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SKELETAL MUSCLE AND EXERCISE |
1 Department of Physiology and Biophysics, Center for Single Molecule Biophysics, State University New York (SUNY) at Buffalo, Buffalo, NY 14214, USA
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Abstract |
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5 mN m–1) that can activate MSCs in the absence of overt stimulation. The inward curvature of patches from mdx mice is eliminated by actin inhibitors. Applying moderate suction to the pipette flattens the membrane, reducing tension, and making the response appear to be stretch inactivated. The pronounced latency to activation in patches from mdx mice is caused by the mechanical relaxation time required to reorganize the cortex from inward to outward curvature. The increased latency is equivalent to a three-fold increase in cortical viscosity. Disruption of the cytoskeleton by chemical or mechanical means eliminates the differences in kinetics and curvature between patches from wild-type and mdx mice. The stretch-induced increase in specific capacitance of the patch, 80 fF µm–2, far exceeds the specific capacitance of bilayers, suggesting the presence of stress-sensitive access to large pools of membrane, possibly caveoli, T-tubules or portions of the gigaseal. In mdx mouse cells the intrinsic gating property of fast voltage-sensitive inactivation is lost. It is robust in wild-type mouse cells (observed in 50% of outside-out patches), but never observed in mdx cells. This link between dystrophin and inactivation may lead to increased background cation currents and Ca2+ influx. Spontaneous Ca2+ transients in mdx mouse cells are sensitive to depolarization and are inhibited by the specific MSC inhibitor GsMTx4, in both the D and L forms.
(Received 16 November 2006;
accepted after revision 18 January 2007;
first published online 25 January 2007)
Corresponding author T. M. Suchyna: Department of Physiology and Biophysics, Center for Single Molecule Biophysics, State University New York (SUNY) at Buffalo, Buffalo, NY 14214, USA. Email: suchyna{at}buffalo.edu
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Introduction |
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The local stimulus for MSCs is not the pipette pressure but membrane stress (Guharay & Sachs, 1984). This stress is a force balance between the hydrostatic pressure and the adhesion energy of the gigaseal that pulls the membrane to the glass (Opsahl & Webb, 1994; Mukhin & Baoukina, 2004; Honore et al. 2006). The sharing of stress between the bilayer and the cortical cytoskeleton affects channel activity. The weakness of muscle membranes in DMD is due to the loss of dystrophin, a membrane-bound reinforcing fibrous protein. Dystrophin buffers membrane tension by cross-linking a group of membrane proteins known as the dystroglycan complex (DGC) (Blake et al. 2002) to the underlying actin cytoskeleton, distributing forces within the cell cortex (Pasternak et al. 1995).
It is possible to assess bilayer stress independently of cortical stress by measuring patch capacitance (Akinlaja & Sachs, 1998). The dynamics of converting pressure to local stress is typified by comparing the rise time of the pressure to the rise time of the patch capacitance. The pressure rise time (0–90%, 2 ms) is much shorter than the 20 ms capacitance rise time observed in astrocytes (Suchyna et al. 2004a), and > 1 s in other cell types (Sokabe et al. 1991; Small & Morris, 1994). Capacitance measurements show that MSC activation correlates with strain, not pressure, and that the strain can be altered by chemical- or mechanical-induced changes in the cytoskeleton (Suchyna et al. 2004a).
We use capacitance and high-resolution video microscopy to investigate whether patch mechanics alone can account for different MSC behaviour in wild-type and mdx muscle cells. The loss of dystrophin increases the tension exerted by actin at rest on the patch membrane, possibly accounting for the SIC behaviour (Franco-Obregon & Lansman, 2002), and increases the viscosity of the patch during pressure application. In addition to higher resting MSC activity, we found that mdx cells have lost the property of fast inactivation. This can increase the steady-state leak of cations. One of the most striking results of comparing patch anatomy and capacitance is that the stretch-induced specific capacitance of both muscle types was of the order of 80 fF µm–2. This is eight-fold higher than expected for biological membranes. This high specific capacitance may arise from tension-sensitive access to membrane pools including T-tubules and caveoli, or to the gigaseal.
The key role of Ca2+ in dystrophy is reflected in its activity during muscle development. Ca2+ transients and the accompanying contractions are important for differentiation of the sarcomeric structure (De Deyne, 2000; Li et al. 2004). Although most of the Ca2+ released during a transient comes from internal stores (Grouselle et al. 1991), the trigger changes over time. In mouse C2C12 myoblasts, the trigger changes from internal to external Ca2+ leaks as the myotube develops (Lorenzon et al. 1997). In both human and mouse myotubes, an increased frequency of transients correlates with a more depolarized resting potential (Lorenzon et al. 1997; Imbert et al. 2001). In DMD, myotubes have an elevated cationic leak that leads to a higher frequency of transients. Some reports attribute the elevated cation leak current to MSCs exposed to increased stress by the loss of dystrophin support (Nakamura et al. 2001; Franco-Obregon & Lansman, 2002). The correlation of the leak with MSCs is supported by the action of GsMTx4, a specific inhibitor of cationic MSCs (Suchyna et al. 2000). GsMTx4 can protect mdx myofibres against the damage caused by eccentric stretch (Yeung et al. 2005). Here we show that GsMtx4 inhibits the spontaneous Ca2+ transients.
