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The Journal of Neuroscience, February 15, 2002, 22(4):1238-1247
Mechanosensitive Ion Channels in Cultured Sensory Neurons of
Neonatal Rats
Hawon
Cho,
Jieun
Shin,
Chan Young
Shin,
Soon-Youl
Lee, and
Uhtaek
Oh
The Sensory Research Center, Creative Research Initiatives, College
of Pharmacy, Seoul National University, Seoul 151-742, Korea
 |
ABSTRACT |
Mechanosensitive (MS) ion channels are present in a variety of
cells. However, very little is known about the ion channels that
account for mechanical sensitivity in sensory neurons. We identified
the two most frequently encountered but distinct types of MS channels
in 1390 of 2962 membrane patches tested in cultured dorsal root
ganglion neurons. The two MS channels exhibited different thresholds,
thus named as low-threshold (LT) and high-threshold (HT) MS channels,
and sensitivity to pressure. The two channels retained different
single-channel conductances and current-voltage relationships: LT and
HT channels elicited large- and small-channel conductance with
outwardly rectifying and linear I-V relationships, respectively. Both LT and HT MS channels were permeable to monovalent cations and Ca2+ and were blocked by gadolinium, a
blocker of MS channels. Colchicine and cytochalasin D markedly reduced
the activities of the two MS channels, indicating that cytoskeletal
elements support the mechanosensitivity. Both types of MS channels were
found primarily in small sensory neurons with diameters of <30 µm.
Furthermore, HT MS channels were sensitized by a well known inducer of
mechanical hyperalgesia, prostaglandin E2, via the
protein kinase A pathway. We identified two distinct types of MS
channels in sensory neurons that probably give rise to the observed MS
whole-cell currents and transduce mechanical stimuli to neural signals
involved in somatosensation, including pain.
Key words:
mechanosensitive channels; cationic; sensory neurons; somatosensation; pain; sensitization
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INTRODUCTION |
The mechanical stimulation of
sensory receptors in vertebrates may generate a variety of sensations,
such as touch, pressure, vibration, proprioception, and pain. Neural
signals of the somatic sensations begin with the excitation of
mechanoreceptors in sensory nerves. Numerous specialized or
encapsulated endings in the skin are mechanoreceptors that are
sensitive to light touch, pressure, or vibration (Winkelmann, 1986
;
Willis and Coggeshall, 1991). In addition, a subset of A
-mechanical
and C-polymodal nociceptors are sensory organs that respond to intense
mechanical stimuli, and therefore, subserve pain sensation. Moreover,
each mechanoreceptor responds uniquely to various temporal and spatial
mechanical stimuli. Combinations of these mechanoreceptors gives rise
to a repertoire of sensations caused by complex mechanical stimuli
(Gardner et al., 2000).
Mechanoreceptor excitation is accomplished by opening or closing ion
channels present in the sensory organs. Many different types of
mechanosensitive (MS) channels have been characterized in a wide
variety of cell types (Garcia-Anoveros and Corey, 1997; Hamill and
Martinac, 2001). These MS channels are involved in various cellular
functions, such as osmoregulation, touch, hearing, or control of
balance (Garcia-Anoveros and Corey, 1997; Hamill and Martinac,
2001). Recently a group of genes, related to mechanosensation was
cloned in genetic mutants of Caenorhabditis elegans or
Drosophila melanogaster. Among these, mec-4 and
mec-10 cloned from Caenorhabditis elegans were
implicated in mechanosensitivity, because mutations in these
genes caused the loss of touch sensitivity in nematodes (Driscoll and
Chalfie, 1991; Mitani et al., 1993; Huang and Chalfie, 1994).
NompC in Drosophila encoding a protein homologous
to transient receptor potential channels is also implicated in
mechanosensitivity because of reduction in bristle mechanoreceptor
currents in null mutants (Walker et al., 2000). In addition, vanilloid
receptor-related osmotically activated channel (VR-OAC), a gene that
shares homology with vanilloid receptor 1 and encodes
osmotically-activated ion channels, has been identified in Merkel cells
surrounding the vibrissae of the rat snout (Liedtke et al., 2000).
Although these genes appear to be related to mechanosensitivity,
evidence that products of these genes are gated mechanically is still
lacking (Tavernarakis and Driscoll, 1997).
Although many MS channels have been identified in a variety of cells,
MS channels in sensory neurons are not well characterized. Recently,
whole-cell currents activated by stretch or pressure in dorsal root
ganglion (DRG) neurons have been identified (McCarter et al., 1999;
Takahashi and Gotoh, 2000). The reversal potential of the
stretch-activated whole-cell current was near zero in
Na+/K+
bi-ionic solution and blocked by gadolinium, a nonspecific MS channel
blocker (McCarter et al., 1999). Although whole-cell currents evoked by
stretch or pressure have been characterized in sensory neurons, ion
channels responsible for the macroscopic mechanically gated currents
have not yet been characterized in sensory neurons. Furthermore,
despite the identification of molecular species for detecting certain
modes of pain sensation (Caterina and Julius, 1999), ion channels
responsible for the excitation of sensory neurons by intense mechanical
force are not well characterized. In this study, we identified the
single-channel currents of two distinct types of MS channels in
cultured rat DRG neurons and characterized the biophysical properties
of these channels.
