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First published online November 2, 2007
Journal of Experimental Biology 210, 3910-3918 (2007)
Published by The Company of Biologists 2007
doi: 10.1242/jeb.009662
Nitric oxide modulation of the electrically excitable skin of Xenopus laevis frog tadpoles
School of Biology, University of St Andrews, St Andrews, Fife, KY16 9TS, UK
* Author for correspondence (e-mail: kts1{at}st-andrews.ac.uk)
Accepted 30 August 2007
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Summary |
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Key words: nitric oxide, tadpole, modulation, skin impulse
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Introduction |
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). This excitability takes the form of an action potential or
`skin impulse,' which resembles a cardiac action potential in duration and
waveform, and propagates from the point of initiation throughout the skin
via electrical connections between neighbouring cells. A range of
anurans, including the South African clawed frog Xenopus laevis
(Daudin), the common frog Rana temporaria, the common toad Bufo
bufo, and salamanders such as Amystoma, have been shown to
display such excitability (Roberts,
1998). One known function of the skin impulse is to trigger escape
behaviour and thereby allow the tadpole to evade predation.
In Xenopus, the skin impulse pathway is one of two sensory systems
in the skin that operate in parallel
(Roberts and Smyth, 1974), the
second one involving a more conventional innervation by mechanosensory
Rohon-Beard (R-B) cells, a subset of extra-ganglionic sensory neurones
(Hughes, 1957). In early
embryos, at stage 27 [about 24 h post-fertilization
(Nieuwkoop and Faber, 1956)],
the skin is already excitable, including in areas yet to be innervated by R-B
cells. Both the skin impulse and the R-B cells can activate neural circuitry
of the spinal cord to initiate trunk flexion in young embryos and rhythmic
swimming movements in older embryos and larvae, but through different routes.
R-B cells directly activate spinal neurons
(Clarke et al., 1984;
Sillar and Roberts, 1988),
while the skin impulse appears to gain access to the central nervous system
(CNS) via a branch of the trigeminal nerve, bypassing primary sensory
R-B neurons in the skin (Roberts,
1996). Thus the electrically excitable epithelium functions as a
bona fide sensory system, which initially precedes and then operates
in parallel with more conventional mechanosensory innervation, before its
excitable properties disappear during later larval life.
Most cutaneous sensory systems are subject to modification under different
circumstances (Sillar, 1989).
The presence of a range of neuromodulatory substances in the skin of
Xenopus raises the possibility that the skin impulse and its
propagation through the epithelium may be subject to regulation, as is the
case for other, more conventional sensory systems. One such modulator, which
is produced by the skin of many vertebrates, including humans
(Weller, 1997), is the free
radical, nitric oxide (NO). NADPH-diaphorase labeling, a marker for the
presence of the NO synthetic enzyme NOS, has been noted in some cells of the
skin of Xenopus embryos at the hatchling stage (37/38), suggesting NO
production in the epidermis (McLean and
Sillar, 2000; McLean et al.,
2001) and a possible role in modulating the skin impulse. Similar
labeling is also found in the skin at equivalent stages of development in
Rana (McLean et al.,
2001).
In the present paper we have investigated the modulatory effects of NO on the duration and rate of propagation of the skin impulse in hatchling Xenopus tadpoles and report a profound effect on both parameters. The NO donor, SNAP, reduces the rate of propagation and increases the duration of the impulse, an effect that is countered by the NO scavenger C-PTIO. SNAP also produces a significant depolarization of the membrane potential of skin cells. The endogenous source of NO has been explored using a range of anatomical techniques, revealing a scattered distribution of NO-producing skin cells over the entire surface of the tadpole. This raises the possibility that the endogenous release of NO modulates the properties of the skin impulse, and the circumstances under which this might occur are discussed.
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Materials and methods |
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)
were obtained by induced breeding of pairs of adults selected from a
laboratory colony. Eggs were collected and reared in aerated trays at
temperatures of approximately 17–23°C to stagger their development
until they had reached the desired stage.
