Department of Neurology, Yale Medical School, New Haven 06510; and Paralyzed Veterans of America/Eastern Paralyzed Veterans Association Neuroscience Research Center and Rehabilitation Research Center, Veterans Affairs Connecticut Healthcare Center, West Haven, Connecticut 06516
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ABSTRACT |
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Renganathan, M., T. R. Cummins, W. N. Hormuzdiar, J. A. Black, and S. G. Waxman. Nitric Oxide Is an Autocrine Regulator of Na+ Currents in Axotomized C-Type DRG Neurons. J. Neurophysiol. 83: 2431-2442, 2000. In this study, we examined whether nitric oxide synthase (NOS) is upregulated in small dorsal root ganglion (DRG) neurons after axotomy and, if so, whether the upregulation of NOS modulates Na+ currents in these cells. We identified axotomized C-type DRG neurons using a fluorescent label, hydroxystilbamine methanesulfonate and found that sciatic nerve transection upregulates NOS activity in 60% of these neurons. Fast-inactivating tetrodotoxin-sensitive (TTX-S) Na+ ("fast") current and slowly inactivating tetrodotoxin-resistant (TTX-R) Na+ ("slow") current were present in control noninjured neurons with current densities of 1.08 ± 0.09 nA/pF and 1.03 ± 0.10 nA/pF, respectively (means ± SE). In some control neurons, a persistent TTX-R Na+ current was observed with current amplitude as much as ~50% of the TTX-S Na+ current amplitude and 100% of the TTX-R Na+ current amplitude. Seven to 10 days after axotomy, current density of the fast and slow Na+ currents was reduced to 0.58 ± 0.05 nA/pF (P < 0.01) and 0.2 ± 0.05 nA/pF (P < 0.001), respectively. Persistent TTX-R Na+ current was not observed in axotomized neurons. Nitric oxide (NO) produced by the upregulation of NOS can block Na+ currents. To examine the role of NOS upregulation on the reduction of the three types of Na+ currents in axotomized neurons, axotomized DRG neurons were incubated with 1 mM NG-nitro-L-arginine methyl ester (L-NAME), a NOS inhibitor. The current density of fast and slow Na+ channels in these neurons increased to 0.82 ± 0.08 nA/pF (P < 0.01) and 0.34 ± 0.04 nA/pF (P < 0.05), respectively. However, we did not observe any persistent TTX-R current in axotomized neurons incubated with L-NAME. These results demonstrate that endogenous NO/NO-related species block both fast and slow Na+ current in DRG neurons and suggest that NO functions as an autocrine regulator of Na+ currents in injured DRG neurons.
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INTRODUCTION |
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Nitric oxide (NO) serves as a cellular mediator
for diverse developmental and physiological processes (Bredt and
Snyder 1994a; Moncada and Higgs 1993
) and is a
putative neuromodulator in the brain, spinal cord, and peripheral
nervous system (Holscher 1997
). Recent evidence suggests
that NO may be involved in central (Machelska et al.
1997
; Moore et al. 1991
; Salter et al.
1996
) and peripheral (Haley et al. 1992
;
Holthusen and Arndt 1995
; Ialenti et al.
1992
; Larson and Kitto 1995
; Lawand et
al. 1997
; Thomas et al. 1996
) pain mechanisms.
It is formed by the enzyme NO synthase (NOS), which generates NO from
the guanidine group of arginine, giving rise to NO and stoichiometric
amounts of citrulline. NO is formed by three different types of NOS
derived from three distinct genes referred to, respectively, as
neuronal NOS (nNOS; NOS-1), macrophage or inducible NOS (iNOS;
NOS2), and endothelial NOS (eNOS; NOS3) (Moncada et al.
1997
). nNOS is localized to discrete populations of neurons in
the embryonic and adult nervous system (Bredt and Snyder
1994b
). Among the neuronal systems positive for NOS-like immunoreactivity are spinal sensory (dorsal root ganglion, DRG) neurons, the highest number of which are found at the thoracic level,
with only a few neurons in the lumbar ganglia in displaying NOS-like
immunoreactivity in uninjured subjects (Aimi et al.
1991
).
After transection of the sciatic nerve in rats, there is a marked
increase in the numbers of lumbar DRG neurons containing NOS mRNA, with
the higher number of cells (45% of the total) detected after 7 days of
injury (Verge et al. 1992). Zhang et al.
(1993)
reported that in normal adult rats, nNOS
is expressed in very few (<4%) small- and medium-diameter DRG
neurons. However, a large percentage (~50%) of DRG neurons contain
nNOS after peripheral nerve damage (Zhang et al. 1993
).
These studies did not definitively establish whether the neurons that
show upregulation of NOS activity are the neurons the axons of which
were transected at the level of sciatic nerve. Because only ~54% of
the cells in the L4 and L5
DRGs project an axon into the sciatic nerve (Devor et al.
1985
; Yip et al. 1984
), it is important to
establish whether NOS is upregulated specifically in the axotomized
lumbar ganglia neurons.