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MATERIALS AND METHODS |
Cell culture. Experiments were performed according to
the Ethical Guidelines of the International Association for the Study of Pain. DRGs were dissected from all levels of the thoracic and lumbar spinal cords of neonatal rats and collected in cold culture medium (4°C). The culture medium, a mixture of DMEM and F-12 solution (Invitrogen, Grand Island, NY), contained 10% fetal bovine serum (Invitrogen), 1 mM sodium pyruvate,
50-100 ng/ml nerve growth factor (Invitrogen), and 100 U/ml
penicillin-streptomycin (Sigma). Ganglia were washed with a mixture of
DMEM and F-12 solution and incubated for 30 min in a warm (37°C)
DMEM-F-12 mixture containing 1 mg/ml of collagenase (Worthington
Biomedical, Freehold, NJ). The ganglia were then washed three times
with Mg2+- and
Ca2+-free HBSS (Invitrogen), and
incubated with gentle shaking for 30 min in warm HBSS (37°C)
containing 2.5 mg/ml of trypsin (Boehringer Mannheim,
Indianapolis, IN). The trypsin-containing solution was then centrifuged
at 1000 rpm for 10 min, and the pellet obtained was washed gently 2-3
times with the culture medium to inhibit the enzyme activity. The
pellet was suspended in the culture medium, gently triturated with a
fire-polished Pasteur pipette, and plated onto round glass coverslips
in small Petri dishes (35 × 12 mm). The glass coverslips were
previously treated with poly-L-lysine (0.5 mg/ml)
and dried. Cells were placed in 37°C incubator in a 95% air and 5%
CO2 atmosphere. Cells were used 2-4 d after plating.
Electrophysiology. As soon as a borosilicate glass pipette
(World Precision Instruments, Saratoga, FL) coated with Sylgard (Dow
Corning, Midland, MI) touched the surface of a cultured sensory neuron,
gentle suction was applied to the pipettes to obtain gigaseals. The tip
resistance of the pipette was ~3 M
for whole-cell and single-channel current recordings. To record whole-cell currents, gigaseals were formed first, and then the membrane in contact with the
pipette was ruptured by applying gentle suction. After a whole cell was
formed, the capacitive transients were canceled. For single-channel
current recording, cell-attached, outside-out, and inside-out patches
were formed. Channel currents were recorded with a patch-clamp
amplifier (Axopatch 200A; Axon Instruments, Foster City, CA) and
filtered at 5 kHz using a low-pass Bessel filter. The output of the
amplifier was digitized at a sampling rate of 94 kilosamples per second
with a digital data recorder (VR-10B; Instrutech, Great Neck, NY) and
stored on videotapes. For chart recording, the output of the amplifier
was filtered at 500 Hz with an eight-pole, low-pass Bessel filter
(Frequency Devices, Haverhill, MA) and then fed to a thermal array
chart recorder (TA240; Gould, Valley View, OH).
The output of the digital data recorder was imported to a personal
computer to analyze open and closed events of single-channel currents
using Fetchex and Fetchan in pClamp (version 6.0.4; Axon Instruments).
The threshold amplitude for taking an opening event was set at 50%
(half-amplitude algorithm in Fetchan). The minimum duration of open
events was set at 100 µsec for measuring channel open probability
(Po) or amplitude histograms.
Po of single-channels was obtained
from the equation (Spruce et al., 1985),
where tj represents the time
spent at each level, j corresponding to 1, 2, ... , N
channels open (N is the maximum number of channels seen in a
patch), and T is the duration of the recording. Channel
activity (NPo) was calculated as the
maximum number (N) of functional channels in the
patch times Po. The maximum number (N) of channels was determined after full activation
of the channels by applying up to 
110 mmHg. The amplitude histograms
were analyzed for patches containing only one or two channels.
Pressure application. Membrane stretch was achieved by the
application of negative (suction) or positive pressures to a patch electrode. Pressures were applied as follows: the end of a 30-cm-high U-tube manometer filled with mercury was connected to a patch pipette
with a polyethylene tube. When negative or positive pressure was
applied to the patch pipette, the pressure was delivered to the
manometer. Difference in the height of mercury columns in the manometer
represented the actual pressure expressed in millimeters of
mercury applied to the pipette. The polyethylene tube was also connected to a pressure transducer (P23XL-1; Gould) to record the
pressure in a chart recorder.
Solution and chemicals. For whole-cell current recording,
the pipette solution contained (in mM): 140 KCl,
2 MgCl2, 5 EGTA, 10 HEPES, 2 ATP, and 0.3 GTP, pH
7.2. The control bath solution for whole-cell recording contained (in
mM): 140 NaCl, 5 KCl, 2 MgCl2, 2 CaCl2, and 10 HEPES, pH 7.2. For single-channel recording, the solution in the bath
and the pipettes contained (in mM): 140 NaCl, 5 KCl, 2 MgCl2, 5 EGTA, and 10 HEPES, pH 7.2, unless described otherwise. Colchicine (Sigma), gadolinium
(Gd3+; Sigma), and dibutyryl cAMP
(Biomol, Plymouth Meeting, PA) were dissolved and stored in distilled
water. Cytochalasin D (Sigma) and prostaglandin
E2 (Sigma) were dissolved and stored in 100% ethanol. Amiloride (Sigma), H-89 (Biomol), and arachidonic acid (Sigma)
were dissolved and stored in 100% DMSO. All other basic agents for
cell culture and the physiological solutions were purchased from Sigma.
Statistics. The paired Student's t test was used
for comparing two means. To compare multiple means, one-way
ANOVA was used followed by a Tukey's post hoc test.
To determine whether cell-size distributions of neurons containing two
different MS channels were different, contingency tables were
constructed, and a Fisher's exact test was used. A p
value < 0.05 was considered significant.
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RESULTS |
Suction protocol to form a gigaseal
Because the magnitude of suction pressure applied to membrane
patches during gigaseal forming affects the mechanosensitivity of
certain MS channels (Hamill and McBride, 1997), we measured the suction
pressure when gigaseals were formed. The suction pressure required was
3.8 ± 0.15 mmHg (n = 10), which was considerably less than the suction pressure that changes the properties of mechanically gated channels present in Xenopus oocytes
(Hamill and McBride, 1997). In addition, there was no indication of
volume change of the cells or blebbing during or after pressure
application, which is also thought to affect the mechanosensitivity of
the MS channels (Hamill and McBride, 1997). As size of the patch
pipettes is also a factor, which affects MS channels (Hamill and
McBride, 1992), the inner diameters of the glass pipettes were measured using a scanning electron microscope. The average inner diameter of the
patch pipettes used for recording whole-cell and single-channel currents was 1.4 ± 0.09 µm (n = 6).