Electrophysiology
For extracellular recordings of the skin impulse and/or ventral root
activity, animals were first immobilized in 12.5 µmol l–1
-bungarotoxin (Sigma, Gillingham, Dorset, UK), before being transferred
to a recording bath filled with recirculating Hepes-buffered saline
(composition in mmol l–1: 115 NaCl, 2.5 KCl, 2.5
NaHCO3, 10 Hepes, 1 MgCl2 and 2 CaCl2,
adjusted to pH 7.4 with NaOH). Saline was gravity-fed from a 100 ml reservoir
into a PerspexTM chamber (c. 5 ml volume) containing a platform with a
SylgardTM (Dow-Corning, Midland, MI, USA) surface onto which the
immobilized animals were secured with fine etched tungsten pins through the
notocord. In some experiments an area of skin on the left side of the flank
was removed using fine etched tungsten needle to allow a ventral root
recording to be made by positioning an extracellular glass suction electrode
over an intermyotomal cleft (Fig.
1A). Initiation of the skin impulse and fictive swimming was
accomplished by stimulating through a second glass suction electrode, placed
on the tail skin, which delivered a 1 ms current pulse via a DS2A
isolated stimulator (Digitimer, Welwyn Garden City, UK). Signals were
amplified using differential AC amplifiers (A-M Systems Model 1700, Carlsborg,
WA, USA), displayed on a digital oscilloscope, digitized using a CED micro
1401 and stored and processed on a PC computer using Spike2 software
(Cambridge Electronic Design v. 3.21).
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.
Signals were amplified with a custom-built DC amplifier (courtesy of Dr Steve
Soffe, University of Bristol, UK). Penetration of skin cells was achieved by a
brief capacity overcompensation, normally revealing a resting potential of
approx. –50 to –80 mV. Cells were recorded for as long as the
penetration could be maintained, but in most cases recordings were stable for
only a few minutes. With unrecoverable cell loss, the electrode was withdrawn
to above the surface of the epidermis, moved a few tens of µm laterally,
and then lowered again to record from another skin cell. The sharp electrode
technique allowed serial recordings of multiple skin cells to be sampled
before, during and after drug applications. In addition, experiments were
performed using the whole-cell patch clamp technique, which allowed stable,
long-term recordings of the skin impulse; the data obtained using either
technique were very similar (Fig.
1B,C). For patching of skin cells, an area of the outer layer of
the two-cell thick epithelium, usually over the yolk sac
(Fig. 1A; `YS'), was carefully
peeled away using fine-etched tungsten needles and watchmaker's forceps. Patch
electrodes (ca. 10 M) were pulled on a Narishige puller (model PP-830,
Willow Way, London, UK) and filled with intracellular solution (in mmol
l–1: 100 potassium gluconate, 2 MgCl2, 10 EGTA, 10
Hepes, 3 Na2ATP, 0.5 Na-GTP, adjusted to pH 7.3 with KOH).
Recordings in whole-cell mode were amplified with an Axoclamp 2B amplifier and
displayed on a PC using Spike 2 software. All reagents were obtained from
Sigma or Tocris Bioscience (Bristol, Avon, UK). SNAP was also provided by the
School of Chemistry, University of St Andrews, Scotland.
Electrophysiological data were analysed using Dataview software (v 4.7c,
courtesy of Dr W. J. Heitler). Extracellular recordings were used to measure
both motor bursts from the ventral root of the spinal cord during fictive
swimming, and the voltage associated with the skin impulse from a population
of epidermal cells. In some experiments, the swimming cycle period and episode
duration were measured as a positive control to confirm that applied drugs
(e.g. SNAP) produced effects on these parameters similar to those previously
documented (McLean and Sillar,
2000). The duration of the intracellular skin impulse was measured
as the interval between the initial rapid depolarization to the point where
the membrane potential returned to rest. Skin impulse delay was calculated as
the interval between the stimulus artefact and the onset of the rising phase
of the impulse. All raw data consisting of measurements from multiple skin
cells were imported into Excel spreadsheets, where data from each period (i.e.
control, drug and wash) were averaged for duration and membrane potential. To
analyze changes in delay, data from the entire periods before, during and
after drug application were fitted to linear regression lines, and the slopes
of these lines were compared using a one-way ANOVA. This method was chosen
because drug-induced changes in delay were gradual, and because in some
experiments they continued for a period during the subsequent wash phase, thus
contaminating the average of that phase of the experiment. Measuring the slope
of the phase reduced this confounding effect. Sharp microelectrode
measurements during an experiment were subject to a skin cell being
penetrated, and were thus made at irregular intervals, but allowed sampling of
multiple cells during each experiment. In contrast, patch microelectrode
recordings were usually maintained for the duration of an experiment, and thus
yielded continuous data, albeit from only one cell per preparation. ANOVAs
were performed on sharp electrode data to determine if there was a
statistically significant difference between periods of a given experiment.
