was much depressed, and the rate of tension development slowed. On the basis of this preliminary study, we elected to investigate the action of cytochalasin D using a concentration of 1 μm.

Figure 3. Original traces illustrating the protocol used to investigate the effect of cytochalasin D on the tension-pCa relationship in skinned cardiac myocytes held at sarcomere lengths of 2.0 μm.

Figure 3

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Myocytes were exposed to solutions with varying pCa values as indicated in the absence of cytochalasin D (control), in the presence of 1 μm cytochalasin D (cyto D), and in the presence of 40 μm cytochalasin D. Cells were exposed to cytochalasin D for 2 min. Because the effect of cytochalasin D was not readily washed out, the effects of the two concentrations of cytochalasin D were investigated in different cells.

The effect of cytochalasin D on the relative tension-pCa relationships from a series of experiments performed at the two sarcomere lengths is shown in Fig. 4. Each cell was only studied at one sarcomere length. At both sarcomere lengths, 1 μm cytochalasin D caused a shift in the tension-pCa curve to the right, indicating a decrease in Ca2+ sensitivity. This shift was not reversed by 6 min of washout of cytochalasin D. Tension-pCa curves from each cell were fitted to the Hill equation and the Hill coefficients and pCa50 obtained are summarised in Table 1. The maximum developed tension (in the presence of saturating pCa 4.5) is also shown in Table 1. In myocytes held at a sarcomere length of 2.3 μm, pCa50 was significantly higher (P < 0.001) than that in myocytes at a sarcomere length of 2.0 μm. There was no significant difference (P > 0.05) in either Hill coefficients or the maximum tension developed at the two sarcomere lengths.

Figure 4. Effects of 1 μm cytochalasin D on the relative tension-pCa relationship in skinned cardiac myocytes held at sarcomere lengths of either 2.0 (± 0.1) μm (A) or 2.3 (± 0.1) μm (B).

Figure 4

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Tension-pCa was measured before cytochalasin D (^), after 2 min perfusion with cytochalasin D (•) and following 6 min washout of cytochalasin D (▾, dotted line). Some data points are superimposed. All data are given as the mean ±s.e.m. of 9 observations. Data from each cell were fitted with the Hill equation, and the resulting parameters are listed in Table 1.

Table 1.

Effect of cytochalasin D on parameters of the Hill equation and maximum tension in skinned cardiac myocytes

SL = 2.0 μm SL = 2.3 μm
pCa50 nH Maximum tension (μN) pCa50 nH Maximum tension (μN)
Control 5.54 ± 0.03 4.81 ± 0.55 9.61 ± 1.36 5.76 ± 0.5††† 4.35 ± 1.17 7.24 ± 1.07
Cyto D 5.45 ± 0.04** 4.06 ± 0.57 8.17 ± 1.27*** 5.55 ± 0.03** 5.59 ± 2.42 5.26 ± 0.75***
Washout 5.43 ± 0.03*** 4.20 ± 0.40 7.67 ± 1.08** 5.62 ± 0.03* 4.92 ± 1.25 4.50 ± 0.83**
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Data from individual cells at sarcomere lengths (SL) of 2.0 (±.0.1) and 2.3 (±0.1) μm were fitted to the Hill equation in the absence of cytochalasin D (Control), following 2 min perfusion with 1 μM cytochalsin D (Cyto D), and following 6 min washout of cytochalasin D (Washout). n%H = Hill coefficient. The width o cells (at slack length) was similar in cells studiedd at a SL of 2.0 μm (24.9 ± 1.3 μm) and those studied at 2.3 μm (23.1 ± 1.2 μm). Values are given as the mean ± S.E.M. of 9 observations.

*

P < 0.05

**

P < 0.01

***

P < 0.001 compared with respective control data (Student’s paired paired t test).

†††

P < 0.001 compared with respective measurement at SL of 2.0 μm (Student’s unpaired t test).

