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. 1999 Aug 15;519(Pt 1):213–221. doi: 10.1111/j.1469-7793.1999.0213o.x

Kinetics of contraction in depolarized smooth muscle from guinea-pig taenia coli after photodestruction of nifedipine

Ulf Malmqvist 1, Anders Arner 1
PMCID: PMC2269492  PMID: 10432352

Abstract

  1. The time course and kinetics of force development following activation by opening of L-type Ca2+ channels was investigated using photodestruction of the Ca2+ channel blocker nifedipine in smooth muscle from the guinea-pig taenia coli.

  2. In muscles activated using high K+ and Ca2+ and subsequently inhibited with nifedipine, photodestruction of the drug using a strong ultraviolet light flash initiated a rapid contraction. The force initiated by photodestruction of nifedipine reached near-maximal levels. This procedure eliminates diffusional delays and can thus be used to investigate the kinetics of depolarization-induced contractions.

  3. The rate of force development of contractions initiated by photodestruction of nifedipine was slower than that observed in maximally thiophosphorylated skinned fibres. This suggests the rate of force development is limited by activation steps in the activation cascade prior to the force generation of the cross-bridge system.

  4. The rate of force development and the plateau force were dependent on the extracellular [CaCl2] suggesting that the intracellular [Ca2+] determines the rate of phosphorylation and force development. The delay between illumination and increase in force was about 300 ms. The delay was similar at low and high extracellular [CaCl2] indicating that buffering by superficial sarcoplasmatic reticulum does not introduce a delay in force development following activation of Ca2+ channels in this muscle.

It is generally accepted that phosphorylation of Ser-19 of the regulatory light chain (RLC) of myosin is the main mechanism for initiation of smooth muscle contraction. The level of RLC phosphorylation is determined by the activities of myosin light chain kinase (MLCK) and the phosphatase(s), which phosphorylate and dephoposhorylate the RLC, respectively. Several factors have been shown to influence the activity of the phosphatase whereas the intracellular calcium concentration is the primary messenger regulating the activity of MLCK. In the cell, calcium binds to calmodulin and this complex activates MLCK leading to phosphorylation of the RLC. Phosphorylation of the RLC results in an increase in the ATPase activity of myosin, cross-bridge cycling and shortening or force development of the muscle.

Two main pathways for activation of smooth muscle have been proposed, electromechanical and pharmacomechanical coupling (Somlyo & Somlyo, 1968). Although the final events, RLC phosphorylation and cross-bridge cycling, are identical the activating pathways leading to an increase in net RLC phosphorylation differ markedly between these two mechanisms for coupling excitation to contraction. Receptor-mediated activation (pharmacomechanical coupling) initiates a complex series of events including Ca2+ release and influx as well as sensitization mechanisms influencing the phosphorylation/dephosphorylation process directly (for review see Arner & Pfitzer, 1998).

Rapid activation using caged compound technology has been used to study the force generation and the kinetics of the different steps in the activation pathway (see Somlyo & Somlyo, 1990). In fully phosphorylated (thiophosphorylated) skinned preparations the force development after photolytic release of ATP, reflecting the force generating transitions, is rapid with a delay of less than 10 ms at room temperature (Somlyo et al. 1988a). After photolytic release of phenylephrine from a caged precursor the delay between activation and the onset of contraction is substantially longer and in the order of 500 ms at physiological temperature and 930–1500 ms at room temperature (Somlyo et al. 1988b; Muralidharan et al. 1993; Walker et al. 1993). The IP3-induced release of Ca2+ from intracellular stores is more rapid and contraction occurs with a delay of 500 ms after photolytic release of IP3 at room temperature (Somlyo et al. 1988b). It has been suggested that the initial production of the second messenger is a comparatively slow process introducing an approximately 1000 ms delay between receptor activation and force development in the pharmacomechanical coupling (Somlyo et al. 1988b). Interestingly, not all receptor-mediated responses are slow; activation of purinoceptors with ATP from caged ATP results in rapid contractions (Sjöblom-Widfeldt et al. 1993) with a delay of about 80 ms at 37°C (Sjuve et al. 1995). These data suggest that a rapid influx or increase of Ca2+ following purinoceptor activation bypasses the slow pathways involved in the adrenergic receptor-mediated contraction.

