Neurosciences, Loeb Health Research Institute, Ottawa Hospital, Ottawa, Ontario, Canada K1Y 4E9
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ABSTRACT |
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Mechanosensitive (MS) channels, ones whose open probability
varies with membrane tension in patch recordings, are diverse and
ubiquitous, yet many are remarkably insensitive to mechanical stimuli
in situ. Failure to elicit mechanocurrents from cells with abundant MS
channels suggests that, in situ, the channels are protected from
mechanical stimuli. To establish what conditions affect MS channel
gating, we monitored Lymnaea neuron
stretch-activated K (SAK) channels in cell-attached patches after
diverse treatments. Mechanosensitivity was gauged by rapidity of onset
and extent of channel activation during a step pressure applied to a
"naive" patch. The following treatments enhanced
mechanosensitivity: actin depolymerization (cytochalasin B),
N-ethylmaleimide, an inhibitor of
ATPases including myosin, elevated Ca (using A-23187), and osmotic
swelling (acutely and after 24 h). Osmotic shrinking decreased mechanosensitivity. A unifying interpretation is that traumatized cortical cytoskeleton cannot prevent transmission of mechanical stimuli
to plasma membrane channels. Mechanoprotection and capricious mechanosensitivity are impediments to cloning efforts with MS channels.
We demonstrate a potpourri of endogenous MS currents from
L-M(TK) fibroblasts;
others had reported these cells to be MS current null and hence to be
suitable for expressing putative MS channels.
tension; actin; snail neuron; fibroblast
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INTRODUCTION |
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DIVERSE MECHANOSENSITIVE (MS) ion channels are widely found by applying suction during patch-clamp recording, but direct evidence for in situ mechanical gating of the channels is rare (42). MS channels in Xenopus oocytes, in Escherichia coli, and in molluscan neurons are examples. For these, when whole cell currents can be elicited mechanically, it is only at near-rupture tensions (1, 16, 32, 50, 51).
Cytoskeletal integrity in situ vs. in patches might explain the general discrepancy between patch and whole-cell MS channel behavior. For molluscan neuron stretch-activated K (SAK) channels, the stimulus history of the patch, neuronal age in culture, and cytochalasin D all affect channel responses (45). In skeletal muscle treated with cytochalasin D, stretch-activated cation channels are hypermechanosensitive (14). In the case of N-methyl-D-aspartate type glutamate channels (38), cytoskeletal damage might be the reason that general cytoplasmic "rundown" coincides with "runup" of mechanosusceptibility. The first rigorous study of the importance of patch history on MS channel behavior dealt with the phenomenon of adaptation by Xenopus oocyte MS channels; adaptation is abolished by repeated stimulation (15).
Even if MS channels are not all mechanotransducers, there are biophysical, physiological, and pathological reasons for examining determinants of their mechanosensitivity. 1) MS channels are integral membrane proteins with gating transitions susceptible to mechanical inputs. The need to deal with mechanical forces would have been factored into the evolutionary design of such proteins and of their links to membrane skeleton. Insofar as mechanical inputs hinder proper function, mechanoprotective features (12) should have evolved. 2) Susceptibility to mechanical inputs should also have created opportunities for physiological signaling; some "mechanosusceptible" channels could be facultative mechanosensors, with responsiveness tuned via cytoskeletal dynamics. In fish embryos, the cytoskeleton is cyclically labile and SAK channels are cyclically active (5). Cell adhesion alters MS channel activity in macrophages (28). In swollen epithelial cells performing volume regulation, actin depolymerization and enhanced stretch-activated channel activity go together (43). In auditory hair cells, mechanoprotection and mechanostimulation of MS channels are exquisitely orchestrated to render mechanical inputs effective exclusively along one axis and provide for adaptation (11). 3) Mechanosusceptible channels may become pathological "leaks" when traumatic or genetic damage compromises the shock-absorbing capabilities of the membrane skeleton. This may be a factor in ischemia (31), it may contribute to the myopathy of dystrophic muscle (29), and, in demyelinated neurons, it may explain axonal hypermechanosensitivity (48).
