2 Division of Gastroenterology and Hepatology and Departments of 1 Physiology and Biophysics and 3 Ophthalmology, Mayo Clinic and Mayo Foundation, Rochester, Minnesota 55905
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ABSTRACT |
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The effects of melatonin on ionic conductances in a cultured
mouse lens epithelial cell line (-TN4) and in cultured human trabecular meshwork (HTM) cells were measured using the amphotericin perforated patch whole cell voltage-clamp technique. Melatonin stimulated a voltage-dependent
Na+-selective current in lens
epithelial cells and trabecular meshwork cells. The effects of
melatonin were observed at 50 pM and were maximal at 100 µM.
Melatonin enhanced activation and inactivation kinetics, but no change
was observed in the voltage dependence of activation. The results are
consistent with an increase in the total number of ion channels
available for activation by membrane depolarization. Melatonin was also
found to stimulate a K+-selective
current at high doses (1 mM). Melatonin did not affect the inwardly
rectifying K+ current or the
delayed rectifier type K+ current
that has been described in cultured mouse lens epithelial cells. The
results show that melatonin specifically stimulated the TTX-insensitive
voltage-dependent Na+ current by
an apparently novel mechanism.
sodium channels; trabecular meshwork; epithelium; electrophysiology
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INTRODUCTION |
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THE SYNTHESIS AND SECRETION of melatonin (N-acetyl-5-methoxytryptamine) by the pineal gland and the retina occur at night, and recent reviews give excellent summaries (10, 13, 16, 17, 21, 27). Circulating melatonin can modulate several physiological processes and may alter the timing of circadian rhythms (4, 8). The effects of melatonin on circadian rhythms are likely mediated by high-affinity receptors located in the suprachiasmatic nucleus (27). High-affinity receptors have also been identified in the inner plexiform layer of the retina as well as the hypophysial pars tuberalis (17). Three specific melatonin receptor subtypes have been identified: Mel1a [dissociation constant (Kd) = 20-40 pM] in the suprachiasmatic nucleus and pars tuberalis, Mel1b (Kd = 160 pM) in the retina, and Mel1c (Kd = 20-60 pM). However, the Mel1c receptor has not been cloned in mammals (27). Receptor activation results in inhibition of adenylate cyclase by a pertussis toxin-sensitive G protein (17, 34). The decrease in intracellular cAMP mediates aggregation of pigment granules in frog skin, photoperiodic regulation of prolactin secretion by the hypophysial pars tuberalis, and vasoconstriction of rat cerebral artery (4, 34). However, the signaling mechanisms coupling melatonin receptor activation to neuronal activity in the suprachiasmatic nucleus or to the inhibition of dopamine release in the retina remain unknown (16).
There is evidence that melatonin modulates ionic conductances in several preparations. Melatonin suppresses the timing of neurons in the suprachiasmatic nucleus in vitro (19). Recent experiments have shown that melatonin stimulates a K+-selective conductance through a receptor-mediated mechanism (15). The activity of several ionic conductances in cultured cells shows circadian patterns, but the signaling mechanisms are unknown. For example, a tetraethylammonium-sensitive K+ conductance in the optic nerve of mollusk is lowest before dawn and increases at dusk (20). A Ca2+-permeable cation-selective conductance in cultured avian pineal gland cells is spontaneously active at night, coincident with melatonin secretion (9).
Melatonin is lipophilic and thus has access to cytosolic,
mitochondrial, and nuclear compartments. In the cytosol, melatonin has
been shown to act as a
Ca2+/calmodulin antagonist and
therefore may directly affect
Ca2+ signaling mechanisms (10,
13). Melatonin-calmodulin binding has been shown to affect tubulin
polymerization and alter the cytoskeleton in Madin-Darby canine kidney
and NIE-115 cells (3). Addition of melatonin has also been shown to
stimulate the Ca2+-ATPase in
cardiomyocytes (6). Melatonin is a powerful free radical scavenger and
may act ubiquitously as an intracellular antioxidant (26). A
high-affinity binding site has been identified in the nucleus
(Kd 10
9 M), where melatonin may
regulate gene transcription or may prevent oxidative damage to DNA (2).
Thus the effects of melatonin may result from binding to high-affinity
receptors in the plasma membrane, cytosol, or the nucleus, leading to
changes in gene expression or altering second messenger levels (cAMP
and Ca2+). The effects of
melatonin may also result from nonspecific interactions related to the
antioxidant activity of melatonin or from direct binding to ion channels.