When smooth muscle is activated via alterations in the membrane potential the increase in cytostolic Ca2+ is due to opening of potential-sensitive calcium channels in the cell membrane (electromechanical coupling). Following electrical stimulation of single isolated smooth muscle cells Ca2+ increases within milliseconds and force development starts after a delay of 265 ms at 22°C (Fay, 1977; Yagi et al. 1988). This is similar to results from skinned smooth muscle preparations where a delay of about 300 ms between Ca2+ increase and force development is observed (Somlyo et al. 1992; Zimmerman et al. 1995). These studies show that the slowest process in the electromechanical coupling is an event late in the activation cascade. It is possible that the pharmacomechanical and electromechanical pathways converge at this point, which could be prephosphorylation reactions or the phosphorylation process itself.

Electromechanical coupling thus seems to activate directly the phosphorylation system via a rapid influx of Ca2+ and, as discussed above, contraction starts after a short delay. This does not necessarily mean that the coupling between the opening of L-channels and the activation of the phosphorylation process cannot be modulated by other cellular systems. It has been proposed that the membrane associated sarcoplasmic reticulum can act as a sink and thereby affect the cytosolic calcium around the myofilaments and the kinetics of the electromechanical coupling (van Breemen, 1977; van Breemen et al. 1995). This superficial buffer barrier system might be more important for activation kinetics after submaximal activation. The kinetics of activation via electromechanical coupling and the influence of the buffer barrier on force development are, however, difficult to study in intact smooth muscle preparations, since rapid and controlled changes in L-channel opening cannot be achieved with high-potassium solutions due to diffusional delays.

The L-type calcium channels can be blocked by dihydropyridines such as nifedipine. Nifedipine contains an o-nitrobencyl moiety which makes it photolabile, and illumination of nifedipine with ultraviolet light results in the destruction of the drug into products which are devoid of calcium antagonist activity (Morad et al. 1983). When nifedipine is illuminated with a strong ultraviolet light flash the reactions leading to photoconversion of nifedipine are completed within 100 μs (Morad et al. 1983). This property of nifedipine has been used to study the excitation-contraction coupling in cardiac muscle (Morad et al. 1983). We have in the present study used photodestruction of nifedipine as a tool to study the kinetics of the electromechanical coupling, and investigated the time course of force development at different degrees of activation in intact smooth muscle from guinea-pig taenia coli.

METHODS

Taenia coli from female guinea-pigs weighing 400–500 g were used. The animals were killed by cervical dislocation and taenia coli muscle strips were dissected and freed from fat and connective tissue. Preparations 5–7 mm long with a diameter of about 200 μm were cut out from the strips. The experiments were carried out according to the guidelines of the local animal ethics committee.

For determination of the dependence of force on calcium, potassium and nifedipine concentration the muscle fibres were mounted in an open organ bath (volume 60 ml). One end of the muscle was connected to a Grass FT03 force transducer and the other end to a stainless steel pin on an adjustable stand. The muscle length was adjusted to give a passive tension of 1–2 mN, which is close to the optimal length for force development (Arner, 1982). The temperature of the organ bath was 37°C. After a 1 h acclimation period in N-Krebs solution (for composition see below) the muscles were stimulated with high-KCl solution for 5 min and thereafter relaxed with a calcium-free N-Krebs solution. When reproducible contractions were obtained the following protocol was used for determination of the [Ca2+] sensitivity: (1) depolarization with high KCl (30, 60 or 120 mM) in calcium-free solution, (2) contraction initiated by adding CaCl2 to the bath and registration of plateau force, and (3) relaxation in calcium-free N-Krebs solution for 10 min. The protocol was repeated from step (1) again with a new CaCl2 concentration. For determination of the potassium sensitivity of force, the muscles were stimulated using the same protocol, but with increasing concentrations of KCl keeping the CaCl2 concentration constant at 2.5 mM. The sensitivity to nifedipine was determined using a similar protocol. Further details are given in the Results section.

