6
Biochem. Soc. Symp. 71, 65–83
(Printed in Great Britain)
© 2004 The Biochemical Society
Nitric oxide signalling: insect
brains and photocytes
Barry A.Trimmer*1, June Aprille† and
Josephine Modica-Napolitano‡
*Department of Biology, Tufts University, Medford, MA 02155, U.S.A.,
†Department of Biology, University of Richmond, Richmond, VA 23173, U.S.A., and
‡Department of Biology, Merrimack College, North Andover, MA 01845, U.S.A.
Abstract
The success of insects arises partly from extraordinary biochemical and
physiological specializations. For example, most species lack glutathione peroxidase, glutathione reductase and respiratory-gas transport proteins and thus
allow oxygen to diffuse directly into cells. To counter the increased potential
for oxidative damage, insect tissues rely on the indirect protection of the
thioredoxin reductase pathway to maintain redox homoeostasis. Such specializations must impact on the control of reactive oxygen species and free radicals
such as the signalling molecule NO. This chapter focuses on NO signalling in
the insect central nervous system and in the light-producing lantern of the firefly. It is shown that neural NO production is coupled to both muscarinic and
nicotinic acetylcholine receptors. The NO-mediated increase in cGMP evokes
changes in spike activity of neurons controlling the gut and body wall musculature. In addition, maps of NO-producing and -responsive neurons make
insects useful models for establishing the range and specificity of NO’s actions
in the central nervous system. The firefly lantern also provides insight into the
interplay of tissue anatomy and cellular biochemistry in NO signalling. In the
lantern, nitric oxide synthase is expressed in tracheal end cells that are interposed between neuron terminals and photocytes. Exogenous NO can activate
light production and NO scavengers block evoked flashes. NO inhibits respiration in isolated lantern mitochondria and this can be reversed by bright light.
It is proposed that NO controls flashes by transiently inhibiting oxygen consumption and permitting direct oxidation of activated luciferin. It is possible
that light production itself contributes to the restoration of mitochondrial
activity and consequent cessation of the flash.
1To
whom correspondence should be addressed (e-mail [email protected]).
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Introduction
Since its discovery as a signalling molecule in the vertebrate circulatory
system, NO has been shown to have important roles in other tissues throughout the Animal Kingdom. This review discusses some of these functions and
concentrates on NO signalling in two insect tissues, the CNS (central nervous
system) of the tobacco-feeding caterpillar Manduca sexta, and the light-producing lantern of the firefly, Photuris sp. These two systems provide insights
into how tissue anatomy, cellular biochemistry and the physical properties of
NO interact to create a diverse and complex signalling system.
NO as a neural signalling molecule
Because NO is synthesized by NOS (nitric oxide synthase) de novo and
can rapidly cross cytoplasmic and membrane compartments, neural NO
release is not confined to synaptic zones [1]. This high diffusibility also
makes NO a potentially far-reaching and rapid-acting transmitter [2].
Calculations of the unrestricted NO diffusion rate in lipid layers show that
NO can travel 200 m in 0.2 ms [3], but the effective range of NO is thought
to be 100 m over several seconds [1,4,5]. For many small insects this diffusion scale would permit NO to signal throughout the entire CNS. The basis
for restricted influence presumably includes the limited expression and sensitivity of effector proteins and the short lifetime of NO due to its chemical
reactivity. NO has a rich chemistry [6] and its biological inactivation is complex; it includes reactions with transition metals in metalloproteins and thiols
[7], and rapid exchanges with molecular oxygen through unstable intermediates to produce the nitrite anion [8,9]. Hence, in the absence of
high-resolution and rapid NO detection in the CNS, much of our understanding about NO’s range is based on mathematical diffusion models
[10–12] in blood vessels [13,14], or well-defined regions such as the mushroom bodies of the insect brain [15] and other idealized neural compartments
[16]. However, these models depend critically on the size and shape of the
release space and usually assume relatively simple and defined interactions
with reactive oxygen species and antioxidants such as GSH or NO-binding
proteins such as haemoglobin.