Activation of MS whole-cell currents
Whole-cell currents activated by pressure were recorded from
cultured DRG neurons. As shown in Figure
1, inward currents were observed after
positive pressures of 15 and 20 mmHg were applied to the whole-cell
pipette. The currents were activated by the pressure with an apparent
1-3 sec delay between currents and the plateaus of pressures as
observed by others (Takahashi and Gotoh, 2000) and returned to the
basal level when the pressures were released. Whole cells were too
fragile to be maintained during pressure application when positive
pressures exceeded 20 mmHg or were prolonged for more than ~5 sec.
The whole-cell currents were largely pressure-dependent: a greater
current was observed when a higher pressure was delivered to patch
pipettes (Fig. 1A,B). Pressure threshold, the lowest
pressure tested to activate currents, varied among neurons, and was as
low as 5 mmHg. These whole-cell currents were activated by 5-20 mmHg
pressure in 81 (28.9%) of 280 DRG neurons. The current-voltage
relationships of MS whole-cell currents were obtained after applying
voltage ramps of 300 msec duration ranging from 100 to +100 mV before
and during pressure application. Each current trace was averaged after
two trials of the voltage ramps. MS whole-cell currents were defined
after subtracting current response to the voltage ramp obtained before pressure application from that obtained during pressure application. As
shown in Figure 1C, the subtraction current reversed at
13.4 ± 1.48 mV (n = 18), suggesting poor cation
selectivity.
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Figure 1.
Activation of MS whole-cell currents by
pressure. A, Top trace, Whole-cell
currents of a cultured sensory neuron activated by pressure.
Bottom trace, Positive pressures applied to the patch
pipette. B, A summary of MS whole-cell currents in
sensory neurons. Because of the fragile nature of whole cells, positive
pressures of <20 mmHg were applied to whole cells. Membrane potential
was held at 60 mV. **p < 0.01 compared with the
mean of whole-cell currents activated by 5 mmHg. Numbers in
parentheses represent the number of experiments.
C, Current-voltage relationship of the whole-cell
current activated by pressure. Voltage ramps (bottom
trace) ranging from 100 to +100 mV with 300 msec duration
were applied before and during pressure application. Current elicited
before pressure (solid line), during pressure
application (dotted line), and difference current
(dashed line).
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Activation of mechanosensitive single-channel currents
To identify single-channel currents activated by pressure,
cell-attached or inside-out membrane patches were formed, and either negative or positive pressure was delivered to the shank of the patch
pipette to activate MS channels. In membrane patches of cultured
sensory neurons, application of capsaicin caused a great activation of
single-channel currents, because of the ubiquitous presence of
capsaicin-activated channels in cultured sensory neurons (Oh et al.,
1996; Jung et al., 1999; Hwang et al., 2000). Single-channel currents
activated by capsaicin, however, were not sensitive to pressure applied
to the pipettes. To prevent the casual openings of capsaicin-activated
channels, 10 µM capsazepine, an antagonist of the
capsaicin receptor, was added to the bath solution throughout the experiments.
We tested a total of 2962 cell-attached, outside-out, or inside-out
membrane patches of DRG neurons to determine the presence of MS
channels. In cell-attached patches with the holding potential of 60
mV, no single-channel currents were observed before applying suction,
except for a few patches, which showed spontaneous openings of
single-channel currents, possibly caused by initial pressures <5 mmHg
applied to the pipettes during gigaseal formation. Two types of
channels were observed most frequently when negative pressures of
various magnitudes were applied to the pipettes (Fig. 2, Table
1).
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Figure 2.
Single-channel currents activated by negative
pressures applied to a cell-attached patch. A, This
channel was classified as an LT MS channel because the channel was
activated by a relatively low pressure.
Ehold = 60 mV. Right,
An amplitude histogram of the single-channel currents. The
mean amplitudes of LT MS channels were best fitted by a Gaussian
distribution. B, Another type of single-channel currents
activated only by negative pressures greater than 80 mmHg. The
channel was classified as an HT MS channel because of its high pressure
threshold. The cell-attached patch contained two HT channels.
Right, An amplitude histogram of the single-channel
currents. C, Pressure-response relationships of the LT
(filled circle; n = 9) and HT
(filled triangle; n = 9) MS
channels. Relative channel activity
(NPo/NPmax)
at each pressure was fitted to the Boltzmann distribution described in
Results. Bars represent SEM.
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One type of MS channel, called a low-threshold (LT) MS channel, was
activated by relatively low pressures, 10 to 20 mmHg (Fig.
2A). Channel activity increased as higher pressures
were applied to the patch and reached a maximum at 80 mmHg.
Interestingly, channel openings continued for a period of time even
after the suction had been released. The residual LT channel activity,
however, slowly declined after the negative pressure was removed (Fig. 2A). The LT channel was also activated by positive
pressure up to +80 mmHg (data not shown). However, positive pressures
broke membrane patches easily. Therefore, most of the single-channel currents were recorded with negative pressures applied to the patch
pipettes. The LT MS channel was observed most frequently (25.7%)
(Table 1) and displayed the largest current amplitude of the MS
channels in the cultured sensory neurons. When the amplitude of the
channel currents was measured in cell-attached patches, the mean
current amplitude of LT was found to be 3.50 ± 0.06 pA
(n = 5) at 60 mV of holding potential.