For determination of the rate of tension development following photolysis of nifedipine aluminium foil was wrapped around both ends of the muscle fibre preparation. The fibre preparation was then mounted horizontally in small cups (volume 550 μl) in an apparatus described earlier (Arner et al. 1987), with one end of the muscle connected to an AME force transducer and the other to a stainless-steel needle mounted on a micrometer screw for length control. The temperature of the cups was 37°C. After a 60 min acclimatization period in N-Krebs solution the muscles were challenged with high-KCl solution as described above and when reproducible contractions were obtained, nifedipine was added at the plateau of contraction to inhibit force. When force had declined to about 5–15 % of maximal, the fibres were transferred to a 50 μl bath equipped with a quartz window and illuminated with an ultraviolet light flash from a xenon lamp (Dr G. Rapp, Optoelektronik, Hamburg, Germany) to photolyse nifedipine. The force transients were recorded on magnetic tape and digitized using an Analog Devices board (ADI-RT800F; Norwood, MA, USA) in a personal computer for subsequent analysis. On each fibre, flash photolysis was performed 3–5 times. The first and last contraction was performed with the same KCl and CaCl2 concentration. Since nifedipine is light sensitive all experiments using this drug were performed in a dark room.

In order to compare the kinetics of force development after opening of calcium channels with the kinetics after purinoceptor activation, we were interested in comparing the force responses after nifedipine photodestruction with those following photolytically released ATP from caged ATP. Since the guinea-pig taenia coli does not have contractile purinergic responses (see Brown & Burnstock, 1981) these experiments were performed in the portal vein obtained from adult Sprague-Dawley rats. The experiments were performed as described above and photodestruction of nifedipine was done in 2.5 mM CaCl2 and 120 mM KCl. ATP was released from 100 μM caged ATP (adenosine-5′-triphosphate, P3-1-(2-nitrophenyl)-ethyl ester; Calbiochem Corp., La Jolla, CA, USA) in N-Krebs solution with 2.5 mM CaCl2.

The normal Krebs (N-Krebs) solution was of the following composition (mM): NaCl, 122; KCl, 4.7; MgCl2, 2.5; NaHCO3, 15.5; KH2PO4, 1.2; EDTA, 0.026; and glucose, 11.5. The Ca2+ concentration was 2.5 mM unless otherwise stated. The high-KCl solutions were made by replacing NaCl in the N-Krebs solution with KCl. The solutions were gassed with 96 % O2-4 % CO2 giving pH 7.4 at 37°C.

Statistics and curve fitting

All values are given as means ± s.e.m., with the number (n) of independent experiments indicated. Curve fitting was performed using a non-linear fitting implemented in SigmaPlot for Windows 3.03 (Jandel Corporation, Germany).

RESULTS

Dependence of force on extracellular [Ca2+] and [K+]

The N-Krebs solution contained 5.9 mM KCl and in this solution the muscles usually exhibit spontaneous contractile activity. Raising the extracellular KCl concentration causes a depolarization of the muscle, and when the muscles were challenged with 10 mM KCl they developed a sustained contraction. The dependence of force on the extracellular KCl concentration is shown in Fig. 1A. The force values are normalized to the maximal force response of each preparation. In six of eight muscles the maximal force was recorded when the muscles were depolarized with 40 mM KCl. The calcium sensitivity of force was determined by depolarizing the muscles with different concentrations of KCl and then adding CaCl2 to initiate the contraction. The relation between extracellular [CaCl2] and force for muscles depolarized using different concentrations of KCl is shown in Fig. 1B. The muscles developed maximal force at 1.6 mM extracellular CaCl2 when depolarized with 30 mM KCl. At higher CaCl2 concentrations force declined again. This is in contrast to the results obtained in 60 or 120 mM KCl where force increased up to 9 mM CaCl2, which was the maximal concentration that could be achieved in the bicarbonate buffered solutions.

Figure 1. Potassium and calcium dependence of active force.

Figure 1

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A, relationship between extracellular potassium concentration and active force. The muscles were depolarized with solutions containing different potassium concentrations. Contractions were initiated by adding CaCl2 to a final concentration of 2.5 mM. Force is expressed relative to the maximal force at optimal potassium concentration for each preparation (n = 8). B, calcium sensitivity of force. The muscles were depolarized with 30 mM (•), 60 mM (▪) or 120 mM KCl (▴) and contractions were initiated by adding different concentrations of CaCl2. Force is expressed relative to the force at optimal calcium concentration at the respective KCl concentration for each preparation (n = 7).