Here we examine the mechanosensitivity of SAK channels in Lymnaea neurons subjected to several treatments often used in cell physiology. The native function of the Lymnaea channel is inferred from Aplysia, whose neurons all have channels with permeation, gating, and pharmacological properties like the Lymnaea SAK channel (55). In identified Aplysia neurons, the channels are characterized both as "serotonin channels" [S channels; voltage-insensitive K channels that modulate neuronal excitability via the neurotransmitters serotonin and Phe-Met-Arg-Phe-NH2 (FMRF-amide)] (17) and as SAK channels (56). Presumably, Lymnaea, Aplysia, and Drosophila (58) SAK channels as well are homologous (see Refs. 46, 47). Why they are mechanosusceptible in patches is unknown, but in the whole cell they activate mechanically only with inflation to near rupture (33, 57).
A poor understanding of the variable mechanosensitivity of MS channel
complicates efforts to clone and functionally express eukaryotic MS
channels. A "null" cell type for heterologous expression of
putative MS channels is required, and
L-M(TK) cells have been
said to lack endogenous MS currents (24). The implication is that MS
currents elicited in
L-M(TK
) cells transfected
with putative MS channel cDNA arise from recombinant channels. In our
hands, as demonstrated here,
L-M(TK
) cells were not MS
current null. We caution that unintentionally greater trauma delivered
to experimental patches (vs. controls) may facilitate the activation of
endogenous MS channels in these patches.
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METHODS |
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Molluscan Neurons
Cell preparation. Central neurons were prepared from mature Lymnaea stagnalis as described previously (43, 57) and cultured in normal saline (NS; in mM: 55 NaCl, 1.6 KCl, 2 MgCl2, 3.5 CaCl2, and 5 HEPES-NaOH adjusted to pH 7.6) with 5 mM glucose and 0.1 mg/ml gentamicin sulfate on plastic culture dishes. Cells were kept 4-6 days in culture before patch-clamp recording.
Patch-clamp recordings. For snail neurons, cell-attached recordings were made with NS in the bath and a high-K solution in the pipette [in mM: 55 KCl, 2 MgCl2, 1 tetraethylammonium (TEA) chloride, and 5 HEPES-KOH adjusted to pH 7.6]. TEA was included to select for SAK over other K channels, particularly Ca-activated K channels; ~50 mM extracellular TEA diminishes single SAK channel currents by only 50% (47).
Pipettes were made from borosilicate glass (ID 1.15 mm, OD 1.65 mm; N51A, Garner Glass, Claremont, CA) on a List L/M-3P-A (Darmstadt, Germany) pipette puller, coated with Sylgard 184 (Dow Corning, Midland, MI) and fire polished. Tip outside diameter was 2-3 µm before fire polishing and 1.5-2.5 µm afterward, with a resistance of 2-4 MMechanical stimulus.
Step changes of pipette pressure were achieved as described previously
(45) using two pressure transducers (Biotek Instruments, Winooski, VT),
one to establish pressure and the other (modified so its output was
suitable for an analog-to-digital converter) to measure the pressure
acting on the patch membrane. The standard pressure used for "first
hits" was 80 mmHg. In some cases, as noted, other pressures
were used. Manual valves initiated and terminated pressure steps (see
Fig. 1A and
1Bi in Ref. 45), the durations of
which varied according to the response. Once channel activity had
plateaued for ~1 s or had reached a maximum and begun to fall, a step
was terminated.
Analysis. Channel currents were recorded using an Axopatch 1D (Axon Instruments, Foster City, CA) connected to a microcomputer via a TL-1 interface (Axon Instruments). pCLAMP (Axon Instruments) was used to digitize and export records to SigmaPlot 5.1 (Jandel Scientific, Corte Madra, CA) for analysis. Current, voltage, and pressure were sampled simultaneously at 9-ms intervals. Ideally, first latency is used to describe delayed channel responses. However, SAK channels often have a nonzero resting activity that contributes sporadic events to the delay period (from initiation of the pressure step to the onset of stimulated activity). Therefore, we quantified delay using a running average of current as previously described (45). Responses to first hits (the very first pressure stimulus delivered to a patch other than the small pressure used for seal formation) were used for all patches unless indicated. The stimulus duration varied, depending on the delay. Once channel activity started, the stimulus was continued for several seconds, as described next.