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Methods |
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Wild-type (C57BL/10SnJ) and mdx (C57BL/10ScSn–Dmdmdx/J) mice from the Jackson Laboratory (aged 6–12 weeks) were euthanized by cervical dislocation and the flexor digitorum brevis foot muscle dissected. Tissue was transferred to a flasle containing 7 ml 0.1% collagenase B (Boehringer Mannheim) in Dulbecco's modified Eagle's medium–F-12 (DMEM/F-12) solution and incubated at 37° on a micro stir plate (wheaton) at (20 r.p.m.) for 45 min. Muscle tissue was dissected from connective tissue and vasculature. Tissue was placed in a fresh collagenase solution for an additional 15 min on the micro stir plate. More connective tissue was dissected. The tissue was transferred to 10 ml 0.25% trypsin-EDTA (Invitrogen) solution and incubated at 37° for 15 min. The solution was replaced with fresh trypsin-containing solution and the tissue was incubated for an additional 15 min. The tissue solution was gently triturated and pre-plated onto 60 mm culture dishes for 30 min. Cells that did not adhere were re-plated onto coverslips coated with mouse laminin in med 89% DMEM/F-12 with 10% fetal bovine serum and 1% penicillin-streptomycin solution (Sigma). After 4–6 days in culture, myotubes started forming without the use of serum starvation. Myotubes for electrophysiological analysis were used between 8 and 15 days. This procedure has been reviewed and accepted by the animal use committee at SUNY Buffalo.
Electrophysiology
An Axopatch 200B (Axon Instruments, CA, USA) was used for patch clamping, and experimental protocols and data acquisition were controlled by Axon Instruments pClamp9 software via a Digidata 1322A acquisition system. Currents were sampled at 10 kHz and low-pass filtered at 2 kHz through the 4-pole Bessel filter on the Axopatch 200B. All potentials are defined as membrane potentials with respect to the extracellular surface. Electrodes were pulled on a HEKA PIP 5 pipette puller (Digitimer, Hertfordshire, UK), painted with Sylgard 184 (Dow Corning Corp., Midland, MI, USA) and fire polished. Electrodes were filled with KCl-containing saline consisting of (mM): KCl 140, EGTA 5, MgCl2 2 and Hepes 10; pH 7.3, and had resistances ranging from 4 to 8 M
. Bath saline consisted of (mM): NaCl 140, KCl 5, CaCl2 2, MgCl2 0.5, glucose 6 and Hepes 10; pH 7.3. Pressure and suction were applied to the pipette by an HSPC-1 pressure clamp (ALA Scientific Instruments, NY, USA) controlled by the pClamp software. Off-line data analysis was performed with Clampfit and Origin 7.0 software. All voltage polarities are indicated with respect to the intracellular surface of the membrane. The resting membrane potential averaged –60 mV as measured upon breaking into whole-cell mode. Thus, in cell-attached mode, –60 mV pipette potential was near the reversal potential for cation-selective channels and was designated 0 mV. A +60 mV pipette potential is designated as –120 mV (i.e. –60 mV to + 60 mV) or a hyperpolarization producing a negative current deflection. A pipette potential of –100 mV is designated +40 mV (i.e. –60 mV to –100 mV) or a depolarization producing a positive current deflection. In outside-out patch mode, the potentials indicated are the pipette potentials and produce current deflections with the same sign as the potential.
Capacitance measurements
Patch capacitance was measured as previously described (Suchyna et al. 2004a). A dual phase lock-in amplifier [phase lock-in amplifier (PLA), EG&G 5207; Princeton Applied Research, Oak, Ridge, TN, USA] applied a 2 kHz carrier signal of 20 mV root mean squared (rms) to the external input of the Axopatch patch-clamp amplifier. To view ongoing channel activity, we first suppressed the carrier signal with the pipette capacitance transient compensation (see below), and then the current from the patch-clamp, front-panel output was low-pass filtered below the carrier frequency to remove visible currents from small imbalances. The unfiltered carrier signal that was modified by the patch circuit (Fig. 1, a dome circuit and seal circuit) was fed back to the PLA for detection of phase shifts. To define the capacitive component of the patch current after gigaseal formation, we first nulled the carrier signal from the patch clamp using the pipette capacitance compensation. Then, to provide a ‘pure capacity’ reference signal, we unbalanced the circuit by turning the fast-transient pipette compensation knob on the Axopatch 200B clockwise as much as possible without overloading the PLA voltage range. The PLA was then locked to that signal, and the signal was again nulled to leave the single-channel current output free of the carrier. The system was calibrated with a ‘two-cent’ parallel plate capacitor. Two pennies (19 mm diameter) were each soldered to gold pins. The gold pin from one penny was inserted into the patch-clamp headstage, and the second penny was mounted face to face 1 mm away attached to the ground wire. With the capacitance nulled, the distance between the pennies was changed in 1 µm increments using the headstage micromanipulator. A 1 µm displacement produced a 2.5 fF change in capacitance. Changes in patch capacitance are designated 
Cp. The in-phase signal (real part of the patch current, i.e. the patch conductance) mirrored the channel activity, but also allowed us to measure channel activity at the reversal potential. In some figures, the conductance is displayed instead of current. Changes in patch conductance are designated 
p and increasing conductance (channel opening) is always an upward deflection, irrespective of the indicated potential. The system phase accuracy was continuously verified by the insensitivity of Cp to channel opening.