Earlier studies from our laboratory established that, after injury to
their axons, C-type DRG neurons display changes in sodium channel
expression that include a downregulation of tetrodotoxin (TTX)-resistant (TTX-R) sodium currents and upregulation of a rapidly
repriming TTX-sensitive (TTX-S) Na+
current (Cummins and Waxman 1997). However,
despite the evidence that axotomy upregulates nNOS in DRG neurons, the
role of NO in the modulation of Na+ currents in
these cells has not been established. NO, when experimentally applied
to tissues via the use of nitric oxide donors, causes reversible
conduction block in both normal and demyelinated axons of the central
and peripheral nervous system (Redford et al. 1997
; Shrager et al. 1998
). In baroreceptor neurons,
endogenous NO as well as exogenously added NO donors have been shown to
inhibit TTX-S and TTX-R Na+ currents (Li
et al. 1998
). These studies suggest that endogenous NO/NO-related activity might alter DRG neuron electrical excitability by modulating Na+ currents.
In this study, we focused on small DRG neurons, which are involved in
nociceptive signaling, and asked whether NOS is upregulated after
axotomy and whether upregulation of NOS blocks
Na+ currents in axotomized neurons. DRG neurons
the axons of which had been transected were identified with a
retrogradely transported fluorescent label, hydroxystilbamine
methanesulfonate. Assay of axotomized C-type DRG neurons for NOS
activity with nNOS specific antibody revealed upregulation of NOS in
60% of cells. Exposure of axotomized type-C DRG neurons to a NOS
inhibitor led to a significantly higher density of fast and slow
Na+ currents, demonstrating that endogenous
NO/NO-related species modulate these Na+ currents
in axotomized C-type DRG neurons and suggesting that NO is an autocrine
regulator of Na+ currents in these cells.
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METHODS |
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Animal care
3-mo old Sprague-Dawley female rats (120 g weight) were used for the study. Animals were fed ad libitum and housed in a pathogen-free area at the Veterans Affairs Medical Center (VAMC), West Haven. Animal care and surgical procedures followed an approved protocol by the Animal Care and Use Committee of Yale University.
Sciatic nerve axotomy
Axotomy of the sciatic nerve was performed as previously
described (Waxman et al. 1994). Animals were
anesthetized with ketamine/xylazine (38/5 mg/kg ip), and the right
sciatic nerve was exposed and a tight ligature was placed around the
sciatic nerve near the sciatic notch proximal to the pyriform ligament.
The nerve was transected immediately distal to the ligature site, and
the proximal nerve stump was fit into a silicone cuff to avoid nerve
regeneration. The cuff contained 4 µl of hydroxystilbamine
methanesulfonate (4% wt/vol in distilled water; Molecular Probes,
Eugene, OR) for retrograde labeling of axotomized neurons. The
fluorescent label clearly identified neurons whose axons were
transected. The incision was sutured and the animal was allowed to
recover. The animals were studied 7-14 days after sciatic nerve ligation.
Culture of DRG neurons
The rats were exsanguinated under ketamine/xylazine anesthesia (38/5 mg/kg ip). The lumbar dorsal root ganglia (L4 and L5) from the right side were used to study axotomy-induced changes and DRG from the left side were used as controls. Briefly, L4 and L5 lumbar DRG were freed from their connective sheaths in sterile calcium-free saline solution. The DRGs then were enzymatically digested for 15 min with collagenase A (1 mg/ml; Boerhinger-Mannheim, Indianapolis, IN) in complete saline solution (CSS) containing 0.5 mM EDTA and 2 mg cysteine, and for 15 min with collagenase D (1 mg/ml; Boerhinger-Mannheim, Indianapolis, IN) and papain (30 units/ml, Worthington Biochemical, Lakewood, NJ) in CSS containing 0.5 mM EDTA and 2 mg cysteine at 37°C. DRGs were removed carefully by Pasteur pipette from the other cellular debris and placed in DRG culture medium (DMEM and F12 in a ratio of 1:1, 10% fetal calf serum, 100 U/ml penicillin, and 0.1 mg/ml streptomycin) containing 1 mg/ml bovine serum albumin (Fraction V, Sigma Chemicals, St. Louis, MO) and 1 mg/ml trypsin inhibitor (Sigma Chemicals). The DRGs were mechanically dispersed (5 strokes up and down) with 1-ml pipette tip and plated on polyornithine and laminin-coated glass coverslips (100 µl suspension per coverslip). Two hours after isolation, the neurons were fed with fresh culture medium. Neurons were placed in a 95% O2-5% CO2 incubator overnight at 37°C and then processed for NOS immunoreactivity or electrophysiological investigation. DRG neurons were studied after short-term culture (16-24 h after isolation). Short-term culture provided cells with truncated (<30 µm) axonal processes that can be voltage-clamped readily and reliably, allowed the cells sufficient time to adhere to the glass coverslips, and was short enough to minimize changes in electrical properties that can occur in long-term cultures.