Melatonin synthesis and secretion by the retina have been well characterized, as well as inhibition by melatonin of dopamine release in the retina, but the target receptors and the mechanisms by which melatonin alters physiological function are not as well understood. Because retinal melatonin does not significantly contribute to circulating melatonin levels, a local effect on ocular tissues, such as modulation of neuronal transmission or excitability, or regulation of intraocular pressure is possible. Intraocular pressure varies diurnally, decreasing at night, and this has been associated with the circadian fluctuations in melatonin concentration in plasma or aqueous humor (18). Plasma melatonin levels have also been correlated with intraocular pressure in humans, and intraocular pressure decreased after oral administration of melatonin (29). However, the basic mechanism for these observations is not known.
We used the patch-clamp technique to measure the effects of melatonin
on ionic currents in two different cell types, a mouse lens epithelial
cell line (-TN4) and a human trabecular meshwork cell line (HTM). We
have found that melatonin specifically stimulates the
Na+ current in cultured mouse lens
epithelial cells by a novel mechanism (33). Voltage-gated
Na+ channels are present in most
excitable cells and can be divided into two types: TTX sensitive and
TTX insensitive. TTX-sensitive Na+
channels, found in nerve and skeletal muscle, have a TTX
Kd of 1-20
nM (12). TTX-insensitive Na+
channels have a TTX
Kd of 0.5-10
µM. This difference in binding affinity results from a change in a
single residue within the TTX binding domain (5). The
Na+-selective channel in mouse
lens epithelial cells is the TTX-insensitive type. Melatonin was also
found to stimulate a Na+ current
in cultured HTM cells, suggesting that the effects were not unique to
the Na+ current in
-TN4 cells,
but that melatonin is Na+ channel
specific. Finally, melatonin did not affect the inwardly rectifying K+ conductance or the
delayed rectifier type K+
conductance in
-TN4 cells but did stimulate an outward
K+ current in HTM cells at very
high doses.
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MATERIALS AND METHODS |
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Cell culture and isolation.
Experiments were performed on two types of cultured cells: an
immortalized transgenic lens epithelial cell line (-TN4) kindly
provided by Dr. Paul Russel, and cultured HTM cells kindly provided by
Dr. Richard Seftor.
-TN4 cells were cultured according to a standard
protocol (28). HTM cells were cultured following a previously published
protocol (30). Because cultured cells may change phenotype after
repeated passages, only cells from the third or fourth passage were
used for these experiments. Cells were dissociated using a trypsin
low-Ca2+ solution (0.5%) and by
gentle trituration with a Pasteur pipette. The cell suspension was
centrifuged at 180 g for 5 min and
resuspended in an enzyme-free Ringer solution. Centrifugation was
repeated, and the cells were resuspended in a Ringer solution
containing 5 mM glucose and stored at room temperature. Experiments
were performed during the next 6 h. Three to six drops of the cell suspension were placed in a 300-µl acrylic recording chamber with a
glass coverslip bottom. The cells were allowed to settle for ~30 min.
The bath was perfused with several volumes of NaCl Ringer solution to
wash away unattached cells and debris.
Patch electrodes for whole cell recordings were made from Kimble KG-12
glass (Garner Glass, Claremont, CA) and pulled on a Sutter Instruments
(Novato, CA) P80 microelectrode puller. Tips were coated with Dow
Corning (Midland, MI) Sylgard no. 184 and fire polished under direct
observation, to a final resistance of 3-5 M. Most whole cell
recordings were made using the amphotericin perforated patch technique,
since this technique results in stable values for the access resistance
and maintains cytoplasmic integrity (23). The electrodes were mounted
in a polycarbonate holder connected to an Axopatch 200 patch
voltage-clamp amplifier (Axon Instruments, Foster City, CA) and
positioned immediately adjacent to the cell membrane. Slight suction
resulted in a gigaohm seal for the majority of cells. Amphotericin
usually partitioned into the membrane isolated in the membrane tip
within 15 min, resulting in access resistance ranging from 6 to 20 M
. Data were recorded with the use of a modified IBM-AT computer
using a TL1 Labmaster interface (Axon Instruments) and driven by pCLAMP
software (version 6.0, Axon Instruments), allowing voltage-clamp
protocols with concomitant digitization of the membrane currents. Whole
cell current records were collected at 10 kHz or faster and filtered at
2 kHz with an 8-pole Bessel filter (Frequency Devices, Haverhill, MA).