Another type of MS channel was activated only by greater negative
pressures than 60 mmHg (Fig. 2B), thus tentatively
called a high-threshold (HT) MS channel. Unlike the LT channel, the HT MS channel showed small current amplitude (Fig. 2B).
In cell-attached patches, the average current amplitude was 0.9 ± 0.04 pA (n = 5) at 60 mV of holding potential.
This channel was also frequently observed in 23.6% of the total number
of patches tested (Table 1). In contrast to the LT MS channel, the HT
channel rarely showed residual activity when suction was removed.
Furthermore, the HT MS channels were not activated by positive
pressures up to +80 mmHg.
Pressure-response relationship
The two types of channels exhibited a pressure-response
relationship, but with different sensitivities (Fig. 2C). To
obtain the pressure-response relationships of MS channels, negative
pressures ranging from 0 to 110 mmHg were plotted against the channel
activity of the MS channels. Membrane patches could not sustain greater negative pressures than 110 mmHg. Relative channel activity
(NPo/NPmax) at each pressure was fitted by a nonlinear regression to a Boltzmann distribution. The channel activity at each pressure was normalized to
the maximum channel activity:
NPo/NPmax = 1/{1 + exp ((p1/2 p)/
)}, where N is the number of functional
MS channels in the patch, Pmax is the
maximum probability of channel opening, p is the suction
pressure, p1/2 is the suction pressure
at which the open probability is 0.5, and , sensitivity to the
suction pressure, represents the pressure change required to cause an
e-fold increase in the relative channel activity (Kim et
al., 1995; Marchenko and Sage, 1997). As shown in Figure 2C,
the half-maximal pressures, p1/2, of
LT and HT MS channels were 60.6 ± 1.2 (n = 9) and
83.1 ± 2.4 mmHg (n = 9), respectively. The
sensitivities () of LT and HT channels to negative pressures were
8.4 ± 1.8 and 4.3 ± 0.9 mmHg, respectively. An interesting
feature of these MS channels was that the maximal channel activity of
the HT channel was much greater than that of the LT channels. For
example, the near maximal, saturating channel activities of LT and HT
MS channels at 100 mmHg were 0.21 ± 0.05 (n = 9) and 0.94 ± 0.09 (n = 9), respectively (see
Fig. 6).
Current-voltage relationship
To determine the I-V relationships of the two MS
channels, membrane potentials were changed from 100 to +100 mV in 20 mV increments after forming inside-out patches. Constant negative pressures were applied to the membrane patches to activate the MS
channels (60 mmHg for LT channels and 80 mmHg for HT channels). The
current amplitude of each channel was measured at each membrane potential in symmetric 140 mM
Na+ solution. The mean amplitudes were
plotted as a function of the membrane potential. As shown in Figure
3, A and B, the
mean amplitudes of LT channels in the symmetrical solution were
3.1 ± 0.04 and +6.5 ± 0.35 pA at
Ehold = 60 and +60 mV, respectively
(n = 13). Thus, the slope conductances of LT MS
channels were 51.0 ± 0.6 and 108.3 ± 5.9 pS at
Ehold = 60 and +60 mV, respectively
(n = 13), indicating that the single-channel currents
were outwardly rectifying. In contrast, the HT MS channel exhibited a
much smaller slope conductance with a nearly linear current-voltage
relationship in the symmetrical solution. The slope conductances of HT
MS channels were 13.7 ± 0.3 and 16.3 ± 0.5 pS at 60
and +60 mV, respectively (n = 11) (Fig.
3A,B).
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Figure 3.
Current-voltage relationships and kinetics of LT
and HT MS channels. A, Traces in the expanded time scale
of single-channel currents of the two MS channels held at different
membrane potentials. Suction pressures of 60 and 80 mmHg were
delivered to pipettes of inside-out patches to activate the LT and HT
MS channels, respectively. B, Current-voltage
relationships of single-channel currents. LT, Filled
circle (n = 13); HT, filled
square (n = 11). To obtain the
current-voltage relationship of LT channels, each data point was
fitted to a single exponential equation. Bars represent SEM. The error
bars are so small that they are included by the circles
or squares. C, Open and closed time
histograms of LT and HT MS channels. LT and HT channels were activated
by 80 and 100 mmHg, respectively. The histograms of open and closed
time durations of the two channels were best fitted by two exponential
functions.
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Channel kinetics
Histograms of open or closed-time durations of both MS channels
activated by suction pressures were best fitted with two exponential functions (Fig. 3C). Therefore, both channels exhibited
short and long openings and closings. Negative pressure of 20 mmHg activated LT MS channels with open time constants of 0.12 ± 0.01 (
O1, n = 5) and 0.82 ± 0.09 msec (O2) and with closed time constants of 0.52 ± 0.06 (C1) and 14.76 ± 1.01 msec (C2). When negative pressure of 80
mmHg was applied to the same channels, the duration of short openings
increased significantly (O1 = 0.16 ± 0.01 msec; p < 0.05; n = 5), but not
that of long openings (O2 = 0.93 ± 0.06 msec). In contrast, at 80 mmHg, the duration of long closings, but
not short closings (C1 = 0.38 ± 0.06 msec; n = 5), decreased significantly
(C2 = 5.82 ± 0.63 msec;
p < 0.001). Similarly, negative pressure of 60 mmHg
activated HT MS channels with O1 and
O2 of 0.11 ± 0.01 and 0.97 ± 0.19 msec (n = 5), respectively, and with
C1 and C2 of
0.34 ± 0.02 and 5.45 ± 0.41 msec, respectively. When the
negative pressure was changed to 100 mmHg,
O2, but not O1,
increased significantly to 1.37 ± 0.25 msec
(p < 0.001; n = 5). In
contrast, both C1 and
C2 decreased significantly (0.13 ± 0.01 and 0.82 ± 0.16 msec, respectively; p < 0.001;
n = 5) at 100 mmHg. Therefore, greater pressures
increased the channel activities of both MS channels largely by
decreasing the duration of long closings.