NOS immunostaining
Coverslips with neurons derived from control or axotomized L4/L5 DRG and maintained in vitro for <24 h were processed for immunocytochemistry as follows: 1) complete saline solution, two times, 1 min each; 2) 4% paraformaldehyde in 0.14 M Sorensen's phosphate buffer, 10 min; 3) PBS, three times, 3 min each; 4) PBS containing 20% normal goat serum, 1% bovine serum albumin, and 0.1% Triton X-100, 15 min; 5) primary antibody (anti-NOS; Transduction Labs, Lexington, KY, 1:250, in blocking solution without Triton X-100), overnight at 4°C; 6) PBS, six times, 5 min each; 7) secondary antibody (goat anti-rabbit IgG-Cy3, 1:3000); and 8) PBS, six times, 5 min each. After the immunocytochemical procedure, the slides were mounted with Aqua-poly-mount and examined with a Leitz Aristoplan light microscope equipped with bright field, Nomarski, and epifluorescence optics. Images were captured with a Dage Dc-330t color camera and Scion CG-7 color PCI frame grabber. Digitized images were processed in Adobe Photoshop with control and axotomized neurons treated in identical manners. Control experiments included incubation without primary antibody; only background levels of fluorescence were detected in the control experiments.
Electrophysiological recordings
Coverslips were mounted in a small flow-through chamber positioned on the stage of a Nikon Diaphot microscope (Nikon) and were perfused continuously with the bath-external solution (see following text) with a push-pull syringe pump (WPI, Saratoga, FL). Hydroxystilbamine-methanesulfonate-labeled neurons were identified by fluorescence emission 510 nm. A short arc mercury lamp was used for the excitation light source and it was defined by broadband (450-490 nm) pass filter (Nikon).
Cells were voltage-clamped via the whole cell configuration of the
patch clamp with an Axopatch-200B amplifier (Axon Instruments, Foster
City, CA) using standard techniques (Hamill et al.
1981). Micropipettes were pulled from borosilicate glasses
(Boralex) with a Flaming Brown micropipette puller (P80, Sutter
Instrument, Novato, CA) and polished by placing them close to a glass
bead on a microforge (Narishige, Tokyo) to obtain electrode resistance ranging from 0.5 to 1.0 M
. To reduce the pipette capacitance, micropipettes were coated with a mixture prepared by mixing
approximately three parts of finely shredded parafilm with one part
each of light and heavy mineral oil (Sigma) and vigorously stirring
over heat for 30-60 min. The pipette solution contained (in mM): 140 CsF, 1 EGTA, 10 NaCl, and 10 HEPES, pH 7.3. The following bath solution
was used (in mM): 140 NaCl, 3 KCl, 1 MgCl2, 1 CaCl2, 0.1 CdCl2, and 20 HEPES, pH 7.3. CdCl2 was used to block
Ca2+ current. The pipette potential was zeroed
before seal formation, and the voltages were not corrected for liquid
junction potential. Capacity transients were cancelled and series
resistance was compensated (>80%) when needed. The leakage current
was digitally subtracted on-line using hyperpolarizing control pulses
of one-sixth test pulse amplitude (
P/6 procedure), and the pulses
were applied before the test pulse. Access resistance was monitored
throughout the recording and the cells were discarded if the resistance
was
5 M
. Cells also were discarded if the leakage current was
>1.0 nA (holding current >1.0 nA at
100 mV). Whole cell currents
were filtered at 5 kHz and acquired at 50 kHz in a computer using
Clampex 8.01 software (Axon Instruments). Digidata 1200B interface
(Axon Instruments) was used for A-D conversion, and the data were
stored on compact disk for analysis. For current density measurements, membrane currents were normalized to membrane capacitance. Membrane capacitance was calculated as the integral of the transient current in
response to a brief hyperpolarizing pulse from
120 mV (holding potential) to
130 mV. The average of the membrane capacitance of the
DRG neurons used for electrophysiological experiments was 23.86 ± 0.89 (n = 106) for control and 32.18 ± 1.54 pF
(n = 126; P < 0.001) for axotomized
DRG neurons.
A Boltzmann function, Availability = 1/{1 + exp[(Vpp Vh)/kh]},
where Vpp is the prepulse potential,
Vh is the midpoint potential, and
kh is the corresponding slope factor
for Boltzmann function, was used to fit inactivation-voltage relationship.
Calculation of fast and slow Na+ current density using prepulse inactivation
Prepulse inactivation takes advantage of the differences in the
inactivation properties of the fast and slow Na+
currents (Cummins and Waxman 1997; MeLean et al.
1988
; Roy and Narahashi 1992
). The currents were
elicited by 20-ms test pulses to
10 mV after 500-ms prepulses to
potentials over the range of
130 to
10 mV. The fast TTX-S
Na+ currents were obtained by subtracting the
current obtained at ~50 mV prepulse (only slow TTX-R
Na+ current) from the current obtained with more
hyperpolarizing prepulses (fast TTX-S and slow TTX-R
Na+ currents). Densities for fast TTX-S
Na+ currents were calculated from cells that
express only TTX-S Na+ currents and from cells
expressing both fast TTX-S and slow TTX-R Na+
currents. Densities for slow TTX-R Na+ currents
were calculated from cells that express only TTX-R
Na+ currents and from cells expressing both fast
TTX-S and slow TTX-R Na+ currents.