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Patches were visualized with differential interference contrast optics on a Zeiss Axiovert 135 inverted microscope (Oberkochen, Germany). The patch dome was aligned perpendicular to the axis of the Wollaston prisms, and the pipette approached the coverslip at 15 deg to minimize the change in focus with patch motion. The optics consisted of a 63 x oil immersion objective and a condenser made of a 40 x water immersion objective (Sokabe & Sachs, 1990). Images were collected with an iXon DV887 camera (Andor, CT, USA), running at 30 frames s–1 mounted on a 4 x lens. Patch motion was analysed using the ‘tracker’ function in ImageJ (rsb.info.nih.gov/ij/) and corrected for the approach angle of the pipette. Patch motion was monitored at the centre of the dome and the contact position where the seal began (Fig. 1B). Figure 1 also defines the conventions for inward and outward curvature. The height of the dome (h) was calculated by subtracting the position of the dome edge from dome centre. The dome radius of curvature (Rc) was calculated using Rc
= (r2
+
h2)/2h, and the dome area was calculated: A
=
(h2
+
r2), where r is the radius of the pipette at the point of patch attachment.
Ca2+ fluorescence imaging
Myotube cultures (7–24 day) were incubated for 15 min in bath saline containing 1 µM acetoxymethyl ester of Fluo-4 (Fluo-4 AM; Molecular Probes) and then in bath saline alone for another 15 min. All experiments were performed at 21°C. Fluo-4 fluorescence was monitored over 510–550 nm. GsMTx4 was applied by whole-bath perfusion. Normal perfusion saline consisted of (mM): NaCl 140, KCl 5, MgCl2 0.5 and Hepes 10; pH 7.3.
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Results |
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Cation-selective MSCs have been characterized in many cell types, including oocytes (Hamill & McBride, 1992), astrocytes (Suchyna et al. 2004a), cardiomyocytes (Suchyna & Sachs, 2005) and myotubes (Franco-Obregon & Lansman, 2002), and they exhibit similar conductance and gating properties. However, the correlation between channel gating and time-dependent tension changes has only been carried out for astrocyte patch membranes. The astrocyte cortex functions in a less mechanically strenuous environment than that of muscle tissue, but the mechanical properties and the MSC response to pressure can provide a useful benchmark for comparison.
MSCs in myotube cell-attached patches, like those in astrocytes, exhibit weak inward rectification (Fig. 2A and Supplemental Figure S1A) and cation selectivity with slightly lower conductance to Na+ (37 pS) than K+ (43 pS) (Fig. 2A and legend to Supplemental Figure S1). Myotube MSCs show the typical fast voltage-sensitive inactivation at hyperpolarized potentials (Fig. 2B), where channel opening is followed by a reduction in opened probability (Po) and increased occupancy of lower conductance states (Suchyna et al. 2004a). Inactivation was never observed in mdx patches. The voltage-sensitive Po, conductance and inactivation properties are elaborated upon in Supplemental Figure S1. In outside-out patches, these channels are sensitive to both the L and D enantiomers of the MSC inhibitor GsMTx4 (Fig. 2C).
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Resting and pressure-induced MSC activity are altered in mdx patch membranes
MSC activity was assessed using a protocol consisting of 10–20 successive 500 ms suction steps separated by 2 s of relaxation (Fig. 3C and example stimulus sequence at top of Supplemental Figure S2A). Each patch was tested by applying three successive protocols (P1–P3) with increasing suction (P1, –30 mmHg; P2, –50 mmHg; P3, –80 mmHg). There was a 1–2 min rest period between successive protocols. We usually observed 1–2 channels per patch corresponding to 1 channel µm–2 (Sachs, 1990). Wild-type and mdx patches displayed similar amounts of MSC activity in cell-attached mode. The percentage increased with suction so that at P3 more than 70% of patches contained MSC current (Fig. 3A). However, the percentage of patches displaying MSC activity in the rest periods between protocols was significantly higher in mdx than in wild-type (Fig. 3B). The resting activity in wild-type patches decreased to 0% at P3 compared with nearly 40% of mdx patches. During the protocol we assessed activation and deactivation by plotting the changes in Po and the average MSC current during the stimulation and at the end of intervening relaxation periods (Fig. 3C–E). Wild-type patches again display a mild ‘mechanoprotective’ (Morris, 2001) property where MSC activity at the end of the relaxation period is slightly reduced with greater strength protocols (Fig. 3D). Initially (at P1), Po and the average current during relaxation were not statistically different between wild-type and mdx patches. However, with further stimulation (during P2 and P3) they differed significantly. As expected, activity in wild-type and mdx patches increased with the magnitude of the stimulus (Fig. 3E). These data suggest that MSC sensitivity to pressure has not been altered in dystrophic membranes, but the rate of deactivation and the ability to ‘adapt’ to pressure stimuli has changed. This behaviour is similar to that observed previously in myotubes (Franco-Obregon & Lansman, 2002), where resting MSC activity increased in mdx patches over a period of several minutes, while in wild-type patches it decreased. Resting MSC activity in mdx patches was also sensitive to previous suction and pressure stimulation (Franco-Obregon & Lansman, 2002) and depolariziation (Gil et al. 1999).