These data were pooled from multiple skin cells from each period and averaged;
either the average value for duration and membrane potential, or the average
slope for delay. Tukey's post-hoc test was used to determine
statistical significance between control, drug and wash periods.
Anatomy
The general topography of the skin surface was studied by immersing stage
37/38 wild-type embryos in a dilute mixture of two vital dyes, Methylene Blue
and Fast Green in tapwater, for 5–10 min. This procedure enhanced
visualization of the boundaries between skin cells. Animals were removed from
the solution, immobilized in MS222 and the flank skin was peeled off, mounted
on a slide in Hepes-buffered saline and topped with a coverslip, ready for
viewing.
NADPH-d histochemistry
The NADPH diaphorase (NADPH-d) histochemical method was applied as
described previously (McLean and Sillar,
2000) but to whole embryos and excised skin patches from stage
37/38 Xenopus. Both wild-type and albino embryos (the latter were
used to enhance contrast) were fixed in 4% paraformaldehyde (pH 7.4, 4°C)
for 2 h on a rocking agitator. The animals were then washed in phosphate
buffer (PB; 3x5 min), transferred to 30% sucrose in 0.1 mol
l–1 PB and stored in the refrigerator until they sank.
Animals were then immersed in 5 ml of NADPH-d staining solution [consisting of
5 mg NADPH (Sigma N-1630), 4.95 ml 0.3% PB-TX, and 50 µl Nitroblue
Tetrazolium salt (NBT; Sigma N-6639) made up from 5 mg NBT dissolved in 0.5 ml
PB-TX] and incubated at 37°C for 2 h. Animals were then washed in 0.1 mol
l–1 PB (3x5 min), dehydrated in acetone/alcohol series
and cleared in xylene before being mounted with DPX in a cavity slide, sealed
and topped with a coverslip, ready for viewing.
nNOS immunofluorescence
Neuronal NOS (nNOS) immunocytochemistry was performed using protocols
developed previously for Xenopus (Ramanthan et al., 2006). Animals
were fixed in 4% paraformaldehyde (pH 7.4) for 2 h at room temperature on a
rocker (Grant-Bio PMR-30, Shepreth, Cambridgeshire, UK), washed in 0.1 mol
l–1 PB (3x5 min), then placed in 5 ml of blocking serum
and primary antibody. This solution consisted of 5% normal goat serum (Jackson
Immuno: 005-000-121, Westgrove, PA, USA); 3% bovine serum albumin in PBS-TX
(pH 7.4; 0.9% PBS plus 0.3% Triton X-100); and primary rabbit polyclonal
antibody [1:100 v/v, NOS-1 (R-20):sc-648, Santa Cruz Biotechnology, Inc.,
Santa Cruz, CA, USA]. Following incubation at 37°C for 24 h, samples were
washed in 0.1 mol l–1 PBS (3x10 min), then incubated in
the dark in diluent and secondary antibody (pH 7.4; AffiniPure Goat
Anti-Rabbit IgG, Jackson Immuno) at 1:100 v/v for 24 h. Samples were then
washed in 0.9% PBS for 24 h, mounted with Citifluor and topped with a
coverslip, as above.
DAF-2 DA fluorescent labelling
Animals were immersed in a solution of 1 µl ml–1 DAF-2
DA (4,5-diaminofluorescein diacetate; Calbiochem, La Jolla, CA, USA) in Hepes
saline, then placed on a rocking agitator for 15–30 min. The staining
solution was replaced with 4% paraformaldehyde in PB (pH 7.4, 4°C) for 2
h. After fixation, animals were washed in 0.1 mol l–1 PB
(3x5 min), mounted in a glass cavity slide, using Citifluor (glycerol
solution, AF2), then coverslips placed on top. The edges of the coverslips
were affixed using clear nail varnish. In some experiments tadpoles were
anaesthetized in MS222 then mounted in the staining solution, placed beneath a
coverslip and viewed using an epifluorescence microscope. This method revealed
that the optimum time for DAF-2DA labeling was 15-30 min; after more than
approximately 2 h the fluorescence of skin cells all but disappeared.
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;
Roberts and Stirling, 1971).
The skin impulse was triggered by a stimulus applied to the caudal tail
[Fig. 1A, Stim (ext)] and
recorded either intracellularly on the head region with sharp electrodes
[Fig. 1A, SKC (sharp);
Fig. 1B, bottom trace] or on
the flank using patch electrodes [Fig.