Inhibition of contraction by nifedipine

The sensitivity of vascular smooth muscle to inhibition of contraction by dihydropyridines increases with membrane depolarization (Nelson & Worley, 1989). We therefore determined the dose-response relationship for nifedipine using two different procedures. In the first, nifedipine was introduced to the muscles for 10 min in N-Krebs solution. The muscles were then activated by 60 mM KCl and 2.5 mM CaCl2, with nifedipine still present. In the second procedure, we depolarized with high K+, then added nifedipine and after 10 min a contraction was initiated by adding CaCl2. The relationship between nifedipine concentration and inhibition of force is shown in Fig. 2. The inhibition of force was more pronounced when nifedipine was introduced to depolarized muscles. The relationship between nifedipine concentration and force can be described by the following equation: F/F* = 1 - N/(N + Kapp), where F* is maximal force before adding nifedipine, F is the force at the concentration N of nifedipine, and Kapp is the apparent inhibition constant. When nifedipine was added to the muscles in N-Krebs solution (first procedure, above), Kapp was 6.74 nM (Fig. 2, open symbols). Nifedipine was more effective when added to muscles in high-KCl solutions (second procedure) with a Kapp of 0.54 and 0.44 nM for muscles depolarized with 60 and 120 mM KCl, respectively (Fig. 2, filled symbols).

Figure 2. Dose-response relationship for nifedipine.

Figure 2

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Open symbols show muscles where nifedipine was introduced in the N-Krebs solution 10 min prior to activation by 60 mM KCl and 2.5 mM CaCl2. Filled symbols show muscles where nifedipine was introduced when the muscle was depolarized with 60 mM (▪) or 120 mM KCl (▴). After 10 min in this solution contractions were initiated by adding CaCl2. Force values (F) are normalized to the force level before adding nifedipine (F*, n = 6). The lines in the figure show the equation F/F* = 1 - N/(N + Kapp) fitted to the mean force values at each nifedipine concentration (N). The Kapp values were 0.54 and 0.44 nM for the 60 and 120 mM KCl groups and 6.7 nM in the group where nifedipine was introduced in N-Krebs solution 10 min prior to activation by 60 mM KCl.

Tension responses after flash photolysis of nifedipine

For determination of the force response after photolytical conversion of nifedipine the muscles were first activated with high-KCl solution. Nifedipine (2 nM) was added at the force plateau to inhibit contraction. In Fig. 3 original recordings of three consecutive contractions from one smooth muscle fibre preparation are shown. The preparation was activated with 60 mM, 120 mM and then again with 60 mM KCl at 2.5 mM CaCl2. After addition of nifedipine force declined. When force was about 5–15 % of maximal the muscle was transferred to the 50 μl bath and illuminated with an ultraviolet light flash. Illumination of the muscles resulted in a rapid contraction. A second illumination only caused a small additional force development. The muscles did not develop any force when illuminated in the N-Krebs solution. In control experiments, an initial and a final flash-induced contraction were performed in 60 mM KCl and 2.5 mM CaCl2. The rate of force development and the amplitude of the force response reached in these contractions were similar (within 10 %). Figure 4 shows, on a faster time base, force transients from a muscle activated with a solution containing 60 mM KCl in 2.5 and 0.4 mM CaCl2. The rate of force development and the amplitude of the force response were higher at high [CaCl2]. Figure 5 shows the early phase after illumination in the muscle activated in 0.4 (lower trace) and 2.5 mM CaCl2 (upper trace). A delay is observed between illumination and the onset of force development. We evaluated the delay by fitting a straight line to the initial linear phase of force development. The time point when this line crossed the base line was considered the end of the delay period. The delay was similar in high and low CaCl2. Figure 6 summarizes the results. As seen in Fig. 6A, the force reached after the light flash was dependent on the extracellular Ca2+ concentration, similar to results where K+ was added by diffusion (cf. Fig. 1B). The half-time for force development was inversely related to the Ca2+ concentration (Fig. 6B). When muscles were activated with 120 mM KCl at 2.5 mM extracellular calcium, both the rate of force development and the plateau force were lower than the values of muscles activated with 60 mM KCl. The delay between illumination and the onset of force was about 300 ms and independent of the extracellular CaCl2 and KCl concentrations used (Fig. 6C). The half-time for force development was inversely correlated with active force (Fig. 6D).