At a sarcomere length of 2.0 μm, cytochalasin D caused a significant decrease (P < 0.01) of 0.09 pCa units in the mean pCa50. At a sarcomere length of 2.3 μm, the effect of cytochalasin D was enhanced and a decrease (P < 0.01) of 0.22 pCa units in the mean pCa50 was seen in the presence of the drug. At neither sarcomere length was the change in pCa50 reversed by washout. The shift in pCa50 at a sarcomere length of 2.3 μm was significantly higher (P < 0.05) than that seen at a sarcomere length of 2.0 μm. At neither sarcomere length was any significant effect (P > 0.05) of cytochalasin D on the Hill coefficient seen. Cytochalasin D significantly reduced (P < 0.001) the maximum tension seen at saturating pCa to 83% of control at a sarcomere length of 2.0 μm, and to 73% of control at a sarcomere length of 2.3 μm. The difference between the reduction in maximum tension at the two lengths was not significant (P > 0.05). The reduction in tension did not arise as a consequence of the shift of the relative tension-pCa relationship with cytochalasin D, as the best ×2 for the curve fit was obtained when it was assumed that the tension at pCa 4.5 in the presence of cytochalasin D was the maximum. The effect of cytochalasin D on maximum tension was not reversed by washout. Although a slight reduction in maximum tension may occur during an experiment of this type with repeated application of pCa 4.5, we saw no significant shift (−5.1 ± 5%; n = 4; P > 0.05) in maximum tension in a number of cells during the second tension-pCa curves performed in the absence of cytochalasin D. Similarly, others have observed no significant depression in maximum force with repeated exposure to a range of pCa values (Cazorla et al. 1999).

DISCUSSION

The present study was conducted in order to determine whether the actin-disrupting drug cytochalasin D interacts with sarcomeric actin within the cardiac cell, and to determine the nature of this interaction. Our experiments were performed with skinned adult cardiac myocytes in which cytochalasin D has free access to the intracellular space and diffuses more easily than in skinned multicellular cardiac preparations. Furthermore, as we used single cells, our data can be interpreted in terms of an action of cytochalasin D on the cardiac myocyte itself. In skinned cardiac muscle preparations employed by some workers, the results may be complicated by the presence of non-muscle cells and connective tissue.

A highly developed actin cytoskeleton has been observed in embryonic, neonatal and cultured adult myocytes. In these cells, electron microscopy and immunocytochemistry have shown clear co-labelling of sarcomeric actin and actin microfilaments (e.g. Sugi & Hirakow, 1991; Messerli & Perriard, 1995; Rothen-Rutishauser et al. 1998). However, these cells are not rod-shaped and can spread over their substrate, an action which appears to be dependent upon the presence of microfilaments. It is in cells such as these that cytochalasin D has been reported to disrupt preferentially cytoskeletal actin (e.g. Sadoshima et al. 1992). In rod-shaped adult myocytes, labelling for actin reveals myofibrils with sarcomeric patterning (e.g. Elbe & Spinale, 1995). In the adult myocyte there is evidence for several different formations of actin microfilaments within the cell – in attachments from the Z-line to discrete regions of the sarcolemma (Sage & Jennings, 1988), in association with leptomeres (dense bars adjacent to the sarcolemma spaced at intervals of 150–200 nm; Hosokawa et al. 1994), and as a sub-sarcolemmal lattice similar to that characterised in erythrocytes (Ganote & Armstrong, 1993). Therefore in the skinned adult cardiac myocytes which we used in the present study, if fluorescent cytochalasin D bound preferentially to non-sarcomeric actin, we would expect to see defined staining patterns characteristic of cytoskeletal actin. The longitudinally striated staining patterns we observed in the centre of the cell are clearly consistent with the structure of the myofibrils and with the binding of cytochalasin D to sarcomeric actin. The punctate labelling we noted randomly spaced along the stripes is not consistent with microfilament structures in the adult myocyte. We cannot identify a structure in the adult myocyte whose cellular organisation is consistent with the random punctate labelling we observed with the fluorescent conjugate of cytochalasin D, and suggest that this staining may be due to accumulations of concentrations of fluorescent dye. We saw no sub-sarcolemmal staining pattern consistent with the organisation of cytoskeletal actin. Cytochalasin D has been shown to depress cardiac contractility (Maltsev & Undrovinas, 1997; Skobel & Kammermeier, 1997; Biermann et al. 1998; Howarth et al. 1998). Until recently, it was assumed that the stability of sarcomeric actin protects it from the effects of the actin-disrupting drug cytochalasin D, and that the actions of cytochalasin D are due to interference with the more dynamic cytoskeletal actin. It has been speculated that this may interfere with force transmission between the sarcomere and sarcolemma, or may modify sarcoplasmic reticulum function (Skobel & Kammermeier, 1997; Biermann et al. 1998; Undrovinas and Maltsev, 1998). However, much evidence regarding the effects of cytochalasin D on cytoskeletal actin has been obtained in neonatal or cultured cells (see above). A recent study by Howarth et al. (1998) provided the first evidence that cytochalasin D may indeed exert at least some of its effects on the contraction of the adult cardiac cell through interaction with sarcomeric actin, effects which were mediated very rapidly (within 2 min). In the present study we observed a significant decrease in pCa50 in skinned rat cardiac myocytes following a short exposure to cytochalasin D. This decrease in the sensitivity of the myofilaments to Ca2+ is consistent with the depression of contractility without a concomitant change in the amplitude of the Ca2+ transient (Maltsev & Undrovinas, 1997; Undrovinas & Maltsev, 1998). These effects have previously been ascribed to an action of cytochalasin D on cytoskeletal actin. Our data are in accord with those of Howarth et al. (1998) who observed a shift in the phase-plane relationship between Ca2+ and contraction in the intact cell following exposure to cytochalasin D. Thus we present, for the first time, direct evidence that the negative inotropic actions of cytochalasin D can be ascribed to an interaction with sarcomeric actin.