Voltage step protocols were repeated twice, and the resulting currents
at each voltage were averaged to produce the final records. Each
current-voltage record was corrected for the offset potential resulting
from a combination of the liquid junction potential and the Donnan
potential produced by the mismatch of pipette and cellular anions.
Previous studies have shown that
K+-selective currents reverse
between 7 and 12 mV when the bath and pipette solutions contain equal
K+ concentrations (23). Therefore,
the current-voltage relationships were corrected by 10 mV. Capacity
transients were adjusted during recording. Whole cell currents were
usually stable during experiments lasting up to 1 h while using the
perforated patch technique or the whole cell technique. The records
were not leak subtracted or modified otherwise. Data were analyzed
using pCLAMP software (Axon Instruments) or using custom-written macros
in Excel (Microsoft, Redman, WA). Final plots were prepared using
SigmaPlot (Jandel, Madera, CA). Data are reported as means ± SE,
and statistical significance was determined using Student's
t-test.
Normal Ringer solution contained (in mM) 4.74 KCl, 149.2 NaCl, 2.54 CaCl2, and 5 HEPES, resulting in a final osmolarity of 293 mosM. For KCl Ringer solution, KCl was substituted for NaCl. The pipette filling solution contained (in mM) 125 KMeSO3, 20 KCl, 5 HEPES, and 2 EGTA, resulting in a final osmolarity of 289 mosM. All solutions were adjusted to a final pH of 7.35. Flufenamic acid, melatonin, fluoxetine (Prozac), quinidine, and diltiazem were purchased from Sigma (St. Louis, MO).
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RESULTS |
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Effects of melatonin on whole cell currents in
cultured mouse lens epithelial cells. Three current
types have been previously described in -TN4 cells: an inwardly
rectifying K+-selective current
(Iir), an
outwardly rectifying K+-selective
current (Idr),
and a Na+-selective
transient inward current
(INa) (7, 22,
33). Typical whole cell currents recorded in NaCl Ringer solution
during a series of voltage steps from
150 to +80 mV are shown in
Fig. 1A.
The three current types are apparent in this experiment.
Hyperpolarizing voltage steps activated
Iir, an inwardly
directed K+ current that activates
rapidly and inactivates very slowly during the voltage command.
Depolarizing voltage steps positive to
60 mV activated
INa. This inward
current reached a maximum very rapidly and completely inactivated
within 20 ms. Depolarizing voltage steps positive to approximately
20 mV activated
Idr. This outward K+ current activated more slowly
compared with INa
or Iir and
persisted for the entire duration of the voltage pulse.
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The effects of bath perfusion with 100 µM melatonin in NaCl Ringer
solution for this experiment are shown in Fig.
1B. An increase in
INa was evident
immediately after bath perfusion with melatonin. Iir and
Idr were not
affected. Current recordings and access resistance were verified to be
stable for 5-10 min after obtaining whole cell access. For some
experiments INa
exhibited rundown, a decrease with time, and these experiments were not
included in the analysis. Bath perfusion with normal Ringer solution
did not alter
INa; in 11 separate experiments
INa averaged
129 ± 38 pA before and
130 ± 39 pA
(P = 0.46) after perfusion with Ringer solution.
The peak inward current and steady-state current-voltage relationships
for this experiment are plotted in Fig. 2.
Figure 2A shows that
INa began to
activate at approximately 50 mV and reached the maximum of
238 pA at
20 mV (circles). Perfusion with 100 µM
melatonin (squares) increased peak
INa to
308
pA. An increase in
INa was observed
in 13 out of 16 cells. The average current was
288 ± 97 pA
in Ringer solution and
374 ± 112 pA
(n = 16, P = 0.003) after bath perfusion with
100 µM melatonin. The effects of melatonin were reversible;
INa averaged
373 ± 176 pA (n = 5, P = 0.33) after the wash with Ringer
solution. Melatonin did not appear to shift the voltage dependence of
INa, because the voltage at the peak current was unaltered. The steady-state
current-voltage relationship is shown in Fig.