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We monitored patch curvature at rest between P1 and P2. The patches had an inward curvature that was much more pronounced for mdx than wild-type myotubes (Fig. 4A, wild-type and mdx). For patches of similar diameter, mdx had half the radius of curvature of wild-type patches (mdx, 4.8 µm; wild-type, 10.7 µm; Fig. 4B). The resting curvature disappeared when cultures were treated with the actin reagents cytochalasin D and latrunculin A (Fig. 4A and B). Actin appears to be pulling normal to the membrane. The force exerted by actin can be estimated by the pressure required to flatten the patch, 5 mmHg or 667 Pa (data not shown). Expressed across a patch area of 5 µm2, this is a net force of 3 nN. If we assume that each actin exerts a force of 5 pN (Nishizaka et al. 2000), then the effective pressure corresponds to a density of 130 actin tethers µm–2. The greater curvature of mdx patches suggests that either the membrane is more compliant, or that there is more actin, which seems unlikely because actin is bound by dystrophin. The higher compliance of mdx patches is consistent with a larger fraction of membrane tension appearing in the bilayer and hence higher resting MSC activity.
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MSC activity in mdx myotubes shows a number of properties related to differences in patch mechanics rather than intrinsic channel kinetics. Figure 5 shows simultaneous recordings of patch capacitance, conductance (channel openings at any voltage produce an increase in conductance or positive deflection), dome height (the axial distance from the middle of the patch to the position where the patch touches the glass) and the calculated area change. Arrows indicate where the frames were taken. The first image shows an mdx patch with an inward curvature at rest (Fig. 5A). Alternating positive and negative 30 mmHg pressure steps were applied for about 6 s.
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Ap) and Cp became positive and an MSC became active (upward deflection in the conductance trace). This suggests that membrane tension in the vicinity of the channel did not increase until Ap and Cp increased. The presence of resting inward curvature may account for published reports of delayed activation (Small & Morris, 1994).
Although the membrane potential was –60 mV, the MSC shown in Fig. 5 did not inactivate with time, even after the patch curvature and Cp reached steady state (Fig. 5C). The channel did not close until the patch returned to zero curvature after the suction step. At 0 mmHg the patch reversibly returned to its inward curvature (Fig. 5D). With positive pressure the curvature increased and Ap and Cp displayed a nearly linear monophasic response, that was larger than the resoponse for suction (Fig. 5E). Positive pressure causes extensive peeling of the membrane away from the glass (Opsahl & Webb, 1994; Mukhin & Baoukina, 2004), in the same manner as tape being pulled back off a surface. It is interesting that even though Ap and Cp were greater with positive than negative pressure, no MSCs were activated. Thus, if we assume that MSCs are gated by bilayer tension, the tension developed from positive and negative pressure are not equivalent and the system is clearly non-linear, as previously reported (Akinlaja & Sachs, 1998). It is unlikely that the channel is responding to changes in membrane curvature because the radii of curvature of the patch are too large to generate much energy over the dimensions of a channel (Markin & Sachs, 2004; Wiggins & Phillips, 2005). The insignificant role of global curvature is also suggested by the fact that channels do not activate when the membrane wrinkles upon passing from inward to outward curvature (Honore et al. 2006).
Comparison of wild-type and mdx MSC pressure sensitivity
MSC sensitivity to steady-state pressure in dystrophic rodent cells has been reported to both increase (Nakamura et al. 2001) and decrease (Franco-Obregon & Lansman, 1994; Franco-Obregon & Lansman, 2002), the latter being described as SIC behaviour. Applying 500 ms suction steps to mdx patches initially produce Cp
0 as the patch changes from inward to outward curvature (Fig. 6, first four steps), mimicking the transient negative response of the 6 s suction step in Fig. 5. During this time, constitutively active currents were sometimes suppressed (see Fig. 6, arrow in first step) and MSCs were rarely activated. With repeated suction steps, the patch reorganized. The outward curvature reached a maximum (Fig. 6, steps 5–6) where Cp increased (indicating local strain) and MSCs activated. This breakdown of the relationship between pipette pressure and tension during restructuring may account for the reported lower pressure sensitivity and SIC behaviour of MSCs from mdx patches (Franco-Obregon & Lansman, 1994). The sudden increase in MSC sensitivity with repeated stimulation is similar to that reported for rat ventricular cells (Bett & Sachs, 2000).
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Cp and conductance (p) from multiple patches. ‘Restructuring’ in mdx patches (Fig. 7A) was marked by conversion from negative to positive Cp and little change in MSC-induced p. Wild-type patches displayed only a brief restructuring: a small positive Cp for the first step, and both Cp and p remaining stable over the remaining steps. After six successive pressure steps, the response in mdx patches reached steady state with Cp and p values similar to those in the wild-type, suggesting that after restructuring the force distribution was similar. However, even after restructuring, the kinetics of Cp was slower for mdx than wild-type patches (Fig. 7B). The loss of dystrophin and the accompanying alteration of other membrane and cytoskeletal components produced a difference in effective viscosity and/or elasticity. The ratio of p to Cp in steady state was 0.075 ± 0.01 (mdx) and 0.086 ± 0.01 pS fF–1 (wild-type), showing that MSCs in mdx are only slightly less sensitive to stress than those in wild-type patches. This agrees with the slight reduction in ‘pressure sensitivity’ of MSCs from mdx myotubes (Franco-Obregon & Lansman, 1994) and suggests that dystrophy does not alter the intrinsic MSC ‘tension sensitivity’.