To obtain an index of channel activation (activation index), the above-baseline current record was integrated from the end of the delay to the end of the stimulus or, if activation showed an obvious peak during the sustained stimulus, to that peak. This resulting charge (current × time) was divided by the pressure × time, giving an activation index in units of picoamperes per millimeters of Hg. Values are given as means ± SE. Paired and unpaired t-tests or nonparametric tests were carried out as indicated, and differences were considered significant when P < 0.05.L-M(TK) Cells
Cell preparation.
L-M(TK) cells from the
American Type Culture Collection (ATCC; CCL-1.3;
passage
19) were grown in DMEM (GIBCO) with
4.5 g/l glucose and 10% fetal bovine serum (FBS). Calcium
phosphate-mediated transfection of cells with DNA coding for a green
fluorescent protein (GFP; pCSnucGFP DNA kindly provided by Adrian
Salic) (18, 53) was carried out as follows. Cells were plated 24 h
before transfection at 2 × 105
cells · 100-mm-diameter
plate
1 · 10 ml medium
1. pCSnucGFP DNA
(10 µg prepared via Qiagen midiprep kit) in 450 µl sterile water
was mixed with 50 µl of 2.5 M
CaCl2; 500 µl of 2×
HEPES-buffered saline (280 mM NaCl, 1.5 mM
Na2HPO4,
and 50 mM HEPES, pH 7.1) were added dropwise to the DNA-Ca solution. Immediately, the DNA precipitate was removed and added dropwise to the
cells. After 18 h, the medium was replaced with fresh medium. After 24 h, cells were trypsinized (0.5% trypsin in PBS + 5 mM EDTA) and
replated at lower density in 35-mm culture plates. Recordings were made
once coexpression of green fluorescent protein was evident by epifluorescence.
Patch recording. The bath and pipette saline was (in mM) 150 NaCl, 2 KCl, 2 CaCl2, 1 MgCl2, 5 HEPES, and 1 glucose (pH 7.3 and osmolarity 312 mosM). Pipettes for cell-attached patch recordings and pressure stimuli were as described above for Lymnaea. Recordings were made at room temperature. Cells expressing green fluorescent protein were located by epifluorescence illumination, but recordings were made under Hoffman modulation optics.
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RESULTS |
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Operational Definition of Mechanosensitivity
Because pipettes and gigaohm seals vary, nominally identical mechanical stimuli differ from patch to patch and over time in a given patch. Characterizing MS channel sensitivity (ease of activation) by a Boltzmann distribution (open probability vs. membrane tension) therefore has limited utility. Here "mechanosensitivity" is used qualitatively: if, in response to a standard step stimulus, channels activate more or less readily than under control conditions, mechanosensitivity increases or decreases, respectively. The two measures of mechanosensitvity used are the delay to the start of activation and extent of activation after the delay ("activation index"). Because delay increases from day 1 to day 4 (45), we used exclusively cells from days 4-6, thus minimizing variance in controls and maximizing latitude for detecting changes.Choosing a Stimulus Protocol
As previously noted (45), SAK channels were ubiquitous in Lymnaea neurons. With NS in the bath and high-K saline in the pipette and pipette voltage of +60 mV, TEA-insensitive inward single channel currents of 4-5 pA were obtained with suction. Naive patches challenged with sufficiently large first-hit stimuli (see sample traces from two cells; Fig. 1, A) exhibited activation of SAK channels, but, notably, activation began after a substantial delay. Delays are not seen in nongentle patches, which, instead, show immediate and sustained activation (e.g., Ref. 45). First-hit stimuli of ~30-40 mmHg occasionally yielded activation after delays of 5-10 s (e.g., Fig. 1A, bottom) but usually produced delays of tens of seconds, making data storage and handling onerous and raising concerns about what elicited activity. Consequently, larger first hits were used for the experiments below.