NOS inhibitor
NG nitro-L-arginine methyl ester (L-NAME), a NOS-specific inhibitor (Sigma Chemicals) was used to investigate whether the upregulation of NOS in axotomized DRG neurons attenuates Na+ current. DRG neurons were incubated overnight with 1 mM L-NAME in the culture medium. L-NAME also was included in the bath and pipette solutions during the recording of Na+ currents.
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RESULTS |
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In this study on the effects of peripheral nerve injury on the
upregulation of NOS and the associated modulation of
Na+ current, we focused on type-C DRG neurons
(<30-µm diam), which have been well characterized after axotomy
(Cummins and Waxman 1997). We selected DRG neurons from
rats 7-14 days post axotomy (DPA), because a higher number of DRG
neurons display a marked increase in NOS mRNA after 7 days of injury
(Verge et al. 1992
), and a dramatic downregulation of
TTX-R Na+ currents occurs after 6 DPA
(Cummins and Waxman 1997
). Neurons cultured from the
uninjured left L4 and L5
DRG neurons of each rat served as controls (106 cells were studied from
12 different cultures). Neurons cultured from the injured right
L4 and L5 DRG of each rat
were used to study the effects of sciatic nerve transection; 126 axotomized cells from 12 different cultures were studied.
NOS activity is upregulated in axotomized C-type DRG neurons
Figure 1A shows the optical image of a representative group of control DRG neurons, and none of these neurons showed NOS activity when assayed by immunocytochemistry (Fig. 1B). Axotomized lumbar ganglion cells the axons of which were transected at midthigh level of sciatic nerve were identified by retrograde fluorescent dye labeling (Fig. 2A). Upregulation of NOS activity in these axotomized neurons was determined by immunochemical reaction (Fig. 2B). To determine whether NOS activity is specifically upregulated in axotomized neurons, images from retrograde fluorescent dye labeling were overlaid on top of the images of the NOS activity; the superimposed images are shown in Fig. 2C. The retrogradely labeled neurons that show upregulation of NOS appear yellow in color, labeled neurons that do not show upregulation of NOS appear green in color, and uninjured neurons that show upregulation of NOS activity appear red in color. Upregulation of NOS activity was seen in 60% of the retrogradely labeled axotomized neurons (87 of 145 neurons from 2 animals).
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We observed NOS activity in 26% of the unlabeled neurons, which did not show fluorescent signal in cultures derived from axotomized rats (12 of 45 neurons). Only ~60% of L4 and L5 DRG neurons project into the sciatic nerve; ~40% contribute to more proximal nerves and are uninjured after sciatic nerve transection. Unlabeled NOS-positive neurons could have been injured but failed to take up the fluorescent dye. Alternatively, NOS might be upregulated in nonaxotomized neurons by a diffusible factor produced in nearby axotomized neurons.
Na+ currents in type-C DRG neurons
Sodium currents were recorded from type-C (<30 µm)
L4 and L5 DRG neurons with
whole cell patch-clamp techniques. Only fluorescent neurons, which we
could identify definitively as axotomized neurons, were studied. The
Na+ currents in control neurons, in cultures from
nonaxotomized DRG, were similar to those previously described in small
DRG neurons (Caffrey et al. 1992; Cummins and
Waxman 1997
; Cummins et al. 1999
; Elliott
and Elliott 1993
; Rizzo et al. 1994
). On the
basis of their inactivation properties, Na+
currents in type-C DRG neurons can be classified into
fast-inactivating, slowly inactivating, and noninactivating
Na+ currents (Cummins and Waxman
1997
; Cummins et al. 1999
), which we refer to
"fast," "slow," and "persistent" Na+
currents, respectively. The fast-inactivating currents in both control
and axotomized DRG neurons were sensitive to nanomolar concentrations
of TTX, whereas the slowly inactivating and the persistent
Na+ currents were resistant to TTX (see following
text and Fig. 6). The observation that fast-inactivating
Na+ currents are sensitive and the
slow-inactivating and persistent Na+ currents are
resistant to nanomolar concentrations of TTX is consistent with earlier
studies (Cummins and Waxman 1997
; Cummins et al.
1999
; Elliott and Elliott 1993
; Roy and
Narahashi 1992
). Prepulse inactivation was used to separate the
fast and slow Na+ currents (Cummins and
Waxman 1997
; MeLean et al. 1988
; Roy and Narahashi 1992
). Prepulse inactivation takes advantage of the differences in the inactivation properties of the fast and slow Na+ currents and is simpler than TTX subtraction.
TTX subtraction and prepulse inactivation have been shown to give
essentially the same results when used to separate fast and slow
Na+ currents (Cummins and Waxman
1997
). It has been demonstrated, using SNS-null mutant mice
that the slow Na+ current in small DRG neurons is
encoded by the SNS sodium channel isoform (Akopian et al.
1999
). However, SNS-null and wild-type DRG neurons also express
a distinct, non-SNS, TTX-R Na+ current
(Cummins et al. 1999
). We refer to this distinct TTX-R Na+ current as the persistent current to
differentiate it from the slow TTX-R Na+ current.
Two criteria were used to separate persistent TTX-R Na+ currents from the fast and slow
Na+ currents. Persistent currents activate at low
threshold depolarizing potentials (about
80 mV) compared with the
fast and slow Na+ currents, which activate at
higher depolarizing potentials (about
40 mV); and at depolarizing
potentials between
80 to
40 mV the persistent currents are
noninactivating (see following text and Fig. 6 and 7).