2B. These currents were quantified as
the average of 20 data points near the end of the voltage step. Neither
Iir, measured as
the inward current evoked during voltage steps more hyperpolarized than
90 mV, nor
Idr, measured as
the outward current evoked by step depolarizations, was affected by melatonin. In six experiments
Iir measured
108 ± 29 and
99.5 ± 28 pA (P > 0.05)
and Idr measured 47.5 ± 18 and
55.5 ± 20 pA (P > 0.05) before
and after 100 µM melatonin, respectively. The resting membrane
potential (Em)
measured
16.0 ± 3.7 mV under control conditions and
25.0 ± 8.5 mV (P > 0.05)
after bath perfusion with 100 µM melatonin. These results show that
K+-selective currents in this
preparation are unaffected by melatonin.
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The site of action for melatonin may be a direct interaction with the
Na+ channel or with specific
melatonin receptors in the plasma membrane. Melatonin stimulates a
Ca2+-dependent ATPase pump in rat
cardiac myocytes with a
Kd of 28 ng/ml
(0.12 pM) (6). If melatonin stimulated the
Ca2+-ATPase pump in -TN4 cells,
then intracellular Ca2+ levels
should decrease. However, fluorescence experiments utilizing the
Ca2+-sensitive ratiometric dye
fura 2 showed that melatonin (100 µM) did not alter the intracellular
free Ca2+ concentration (data not
shown, n = 4). Melatonin was found to selectively stimulate
INa in
-TN4
cells, suggesting a tightly coupled mechanism, as opposed to a more
general cell signaling mechanism that would be expected to modulate
several ionic conductances in different cell types.
Stimulation of
INa by melatonin
may result from a change in the time course of activation or
inactivation. Several toxins such as TTX specifically alter
Na+ currents, and these effects
may be characterized by alterations in channel gating. The kinetics of
INa were examined
using a faster sampling rate (20 kHz) and a pulse protocol that focused
on the voltages where
INa was activated
(Fig. 3). Close inspection of the current
records in Fig. 3 shows that activation and inactivation kinetics were
faster after melatonin (Fig. 3B)
compared with control records (Fig.
3A). This is more clearly seen in
Fig. 3, inset, where the peak
currents, before and after melatonin, were plotted on an expanded time
scale. Activation time was measured as time from the start of the
depolarizing voltage step to time when
INa peaked. The
inactivation time course was well fit with a single exponential
function [inactivation time constant
(inactivation)]. The time to peak and
inactivation were determined at
test voltages where
INa was well
resolved for four experiments (Fig. 4).
Melatonin (100 µM, squares) decreased activation time as well as
inactivation (Fig.
4B).
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Dissociation constants for melatonin binding receptors range from 20 to
160 pM (27). The dose dependence of melatonin on INa is shown in
Fig. 5. These data show that melatonin
stimulated INa at
doses consistent with receptor binding. Figure 5 also shows further
stimulation of
INa by melatonin
at doses four orders of magnitude greater than the reported
dissociation constants. A maximal effect was observed at 100 µM, and
1 mM melatonin inhibited INa.
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Whole cell currents measured in HTM
cells. Whole cell currents were also measured in
cultured HTM cells. Typical whole cell currents recorded in NaCl Ringer
solution are shown in Fig.
6A. Cells
were held at 90 mV and stepped from
120 to 30 mV for 40 ms in 10-mV increments. The most prominent current under these conditions was a rapidly activating inward current that completely inactivated within 20 ms. The peak inward current and steady-state current-voltage relationships for this experiment are shown in Fig.
6B. Inward current began to activate
at approximately
20 mV and reached the maximum of
384 pA
at 10 mV in this cell.
INa averaged
538 ± 79 pA in seven separate experiments. The observed current activated at voltages of ~20 mV depolarized compared with INa measured in
-TN4 cells. The steady-state current-voltage relationship,
quantified as the average of 20 data points near the end of the voltage
step, was very small compared with
INa. Input
resistance, calculated as the slope of a straight line fit to the four
steady-state current values recorded during hyperpolarizing voltage
steps and constrained to go through the origin, was 11.6 ± 2.0 G
(n = 18).
Em was
39.2 ± 5.2 mV (n = 15). The
peak inward current was identified as
Na+ selective based on several
criteria. First, the only ions that have electrochemical gradients that
would drive inward current from
20 to 40 mV are
Na+ and
Ca2+. At these potentials
K+ would move outward
(Ek =
87.9
mV) and Cl
would move
inward (ECl =
53 mV), both resulting in outwardly directed current records.