Figure 3. Original recordings of a muscle activated by photolysis of nifedipine.

Figure 3

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Three subsequent contractions from the same muscle fibre preparation are shown. The muscle was activated with high KCl and at the force plateau 2 nM nifedipine was added. When force was inhibited to about 80–95 % a strong light flash resulted in a rapid force development. A second light flash gave a small additional force development. The times for the light flashes are indicated by arrows.

Figure 4. Force transients after photolysis of nifedipine.

Figure 4

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The muscle was activated with high KCl and force was inhibited with 2 nM nifedipine. The muscle was activated (at time = 0) by flash photolysis of nifedipine in the presence of 2.5 mM (upper trace) or 0.4 mM CaCl2 (lower trace).

Figure 5. Delay of force development after activation by photolysis of nifedipine.

Figure 5

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Same experiments as in Fig. 4 are shown using a faster time base. The flash was performed at time = 0. The upper trace shows activation at 2.5 mM CaCl2 and the lower trace at 0.4 mM CaCl2.

Figure 6. Calcium dependence of active force, half-time for force development and delay after activation by photodestruction of nifedipine.

Figure 6

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A, dependence of active force on extracellular CaCl2. B, dependence of half-time for force development on extracellular CaCl2. C, delay between flash and onset of force development. D, the relation between the half-time for force development and active force. Open and filled symbols show muscles activated using 60 and 120 mM KCl, respectively (n = 5–9). Force is expressed relative to the force at 9 mM CaCl2 and 60 mM KCl.

Following nifedipine photodestruction in the rat portal vein, force developed after a delay of 0.185 ± 0.014 s (n = 5). The delay following ATP release was significantly shorter, 0.048 ± 0.002 s (n = 5). These results show that in this tissue, purinoceptor activation with ATP results in a significantly shorter delay compared with that after nifedipine photodestruction.

DISCUSSION

Nifedipine is a dihydropyridine that blocks the potential-sensitive L-type calcium channel in cardiac and smooth muscle, as well as in non-muscle cells (see McDonald et al. 1994). The smooth muscle cells of taenia coli are of the phasic type and have spontaneous contractile activity in physiological salt solutions. Increasing the extracellular potassium concentration caused an increase in the frequency of contractions and above 10 mM KCl the contractions became sustained. These contractions could be completely inhibited by nifedipine showing that the major pathway for influx of calcium during high-potassium activation is through the L-type calcium channels.

The L-type calcium channel is formed by at least three subunits and the dihydropyridines bind to the α-subunit. This is the major pore forming unit of the channel, but the exact molecular mechanisms for how the dihydropyridines block the channel are not yet resolved (see McDonald et al. 1994). Earlier studies have shown that nifedipine binds with a higher affinity to the channel when it is in its inactive state than in the closed and open states (see McDonald et al. 1994). Consistent with earlier studies (Nelson & Worley, 1989) we find that the Kapp of nifedipine for inhibition of force was lower when nifedipine was added to muscles depolarized with potassium. This is likely to reflect the channels in their open or, maybe more likely, in their inactive states, compared with the situation when nifedipine was added to muscles in normal salt solution, when the channels are mainly in their closed states.

Illumination of depolarized muscles inhibited with nifedipine leads to a rapid development of force to the preinhibition level. This shows that we can, by using photodestruction of nifedipine, simultaneously activate all the smooth muscle cells in the preparation and thereby eliminate diffusional delays. Thus photodestruction of nifedipine can be used as an experimental tool to investigate the activation kinetics of contractions induced by opening of L-type calcium channels. The rate of force development and force amplitude of an initial and a final contraction, performed at 60 mM KCl and 2.5 mM CaCl2, were similar showing that no deterioration of the contractile performance occurred during the experiments. In addition, the muscle did not develop force when illuminated in the N-Krebs solution showing that the ultraviolet light in itself does not lead to activation of contraction.