Fast and slow Na+ current density are reduced in axotomized neurons
To quantitate fast and slow Na+ currents
expressed in each neuron, prepulse inactivation (MeLean et al.
1988; Roy and Narahashi 1992
) was used, and the
current amplitudes were measured. The fast and slow current amplitudes
were 25.40 ± 2.61 and 26.17 ± 2.83 in control neurons and
19.77 ± 3.81 (P < 0.04) and 6.41 ± 1.39 (P < 0.0001) in axotomized neurons. To compensate for
differences in cell size, currents were normalized to cell capacitance
and expressed as current density. The fast TTX-S and slow TTX-R
Na+ current densities in control neurons were
1.08 ± 0.08 and 1.03 ± 0.10 nA/pF, respectively. Axotomy
significantly reduced the fast and slow Na+
current densities to 0.58 ± 0.05 (P < 0.01) and
0.20 ± 0.05 nA/pF (P < 0.001), respectively (Fig.
3, A and B). The
cell capacitance in axotomized cells was 32.06 ± 1.43 pF, 32%
higher than in control cells where it was 23.86 ± 0.89 pF (Fig.
3C, P < 0.01). An increase in capacitance of
axotomized neurons is expected because axotomized neurons sprout
neurites more rapidly than control neurons in culture (Lankford
et al. 1998
). The increase in cell capacitance (32%), which
reflects increased cell membrane area in axotomized neurons, may be
responsible for part of the observed reduction in fast Na+ current density (which was reduced by 46% in
axotomized cells compared with controls) and slow
Na+ current density (which was reduced by 81% in
axotomized cells compared with controls).
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L-NAME increases fast and slow Na+ current densities in axotomized neurons
To investigate whether upregulation of NOS in axotomized DRG neurons attenuates Na+ currents, DRG neurons were incubated with 1 mM L-NAME, a NOS inhibitor, overnight in the culture medium. Although, acute application of NOS inhibitor to DRG cells via bath and pipette solutions slightly increased fast TTX-S and slow TTX-R Na+ current density, it was twofold lesser than the increase observed in overnight incubation of cultured DRG cells with L-NAME and including L-NAME in bath and pipette solutions. Incubation of control DRG neurons with L-NAME did not increase the density of fast or slow Na+ currents (Fig. 3, A and B); in these neurons the fast or slow Na+ current densities were 1.02 ± 0.15 and 0.81 ± 0.08 nA/pF (P > 0.05), respectively. In contrast, incubation of axotomized neurons with L-NAME significantly increased both fast and slow Na+ current density to 0.82 ± 0.08 (P < 0.01) and 0.34 ± 0.04 nA/pF (P < 0.05), respectively (Fig. 3, A and B) and the respective current amplitudes to 25.96 ± 2.86 (P < 0.04) and 11.20 ± 1.25 (P < 0.03). The capacitance of axotomized neurons incubated with L-NAME was 31.77 ± 1.87 pF, similar to the capacitance of untreated axotomized neurons (Fig. 3C). Taken together these results suggest that upregulation of NOS results in block of part of the fast and slow Na+ currents in axotomized but not in control neurons.
L-NAME increases the number of cells coexpressing fast and slow Na+ currents in axotomized neurons
The proportion of cells expressing fast and slow
Na+ currents in control cultures were ~85 and
70%, respectively, with 60% of control cells coexpressing fast and
slow Na+ currents. Axotomy reduced the number of
neurons expressing slow Na+ currents to ~20%
and increased the number of neurons expressing fast
Na+ currents to >90% with 21% of cells
coexpressing fast and slow Na+ currents (Table
1). These results are consistent with an
earlier study, which demonstrated that peripheral nerve injury
downregulates slow Na+ currents and upregulates
fast Na+ currents in type-C neurons
(Cummins and Waxman 1997). The incubation of axotomized
neurons with L-NAME increased the proportion of axotomized
cells coexpressing fast and slow Na+ currents to
~40%. However, incubation of control neurons with L-NAME
did not significantly increase the number of cells coexpressing fast
TTX-S and slow TTX-R Na+ currents (Table 1). The
increase in the proportion of axotomized neurons expressing slow
Na+ channels after exposure to L-NAME
provides additional evidence that upregulation of NOS in axotomized
neurons blocks Na+ currents.
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Voltage dependence of steady-state inactivation of Na+ currents is not altered by axotomy
A majority of the control neurons coexpress fast and slow
Na+ currents (Table 1; Fig.
4A), and the voltage
dependence of inactivation of the Na+ currents in
these neurons showed a bimodal distribution because of the different
inactivation properties of the fast and slow Na+
currents (Fig. 4B). The midpoint of steady-state
inactivation for fast and slow currents in control neurons was about
74.6 ± 7.4 and
33.7 ± 6.5 mV, respectively (Table
2). These values are similar to the
values reported in earlier studies (Caffrey et al. 1992
;
Cummins and Waxman 1997
; Elliott and Elliott
1993
; Rizzo et al. 1994
; Roy and
Narahashi 1992
; Rush et al. 1998
). Axotomy
reduced the amplitude of slow Na+ currents (note
the absence of slow-inactivating current in Fig. 4D). The
voltage dependence of inactivation of the Na+
currents in most axotomized neurons could be fit with a single Boltzmann distribution indicative of the presence of only one type of
Na+ current, i.e., fast Na+
current (Fig. 4E). However, slow Na+
current was observed in 25% of the axotomized neurons (Table 1).