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To better quantify the differences in Ap and Cp between mdx and wild-type patches (Fig. 7B), we selected data from patches with similar diameter, pressure and stimulation history (i.e. the number of protocols applied before the recording, Fig. 8). These averages revealed significant differences in the kinetics of Cp and Ap (Fig. 8A and B). Cp did not reach steady state within 500 ms so we modelled the response with a single exponential and extrapolated that to estimate the maximum Cp at steady-state (Cpmax, Table 1). We did not use longer steps because they caused patch creep.
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Cp to a viscoelastic model for a Kelvin body under constant stress (Fung, 1981; Bausch et al. 1998) (Fig. 8A dashed lines). We analysed the kinetics of Cp rather than Ap because the resolution of Cp was greater. The Cp response of wild-type and mdx were proportional to Ap in both rate and magnitude (linear relationship in Fig. 8C). However, the average slope in Fig. 8C was an incredible 85 fF µm–2; far too large for biological membranes (10 fF µm–2), but comparable with the reported apparent muscle membrane capacitance (see below for discussion of the specific capacitance). Cp was converted to Ap using 85 fF µm–2, resulting in solutions of the Kelvin model as:
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| (1) |
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| (2) |
A/A is the relative area change, Tss is the steady state tension, 
and 
are the time constants for relaxation under constant strain and stress, respectively. k1 is the stiffness of the spring in series with the dashpot 
1, and k0 is the parallel spring constant.
The responses of wild-type and mdx patches do not show the rapid initial elastic deflection expected from a series elastic component (but see Fig. 8F for an example of the initial elastic response in an outside-out patch that appears to reflect simply bilayer stretching). In eqn (1) a step stimulus is assumed, but the radius of curvature of the patch changed with time, so that the actual stimulus was not a step function. As a compromise, we replaced the driving force with the predicted steady-state tension, Tss
= (P
x
RCmax)/2, where P is the applied pressure and RCmax is the maximum patch radius of curvature determined by fitting the Ap kinetics with an exponential. Although this model does not account for the three-dimensional nature of the patch (including the membrane–cytoskeleton complex and the changing stimulus) it suggests the relative contributions of viscosity and elasticity.
Table 1 lists the values for the Kelvin parameters from wild-type and mdx patches with, and without, actin inhibitors. Wild-type and mdx untreated patch dynamics were fit well by the single exponential form of the Kelvin model. The maximal area changes were similar: mdx:wild-type = 1.08, as expected from the stiffness ratio of the parallel spring elements (k0–
mdx: k0– wild-type = 1.17) that bear the entire load at steady state. In mdx patches, the series elements 1 and k1 both increased by similar amounts (mdx:wild-type = 3.4 and 4.1, respectively). An increase in viscosity produces a proportional increase in the stress relaxation time constant. However, the effect of k1 on the time constant is cross-correlated with that of k0. k1 contributes less to the time constant, but more to the initial elastic deflection if it is stiffer than k0. Here k1
2 orders of magnitude stiffer than k0 so that the difference between the k1 elements of mdx and wild-type has little effect on the stress relaxation rate. Thus, loss of dystrophin, and other defects linked to its absence, cause an increase in the relaxation time mainly through changes in viscosity. This viscosity probably reflects the difference in time to make and break cytoskeletal crosslinks rather than lipid flow (see Discussion).
If the viscosity arises from cytoskeletal interactions, the relaxation times should decrease with cytoskeletal disruption. When myotubes are treated with actin inhibitors, the differences between mechanics of mdx and wild-type disappear and the rising phases are more rapid and biphasic showing at least two distinct contributions (Fig. 8D and E, and Table 1). Forming an outside-out patch disrupts the cytoskeleton and eliminates the energy sources for active remodelling. Outside-out patches from wild-type and mdx myotubes had nearly identical Cp kinetics (Fig. 8E). Cp in outside-out patches displays three phases of relaxation: a fast elastic deflection, a slow relaxation and linear creep. We were able to visualize a few outside-out patches and the relationship between Cp and Ap was linear with a slope of 80 fF µm–2 (data not shown). Because the effect of peeling is much smaller in outside-out patches than in cell-attached or inside-out patches (the stress is parallel to the glass instead of anti-parallel), the large specific capacitance implies that the tension-gated areas are part of the membrane structure and not a result of peeling of the gigaseal.
Contributing factors to the patch capacitance
Capacitance is a measure of the electrically accessible membrane area, but this may not match the histologically visible area. The dome is ‘nominally’ a spherical cap, so measuring Ap only requires measurement of the planar patch diameter and height of the dome. The dome height was monitored at video rates by subtracting the position of the patch edge from the position of the dome centre (see Fig. 1B). Ap changed proportionately to Cp, and the specific capacitance (Cs
=
Cp/Ap) did not change with pressure (Fig. 8C). However, Cs was much larger than the 10 fF µm–2 (or 1 µF cm–2) as expected for bilayers (Sokabe & Sachs, 1990; Solsona et al. 1998; Akinlaja & Sachs, 1998). We consistently observed Cs of 80 fF µm–2 in both cell-attached and outside-out patches. Note that Cs is not that of the resting membrane, but the Cs of the stretch-induced component. A high value of Cs implies that the relevant anatomical area is being underestimated. Where is this pool of membrane? There are two possible sources: (1) peeling of membrane from the seal (Opsahl & Webb, 1994; Mukhin & Baoukina, 2004); or (2) opening pores to subsarcolemmal membrane structures such as caveoli or to T-tubules, where the pore conductance (Lollike et al. 1998; Sombers et al. 2004) is modulated by membrane tension (Dulhunty & Franzini-Armstrong, 1975).