Reducing the concentration of Na+
in the bath solution from 149.2 to 75 mM (while maintaining constant osmolarity with mannitol replacement) resulted in a decrease in the
inward current from 465 ± 141 to
228 ± 64 pA
(n = 7, P = 0.014), and replacing
extracellular Na+ with
K+ completely abolished the
current (data not shown). Extracellular Ca2+ concentration was constant
during these experiments, indicating that the inward current was not
carried by Ca2+. Furthermore,
replacing extracellular Ca2+ with
80 mM Ba2+, which enhances inward
current through L type Ca2+
channels, blocked the fast inward current (data not shown), indicating that the fast inward current is not carried by
Ca2+. Depolarizing the holding
potential reduced the inward current, identical to voltage-dependent
inactivation of
INa described in
-TN4 cells (33). The inward current in HTM cells also shared pharmacological properties with
INa that we have
observed in the
-TN4 cell line: fluoxetine, diltiazem, flufenamic
acid, and quinidine all blocked
INa (data not
shown). These data are all consistent with the hypothesis that the fast
inward current is Na+ selective.
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The effects of bath perfusion with melatonin in NaCl Ringer solution
are shown in Fig. 7. Bath perfusion with
100 µM melatonin (Fig. 7B)
stimulated INa,
similar to the effects observed in -TN4 cells. Higher levels of
melatonin (1 mM) also activated a prominent outward current
(Iout)
in this preparation (Fig. 7C).
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The current-voltage relationship for this experiment is shown in Fig.
8.
INa (Fig.
8A) activated at 20 mV and
reached a maximum of
720 pA at 0 mV. Bath perfusion with 100 µM melatonin increased INa to
1,044 pA. Melatonin increased
INa in six out of
six separate experiments from
500 ± 82 pA to
620 ± 110 pA (P = 0.04). Bath perfusion with 1 mM melatonin resulted in a decrease in
INa to
469
pA for the experiment shown and INa averaged
415 ± 91 (n = 7). The
effects of melatonin were reversible:
INa averaged
505 ± 84 (P = 0.44) after
bath perfusion with Ringer solution after melatonin.
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It is difficult to determine whether the decrease in the peak inward
current after 1 mM melatonin resulted from inhibition of
INa or from
stimulation of
Iout. Close
inspection of the current records in Fig. 7 shows that activation of
Iout overlaps
with INa.
Therefore, an increase in
Iout would be
reflected as a decrease in
INa. The effects
of melatonin on
Iout are shown in
Fig. 8B. Bath perfusion with 100 µM
melatonin increased outward current from 12 to 176 pA and
hyperpolarized Em
from 50 mV to
68 mV. Perfusion with 1 mM melatonin
increased Iout to
685 pA and further hyperpolarized
Em to
75
mV. These results suggest that
Iout was highly
K+ selective because the reversal
potential for K+ under these
conditions was calculated to be
86 mV.
The apparent similarity between Iout in HTM cells and the K+ current (IK) in rabbit corneal epithelial cells (see Fig. 1 in Ref. 11) led us to test the effects of agents that selectively inhibit IK in the corneal epithelium such as diltiazem (1 mM), quinidine (1 mM), and fluoxetine (100 µM) (11, 24, 25). These agents were all effective blockers of Iout: quinidine resulted in an 81 ± 8% (n = 3) decrease, fluoxetine resulted in a 92% (n = 1) decrease, and diltiazem resulted in a 39 ± 27% (n = 2) decrease in Iout. These results are consistent with the presence of a delayed rectifier type K+ current in HTM cells.