In fact, insect cells derive their oxygen by tracheal diffusion and most
insects lack oxygen-carrying proteins. This direct penetration of oxygen into
tissues could subject them to chronic oxidative stress [17]. Furthermore,
insects lack glutathione peroxidase and glutathione reductase [18], which are
essential enzymes for protection against reactive oxygen species in bacteria,
fungi, nematode worms and mammals [18,19]. It appears that insects can
maintain sufficient GSH in cells using an indirect thioredoxin reductase system [18,20] (Figure 1). Insects express many of the other well-described
redox protective enzymes including mitochondrial MnSOD (manganesedependent superoxide dismutase) and cytosolic/nuclear CuZnSOD (copper-/
zinc-dependent SOD) [17,21], catalase [22], ascorbate peroxidase [23],
glutathione S-transferase [24], iron-regulating transferrin and ferritin [25],
and the peridoxins [20]. The expression of these enzymes has been studied
© 2004 The Biochemical Society
Nitric oxide signalling: insect brains and photocytes
67
Perineural sheath
P-glycoprotein
Cytochrome P450dependent PSMO
Sensory neurons
ACh
Interneurons
NO
Muscle
ACh
VUM
ACh
NO producers
Motoneurons
Periphery
CNS
Neuropile
Trx, SOD, CAT,
GST, APOX,
transferrin, ferritin?
Figure 1 Factors involved in NO/cGMP signalling in the insect nervous system The primary afferent neurotransmitter ACh is released centrally
by sensory stimulation. Many motoneurons and interneurons respond to ACh
through both ionotropic nicotinic receptors and metabotropic muscarinic
receptors. These receptors are coupled to the production of NO by NOS in
cells such as the ventral unpaired median neurons (VUM). The activity of
interneurons such as IN 703, and various motoneurons, is then altered by NOdependent activation of soluble guanylate cyclase. These changes in turn
control motor activity of the body wall and gut. The distance over which NO
acts in the CNS is presently unknown but is certainly influenced by oxidation
protection enzymes such as thioredoxin reductase (Trx), superoxide dismutase
(SOD), catalase (CAT), glutathione S-transferase (GST), ascorbate peroxidase
(APOX), transferrin and ferritin. In insects, these redox pathways are expected
to be critical in preventing damage caused by the direct supply of oxygen
through tracheal air tubes. Furthemore, P-glycoprotein transporters and
cytochrome P450-dependent polysubstrate mono-oxygenase (PSMO) in the
perineural sheath help protect against systemic radicals and xenobiotic toxins.
See the main text for details and references.
extensively in insects with particular emphasis on their roles in processing
ingested toxins [26] and their importance in aging [27]. However, with the
exception of SOD [28] and glutathione S-transferase [24], the function of
these enzymes in the insect CNS has not been studied directly. Therefore,
while idealized models of NO diffusion are helpful in defining the potential
range of neural NO signalling, they are unlikely to describe the actual duration or distance over which NO normally signals in insects.
© 2004 The Biochemical Society
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
NO release
The production of NO by neuronal NOS is regulated by Ca2 in both
vertebrates [29,30] and invertebrates [31–35], and includes Ca2 entry through
ligand-gated channels [1,36] and voltage-gated channels [37–39], and by InsP3mediated Ca2 release [36,40–44]. Ca2 entry through NMDA (N-methyl
D-aspartate) receptors is particularly effective because mammalian NOS is
anchored to the receptor through post-synaptic density proteins containing the
PDZ motif [45]. In insects, ACh (acetylcholine) performs many of the roles of
glutamate in the vertebrate CNS; it is the primary afferent transmitter and the
predominant excitatory signal [46] (Figure 1). ACh can activate NO production
through both mAChRs (muscarinic ACh receptors) [47,48] and ionotropic
nAChRs (nicotinic ACh receptors) [32,49,50]. Despite this close functional
association, none of the cloned insect NOS sequences contain PDZ domains
[34,35,51]. If NOS and nAChRs are physically associated it must occur through
an alternative protein–protein or lipid-binding site [52]. Despite detailed information on the interaction of Ca2 with NOS [53], and evidence that NO release
depends on firing frequency [54], relatively little is known about the relationship between NO production and cellular activity in any nervous system.