We can estimate the magnitude of the peeling effect if we treat the pipette and seal as a cylinder and ignore the distributed access resistance of the seal (Suchyna et al. 2004a). Peeling the seal by a distance z increases the capacitance by 2rzCbilayer where r is the radius of the pipette and Cbilayer
= 10 fF µm–2. Assuming r
= 1.25 µm, we would have to expose the seal for only 0.4 µm to account for the Cpmax in normal and mdx patches of 30 fF. The accuracy of this estimate can be gauged with the deliberate peeling experiment shown in Fig. 5, where we applied positive pressure to a 2.5 µm diameter patch producing a Cp of 37 fF. The cylindrical peel model predicts that z
= 0.47 µm, and the optically measured z value was 0.35 µm. This result implies that the peeled membrane has a Cs of 10 fF µm–2, in close agreement with the accepted Cs of biological membranes. Analysis of the edge motion during suction steps actually shows the edge moving upwards by approximately 0.35 µm instead of downwards (see Supplemental Figure S2 for thorough analysis of pressure, voltage and cytoskeletal effects on patch-edge motion). However, seal-thickness changes below the optical resolution could account for a significant drop in access resistance exposing new membrane (see Discussion below, and appendix in Suchyna et al. (2004a)).
Another contributing factor to the large Cs may be the inner surface of myotube membranes that are extensively fenestrated with caveolae and developing T-tubules (Kelly, 1971; Parton et al. 1997). In differentiated myofibres, surface openings of caveolae are known to be sensitive to tension (Dulhunty & Franzini-Armstrong, 1975). Fusion or opening of a 75 nm diameter caveolus would produce a capacitance change of 0.2 fF. Examination of single Cp traces did not reveal any obvious quantized changes in capacitance due to the relatively high noise level (5–10 fF) of myotube patches. In addition, T-tubules and caveolae can form deep tubular/reticular structures in developing myotubes that generally lie tangential to the sarcolemma and have multiple connecting points to the surface (Kelly, 1971; Parton et al. 1997) that may not be simple quantized steps in capitance. see Discussion for a further explanation of how these sources may contribute to Cp.
Inactivation and the cytoskeleton
About 60% of both wild-type and mdx cell-attached patches displayed MSC activity at –50 mmHg (Fig. 3A). However, > 95% of outside-out patches showed MSC activity. This correlates with data from astrocytes where MSC activity was 17% greater in outside-out patches than in cell-attached patches (Suchyna et al. 2004a). This increase in activity is likely to be the result of disrupting the mechanoprotective cytoskeleton. MSCs also show time-dependent inactivation following a step of tension (Hamill & McBride, 1992; Niu & Sachs, 2003; Suchyna et al. 2004a; Akitake et al. 2005; Honore et al. 2006) that is also sensitive to cytoskeletal integrity (Hamill & McBride, 1992; Suchyna et al. 2004a). We observed inactivation in 10% of wild-type cell-attached patches, but never in mdx patches. The frequency of observing inactivation in wild-type cell-attached patches is similar to that in astrocytes (13%). It is quite remarkable that, given the sensitivity to cytoskeletal integrity, 50% (n
= 20) of wild-type outside-out patches showed inactivation (Fig. 9), but again mdx patches never showed inactivation (n
= 16). Apparently the reported strong link between cytoskeleton and membrane created by dystrophin (Campbell & Kahl, 1989; Rybakova et al. 2000), plays a key role in inactivation.
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The activity of MSCs in resting cells cannot be determined using a patch clamp because patches are always under tension due to adhesion of the gigaseal. However, MSC activity in resting cells can be assessed by applying the specific inhibitor GsMTx4. GsMTx4 inhibits MSCs in both wild-type and mdx cultured myotubes, and has been shown to inhibit the rise in [Ca2+]i associated with eccentric contractions (Yeung et al. 2005). Both the D (Fig. 2C) and L (Yeung et al. 2005) enantiomers of GsMTx4 inhibit MSCs in mdx and wild-type cultures.
Myotubes exhibited spontaneous Ca2+ transients that were potentiated by shear stress from local perfusion and inhibited by GsMTx4 (See video in Supplemental Figure S3). The incidence of Ca2+ transients have been correlated with increased sarcolemmal leak currents (Lorenzon et al. 1997). In contrast to the spontaneous transients, Ca2+ transients initiated by voltage-clamp depolarizations were unaffected (Fig. 10A and B). We found that at later stages in development (after 18 days in culture) transients in 65% of the wild-type or mdx patches were completely inhibited by GsMTx4. The remaining 35% showed a decreased frequency.
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Discussion |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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The MSCs of myotubes are similar to those of the well-characterized rat astrocytes (i.e. weak inward rectification, 40 pS conductance, voltage-sensitive Po, voltage-sensitive fast-inactivation and GsMTx4 sensitivity). The voltage-sensitive Po is similar to that already reported for myotube MSCs (Franco-Obregon & Lansman, 2002), but our unitary conductance was 17 pS larger under similar ionic conditions. Franco-Obregon & Lansman (2002) did not characterize the fast inactivation or GsMTx4 sensitivity.