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DISCUSSION |
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Voltage-gated Na+ channels are
found in nerve cells, skeletal muscle, and smooth muscle and support
excitability in these cell types. Cardiac muscle, Purkinje fibers, and
denervated skeletal muscle also express voltage-gated
Na+ channels that support impulse
propagation. Na+ channels from
these tissues are distinct in that one type is TTX sensitive
(Kd 10
9 M) and the other is TTX
insensitive (Kd
10
6 M). Functional
expression of TTX-insensitive Na+
channels has also been observed in the lens epithelium and corneal endothelium (33). We have shown that melatonin stimulated
INa in cultured
human lens epithelial cells and in cultured HTM cells at doses
comparable with the reported
Kd for receptor
binding (20-160 pM) (17). Melatonin did not affect the other ionic
conductances that have been characterized in lens epithelial cells,
including an inward rectifying
K+-selective current
(Iir) and an
outward delayed rectifier type K+-selective current
(Idr) (7). We
also did not observe any effects on the delayed rectifier
K+-selective currents in two types
of freshly dispersed cells that are commonly used in the laboratory:
rabbit corneal epithelial cells or smooth muscle cells from the canine
jejunum (unpublished results). At high doses (1 mM) melatonin also
stimulated an outward K+ current
in cultured HTM cells.
Melatonin increased the peak Na+
current amplitude and the kinetics of activation and inactivation
(Figs. 3 and 4). The current-voltage relationship appears unaffected.
The most simple gating diagram for
Na+ channels exhibits three
connected conformational states: a nonconducting resting state, a
conducting open state, and a nonconducting inactivated state. A number
of toxins specifically alter Na+
channel gating. The most familiar may be the guanidinium toxins, TTX
and saxitoxin, commonly called Na+
channel blockers, which act by stabilizing the inactivated state (31).
Several other toxins increase
INa. The effects
of melatonin are most similar to -scorpion toxin, a small
water-soluble polypeptide, which prolongs action potential duration
(31). Lipophilic toxins, which are thought to act at sites within the
lipid layer, increase the activation rate like melatonin (31). However,
lipophilic toxins also change the voltage dependence of
activation. In cultured neuroblastoma cells
-toxin slowed the rate
of inactivation and thereby increased peak
INa (12). It is
possible that melatonin increased peak
INa due to
enhanced recovery from an inactivated state, thereby recruiting
channels for activation.
HTM cells function to maintain the composition of the aqueous humor outflow pathway in the juxtacanicular region of the eye. Regulation of the outflow facility through the trabecular meshwork is likely to play an important role in determining intraocular pressure. This hypothesis is supported by studies that correlate relaxation of the trabecular meshwork with an increase in ocular outflow (35). Cultured bovine trabecular meshwork cells have recently been shown to express maxi-K channels that are activated by cGMP (32). In smooth muscle cells, activation of K+-selective channels leads to membrane hyperpolarization and relaxation (14). However, little is known about ion channels in HTM cells. We have advanced these results and shown that cultured HTM cells also express Na+ channels and delayed rectifier type K+ channels. Ionic currents with similar pharmacology have been characterized in other ocular tissues, but molecular studies are necessary to unambiguously identify these channels (22).
The lens epithelium transports fluid and salt to preserve lens integrity. The physiological role of INa is not clear. One possibility, assuming that a small window current exists at the resting membrane potential, is that INa provides a Na+ inflow pathway that interacts with the Na+-K+ pump or that INa may serve as a cellular signal in response to Ca2+ depletion (33). The physiological role for Na+ channels in HTM cells must also depend on a small but significant channel open probability at the resting membrane potential. In this case INa may be involved in modulating the tone of HTM cells and thereby modulate outflow resistance and intraocular pressure.
Melatonin appears to be a novel activator of voltage-gated
TTX-insensitive Na+ channels. We
have shown that melatonin increases peak
INa and speeds
activation and inactivation kinetics and increases peak INa in two types
of cultured cells, -TN4 and HTM. Because melatonin was effective at
doses that are similar to the reported
Kd for the cloned
melatonin receptors, it is possible that these effects are receptor
mediated, but a direct interaction between melatonin and
Na+ channels cannot be ruled out.
Although a great deal is known about the regulation of melatonin
synthesis and the diurnal variation of melatonin in the bloodstream,
much less is known about its physiological roles. This study shows that
alterations in ion channel gating may be considered as a final effector
of the actions of melatonin.
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ACKNOWLEDGEMENTS |
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We thank Jan Applequist for secretarial help and Joan Rae for software development.
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FOOTNOTES |
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This work was supported by National Eye Institute Grants EY-O3282 and EY-O6005.
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 and other correspondence: A. Rich, Dept. of Physiology and Biophysics, Mayo Clinic and Mayo Foundation, 200 First St. SW, Rochester, MN 55905 (E-mail: rich.adam{at}mayo.edu).
Received 9 October 1998; accepted in final form 8 January 1999.
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