When CaCl2 was added directly in the organ bath to depolarized muscles, the half-time for force development was about 15 s. The rate of force development increased more than 3-fold when muscles were activated by photodestruction of nifedipine showing that diffusional delay of CaCl2 can influence the activation kinetics. The shortest half-time was about 4.5 s (Fig. 6B), which corresponds to an apparent maximal rate of force development of about 0.15 s−1. This is about 10-fold slower than that observed for force development in maximally and irreversibly thiophosphorylated skinned muscles activated with release of ATP from caged ATP at 37°C (Jaworowski & Arner, 1998). Previous work on skinned muscles has shown that the rate of force development is slower when the muscles are activated by ATP from caged ATP in the presence of Ca2+ or by release of Ca2+ from caged Ca2+ in the presence of ATP (Horiuti et al. 1989) than that observed in thiophosphorylated muscles. The rate of force development in the caged Ca2+ experiments correlated with the rate of phosphorylation (Zimmermann et al. 1995). This shows that the rate of the force development following a rapid increase in Ca2+ is controlled by the rate of phosphorylation. Thus the slow rate of force development found in the present study on intact smooth muscle cells is most likely not rate limited by the cross-bridge kinetics, but rather by the rate of calcium increase or the phosphorylation process itself.

We observed a delay, of about 300 ms, between illumination of the preparation and the onset of force. This delay is not due to an initial internal shortening, taking up slack, since force prior to illumination was between 5 and 15 % of maximal. The physiological events in the activation cascade, from the cell membrane to the myofilaments, that can be responsible for the delay are the opening of the L-type calcium channels, the influx of calcium, intracellular diffusion and buffering of calcium, Ca2+ binding to calmodulin, Ca2+-calmodulin binding to the myosin light chain kinase (MLCK), isomerization of the Ca2+- calmodulin-MLCK complex and phosphorylation of the regulatory light chain on myosin. The kinetics of several of these steps have been determined in earlier studies. The photoconversion of nifedipine is completed within 100 μs (Morad et al. 1983) and the lower time limit for detection of flash-induced current after long depolarizations of skeletal muscle has been found to be about 10 ms (Feldmeyer et al. 1995). This shows that the photodestrucion of nifedipine and the subsequent influx of calcium through the L-channel are not rate limiting for the delay. In isolated smooth muscle cells from the stomach of Bufo marinus it has been found, using fura-2 and a derivative of fura-2 that measures membrane near alterations in [Ca2+], that the initial rate of Ca2+ increase after electrical stimulation was between 1 and 7 μM s−1 (Etter et al. 1996; Yagi et al. 1988). The diffusion of calcium within in the cell cytoplasm is comparatively slow, and a 5-fold increase in bulk cytoplasmic calcium has been observed 100 ms after electrical stimulation of isolated single smooth muscle cells (Kargacin & Fay, 1991). We have at present no data on the rate at which [Ca2+] in the cytosol increases after photodestruction of nifedipine, but the previous results, obtained at room temperature, reviewed above suggest that the photoconversion, the opening of the channels and influx of Ca2+ are not the major contributors to the 300 ms delay at 37°C reported here. This conclusion, that the influx is not the major factor determining the delay, is further supported by the finding that the delay was independent of the extracellular [CaCl2].