The loss of dystrophin did not appear to affect the inherent channel sensitivity to tension (Figs 3E and 7B). However, the mechanoprotective adaptation of wild-type cells, where resting MSC activity decreased with repeated stimulation and the rate of MSC relaxation increased, did not appear in mdx cells. In fact, mdx patches showed the exact opposite behaviour. The higher resting activity in mdx patches may reflect a shift in cortical tension from dystrophin to the bilayer and/or a change in the number of active MSCs. This is similar to the stimulation-induced increase in MSC activity that caused the conversion of SACs to SICs (Franco-Obregon & Lansman, 2002).
During outside-out patch formation that causes cytoskeletal disruption, wild-type channels frequently maintain the ability to rapidly inactivate, while mdx patches never do. The loss of MSC inactivation in mdx patches correlates with the effect of cytoskeletal disruption observed in non-muscle cell types that possess a less-resilient cortex (Hamill & McBride 1992; Suchyna et al. 2004a). Although we cannot directly relate the observed mechanoprotection, deactivation and fast inactivation in patches to the whole cell, these processes are all biased towards increased MSC activity in the mdx cells.
The sequence identity of the cationic MSCs in myotubes and fibres is currently unknown, but they may be members of the transient receptor potential (TRP) family (Maroto et al. 2005) which are blocked by GsMTx4 (Spassova et al. 2006). TRPC1 has been implicated in the elevated Ca2+ influx in dystrophic myofibres (Vandebrouck et al. 2002) and is associated with caveolae (Brazer et al. 2003). TRPV2 has also been shown to be sensitive to stretch (Muraki et al. 2003) and redistributed to the sarcolemma in dystrophic muscle (Iwata et al. 2003).
Electroanatomy of the patch
MSCs are experimentally stimulated by pressure steps, but the conversion of hydrostatic pressure to local stress is a function of the constitutive mechanical properties of the cell cortex and the patch geometry. A striking feature of mdx patches is the resting inward curvature which is nearly twice that of wild-type patches. The curvature is removed by disrupting actin, but how much of this internal stress is active in vivo remains to be determined.
Why might the loss of dystrophin exaggerate the pull of actin? Dystrophin could increase the membrane bending stiffness, either by having a bending resistance of its own (Saadat et al. 2006), or if it were under tension, acting as a ‘clothesline’. We measured the suction (667 Pa) required to make the patch flat, and using Laplace's law estimated the normal force produced by actin as 1.6 mN m–1 for mdx and 3.6 mN m–1 for wild-type. If we assume that the force exerted by actin is the same in wild-type and mdx, then the bending resistance of the wild-type cortex is more than twice that of mdx. This is in agreement with stiffness measurements by Pasternak et al. (1995) who showed a three- to four-fold decrease in mdx cortex stiffness assessed by indentation with a glass filament. This decreased bending stiffness may lead to increased tension and increased MSC activity.
The initial concave patch curvature and the increased viscoelastic relaxation time caused a delay in generating patch tension in response to a step of suction. Initially, the suction step causes the dome to move upwards and resting tension to decrease as the area of the initial concave spherical dome is larger than the disk required to span the pipette. The patch may actually become wrinkled as it passes through the region of zero mean curvature (Fig. 11) (Honore et al. 2006). With this loss of tension, constitutively active channels can deactivate showing that local curvature at the level of optical resolution is not a stimulus for MSC activation (Markin & Sachs, 2004; Wiggins & Phillips, 2005). In addition, the capacitance does not change because the effective area has not changed. Although we did not observe significant levels of SIC activity, the mechanical pre-stress to inward curvature of mdx membranes and the delay in generating patch tension may create the conditions for the SIC response (Morris & Sigurdson, 1989; Franco & Lansman, 1990; Franco-Obregon & Lansman, 2002). After passing though zero curvature, the loose membrane re-anneals to the glass and decreases patch capacity.
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Cp reaches a steady state, but the kinetics to reach this steady state are three times slower in mdx patches. This difference appears to be a result of increased viscosity rather than reduced elasticity. In both types of muscle, Cp kinetics are significantly slower than in non-contractile cells (Suchyna et al. 2004a).
The most striking finding from Cp was that muscle cell patches show a stretch-activated Cs nearly eight times greater than the resting Cs of a bilayer. This implies a lack of optical resolution for detection of the electrically exposed membrane. For comparison, in astrocyte patches Cpmax is 0.5–1 fF, with a rise time composed of fast (
= 0.014 s–1) and slow (
= 0.16 s–1) components (Suchyna et al. 2004a). The fast rate is 20 times faster than in myotubes, and Cpmax is 20 times less. The stretch-induced Cs of astrocyte patches is close to the nominal bilayer value of 10 fF µm–2 (i.e. the stretch-induced change in patch area involves a flow of new bilayer into the patch; Sokabe et al. 1991). When actin polymerization is inhibited, the myotube membrane becomes more like the astrocyte membrane, with smaller Cp and biphasic kinetics. Where does the huge Cp of muscle cells originate? There appear to be only two, non-exclusive sources: peeling of the seal and opening of vesicles or T-tubules. These mechanisms are illustrated in Fig. 11 and discussed below.