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The duration of delay depended on stimulus intensity (Fig.
1B) and patch history (Fig.
2). As shown in Fig.
1B, first hits at 60 mmHg
yielded delays more than twice as long (significant) as first hits at
80 mmHg, although activation index (Fig.
1B) was not
significantly different at these two pressures (unpaired t-tests). The delays were impressive;
channels were oblivious to
60 mmHg for ~10 s and to
80
mmHg for ~3 s. For subsequent experiments, we chose to use
80
mmHg on the grounds that the delay was neither impracticably long nor
too short to accurately detect any decreases and that activation was
not near saturation levels so that increases would be readily
detectable. Near saturation, four to eight channels were generally seen
per patch.
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Attempts to Recover From Exercising Patch
Repeated stimulation ("exercise") affected SAK channels. Figure 2A shows sample traces for an exercised patch. The delay and activation index for this example are plotted in Fig. 2B, along with the summary data for 16 patches. The first and second hits were separated by 10 s. After this interval, delay fell and the activation index increased (significant by paired t-tests; Fig. 2B). Although channels were always active during the last part of the first hit, they did not reactivate immediately when the second-hit step was applied. A measurable delay, albeit a reduced one, was obtained with the second hit. Thus during the 10-s interstimulus interval, some process occurred that partially restored the patch to its initial large-delay state.Given a considerably longer interstimulus interval than 10 s, would patches recover more fully from stimulation-induced changes? To test this, we followed the second hit with a 10-min rest and then restimulated (third hit). After the rest, neither mechanosensitivity parameter was significantly different than at the second hit. Thus, although the patch features responsible for the rapid (<10 s) partial recovery that occurred between the first and second hits had not deteriorated, the stimulus-induced change (damage?) that prevented full recovery was not, evidently, rectified during 10 min of rest. Attempts to provide a longer (30 min) rest period before the third hit were unsuccessful because gigaohm seals deteriorated.
Location of Patch
If gentle seals on Lymnaea neurons (see METHODS) formed tens of microns into the pipette, as can happen (49), then, even before a first hit, major cytoplasmic trauma may have been incurred. With gentle seals and first hits at
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To confirm that gentle patches were indeed nearer the pipette tip, we
used fluorescence. With Lucifer yellow (1 mM) in the pipette (not
shown), the patch position (i.e., the fluorescence front) did not
detectably change between first touching the cell and formation of a
gentle gigaohm seal or with a first hit to 80 mmHg. With
overstimulation (as in Fig. 3, A and
B), however, the front moved as
nonfluorescent cytoplasm displaced pipette solution. In another test,
Lucifer yellow (1 mg/ml) was dialyzed into the cytoplasm by
whole cell recording (Fig. 3
D-G).
Once fluorescent, neurons were repatched (gentle seal, cell attached) at a new location and, despite low signal, the patch location was
evident near the pipette tip and did not relocate during a
80
mmHg hit.
Thus, in experiments below involving gentle seals and 80 mmHg
first-hit stimuli, but viewed by Hoffman optics, patches were evidently
obscured because they formed stable seals close to the pipette tip.
Effects of Cytochalasin B and N-ethylmaleimide
In the following experiments, cells were exposed to agent before gigaohm seal formation, and agents were present until experiments ended. All the data were obtained from first hits that were applied within ~10 s of obtaining a gentle seal.Pretreatment with 25 µM cytochalasin B (1-2 h) significantly
decreased the first-hit delay and increased the activation index (Fig.
4, top).
Controls for the vehicle, 0.5% DMSO, showed no effect on either
mechanosensitivity parameter.
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Pretreatment (20-30 min) of cells with the sulfhydryl reagent N-ethylmaleimide (NEM; 1 mM) also had a major impact on mechanosensitivity; delay fell to 4% of control values, and activation index increased 3.8-fold. NEM impairs many proteins, but visual inspection of single-channel events for amplitudes and flickeriness (Fig. 4, bottom) revealed no apparent inhibitory effect of NEM pretreatment on SAK channel permeation characteristics. Clearly, NEM did not prevent the channels from activating in response to mechanical stimuli. Thus NEM, which chemically disrupts many cellular functions by abolishing ATPase and GTPase activities, unequivocally enhanced rather than diminished the mechanosensitivity of SAK channels.