It is well documented that an increase in sarcomere length increases the sensitivity of the myofilaments to Ca2+ through an increase in the affinity of troponin C for Ca2+ (Hofmann & Fuchs, 1987; for a review, see Calaghan & White, 1999). Therefore, it is interesting to note that in the present study the effect of cytochalasin D at the longer sarcomere length was significantly more pronounced (P < 0.05) than at the shorter sarcomere length. In fact, the pCa50 at a sarcomere length of 2.3 μm in the presence of cytochalasin D was the same as that at a sarcomere length of 2.0 μm in the absence of cytochalasin D. As the shift in pCa50 is more pronounced at a longer length, we propose that the same mechanism (i.e. a change in the affinity of troponin C for Ca2+) underlies the effect of cytochalasin D and the effect of stretch on myofilament Ca2+ sensitivity. A decrease in the affinity of troponin C for Ca2+ is also consistent with the slowing of the kinetics of the Ca2+ transient with cytochalasin D observed by Undrovinas & Maltsev (1998). For example, the decrease in myofilament sensitivity seen when cardiac muscle length is shortened is also associated with a prolongation of the Ca2+ transient duration (Allen et al. 1988).

As well as a decrease in the sensitivity of the myofilaments to Ca2+, we also observed a significant depression of the maximum tension recorded in the presence of saturating Ca2+ with cytochalasin D. This depression of maximum tension was independent of the effect of cytochalasin D on myofilament sensitivity, suggesting that cytochalasin D is not acting as a simple Ca2+ desensitiser. In the two-state crossbridge model similar to that proposed by Huxley (1957), tension can be represented by Fn(f/f+g) where F is the force per crossbridge, n is the number of cycling crossbridges and transition from detached non-force-generating crossbridges to force-generating crossbridges is governed by a rate constant f and the rate of detachment by a rate constant g (Wolff et al. 1995). Therefore the effect of cytochalasin D on maximum tension suggests that cytochalasin D modulates the number of cycling crossbridges, the rate of cycling of crossbridges, or the force produced per crossbridge. An action of cytochalasin D on crossbridges might also account for the observed decrease in myofilament Ca2+ sensitivity, acting co-operatively to decrease the affinity of troponin C for Ca2+ (Rice et al. 1999).

It seems unlikely that the actions of cytochalasin D observed within the short time scale of the present experiment are due to its binding to the barbed end of the actin filament, thus preventing addition of monomers. The rapid effects of cytochalasin D on the skinned cardiac cell may arise as a result of binding elsewhere within the actin filament. The confocal images that we have obtained using a fluorescent conjugate of cytochalasin D suggest that cytochalasin D may bind along the length of the thin filament.

In the present study, we have shown for the first time that cytochalasin D can interact with the contractile machinery to decrease both the sensitivity of the myofilaments to Ca2+ and the maximum tension produced. Our data are consistent with depression of contractility and slowing of the intracellular Ca2+ transient seen by others, yet allow us to conclude that such effects may be due to an interaction of cytochalasin D with sarcomeric, rather than cytoskeletal, actin.