Despite the small size of these currents (less than
5 nA in most
cells) we were able to measure their inactivation to the nearest 0.5 mV. The midpoint of steady-state inactivation for fast and slow
currents in these neurons was
70.0 ± 6.8 and
34.0 ± 5.2 mV, respectively, similar to the values seen in control neurons (Table
2). These values are similar to the values reported in an earlier study
for axotomized type-C neurons (Cummins and Waxman 1997
).
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Voltage dependence of steady-state inactivation of Na+ currents is not altered by L-NAME
After incubation with L-NAME, 40% of the axotomized
neurons showed a bimodal distribution for the voltage dependence of
inactivation, indicating the presence of both fast and slow
Na+ currents in these neurons (Table 1 and Fig.
5, A and B). The remaining 60% of the neurons had midpoint potentials similar to those
of fast Na+ currents (Fig. 5E). The
midpoint of steady-state voltage-dependent inactivation for fast and
slow Na+ currents in axotomized neurons incubated
with L-NAME was 73.3 ± 7.6 and
34.4 ± 4, respectively (Table 2). The voltage dependence of inactivation for
control neurons incubated with L-NAME was
72.7 ± 5.5 and
33.0 ± 6.1 mV, respectively, for fast and slow Na+ currents (Table 2). The small differences in
the midpoints of steady-state voltage-dependent inactivation for fast
and slow Na+ currents in control, axotomized,
L-NAME-incubated axotomized neurons were not statistically
significant, indicating that exposure to NOS does not alter
steady-state inactivation. These results, therefore suggest that the
same populations of fast and slow Na+ channels
are subject to potential modulation by NOS in control and axotomized
DRG neurons.
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Recovery from inactivation
We previously have shown that axotomy of small DRG neurons is
followed by the emergence of a distinct fast TTX-S
Na+ current that shows a rapid recovery from
inactivation (Cummins and Waxman 1997). Therefore we
examined the recovery from inactivation at
100 mV of the fast
Na+ channels in control, axotomized, and
L-NAME-treated control and axotomized neurons.
Recovery-from-inactivation curves for typical control and axotomized
neurons are shown in Fig. 4, C and F. Control neurons treated with L-NAME displayed recovery from
inactivation that was similar to that of untreated control neurons. The
time course of recovery from inactivation in control neurons could be
fit with two exponentials. The rapidly recovering component showed
complete recovery within 10 ms and the slowly recovering component
showed recovery in 100 ms (Fig. 4C). TTX was used to determine that the rapidly recovering component corresponds to slow
TTX-R Na+ current and the slowly recovering
component to a fast TTX-S Na+ current (results
not shown). These results are similar to the results reported earlier
(Cummins and Waxman 1997
; Elliott and Elliott
1993
). Eighty percent of the axotomized neurons expressed only
fast TTX-S Na+ current, and in these neurons,
>90% of the recovery from inactivation was seen in ~10 ms (Fig.
4F). Because these neurons had only TTX-S Na+ current, the results suggest that the
recovery from inactivation of TTX-S Na+ current
in axotomized neurons is rapid and faster than in the uninjured
neurons, consistent with the results obtained in an earlier report
(Cummins and Waxman 1997
). Twenty percent of the axotomized neurons expressed both fast TTX-S and slow TTX-R
Na+ currents, and these currents recovered from
inactivation in ~10 ms.
Both fast TTX-S and slow TTX-R Na+ current were seen in 40% of L-NAME-incubated axotomized neurons, and complete recovery from inactivation also was seen in ~10 ms in these neurons (Fig. 5C). The other 60% of the axotomized neurons incubated with L-NAME, which expressed only the fast TTX-S Na+ currents, also showed >85% recovery from inactivation in ~10 ms (Fig. 5F). The rapid recovery from inactivation of the fast TTX-S currents in axotomized neurons compared with control neurons suggests that fast TTX-S Na+ current in axotomized neurons may arise from a different type of Na+ channel that is expressed in these cells. The similar time course for recovery of the Na+ currents from axotomized neurons incubated with L-NAME suggests that the channels, which underlie these currents, are similar to those in untreated axotomized neurons. L-NAME thus appears to have little effect on recovery from inactivation of the fast TTX-S and slow TTX-R channels.
Persistent TTX-R Na+ currents are decreased after axotomy
As described by Cummins et al. (1999) in wild-type
and SNS-null mutant mice, when control DRG neurons were held at
130
mV, persistent TTX-R currents (defined as the current remaining at the
end of
40 ms depolarizing pulses) were seen in 24 of 40 cells, with
current amplitudes ranging from about
40 to about
5 nA. The
amplitude of the persistent current in control DRG neurons was 10-50%
of the amplitude of fast TTX-S Na+ current and
50-100% of the amplitude of slow TTX-R Na+
current (Fig. 6, A and
B). In these experiments, the bath solution contained 100 µM Cd2+, a concentration high enough to block
low-voltage-activated T-type Ca2+ currents.