Effects of Hyposmotic and Hyperosmotic Conditions
Under perforated patch, swelling Lymnaea neurons develop a TEA-insensitive K conductance that may be mediated by SAK channels (57). Whether these or the many other swelling-activated channels reported in the literature gate by membrane tension is a major unresolved issue, so it is particularly interesting to determine whether osmotic perturbations affect channel mechanosensitivity.To test the effect of swelling, neurons were exposed to 0.6×
normal osmolarity (some NaCl removed from NS) for 15 min before patching. Although this is physiologically extreme,
Lymnaea neurons not only withstand but
remain healthy (rearborize processes) after far more extreme and
prolonged exposures (36, 57). Gentle seals were obtained on swollen
neurons, and then 80 mmHg first hits were done. Delay was only
11% of the control value, and, after the shortened delay, a
significantly greater level of channel activity was obtained, as
indicated by the 4.7-fold increase in the activation index (Fig.
5).
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An additional experiment showed that for both the parameters the swelling-induced changes were persistent (Fig. 5). Neurons were exposed for 15 min to 0.6× NS, returned to NS for 24 h, and then tested. As expected (41), the swollen neurons reshrank and made transient vacuole-like dilations when switched back to NS. Although neurons appeared as healthy and as well arborized as neurons in unperturbed control dishes when patched 24 h later, their plasma membranes were evidently mechanically different, since delay and activation had not recovered.
Neurons were also tested after exposure to hyperosmotic solution (1.9× normal; sucrose added to NS) for 15 min. The cells were clearly shrunken before formation of gentle seals. Hyperosmia almost doubled the first-hit delay (Fig. 5) but had no effect on activation index. Thus, in shrunken cells, the channels were able to avoid the stimulus for an extended period, but, once felt, the stimulus was as effective as in controls.
Effects of Elevated Intracellular Ca
Because many physiological and pathological events elevate intracellular Ca, we tested whether elevated Ca affects SAK channel mechanosensitivity. Intracellular Ca in cultured Lymnaea neurons bathed in NS is ~0.1 µM (21). We chose to clamp Lymnaea neuron cytoplasmic Ca to a level between rest (~0.1 µM) and 1.0 µM [attained in squid axons during a train of action potentials (2)]. Ca at 0.5 µM is more than sufficient to activate the actin-depolymerizing enzyme gelsolin (39), so neurons were incubated in saline containing 0.5 µM free Ca (NS but with Ca 1.9 mM and EGTA 2 mM, yielding 0.5 µM free Ca) and 20 µM A-23187, a Ca ionophore (final concentration 0.3% in DMSO). When incubated for >5 min in this solution, Lymnaea neurons swelled and developed spherical blebs. We therefore patched the cells as rapidly as possible after adding the ionophore solution (by ~2 min), before blebbing began. Figure 5 shows the results from first hits to 14 cells. The delay of channel activation was decreased to 16% of controls, and the activation index increased sevenfold. Thus elevated Ca strongly increased SAK channel mechanosensitivity.MS Currents in Fibroblasts
Figure 6 illustrates a collection of MS currents (plus an example of unusual spontaneous activity) elicited from L-M(TK
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There was a dismaying variability in the mechanocurrents elicited, and
the variations showed no consistent correlation with transfection
status. Although a greater proportion of MS-current-positive patches
was observed among the transfected cells, we judged it nearly
impossible to ascertain whether there might be "signal" hidden in
the high background. As it was not our goal to study the endogenous MS
channels of L-M(TK)
cells, we characterized the currents no further. It is worth pointing
out that we turned to
L-M(TK
) cells because, in
our hands, HEK-293 cells were even more prone than
L-M(TK
) cells to exhibit
endogenous MS currents.