Ion selectivity
To determine whether the two MS channels were permeable to cations
or anions, 140 mM of Na+ in
bath solution was replaced by
N-methyl-D-glucamine in inside-out patches under constant pressures (60 and 80 mmHg for LT and HT
channels, respectively). When 140 mM
N-methyl-D-glucamine in the bath was
substituted for 140 mM
Na+, inward currents with similar
amplitudes to those observed for the two MS channels in the control
bath solution were observed (n = 3 for each LT and HT
MS channel) when the membrane potential was held at 40 mV or lower.
However, no detectable outward currents were observed for either type
of MS channels when the membrane potential was held at +40 mV or
greater (Fig. 4A).
These results indicate that the two MS channels are permeable to
cations but not to anions.
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Figure 4.
Ion selectivity of LT and HT channels.
A, The N-methyl-D-glucamine
substitution. No outward channel currents were observed for the two
types of MS channels when the 140 mM Na+
bath solution was replaced by a solution containing 140 mM
N-methyl-D-glucamine (n = 3 for each MS channel). B, C, Cation
selectivity of LT (B) and HT
(C) MS channels. Current-voltage relationships
of each type of MS channels were obtained after the symmetrical 140 mM Na+ bath solution of inside-out
patches was replaced by solutions containing equimolar
K+, Cs+, or
Li+. The I-V relationships of the MS
channels in the symmetrical 140 mM Na+
salt condition are identical to those shown in Figure 3.
D, E, Permeability of LT
(D) and HT (E) MS channels
to Ca2+. Current-voltage relationships were
obtained from inside-out patches that contained 100 mM
Ca2+ in the pipette solution and 140 mM
Na+ in the bath solution.
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We further determined the ion selectivity of the channels among
cations, by testing changes in reversal potential in inside-out patches
under bi-ionic salt conditions. Current amplitudes, at various holding
potentials, were measured before and after the bath
Na+ solution was replaced with equimolar
Li+, K+, or
Cs+. As shown in Figure 4, B
and C, the replacement of Na+
with other monovalent cations did not shift the reversal potentials of
the MS channels from 0 mV, indicating that the two types of MS channels
could discriminate these monovalent cations poorly. Selectivity over
Ca2+ was also assessed by measuring the
reversal potential when 140 mM
Na+ in the pipette was replaced by 100 mM Ca2+ (Fig.
4D,E). In the bi-ionic system (100 mM
[Ca2+]0/140
mM
[Na+]i), the
reversal potentials of the LT and HT MS channels were 0.78 ± 0.8 (n = 5) and 2.38 ± 0.02 mV (n = 3), respectively, indicating that the MS channels are also permeable to
Ca2+. The permeability ratios
(Pcation/PNa)
for both LT and HT channels for Na+ and
monovalent cations or Ca2+ were calculated
from the modified constant-field equation (Fatt and Ginsborg, 1958) and
are shown in Table 1.
Block by gadolinium
The gadolinium ion (Gd3+) has been
reported to block various types of MS channels (Yang and Sachs, 1989;
Hamill and McBride, 1996). Thus, we investigated whether
Gd3+, the nonselective blocker of many MS
channels, could block the LT and HT MS channels present in sensory
neurons. To apply Gd3+ to the
extracellular surface of the channels, outside-out patches were formed.
Positive pressures ranging from +60 to +100 mmHg were delivered to the
pipettes of outside-out patches before and after 10 µM
Gd3+ was applied. Because EGTA,
which chelates Ca2+ also reduces free
Gd3+ (Boland et al., 1991; Hamill and
McBride, 1996; Caldwell et al., 1998), EGTA was removed from both the
control and the Gd3+-containing solution.
Bath application of the Gd3+ greatly
blocked activity of the LT channel (Fig.
5A,C) at +60 and +80 mmHg by
93.5 ± 0.6 and 92.4 ± 0.4%, respectively. Activity of the
HT MS channel was also blocked by Gd3+, as
shown in Figure 5, B and D: 10 µM Gd3+ reduced
activity of the HT channel activated by +80 and +100 mmHg by 97.9 ± 0.5 and 92.0 ± 8.6%, respectively.
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Figure 5.
Block of MS channels by Gd. A,
B, Example traces depicting the effects of 10 µM Gd on activities of LT (A) and
HT (B) MS channels in outside-out patches. The
membrane potential was held at 60 mV throughout the experiments.
C, D, Summaries of the effects of Gd on
activities of LT (C) and HT
(D) MS channels. Numbers on the
bars represent the number of experiments.
**p < 0.01
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Effect of the disruption of cytoskeletal elements
Many types of MS channels are often sensitive to conformational
changes in cytoskeletons (Small and Morris, 1994; Hamill and McBride,
1996; Maingret et al., 1999). Thus, the integrity of cytoskeletal
elements near the cell membrane appears to be important for the
activation of MS channels. Therefore, we examined whether the MS
channels of DRG neurons were affected by cytoskeletal disruption in the
cell. Colchicine and cytochalasin D are known to disrupt the assembly
of cytoskeletons and are considered to be cytoskeleton-disrupting agents (Cooper, 1987; Sadoshima et al., 1992). To determine the effects
of colchicine or cytochalasin D on the MS channels, channel activities
activated by various pressures were obtained and compared before and
after treatment with cytoskeleton-disrupting agents. Repeated
applications with 20 min intervals of negative-pressures up to 110
mmHg did not change the overall pressure-activity relationships of the
LT and HT MS channels in cell-attached patches (Fig.
6C).
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Figure 6.