The binding of Ca2+ to calmodulin and of the Ca2+-calmodulin complex to MLCK are very fast and will contribute less than 2 ms to the delay between activation and onset of force at 22°C (Kasturi et al. 1993; Török & Trentham, 1994). The equilibrium constant, in vitro, for interaction of the Ca2+-calmodulin complex with MLCK is less than 1 nM suggesting that small increases in cytosolic calcium should be able to maximally activate the kinase (Walsh, 1994). However, not all of the calmodulin in the cell is available for activation of the myofilaments and mobilization or diffusion of calmodulin to the myofilaments might contribute to the delay between flash and force development (Rüegg et al. 1984; Tansey et al. 1994; Zimmerman et al. 1995). In vitro studies have suggested an isomerization of the Ca2+-calmodulin-MLCK complex at approximately 1 s−1 before MLCK is active (Török & Trentham, 1994). Evidence has been provided that this isomerization is not limiting for skeletal muscle MLCK (Bowman et al. 1992), but the issue is still unresolved in smooth muscle. Using skinned muscle fibres and photorelease of Ca2+ it has been shown that the delay between increase in [Ca2+] and onset of force is about 560 ms at 22°C (Zimmermann et al. 1995), which is comparable to the delay observed in the present study considering the difference in temperature. When skinned smooth muscle is pretreated with Ca2+-calmodulin and activated by release of ATP the delay is much shorter, about 50 ms (Zimmermann et al. 1995). These authors concluded that in skinned smooth muscle the two major steps responsible for the delay of activation were the recruitment of Ca2+-calmodulin and the isomerization of the Ca2+-calmodulin-MLCK complex. We have at the moment no data regarding the kinetics of these two processes and they could tentatively be responsible for part of the delay following flash-induced activation in the present study. When intact smooth muscle is activated by purinoceptors using ATP released from caged ATP, force development starts within 80 ms (Sjuve et al. 1995) or 50 ms (rat portal vein, present study). Since the guinea-pig taenia coli does not have contractile responses to ATP (Brown & Burnstock, 1981) we could not perform similar caged ATP experiments on this tissue. However, our experiments on the rat portal vein clearly show a difference in delay between nifedipine photodestruction and ATP release suggesting that these two modes of activation utilize different pathways. Although we cannot exclude a slow activation of MLCK by Ca2+-calmodulin as a cause for the delay in activation of the guinea-pig taenia coli, the caged ATP data suggest that the isomerization process and the phosphorylation of the RLC can be initiated within 50–80 ms after activation. The shorter delay in force development after opening of purinoceptor-activated channels (following release of ATP) compared with the delay after opening of L-type channels (after photoconversion of nifedipine) could reside in the localization of the channels in the cell membrane. The ATP-dependent channels could be more homogeneously distributed over the cell surface whereas the L-channels might have a more restricted localization. Since the delay was Ca2+ independent in the nifedipine experiments we suggest that the major contributor to the delay after flash-induced opening of the L-type Ca2+ channels is the diffusion or mobilization of the Ca2+-calmodulin complex to the myofilaments.

It has been reported that the amplitude and rate of force development during potassium-induced contractions depend more on the rate at which [Ca2+] in the cytosol increases than on the net amount of Ca2+ that enters the cell (van Breemen, 1977). It has been suggested that when the rate of Ca2+ influx is high, most of the Ca2+ is available for activation of the myofilaments, but when the rate is low a part of the Ca2+ is buffered by the superficial sarcoplasmatic reticulum, which decreases the amount of Ca2+ available for activation of the myofilaments (van Breemen et al. 1995). We find, in accordance with earlier studies, that both active force and the rate of force development are dependent on the extracellular [CaCl2] (see van Breemen et al. 1995). The rate of force development correlated with the final level of plateau force. It seems, however, unlikely that the rate of Ca2+ influx directly determines the final force since the relation between extracellullar CaCl2 and the force after flash activation (Fig. 6A) was similar to that obtained by adding CaCl2 to muscles depolarized with high-KCl solutions (Fig. 1B), where the rate of force development is limited by diffusion and 3-fold slower.

As discussed above, the rate of force development is most likely to be controlled by the rate of RLC phosphorylation, which is determined by the amount of Ca2+-calmodulin available at the myofilaments for activation of the MLCK. The activation process is tightly coupled to the intracellular [Ca2+] since rapid changes in the intracellular [Ca2+] can alter the rate of force development (Arner et al. 1998). In the nifedipine experiments the rate of force development was dependent on the extracellular CaCl2 concentration. This indicates that there is a direct coupling between calcium entry and MLCK activity during both high and low rate of Ca2+ influx. In contrast, the delay was similar at high and low [Ca2+] showing that the delay has a different Ca2+ dependence to the rate of force development. If the delay is dependent on Ca2+ buffering by a superficial sarcoplasmatic reticulum it is expected that it would be sensitive to the Ca2+ gradient. Since this was not the case our results suggest that, in this type of muscle, buffering by superficial sarcoplasmatic reticulum does not explain the delay in the force development.

Acknowledgments

This work was supported by grants from the Swedish Medical Research Council (04X-12584 and 04X-8268) and the Medical Faculty of Lund University.

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