1A, SKC (patch); C, bottom trace]. In some experiments the skin
impulse and/or ventral root activity were also recorded extracellularly
[Fig. 1A, SKC (ext), VR (ext);
B,C top traces].
Intracellular recordings with either sharp or patch electrodes reveal an
overshooting skin impulse with a characteristic waveform that superficially
resembles a cardiac action potential (Fig.
1B,C). The only obvious difference between the two recording
techniques is that the patch recordings have a sharper rising phase (time to
peak ca. 5 vs 30 ms), presumably due to the lower electrode
capacitance. There is a relatively rapid rising phase, which is followed by a
longer plateau phase (ca. 60 ms to 150 ms) and then by a slow decay back to
rest. The total duration of the impulse varied with different recordings, but
under control conditions was usually in the range of 150 to 200 ms. The onset
of the rising phase of the skin impulse had a delay from the stimulus of 60 to
100 ms (Fig. 1B,C) which, with
a separation between stimulating and recording electrodes of approximately 4
mm (Fig. 1A), equates to a
conduction velocity of approximately 4–7 cm s–1,
similar to that reported previously [5–11 cm s–1
(Roberts, 1971)]. The measured
conduction delay varied within and between different preparations and depended
upon a range of parameters including stimulus frequency, with higher
frequencies increasing delay (not illustrated). The extracellular recordings
showed a complex, multi-phasic waveform presumably reflecting the relatively
rapid rising phases of several skin cells located beneath the recording
electrode. The subsequent slower phases of the impulses were attenuated due to
the high-pass filter characteristics of the extracellular amplifier. A
supra-threshold stimulus that initiated a skin impulse usually also initiated
an episode of fictive swimming, as recorded using a suction electrode
positioned over a ventral root (Fig.
1C, top trace).
The effects of the NO donor, SNAP
The NO donor, SNAP [200–500 µmol l–1 (see also
McLean and Sillar, 2000)] was
bath-applied to investigate the potential modulatory effects of NO on the skin
impulse. SNAP had three consistent and highly significant (P<0.01)
effects (Figs 2,
3): (1) it increased the
duration of the skin impulse (Fig.
2Ai,Bii, Fig. 3A);
(2) it increased the delay from the stimulus to the skin impulse
(Fig. 2Aii,B,
Fig. 3B); and (3) it caused the
membrane potential to depolarize (Fig.
2Aiii,B, Fig. 3C).
These effects were apparent in both patch
(Fig. 2; N=5) and
sharp-electrode (Fig. 3;
N=27) recordings. The increase in duration became clear shortly after
the application of SNAP, and its rise to a stable maximum was relatively rapid
(e.g. Fig. 2Ai). The duration
increase reversed significantly towards baseline levels with wash
(Fig. 3A; P<0.05).
In contrast, the delay consistently showed a brief but significant
(P<0.05) decrease immediately following SNAP application (down
arrow in Fig. 2Aii), followed
by a slow increase. This increase frequently continued even in the initial
stages of the wash, although it usually started to gradually reverse with
sustained washing (Fig. 2Aii).
The delay did not usually completely return to baseline within the time period
of an experiment, although the reversal was significant (P<0.05).
Similarly, the effect on membrane potential was gradual in onset and only
partially reversed on washout (Fig.
2Aiii, Fig. 3C). To
control for vehicle-related effects, 0.5% dimethyl sulfoxide (DMSO) was
applied on its own. In pooled data for six cells, no significant changes were
observed for delay, duration or resting potential (not illustrated). In
summary, the delay and duration of the skin impulse and the resting potential
of skin cells are significantly affected by SNAP; these effects are not
vehicle related and therefore are presumably mediated by NO.
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) and
Rana (McLean et al.,
2001). To confirm this finding, skin samples from both wild-type
(Fig. 5B) and albino
(Fig. 5C) stage 37/38
Xenopus embryos were stained using the NADPH-d technique. In both
wild-type (N=6) and albino embryos (N=9), a proportion of
skin cells, approximately one in five, displayed positive staining and
although the intensity of labeling was rather variable, this punctate pattern
was observed over the entire surface of the body.