Figure 6A shows the presence of fast TTX-S
Na+ current and slow TTX-R
Na+ current together with a persistent current
that is present at the end of 40-ms test pulses; the persistent
currents manifest as tail currents on returning to a holding potential
of
130 mV. Even when measured at the end of 100 ms and in the
presence of 300 nM TTX, these persistent currents can be seen (Fig.
6B). Activation of fast TTX-S Na+
currents and slow TTX-R Na+ currents occurs at
about
40 mV (Fig. 7, B and
C) consistent with the results obtained in earlier studies
(Elliott and Elliott 1993
; Rush et al.
1998
). In contrast, activation of the persistent Na+ currents is seen at about
80 mV (Fig.
7D) similar to the TTX-R persistent current observed by
Cummins et al. (1999)
in wild-type and SNS-null mutant
mice. Further, at test potentials between
80 and
40 mV, the
persistent currents are noninactivating, similar to the observation by
Cummins et al. (1999)
in wild-type and SNS-null mutant
mice. Therefore the Na+ currents observed between
80 and
40 mV are TTX-R persistent currents.
|
|
The peak current amplitude of the current traces shown in Fig. 6,
A and B, was measured and depicted as
current-voltage plot in Fig. 8,
A and B (). Activation of inward
Na+ current is seen at
80 mV, reaches a plateau
or peak at
40 mV (note the hump at
40 mV), with a peak at
15 mV.
Axotomy almost completely abolished the persistent current as seen in
the traces in Fig. 6, C and D (note the absence
of tail currents), and in the loss of amplitude of the persistent
current at test potentials between
80 and
40 mV (Fig. 8,
A and B,
). Very few cells (<10% of the
cells) showed persistent current after axotomy, and the maximum
amplitude of the persistent current seen in these cells was <5 nA
(Fig. 6D). These results are similar to the results obtained
in an earlier study (Cummins and Waxman 1997
). After incubation with L-NAME, the axotomized neurons show an
increase in fast and slow Na+ current amplitude
but did not show any significant change in the amplitude of the
persistent current (Fig. 6, E and F). This also
can be seen in Fig. 8, A and B (
), where
incubation in L-NAME results in an increase in the currents
activated positive to
40 mV, but not in the persistent currents
activated between
80 and
40 mV. These results suggest that the
decrease in the persistent Na+ current amplitude
in axotomized neurons is not due to the upregulation of NOS (see
DISCUSSION).
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DISCUSSION |
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The present study reports the first evidence that upregulation of NOS is seen predominantly in axotomized C-type neurons and supports the hypothesis that NO derived from NOS is a modulator of Na+ currents in these cells after axotomy. This conclusion is supported by our findings that the NOS inhibitor L-NAME increased Na+ current density of both fast TTX-S and slow TTX-R Na+ current in axotomized C-type DRG neurons but not in control nonaxotomized neurons. These results indicate a role for NO in suppressing fast and slow Na+ currents in damaged neurons at the ganglionic level after axotomy and suggest that NO may be an autocrine regulator of Na+ currents in C-type DRG neurons.
Our observation that NOS activity was increased in the ipsilateral DRG
at the level of nerve injury is consistent with previous observations
suggesting a peripheral role for NO in altering the excitability of
injured DRG neurons (Choi et al. 1996; Ferreira et al. 1991
; Fiallos-Estrada et al. 1993
;
Steel et al. 1994
; Verge et al. 1992
).
Using fluorescent dye to identify the axotomized neuron and
nNOS-specific antibody to identify NOS activity, we demonstrate that
~60% of retrogradely labeled C-type neurons within L4 and L5 DRG show
upregulation of NOS activity after axotomy within the sciatic nerve.
Axotomy results in dramatic and complex changes in the sodium
currents expressed in C-type DRG neurons (Cummins and Waxman 1997). Our observation that axotomy decreased the
density of slow Na+ current density to 0.2 nA/pF
in C-type DRG neurons is consistent with the previous observations that
axotomy triggers decreases in slow Na+ current
density (Cummins and Waxman 1997
) and in the
steady-state levels of alpha-SNS mRNA, which encodes a slow TTX-R
current in these cells (Dib-Hajj et al. 1996
). The
present results are also consistent with earlier studies that showed
loss in axotomized neurons of TTX-R-persistent currents (Cummins
and Waxman 1997
) and of the NaN transcript (Dib-Hajj et
al. 1998
), which appears to encode the channels that produce
this current (Cummins et al. 1999
). However, our
observation that axotomy reduced fast TTX-S Na+
current density to 0.6 nA/pF differs from an earlier study
(Cummins and Waxman 1997
), which did not report a
significant reduction in TTX-S Na+ current
density after axotomy. This difference may be attributed to the
difference in cell capacitance of axotomized neurons observed between
this study and the earlier study (Cummins and Waxman
1997
). We examined DRG neurons 16-24 h after dissociation and
observed a 30% increase in cell capacitance after axotomy; however,
the earlier study, which examined DRG neurons 10-16 h after
dissociation, did not find a significant change in cell capacitance
after axotomy (Cummins and Waxman 1997
). It recently has
been shown that sciatic nerve ligation significantly can enhance
neurite growth in isolated DRG neurons (Lankford et al.