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DISCUSSION |
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Equivocal Mechanosensitivity of SAK Channels
The designation "MS channels" applies to Lymnaea SAK channels because in exercised patches, small suctions of, say,Results presented here demonstrate that even at the most fundamental
quantitative level (i.e., scoring 0 or 1 for presence or absence of MS
channels in the cells) there can be problems. With a small change in
our stimulus protocol, it would have been quite legitimate to decide
that Lymnaea neurons are "MS
channel null". Assume that a gentle seal was obtained and then a
large first hit of, say, 60 mmHg for 2 s was applied. No
channels would have been activated by this substantial mechanical
stimulus. The standard protocol that we arrived at (
80 mmHg for
as many seconds as was required to see activation) was chosen
empirically because, bluntly put, it was sufficiently brutal to
routinely activate the channels. Thus, although the molluscan SAK
channels are certainly mechanosusceptible, they must be regarded as
poor mechanotransducers unless, in vivo, unknown factors intervene and
boost their mechanosensitivity. Admittedly, such channels might serve
to signal that a dangerously high membrane tension had developed (3,
34), but even that signal might come too late to be of use.
Delay, Activation Index, and Membrane Mechanical Properties
Because pressure steps had rise times in the 100-ms range, delays of several seconds were a feature not of the stimulator but of the membrane patch. On a typical healthy patch of control membrane, therefore, a large mechanical stimulus (first hit) failed for several seconds to activate MS channels. The implication is that stimulus energy was dissipated for those several seconds before affecting channel gating. This suggests that significant mechanical forces were felt by channels only when the cortical skeleton had become distended, disorganized, or damaged.Overall, then, treatments that sped activation on a first hit also yielded a greater extent of activation; this is consistent with the idea that, as for MS channels from E. coli (51), tension applied to naked channels in the bilayer sufficed to elicit MS gating in mechanosusceptible channels.
SAK channel responses to multiple stimuli can be viewed as involving a combination of elasticity, viscoelasticity, and plasticity. The elastic component was the fundamental tension-induced channel activation (see Ref. 42). Viscoelasticity was seen in the fact that second-hit delay, although reduced, was not zero but instead remained on the order of seconds. Between hits (time frame 10 s), partial viscoelastic recovery may occur as a result of corrective adjustments of the cortical actin cytoskeleton (12, 39) and, more speculatively for neurons, of the skeleton (see Ref. 54 for mechanical effects on the spectrin skeleton in erythrocytes). Finally, plasticity (strain) is evident in the failure of delay to relengthen even partially toward its control (first-hit) level after a 10-min rest. Activation index, too, was also unchanged after 10 min, indicating that the fundamental mechanotransduction machinery did not deteriorate as the patch became more traumatized. This is consistent with the idea that the unadorned channel in the bilayer is all that is required for a mechanosusceptible SAK channel (see Ref. 13).
Cortical Cytoskeleton
Cytochalasins interfere with actin polymerization, resulting in a net loss of filamentous actin. They promote activation of many channels (26, 27, 37, 43), suggesting a broad regulatory role for filamentous actin on membrane proteins.Like cytochalasin D (45), cytochalasin B greatly enhanced SAK channel mechanosensitivity. In Lymnaea growth cones (whole cell recording), cytochalasin B does not facilitate activation of the channels (33). Even with the B form, therefore, there continues to be a discrepancy between whole cell and patch effects of cytochalasins on channel mechanosensitivity. The present finding confirms that "adequate" mechanical stimuli are more readily delivered to channels in patches than to channels in situ. Cytochalasin-resistant elements of the membrane skeleton may be more fragile in patches than in situ and therefore less able to provide any mechanoprotection. For bacterial MS channels, the channel protein alone (linked to no network) is the basic mechanosensitive unit (51); if naked SAK channels too are mechanosensitive, then, in situ, even damaged membrane skeleton may protect channels from mechanical loads.