Inhibition of activities of the LT and HT
channels by cytoskeleton disrupters, cytochalasin D, and colchicine.
A, B, Example traces of the effect of
cytochalasin D on activities of LT (A) and HT
(B) MS channels. Negative pressures were applied
to patch pipettes to activate the MS channels after forming
cell-attached patches. The cell-attached patches were incubated with 10 µM cytochalasin D for ~20 min. The negative pressures
were repeated after the cytochalasin D application. C,
Summaries of the effects of repeated applications of suction pressures
on the pressure-activity relationships of LT (n = 4-6) and HT (n = 4-7) MS channels. Note that
repeated applications of suction pressures do not change the overall
responses of the MS channels to pressures. D, Summaries
of the effects of 10 µM cytochalasin D on the pressure
responses of LT (n = 4-6) and HT
(n = 3-7) MS channels. E, Summaries
of the effects of 500 µM colchicine on the pressure
responses of LT (n = 4-5) and HT
(n = 3-6) MS channels.
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|
In contrast, the application of 10 µM of cytochalasin D
to the bath of cell-attached patches for ~20 min significantly
suppressed the channel activities of LT and HT channels over the range
of pressures tested (Fig. 6D). Cytochalasin D (10 µM) significantly attenuated the maximal
activity of LT and HT channels by 84.6 ± 2.6 (n = 6; p < 0.001) and 59.8 ± 3.2%
(n = 7; p < 0.001). A similar response
pattern was observed after treatment with another cytoskeleton
disrupter, colchicine. Colchicine (500 µM)
treatment reduced the maximal activities of the LT and HT channels
significantly by 88.4 ± 3.2 (n = 5;
p < 0.001) and 50.9 ± 6.9% (n = 6; p < 0.001), respectively (Fig.
6E). Treatments with colchicine and cytochalasin D
shifted the pressure-activity curve of HT channels rightward, increasing p1/2 of HT channels significantly from
80.8 ± 2.8 to 91.4 ± 3.3 mmHg (n = 6;
p < 0.05) and from 79.8 ± 0.76 to 87.7 ± 2.03 mmHg (n = 7; p < 0.01),
respectively. The cytoskeletal disrupters, however, did not change the
p1/2 of LT channels.
Effect of excision
Because excision of the cell membrane often changes
activity of certain MS channels (Marchenko and Sage, 1997; Maingret et al., 1999), we also examined the effect of excision on activity of the
MS channels. Excision of the cell membrane from cell-attached patches
to make inside-out patches greatly reduced the maximal current
responses of the LT and HT channels. After forming inside-out patches,
activity of LT MS channels at 100 mmHg reduced to 0.028 ± 0.005 by 87.5 ± 4.7% (n = 5) from that measured when
cell-attached patches were formed (0.21 ± 0.005;
n = 5). In HT MS channels, excision of the patch
membrane reduced the channel activity activated by 110 mmHg from
1.02 ± 0.09 to 0.42 ± 0.07 (57.8 ± 7.5% reduction; n = 4).
Effect of amiloride and arachidonic acid
A diuretic, amiloride, and its analogs are known to block many MS
channels, including MS currents in sensory neurons and epithelial sodium channels (Lane et al., 1991; Awayda et al., 1995; Hamill and
McBride, 1996; McCarter et al., 1999). Thus, it appeared likely that
amiloride would block the activity of the MS channels present in
sensory neurons. However, the application of amiloride up to 200 µM in the bath of outside-out patches (n = 5) or inside-out patches (n = 6) did not affect
activities of LT and HT MS channels. Arachidonic acid and other
unsaturated fatty acids are known to activate certain types of
mechanically-activated K+ channels (Kim et
al., 1995; Maingret et al., 1999). To test whether arachidonic acid
affects the two types of MS channels, arachidonic acid was applied to
the bath of inside-out patches containing each type of the MS channels.
When applied to the bath of inside-out patches, 10 µM of arachidonic acid failed to change the
activities of LT or HT MS channels (n = 4 for each type
of the MS channels).
Effect of prostaglandin E2
A unique action of prostaglandin E2
(PGE2) on the pain sensory system is accounted
for by its ability to cause hyperalgesia or sensitize nociceptive
sensory neurons to mechanical stimuli, possibly via the protein kinase
A pathway (Martin et al., 1987; Taiwo et al., 1989; Taiwo and Levine,
1991; Ouseph et al., 1995; Cunha et al., 1999; Chen et al., 1999).
Thus, we examined whether PGE2 sensitized the MS
channels present in sensory neurons. To determine the effects of
PGE2 on the activities of the LT and HT channels,
10 µM PGE2 was administered to the
bath of cell-attached patches containing the MS channels for 5 min
before a second series of negative-pressures were applied. As shown
above (Fig. 6C), under control conditions repeated
applications of suction pressures in cell-attached patches did not
change the overall pattern of pressure-activity relationships of the MS
channels. However, pretreatment of 10 µM
PGE2 for 5 min shifted the pressure-activity
curve to the left, decreasing significantly the mechanical threshold
(60 vs 40 mmHg) and the p1/2 (81.6 ± 5.0 vs
56.9 ± 2.3 mmHg; p < 0.001; n = 14) of the HT MS channel (Fig.
7A). The
PGE2-induced sensitization of the HT MS channels
was presumably mediated by the cAMP-dependent protein kinase pathway,
because the pretreatment of sensory neurons with H-89 (10 µM), a membrane-permeable inhibitor of protein
kinase A, completely blocked the sensitization of the channel by
PGE2 (Fig. 7B,C). Furthermore,
application of 100 µM dibutyryl cAMP, a
membrane-permeable analog of cAMP, also shifted the pressure-activity relationship to the left (Fig. 7D), reducing
p1/2 from 83.6 ± 5.3 to 59.1 ± 2.9 mmHg (p < 0.001; n = 10). In
contrast to the HT channels, however, PGE2 failed
to change the overall pattern of the pressure-activity relationship of
LT channels (p1/2 = 62.6 ± 2.0 vs
59.0 ± 2.1 mmHg; n = 6).