1998
). This enhancement probably accounts for the increased
capacitance and decreased TTX-S current density observed in axotomized
neurons studied at 16-24 h in vitro (this study) but not at 10-16 h
in vitro after axotomy (Cummins and Waxman 1997
). In
this study, DRG neurons were studied 16-24 h after dissociation to
give the neurons a longer time to better withstand the repeated
washings involved in the immunocytochemical measurement of NOS upregulation.
Axotomized neurons incubated with L-NAME, a specific
inhibitor of NOS, had increased fast and the slow
Na+ current densities compared with untreated
axotomized neurons. NO may block Na+ currents
through generation of cGMP (Moncada 1999) and/or
through mechanisms independent of cGMP, i.e., via direct covalent
interaction with susceptible thiol groups to form
S-nitrosothiols that modify the protein function (Jia
et al. 1996
). Li et al. (1998)
have reported
that NO blocks Na+ currents in nodose ganglia via
nitrosylation rather than through the generation of cGMP. However,
whether the NO-mediated Na+ current block in
small type C neurons is via cGMP and/or nitrosylation has not been determined.
The much smaller effect, if any, on the persistent TTX-R
Na+ current on L-NAME incubation
suggests that the channels that produce persistent current are present
at very low levels in axotomized neurons. NaN transcript levels that
are likely to encode persistent TTX-R channels (Cummins et al.
1999) are reduced significantly 7 days post axotomy in DRG
neurons (Dib-Hajj et al. 1998
; Tate et al.
1998
). In our experiments, axotomized neurons were incubated with L-NAME for 12 h before the observed increase in
fast TTX-S and slow TTX-R Na+ current density, a
time period probably not sufficient enough to modulate the
transcription and translation of channels. These results suggest that
gene regulation rather than modulation of channels by NOS may account
for most of the decrease observed in persistent TTX-R
Na+ current in axotomized DRG neurons.
Consistent with a role of NOS in modulating the excitability of
axotomized DRG neurons, Wiesenfeld-Hallin et al. (1993)
have shown that NOS inhibitors suppress the ongoing discharges in DRG neurons after sciatic nerve injury. Our results show that exposure of
axotomized neurons to L-NAME increased the number of cells expressing both fast Na+ current and slow
Na+ current to 40% (from 21% in untreated
cells) and increased the current density of fast and slow
Na+ current density by 37 and 70%, respectively.
These results suggest that NOS upregulation has effects on both fast
TTX-S and slow TTX-R Na+ currents, possibly with
a larger effect on slow Na+ current. At first
sight, it might be expected that the downmodulation of both fast and
slow Na+ current by NOS upregulation would
decrease rather than augment DRG hyperexcitability. However, threshold
and excitability of DRG neurons appear to be complex functions and
depend on the densities of various types of Na+
channels (Cummins and Waxman 1997
; Elliot
1997
; Schild and Kunze 1997
), possibly with
optima at specific densities, above and below which excitability is
altered. In SNS-null mutant mice, which do not express slow TTX-R
Na+ channels but express fast TTX-S
Na+ channels, compound action potentials recorded
from the L4 dorsal roots had a lower electrical
threshold and the recruitment curve was shifted significantly to the
left (Akopian et al. 1999
). TTX-R Na+ currents paradoxically might play a role in
limiting the excitability of sensory neurons. The downmodulation of
slow Na+ current to a larger extent than the fast
Na+ current presents a possible scenario where
the fast Na+ current, which has a rapid recovery
from inactivation in axotomized DRG neurons (Cummins and Waxman
1997
), might play a major role in determining the excitability
of the axotomized neuron. Alternatively, NO also might affect
excitability by altering K+ conductance.
In conclusion, the present results demonstrate NOS-related changes in fast and slow Na+ channel function in axotomized but not in normal C-type DRG neurons. These findings suggest that activity of endogenous NO/NO-related species due to the upregulation of NOS could block both fast and slow Na+ currents in axotomized DRG neurons. NOS thus appears to play an autocrine role in regulating the Na+ currents of small C-type DRG neurons after injury.
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ACKNOWLEDGMENTS |
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We thank the Eastern Paralyzed Veterans Association and the Paralyzed Veterans of America for support. We thank B. Tuftness for help with the figures and Dr. Sulayman Dib-Hajj and M. Baccei for helpful discussions.
This work was supported by grants from the Medical Research Service and Rehabilitation Research Service, Department of Veterans Affairs and from the National Multiple Sclerosis Society.
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FOOTNOTES |
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Address for reprint requests: S. G. Waxman, Yale University School of Medicine, Dept. of Neurology, 707 LCI, 333 Cedar St., New Haven, CT 06510.
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. Section 1734 solely to indicate this fact.
Received 10 November 1999; accepted in final form 30 December 1999.
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REFERENCES |
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