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Although the NADPH-d technique is a reliable marker for NOS in older stage
Xenopus tadpole CNS neurons
(Ramanathan et al., 2006), we
performed NOS immunocytochemistry to confirm this with skin of hatching stage
Xenopus embryos. Since the skin and the nervous system share a common
embryological origin, an nNOS antibody was used on excised pieces of skin from
three stage 37/38 embryos (N=3;
Fig. 5D). nNOS-positive
labeling displayed an irregular patterning in all skin patches with a
distribution that was very similar to that found using the NADPH-d technique.
In control preparations, where the primary antibody was omitted from the
incubation schedule, no fluorescence was observed
(Fig. 5E; N=3). This
suggests that the staining observed in the skin cells is specific to nNOS.
The experiments using NADPH-d and nNOS immunohistochemistry strongly
suggest that a sub-population of skin cells possess the enzyme necessary for
generating NO. To investigate further whether NO is actually produced by cells
in the skin, as has been shown at later premetamorphic stages
(Wilding and Kerschbaum,
2007), stage 37/38 Xenopus embryos were treated with the
fluorescent probe DAF-2 DA, a cell-permeable molecule converted into the
cell-impermeable DAF-2 by intracellular esterases. Within NO-generating cells,
NO binds DAF-2 to form fluorescent triazolofluorescein. Green fluorescence,
indicating the presence of NO, was found in about 20% of cells, dispersed over
the entire surface of the animal in whole-mount preparations
(Fig. 5F; N=12). This
punctate pattern of staining in the skin was observed over the entire surface
of the body, although the clearest fluorescent signal was found in the fin of
the tadpole, where only skin cells are present. Thus, the appearance of DAF
fluorescence in the skin suggests that skin cells are not only NOS positive,
but actually produce NO from these early stages of development.
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Discussion |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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;
Wilding and Kerschbaum, 2007).
NO-producing cells are distributed apparently randomly and scattered
throughout the skin of hatchling Xenopus tadpoles. Given this spatial
organization of NOS, global upregulation of NO produced in the skin may
modulate the skin sensory pathway over the entire body surface, or local
increases in NOS activity could occur in response to a focal challenge,
producing a spatially restricted response. However, the circumstances
responsible for triggering NO release have not been investigated.
Modulation of the skin impulse by NO
Several neuroactive substances produced in the skin of amphibians,
including NO (McLean and Sillar,
2001; Wilding and Herschbaum,
2007; Zaccone et al.,
2006), biogenic amines and peptides
(Erspamer, 1971), could in
principle modulate the skin impulse, either by acting on the biophysical
properties of individual skin cells, or on the propagation of the impulse
via direct gap junctional coupling between cells, or both. In this
paper we have addressed the possibility that NO functions in this capacity.
Experiments with the NO donor, SNAP, have led to three new findings: (1) SNAP
produces a fast, large and reversible increase in the duration of the skin
impulse; (2) SNAP reversibly decreases the conduction velocity of the skin
impulse as it propagates across the epithelium; and (3) SNAP depolarizes skin
cells. The effects of SNAP on each of these parameters can be reversed by the
NO scavenger, C-PTIO.
The skin impulse comprises a fast rising phase that is thought to be
Na+-dependent (Roberts,
1971), a slower plateau phase, and a relaxation phase that returns
the membrane potential back to rest. The ionic bases of the plateau and
relaxation phases are not known, but the increase in duration in the presence
of SNAP is likely to be due to effects on these later phases, because the rise
time of the impulse is not markedly affected by SNAP
(Fig. 2B). The SNAP-released NO
could, for example, modulate cyclic nucleotide-gated channels (CNG)
(Hofmann et al., 2005) by
acting via a 2nd messenger such as cGMP or cAMP. Such channels could
contribute to the repolarizing phase of the impulse, and/or could also
underlie the SNAP-induced depolarization of skin cell resting potentials.
Given the ability of NO to uncouple gap junctions (GJs) in other systems
(Fessenden and Schacht, 1998;
Kameritsch et al., 2003;
Kameritsch et al., 2005;
Yao et al., 2005;
Patel et al., 2006), the SNAP
effects on delay could be explained if NO reduces the junctional coupling
thought to be responsible for propagation of the impulse between the cells of
the skin (Roberts, 1971). Such
a reduction could increase the effective path length as the skin impulse
follows a more circuitous route from the point of initiation to the recording
site and/or it could reduce the rate of depolarization of successive cells in
the pathway thus producing a cumulative increase in propagation delay.