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Figure 7.
Sensitization of HT MS channel by PGE2
via the protein kinase A pathway. A, Negative pressures
were repeated in cell-attached patches at a holding potential of 60
mV. Before and during the second application of suction pressures, the
cell-attached patches of sensory neurons were incubated with 10 µM PGE2 for ~5 min. Right,
The pressure-activity relationships of HT MS channels before
(circle; n = 6-14) and after
PGE2 (triangle; n = 6-14) treatment. Channel activity
(NPo) at each pressure was fitted to
the Boltzmann distribution described in the text. Bars represent SEM.
B, Block of the PGE2-induced sensitization
of HT channels by H-89, an inhibitor of protein kinase A. H-89 (10 µM) was incubated before and during PGE2
application. C, A summary of the effect of coapplication
of H-89 and PGE2 on the pressure-response relationship of
HT MS channels. Control (circle; n = 5-10), after H-89 treatment (triangle;
n = 5-10). D, A summary of the
effect of the application of dibutyryl cAMP (dbcAMP), a
soluble cAMP analog, on the pressure-response relationship of HT MS
channels. Control (circle; n = 5-10),
after dbcAMP treatment (triangle; n = 5-10).
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Cell-size distribution
The size of sensory neurons is one way of determining their
sensory modalities, such as touch or pain (Winkelmann, 1986; Willis and
Coggeshall, 1991). Therefore, we measured the diameter of the soma of
cultured DRG neurons in which MS channels were detected. Figure
8 shows distribution of cultured sensory
neurons having each type of MS channel. Proportions of 1461 neurons
that elicited LT (n = 761) and HT (n = 700) single-channel currents were constructed according to neuron size.
As shown in Figure 8, the LT and HT MS channels were found mainly in
small to medium-sized DRG neurons, whereas, LT and HT MS channels were
rarely found in relatively large (>30 µm in diameter) sensory
neurons. We tested the presence of the MS single-channel currents in
190 additional cell-attached patches from 190 neurons having diameters
>30 µm up to 40 µm, none of which elicited the MS single-channel
currents. The distribution of cells having HT MS channel currents was
skewed (88%) toward smaller cells with diameters of 10-17.5 µm and
significantly (p < 0.0001) different from that
of cells having LT channels. LT MS channels appeared to be distributed
throughout cells with diameters of 10-30 µm: only 48% of cells with
diameters of 10-17.5 µm elicited the LT MS channel currents.
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Figure 8.
Cell-size proportions of cultured sensory neurons
eliciting each type of MS channel or whole-cell currents in cultured
DRG neurons.
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Figure 8 also depicts the cell size distribution of sensory neurons
eliciting whole-cell currents. Consistent with another report
(Takahashi and Gotoh, 2000), cells eliciting MS whole-cell currents had
a diameter >20 µm. The distribution of cells eliciting MS whole-cell
currents overlapped substantially with that of LT MS channels. However,
analysis with a Fisher's exact test revealed that distribution of
cells eliciting the whole-cell currents was different from those of
cells having LT (p < 0.0002) and HT
(p < 0.0002) channel currents.
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DISCUSSION |
DRG neurons are primary afferent neurons that carry sensory
signals from the skin, muscles, joints, and visceral organs to the
spinal cord. The nerve endings of DRG neurons possess a variety of
sensory receptors activated by mechanical, thermal, chemical, and
noxious stimuli. Various mechanical, thermal, or noxious stimuli are
known to generate action potentials at peripheral nerve endings of the
DRG neurons (Winkelmann, 1986; Willis and Coggeshall, 1991). Ion
channels thought to be responsible for somatosensory transduction have
been identified in sensory neurons, and some of their molecular species
have now been cloned (Caterina et al., 1997; Caterina and Julius,
1999). Although ion channels that account for the neural signals of
various sensory modalities are now known, little is known of their
channels for mechanosensation. In the present study, we identified two
novel cationic channels in cultured sensory neurons that were activated
by mechanical stimulus. The two MS channels appeared distinct from
other MS channels found in various cell types or newly cloned channels
because of their difference in biophysical properties such as their
conductance, ion selectivity, or current-voltage relationship as well
as sensitivity to pressure (Hamill and McBride, 1996; Hamill and
Martinac, 2001). Because DRG neurons are neural substrates for
mediating somatic and visceral sensation as well as proprioception, the
two types of MS channels in sensory neurons implicate in
mechanosensation, proprioception or possibly pain caused by noxious
mechanical stimuli.
Pressures applied to membrane patches or whole cells would develop
different membrane tensions that would be more relevant stimuli than
pressures. The membrane tension (T) caused by
pressure can be calculated from Laplace's Law:
where P is the pressure applied to a patch, and
r is the radius of the curvature of a membrane patch
(Guharay and Sachs, 1984). If the geometry of the patch is assumed to
be hemispheric with a diameter equal to that of the pipette (1.4 µm), pressures of 10-110 mmHg applied to the membrane patch would
produce tensions of 0.47-5.1 mN/m. However, the geometry of a patch in
a glass pipette is known to be much flatter than a hemisphere, having a
greater radius than that of a hemisphere (Sokabe et al., 1991; Opsahl
and Webb, 1994; Akinlaja and Sachs, 1998). Therefore, it is highly
likely that the actual tensions of the patch caused by the pressures
would be greater than the assumed tensions. Furthermore, the cell
membrane consists of lipid bilayer and cytoskeletons underlying the
lipid bilayer (Hamill and McBride, 1997; Hamill and Martinac, 2001).