Seal peeling
Peeling of the gigaseal occurs as the component of tension normal to the pipette wall exceeds the adhesion energy. The membrane–glass adhesion energy is estimated from Opsahl & Webb (1994):
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| (3) |
Apmax in Table 1. At –50 mmHg the adhesion energy is 5.5 mN m–1, about half of the nominal accepted lytic tension (Akinlaja & Sachs, 1998). For pure lipid bilayers, Opsahl & Webb (1994) reported adhesion energy of 0.5–4 mN m–1. The higher adhesion energy we observe may result from membrane proteins denaturing against the glass.
What is the tension in a patch? At the onset of a pressure step, Laplace's law predicts that a flat patch of elastic membrane will undergo a rapid increase in tension followed by relaxation as the radius decreases. In wild-type patches the mean tension relaxed to 8 mN m–1 within 500 ms. However, due to the slower kinetics, it never fell below 10 mN m–1 in mdx patches. These estimates are the upper limit because they assume that the patch is a two-dimensional elastic sheet and not a three-dimensional viscoelastic gel. The lytic tension of a membrane is not fixed, but depends upon the duration of the stimulus; a short stimulus requires more tension to lyse the membrane than a longer one (Evans et al. 2003). Bacterial MSCs require near lytic bilayer tension for activation (Sukharev et al. 1999). If this were true for eukaryotic MSCs, it may account for why they have been so difficult to activate in whole-cell preparations (Morris & Horn, 1991; Zhang & Hamill, 2000). Cells are much larger than a patch and are more likely to contain mechanical defects. Although we do not know how the apparent viscosity increase in mdx patches is manifested in situ, it is known that mdx cells rupture more easily than wild-type cells (Petrof et al. 1993; McCarter & Steinhardt, 2000).
The magnitude of stress does not address the stress-sensitive Cs. If Cp is due to membrane peeling from the seal, why do we not see that in the images? There are a number of possible explanations. Minor changes in seal thickness (far below optical resolution) could lead to a large drop in access resistance (See Supplemental Figure S2 and discussion in appendix of Suchyna et al. (2004a)). Even if we assume that the seal has a discrete boundary, a change in patch position of a few tenths of a micrometer could account for Cp. The peeled area would be 2rz, where r is the radius of the pipette and z is the change in seal position. The seal, however, is not discrete but distributed. We have observed channel activity where the real part of the impedance is out of phase with the single-channel currents. That can only be explained if the carrier signal has changed phase at the location of the channel; this is what happens to signals as they pass through a dissipative cable. The seal resistance forms an annular electrotonic cable with the membrane capacitance (Suchyna et al. 2004b).
Tension-sensitive vesicle fusion and T-tubules
An alternative explanation for the large Cp is the presence of vesicular or tubular structures that have a stretch-dependent access resistance (Lollike et al. 1998). Cytoskeletal dynamics could affect the kinetics of opening in the same way as MSCs. While we do not observe vesicular structures in the patch using light microscopy, electron microscope images of adult myofibres show a high density of caveolae with 19 openings µm–2 (Dulhunty & Franzini-Armstrong, 1975; Bonilla et al. 1988). Caveolae are vesicles 25–150 nm in diameter that function as a scaffolding for many signalling functions (for review see Anderson, 1998). Caveolae in combination with the T-tubules create an apparent plasmalemma-specific capacitance of 90 fF µm–2 in adult myofibres (Garcia et al. 1992).
Caveolae are connected to the sarcolemma by a 10–20 nm diameter stalk, and about 50% are accessible to the extracellular solution, as judged by ruthenium red staining (Thorn et al. 2003). Multiple caveolae can bud off a primary one, so that in adult fibres there is an average of two caveolae connected in series per stalk; this effectively doubles the surface area (Dulhunty & Franzini-Armstrong, 1975). However in myotubes after 5–11 days in culture, a much more extensive system of tubular/reticular elements connect to the surface and penetrate deep into the cytoplasm (Kelly & Price, 1997; Parton et al. 1997). Although the surface area of this network has not been accurately quantified, it is significantly larger than the two-fold increase in surface area expected from caveolae in adult myofibres. These structures have been identified as networks of fused caveolae that are the genesis of the T-tubules (Parton et al. 1997; Galbiati et al. 2001a; Lee et al. 2002). These immature T-tubules are tangential to the plasmalemma and make numerous connections to the surface.
In adult fibres, the diameter of caveolae stalks are tension sensitive and at tensions above a critical sarcomere length t
) or KCl (
) as the primary charge carriers in the pipette solution. A weak voltage-sensitive inward rectification, and weak selectivity of K+ over Na+ are evident. For comparison, astrocyte MSC unitary current data from 
; KCl, 
). B, single-channel current records from a cell-attached patch at hyperpolarized (–100 mV) and depolarized (40 mV) potentials showing voltage-sensitive fast inactivation at hyperpolarized potential via both decreased opening probability (Po) and increased occupancy of substates. C, average MSC current from an outside-out patch at –60 mV, before (control black trace, 13 pressure steps) and during the application of the D-enantiomer of GsMTx4 (grey trace, 23 pressure steps) showing > 90% inhibition of the MSC current. The data are an example of the non-inactivating currents that can be seen in outside-out patches.
P2
= 0.05 (wild-type versus mdx at P2, P
= 0.04; at P3, P
= 0.03). The MSC activity elicited during the 500 ms suction step (E) shows the change in Po (