Connexins and their resulting intercellular GJs are present in the early
embryo (reviewed in Mackie,
1970; Warner,
1985; Kandler and Katz,
1995; Levin and Mercola,
2000; Landesman et al.,
2003) and could be retained through the late embryo stage. This is
important because the presence of GJs beginning at stage 22 when the skin
impulse is first found (Roberts,
1971), would support Roberts' theory that the impulse propagates
through the skin by direct current flow via low-resistance
junctions.
NO is present in the skin
NADPH-d histochemistry is a technique used to localize putative
NOS-containing cells. The NADPH-d reactivity found in cross sections of skin
(McLean and Sillar, 2001), and
the further identification of labeling in whole-mount skin samples presented
here, support the conclusion that NOS is localised to a sub-population of skin
cells. Staining was found in both wild-type and albino skin, with staining
being much clearer in the latter due to the lack of pigment obscuring the
labelling. However, NADPH specificity for NOS can be unreliable (cf.
Vincent, 1995). The appearance
of nNOS immunofluorescence in excised skin patches confirmed the presence of
NOS in skin cells of Xenopus embryos at stage 37/38, with the pattern
of staining being similar to the punctate distribution of NADPH-d staining.
During vertebrate development, the skin is derived from the ectodermal tissue.
This early germ layer also differentiates into nervous tissue. Thus, nNOS
present in skin cells at stage 37/38, may be retained from synthesis at
earlier stages in development. During the time of gastrulation, cells could
express the NOS enzyme before diverging to become either part of the CNS where
NOS is expressed in brainstem neurons at early embryonic stages
(McLean and Sillar, 2001), or
part of the epidermis, as indicated by the nNOS immunofluorescence reported
here.
DAF-2 fluorescence was used to visualize the endogenous production of NO in skin cells and a punctate pattern of staining was again found, suggesting that NO is produced by about 20% of skin cells, similar to the distribution of NADPH-d and nNOS staining. Taken together these results suggest that NOS and NO colocalize in the same skin cells. The identification of endogenous NO through DAF-2 labelling suggests that NO is being released as an autocrine or paracrine messenger. The punctate distribution of NOS across the skin, together with the ease with which NO can diffuse through tissue, suggests that NO can affect the entire surface of the tadpole.
A similar distribution of NADPH diaphorase labeling and NO production has
been described in the skin at later stages of Xenopus tadpole
development (Wilding and Kerschbaum,
2007), where NO has been implicated in the regulation of ammonium
release. In a related amphibian species, Triturus italicus, the
endothelial isoform of NOS (eNOS) is expressed strongly in the larval skin
(Brunelli et al., 2005), but
this begins to disappear in pre-metamorphic and metamorphic periods coincident
with a simultaneous rise in expression of the inducible isoform of NOS (iNOS).
This suggests that in Triturus, a switch occurs in the developmental
cycle, and NOS-derived NO may be responsible for the remodeling of skin cells
during the metamorphic period.
Functional considerations
Given that NO appears to be produced and released by a proportion of skin
cells and is capable of modulating skin physiology, what might be the role of
this phenomenon for the hatchling tadpole? One possibility is that the NO
alters the properties of the skin pathway for eliciting escape responses. The
threshold stimulus to evoke the skin impulse did not appear to change during
the bath application of SNAP (not illustrated), however, suggesting that NO
does not normally modulate the sensitivity of this sensory pathway. The
increased duration of the skin impulse might enhance the efficacy of its
transmission to the central nervous system, although the behavioural advantage
of slowing skin impulse propagation is unclear. Another possibility is that NO
is involved in the response to tissue damage. Noxious stimuli to the skin
could increase NO production, and it is known that NO can enhance repair
processes (reviewed in Witte and Barbul,
2002). NO-induced gap junction uncoupling could reduce the leakage
of intracellular contents from intact cells through damaged tissue, while the
increased duration of the skin impulse will prolong activation of
voltage-dependent channels and could thus have profound effects on
intracellular second messengers, including those potentially involved in wound
healing.
We have shown that a widespread but punctate population of skin cells contain the crucial enzyme for producing NO and that a similar population do indeed have elevated levels of NO. This population could thus act as a source to trigger the significant NO effects on skin cell properties that we have documented, notably the changes in the rate of propagation and waveform of the skin impulse. Taken together, our data point to NO as a potent modulator of the skin impulse in Xenopus laevis.
List of abbreviations
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Acknowledgments |
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TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
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