Pressures applied to the patch membrane would act on the lipid bilayer
and the cytoskeletons (Hamill and McBride, 1997). Therefore, the
assumption that pressures cause the estimated tensions along the lipid
bilayer might be oversimplified.
Recently, nonselective cation currents activated by stretch or pressure
applied to whole cells was reported in cultured DRG neurons (McCarter
et al., 1999; Takahashi and Gotoh, 2000). The MS whole-cell currents
were generally consistent with the inward whole-cell currents observed
in the present study. Because membrane geometry of whole cells is
different from those of membrane patches, it is difficult to compare
pressure sensitivities of the two types of MS channels with those of
the whole-cell currents. However, some aspects of the MS whole-cell
current possess similarities with the LT and HT MS channels. First, the
two MS channels were the most frequently encountered MS channels in
sensory neurons. Second, the whole-cell and the MS single-channel
currents were cationic. Therefore, the two MS channels would represent
in part the macroscopic currents activated by pressures. However, we
cannot exclude the possible presence of other MS channels in sensory neurons that contribute to the MS whole-cell currents, partly because
distributions of neurons eliciting the two MS single-channel currents
did not overlap with that of neurons having the MS whole-cell currents
(Fig. 8).
Several lines of evidence in the present study suggest that HT MS
channels are related to mechanosensation in a noxious range. First, HT
channels were observed mainly in small DRG neurons and rarely found in
large cells (>30 µm in diameter). Direct measurements of cell body
size and the conduction velocity of primary afferent fibers demonstrate
that most of the large sensory neurons are low-threshold
mechanoreceptors and detect innocuous stimuli, whereas most smaller
neurons are nociceptors, which detect noxious mechanical, thermal, or
chemical stimuli (Harper and Lawson, 1985; Winkelmann, 1986; Willis and
Coggeshall, 1991; Perl, 1996). Thus, the presence of the HT channels in
small DRG neurons indicates their possible role in the mediation of
mechanical pain in nociceptive neurons. Second, the majority of HT
channels were activated by pressures over 60 mmHg. This channel had the
highest-pressure threshold found in sensory neurons. Furthermore, the
channel was sensitized by PGE2.
PGE2 released in response to tissue injury and
inflammation is known to cause mechanical hyperalgesia and sensitize
sensory neurons to mechanical stimuli specifically via the protein
kinase A pathway (Martin et al., 1987; Schaible and Schmidt, 1988;
Taiwo and Levine, 1988; Ouseph et al., 1995; Chen et al., 1999; Cunha et al., 1999). In the present study, PGE2 lowered
the threshold of HT, but not that of LT MS channels. The sensitization
of HT channels by PGE2 was mediated by the
protein kinase A pathway (Fig. 7). Thus, our results indicate that the
HT MS channel is a likely candidate for transducing mechanical stimuli
to nociceptive neural signals.
Many MS channels in various cells are often coupled to the cytoskeleton
(Hamill and Martinac, 2001). Thus, cytoskeletons may be an important
factor for regulating MS channel function. Colchicine, an antimitotic
drug that disrupts microtubules (Borgers et al., 1975), abolishes
mechanotransduction in Caenorhabditis elegans (Chalfie and
Thomson, 1982). The activity of TRAAK, a mammalian neuronal
two-pore K+ channel that is sensitive to
mechanical stimulation, is enhanced by colchicine treatment (Maingret
et al., 1999), and cytochalasin D increases the sensitivity of MS
channels to stretch in chick skeletal muscle and in Lymnaea
neurons (Guharay and Sachs, 1984; Small and Morris, 1994). Thus, MS
channel activity appears to be dependent largely on the integrity of
the cytoskeleton. In the present study, disruption of cytoskeletons by
colchicine or cytochalasin D greatly reduced the activity of MS
channels. In addition, although it is not clearly defined, excision of
the cell membrane to make inside-out or outside-out patches is likely to affect the membrane-cytoskeleton interaction (Hamill and McBride, 1997; Marchenko and Sage, 1997; Maingret et al. 1999). We also observed
that excision of the cell membrane greatly reduced the current
responses to pressures. Especially, when outside-out patches were
formed, a substantial portion of the membrane was pulled and
re-annealed. After forming outside-out patches, however, the integrity
of cytoskeleton was not lost completely because activities of MS
channels remained (Fig. 5). Thus, unlike other MS channels, whose
activity was enhanced after cytoskeletal disruption, cytoskeleton and
associated proteins seem to be functionally connected to the MS
channels in sensory neurons as positive regulators.
In summary, we report here on two distinct types of MS channels present
in cultured sensory neurons. These cationic channels have different
pressure thresholds as well as distinct biophysical properties. Thus,
the channels may account in part for macroscopic MS currents observed
in sensory neurons and subserve molecular transducers in sensory
neurons for mechanosensations such as somatosensation or
proprioception. Because the HT channels are present in small sensory
neurons, activated only by high pressures, and sensitized by
PGE2, the HT MS channels are also likely to be
implicated in generation of nociceptive neural signals. However, the
precise functional roles of the channels remain to be studied until
molecular species for the channels are identified.
| |
FOOTNOTES |
Received Aug. 13, 2001; revised Nov. 27, 2001; accepted Nov. 29, 2001.
The present study was supported by Creative Research Initiatives of the
Ministry of Science and Technology of Korea and in part by a BK21
program. We thank Dr. Donghee Kim for his critical reading of this manuscript.
Correspondence should be addressed to Dr. Uhtaek Oh, Sensory Research
Center, CRI, Seoul National University, College of Pharmacy, Kwanak,
Shinlim San 56-1, Seoul 151-742, Korea. E-mail: utoh{at}plaza.snu.ac.kr.
| |
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ABSTRACT
INTRODUCTION