Acknowledgments

This work was funded by a Wellcome Trust Biomedical Research Collaboration grant and the British Heart Foundation.

References

  1. Allen DG, Nichols CG, Smith GL. The effects of changes in muscle length during diastole on the calcium transient in ferret papillary muscle. Journal of Physiology. 1988;406:359–370. doi: 10.1113/jphysiol.1988.sp017385. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Bennett V. Spectrin-based membrane skeleton: a multipotential adaptor between plasma membrane and cytoplasm. Physiological Reviews. 1990;70:1029–1065. doi: 10.1152/physrev.1990.70.4.1029. [DOI] [PubMed] [Google Scholar]
  3. Biermann M, Rubart M, Moreno A, Wu J, Josiah-Durant A, Zipes D. Differential effects of cytochalasin D and 2,3 butanedione monoxime on isometric twitch force and transmembrane action potential in isolated ventricular muscle: Implications for optical measurements of cardiac repolarization. Journal of Cardiovascular Electrophysiology. 1998;9:1348–1357. doi: 10.1111/j.1540-8167.1998.tb00110.x. [DOI] [PubMed] [Google Scholar]
  4. Brenner SL, Korn ED. Substoichiometric concentrations of cytochalasin D inhibit actin polymerisation. Journal of Biological Chemistry. 1979;254:9982–9985. [PubMed] [Google Scholar]
  5. Bukatina AE, Fuchs F, Brandt PW. Thin filament activation by phalloidin in skinned cardiac muscle. Journal of Molecular and Cellular Cardiology. 1995;27:1311–1315. doi: 10.1016/s0022-2828(05)82393-4. [DOI] [PubMed] [Google Scholar]
  6. Calaghan SC, White E. The role of calcium in the response of cardiac muscle to stretch. Progress in Biophysics and Molecular Biology. 1999;71:59–90. doi: 10.1016/s0079-6107(98)00037-6. [DOI] [PubMed] [Google Scholar]
  7. Cazorla O, Vassort G, Garnier D, Le Guennec J-Y. Length modulation of active force in rat cardiac myocytes: is titin the sensor? Journal of Molecular and Cellular Cardiology. 1999;31:1215–1227. doi: 10.1006/jmcc.1999.0954. [DOI] [PubMed] [Google Scholar]
  8. Cooper JA. Effects of cytochalasin and phalloidin on actin. Journal of Cell Biology. 1987;105:1473–1478. doi: 10.1083/jcb.105.4.1473. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Elbe DM, Spinale FG. Contractile and cytoskeletal content, structure and mRNA levels with tachycardia-induced cardiomyopathy. American Journal of Physiology. 1995;268:H2426–2439. doi: 10.1152/ajpheart.1995.268.6.H2426. [DOI] [PubMed] [Google Scholar]
  10. Gannier F, Jacquemond V, Bernango J-C, Garnier D. Measurement of sarcomere dynamics simultaneously with auxotonic force in isolated cardiac cells. Institute of Electronics and Electrical Engineering. 1993;40:1226–1232. doi: 10.1109/10.250578. [DOI] [PubMed] [Google Scholar]
  11. Ganote C, Armstrong S. Ischaemia and the myocyte cytoskeleton: review and speculation. Cardiovascular Research. 1993;27:1387–1403. doi: 10.1093/cvr/27.8.1387. [DOI] [PubMed] [Google Scholar]
  12. Hilgemann DW. Cytoplasmic ATP-dependent regulation of ion transporters and channels: Mechanisms and messengers. Annual Review in Physiology. 1997;59:193–220. doi: 10.1146/annurev.physiol.59.1.193. [DOI] [PubMed] [Google Scholar]
  13. Hofmann PA, Fuchs F. Effect of length and cross-bridge attachment on Ca2+ binding to cardiac troponin C. American Journal of Physiology. 1987;253:C90–96. doi: 10.1152/ajpcell.1987.253.1.C90. [DOI] [PubMed] [Google Scholar]
  14. Hosokawa T, Okada T, Kobayashi T, Hashimoto K, Seguchi H. Ultrastructural and immunocytochemical study of leptomeres in the mouse cardiac muscle fibre. Histology and Histopathology. 1994;9:85–94. [PubMed] [Google Scholar]