Although NEM inhibits many proteins, there can be no suggestion that the NEM-induced increase in mechanosensitivity observed here related to any inhibition of channel function. There were no obvious effects of NEM on single-SAK channel properties and, supplied intracellularly (6), NEM (0.3 mM) does not block hair cell mechanotransducer channels. External NEM (1 mM, as used here) abolishes cytoplasmic and filopodial motility in Lymnaea neurons and renders them osmotically fragile (41, 57), effects likely to involve actomyosin. The profound enhancement of SAK channel mechanosensitivity by 1 mM NEM probably depended on disruption of disulfide-bearing enzymes, including myosin (23), in the cortical cytoskeleton.
The actions of cytochalasin and NEM suggest that the "tonus" of the cortical cytoskeleton normally protects ion channels from unacceptable mechanical loads. If dynamic actomyosin cross bridges contribute to the load-bearing strength of the membrane, then increased osmotic fragility (57) and increased SAK channel mechanosensitivity are both predictable outcomes of NEM or cytochalasin treatment.
Elevated Intracellular Ca
Clamping intracellular Ca at 0.5 µM (see Ref. 21 for an account of fura 2 measurements of intracellular Ca under conditions similar to those used here) produced a liquefied unhealthy-looking cytoplasm, and its effect on SAK channel mechanosensitivity was similar to cytochalasin: unequivocally increased mechanosensitivity. Ca activates the actin-depolymerizing enzyme gelsolin (40) plus various Ca-dependent proteases, including ones that cleave the spectrin skeleton (31). Clearly, SAK channel mechanosensitivity is not dependent on a well-organized cortical cytoskeleton. Interestingly, volume-sensitive Cl currents in astrocytes are potentiated by both intracellular Ca and cytochalasin (26). In cells in which Ca-permeant MS channels appear to be mechanotransducers, such as in baroreceptors (52), it should be interesting to determine whether mechanosensitivity is Ca enhanced. Assuming that persistent pressure changes are of primary interest, auto-upregulation of mechanoresponsiveness by the Ca-permeant MS channels might enhance low-pass mechanosensory filtering by baroreceptors.Osmotic Perturbations
Swelling and shrinking both alter cortical architecture, but only swelling, which disorganizes actin filaments (7, 19), increased SAK channel mechanosensitivity. In this case, the change was not secondary to Ca effects, since in Lymnaea neurons 50% dilution has a negligible effect on intracellular Ca (20). For swelling (as for cytochalasin, NEM, and high intracellular Ca) the simplest explanation for enhanced SAK channel mechanosensitivity is a nonspecific degradation of the submembrane shock absorber. In 50% medium, membrane tension rises approximately threefold above controls (9), and gradually (over ~20 min) a macroscopic K current develops (57). Whether tension is what eventually activates the SAK channels in swollen neurons is unresolved. In fact, what stimulates any "volume-activated" conductance (see Refs. 26, 27) is an open question. Nevertheless, it is intriguing to wonder whether the mechanosensitivity of volume-activated channel increases when hyposmia impairs the cortical shock absorber.Hyperosmolarity, which causes F-actin to reorganize at membranes in Lymnaea neurons (20), may increase delay by augmenting the robustness of the cortical cytoskeleton.
"Osmomodulation" of MS channel mechanosensitivity might be able to tune osmoreception by MS channels. Consider stretch-activated channels supporting a net osmolyte efflux: osmomodulation by swelling would augment the loss of osmolytes, favoring regulatory volume decrease. Or, for stretch-inactivated channels permeable to depolarizing ions (e.g., Ref. 4), osmomodulation during swelling (increased ease of stretch-inactivation) would facilitate stretch-induced repolarization, thereby bolstering the sensory response.
Overall, then, treatments that sped activation on a first hit also yielded a greater extent of activation; this is consistent with the idea that, as for MS channels from E. coli (51), tension applied to naked channels in the bilayer sufficed to elicit MS gating in mechanosusceptible channels.
Persistent Effects
The enhanced SAK channel mechanosensitivity produced by 15 min of hyposmia persisted in neurons patched 24 h later. Given the evident health of the rearborizing neurons at 24 h, one cannot necessarily infer long-term damage to the cortical cytoskeleton. The persistent effect may, instead, reflect the dynamism of the cytoskeleton as the neurons remodeled (see Ref. 30) after the perturbation.If a 15-min bout of osmotic swelling is a mechanical trauma yielding
persistent posttrauma effects, so too is isolation of neurons from the
central nervous system: SAK channels in neurons 24 h postisolation are
significantly more mechanosensitive than those 4-6 days
postisolation (45). In a cellular model of mechanical brain trauma
(59), nonlethal (31%) stretching of neurons for 50 ms yields, after
some minutes, sustained (6 h) protein kinase C-dependent
changes in channel properties. Clearly, caution is needed when
extrapolating about channels from acutely isolated (read, "recently
traumatized") neurons, and MS channels are no exception.
Hyperosmia, Neuroprotection, and Ischemia
Hyperosmia and reshrinkage of previously swollen cells (including Lymnaea neurons) both cause cortical F-actin to rapidly organize (7, 19, 21), and so in neurons exposed to hyperosmia the cortical cytoskeleton should be strengthened. Because disrupters of F-actin increase MS channel mechanosensitivity (Refs. 14, 45, and the present study), "enhancers" of F-actin like hyperosmia should decrease MS channel mechanosensitivity. In line with this, the F-actin stabilizing reagent phalloidin inhibited whole cell mechanosensitive currents from putative baroreceptor neurons in which the MS current is thought to arise from channels residing in the soma (8).The mild but significant reduction of SAK channel responsiveness with hyperosmia (delay almost doubled) may have clinical resonances, since hyperosmotic media are broadly neuroprotective in the ischemic brain (22). Glutamate channel activity (38) and voltage-gated Ca channel activity (25) can both be enhanced by membrane stretch. Because excessive activity of such channels spells excitotoxic damage in stroke, a trauma accompanied by neuronal swelling and by degradation of the spectrin skeleton (31), the neuroprotection afforded by hyperosmia might occur in part because it counteracts spurious MS channel activity that is only of significance after loss of membrane mechanoprotection. Although any clinically applied hyperosmia would be of more modest magnitude than that used here, duration might compensate.
L-M(TK) Cell Currents
Trauma and Mechanotransduction
A recurrent finding was that trauma facilitated mechanosensitivity in channels that are mechanosusceptible but not normally used for mechanotransduction. The native channel microenvironment evidently provides a mechanoprotective shock absorber. Even in auditory hair cells, cytoskeletal mechanoprotection seems to prevail, relaxed only on the single axis along which minute vibrational stimuli are detected; in that axis, moreover, actomyosin-based mechanoprotection of the channel provides for adaptation (11). Conceivably, there may be cases in which trauma-induced enhancement of MS channel mechanosensitivity is used physiologically. Hypermechanosensitivity of somatic sensory fields following trauma (e.g., in Aplysia; see Ref. 10) might, for instance, partly depend on trauma-induced lowering of the threshold for nociceptive MS membrane proteins. ![]() |
NOTE ADDED IN PROOF |
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Patel et al. (A. J. Patel, E. Honoré, F. Maingret, F. Lesage, M. Fink, F. Duprat, and M. Lazdunski. A mammalian two pore domain mechano-gated S-like K+ channel. EMBO J. 17: 4283-4290, 1998) recently showed that a cloned mammalian K channel, TREK-1, is mechanosensitive and a member of the wider family to which the stretch-activated K channel described here belongs. They also showed that TREK-1 can be activated by amphipathic crenators, strong evidence that mechanical energy is conveyed to the channel directly via the bilayer. This is fully consistent with our contention that membrane skeleton is mechanoprotective for the channel.
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ACKNOWLEDGEMENTS |
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This work was supported by Grant ST2716 from the Heart and Stroke Foundation of Ontario and by equipment and research grants from the Natural Sciences and Engineering Council, Canada.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: C. E. Morris, Loeb Health Research Institute, Ottawa Hospital, 725 Parkdale Ave., Ottawa, ON, Canada K1Y 4E9.
Received 13 May 1998; accepted in final form 4 November 1998.
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