  15. Howarth FC, Boyett MR, White E. Rapid effects of cytochalasin-D on contraction and intracellular calcium in single rat ventricular myocytes. Pflügers Archiv. 1998;436:804–806. doi: 10.1007/s004240050706. [DOI] [PubMed] [Google Scholar]
  16. Huxley AF. Muscle structure and theories of contraction. Progress in Biophysics. 1957;7:255–318. [PubMed] [Google Scholar]
  17. Maltsev VA, Undrovinas AI. Cytoskeletal involvement in excitation-contraction coupling in rat ventricular cardiomyocytes (abstract) Biophysical Journal. 1997;72:TU154. [Google Scholar]
  18. Messerli JM, Perriard JC. Three-dimensional analysis and visualization of myofibrillogenesis in adult cardiomyocytes by confocal microscopy. Microscopy Research and Technique. 1995;30:521–530. doi: 10.1002/jemt.1070300609. [DOI] [PubMed] [Google Scholar]
  19. Messina DA, Lemanski LF. Immunocytochemical studies of spectrin in hamster cardiac tissue. Cell Motility and the Cytoskeleton. 1989;12:139–149. doi: 10.1002/cm.970120303. [DOI] [PubMed] [Google Scholar]
  20. Moses RL, Delcarpio JB. Sarcoplasmic reticulum and intermediate filament organization in cultured neonatal cardiac muscle cells. Cell and Tissue Research. 1983;228:489–496. doi: 10.1007/BF00211470. [DOI] [PubMed] [Google Scholar]
  21. Pucéat M, Clément O, Lechene P, Pelosin JM, Ventura-Clapier R, Vassort G. Neurohumoral control of calcium sensitivity of myofilaments in rat skinned heart cells. Circulation Research. 1990;67:517–524. doi: 10.1161/01.res.67.2.517. [DOI] [PubMed] [Google Scholar]
  22. Rice JJ, Winslow RL, Hunter WC. Comparison of putative cooperative mechanisms in cardiac muscle: length dependence and dynamic responses. American Journal of Physiology. 1999;276:H1734–1754. doi: 10.1152/ajpheart.1999.276.5.H1734. [DOI] [PubMed] [Google Scholar]
  23. Rothen-Rutishauser BM, Ehler E, Perriard E, Messerli JM, Perriard J-C. Different behaviour of the non-sarcomeric cytoskeleton in neonatal and adult rat cardiomyocytes. Journal of Molecular and Cellular Cardiology. 1998;30:19–31. doi: 10.1006/jmcc.1997.0596. [DOI] [PubMed] [Google Scholar]
  24. Sadoshima J-I, Takahashi T, Jahn L, Izumo S. Role of mechano-sensitive ion channels, cytoskeleton, and contractile activity in stretch-induced immediate-early gene expression and hypertrophy of cardiac myocytes. Proceedings of the National Academy of Sciences of the USA. 1992;89:9905–9909. doi: 10.1073/pnas.89.20.9905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Sage MD, Jennings RB. Myocyte swelling and plasmalemmal integrity during early experimental myocardial ischaemia in vivo. Scanning Microscopy. 1988;2:477–484. [PubMed] [Google Scholar]
  26. Skobel E, Kammermeier H. Relation between enzyme release and irreversible cell injury of the heart under the influence of the cytoskeletal modulating agents. Biochimica et Biophysica Acta. 1997;1362:128–134. doi: 10.1016/s0925-4439(97)00060-4. [DOI] [PubMed] [Google Scholar]
  27. Sugi Y, Hirakow R. Cytoskeletal filaments in embryonic chick myocardial cells as revealed by the quick-freeze deep-etch method combined with immunocytochemistry. Cell and Tissue Research. 1991;263:459–469. doi: 10.1007/BF00327279. [DOI] [PubMed] [Google Scholar]
  28. Undrovinas AI, Maltsev VA. Cytochalasin D alters kinetics of Ca2+ transient in rat ventricular cardiomyocytes: An effect of altered actin cytoskeleton? Journal of Molecular and Cellular Cardiology. 1998;30:1665–1670. doi: 10.1006/jmcc.1998.0715. [DOI] [PubMed] [Google Scholar]
  29. Wolff MR, McDonald KS, Moss RL. Rate of tension redevelopment in cardiac muscle varies with the level of activator calcium. Circulation Research. 1995;76:154–160. doi: 10.1161/01.res.76.1.154. [DOI] [PubMed] [Google Scholar]

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