Cellular actions of NO/cGMP
The best-described target of NO is the activation of sGC (soluble guanylate cyclase) to increase cGMP production [55–61] but several other NO
targets have been demonstrated in the CNS. Hence, the inhibition of NMDAtype glutamate receptors [62], and the slow inactivation of Na channels in
crayfish axons [63,64] and sciatic and vagus nerves [65] are mediated by direct
generation of S-nitrosothiols and subsequent formation of disulphide bonds.
Other NO effects include acute inhibition of mitochondrial function (see
below; [66]), the regulation of enzymes in the ADP-ribose pathway [67,68], the
inhibition of L-type Ca2 channels in the carotid body [69] and the direct activation of Ca2-dependent K channels [70]. Some of these effects might be
pathological since they have only been demonstrated at relatively high NO
concentrations (1–10 M), such as those expected during inflammation reactions (see calculations in [65]) or following brain ischaemia [71]. Despite the
specialized biochemical protections that insects employ against oxidative stress
[18], and their probable importance in insecticide resistance [72,73], most of
these alternative mechanisms have not been examined in insects.
Neural function of NO/cGMP
In addition to its role in the developing CNS of vertebrates and invertebrates [16,74–77], NO is implicated in synaptic modulation such as long-term
potentiation [39,78–80], long-term depression [81,82], the regulation of neural
activity [83,84] and the modulation of NMDA receptors by modifying critical
cysteine residues [62]. In molluscs, NO can evoke excitatory post-synaptic
potentials (e.g. in the pond snail Lymnaea [85] and Aplysia [86]), and in
Drosophila it can increase vesicle release from motoneurons [87]. NO can also
activate or augment specific conductances in crustacean muscle [88,89].
© 2004 The Biochemical Society
Nitric oxide signalling: insect brains and photocytes
69
Several behavioural functions of NO have been resolved in invertebrate
model systems. In molluscs NO signalling is implicated in feeding [31,90–92],
appetitive classical conditioning. [93], long-term memory formation (Octopus
vulgaris [94]) and rotational behaviour [95]. In insects, NO affects Drosophila’s
responses to hypoxia [96] and Malpighian tubule secretion [97,98], eclosion in
Bombyx [99], flash control in fireflies [100], and appetitive habituation [101]
and the transfer of short- to long-term memory in bees [102]. From the expression patterns of NOS and sGC in the CNS it has also been inferred that NO is
involved in olfaction and vision (for a review see [103]).
In keeping with its toxic actions in the immune system [104], large
amounts of NO generated in the mammalian CNS by inflammation and
ischaemia can be very damaging to neurons [30]. NO is also implicated in
insect immune responses [105,106] but there have been no systematic reports of
its central toxic effects in the insect CNS. This could be a very interesting area
of study because insects are increasingly important as models for understanding the effects of reactive oxygen species in chemical resistance [18,20,107] and
aging [23,108,109].
NO/cGMP signalling in Manduca
Mapping the NO/cGMP transmitter system
NO-producing neurons
Using NADPH-diaphorase staining and an antibody directed against a
highly conserved region of NOS, several NOS-containing neurons have been
identified in the caterpillar Manduca [110]. These include the ventral unpaired
midline neurons (Figure 1), three pairs of lateral neurons (possibly cells 24, 25
and 26) [111,112] and a pair of dorsal midline neurons (PM2s) in each of ganglia
A5, A6 and A7 [112]. The strongly staining neurons express NOS throughout
the cell body, dendrites and axons. The ventral unpaired midline neurons are
well-described octopaminergic cells that project to internal longitudinal muscles where they modulate muscle contraction and metabolism [113–116]. The
PM2 neurons are immunoreactive for allatostatin in some segments and project
through the proctodeal nerve to innervate posterior portions of the midgut
[112]. Other NOS-containing neurons are found in the frontal and suboesophageal ganglia, suggesting functions in gut control or feeding [110].
NO-receptive neurons
Some potentially NO-sensitive neurons have been mapped by staining
the nervous system with antibodies to cGMP after it has been exposed to NO
donors (Figure 2) [110,117–119]. These include a pair of interganglionic
interneurons (IN 505) in each abdominal ganglion, interneurons 702 or 703 in
the ganglial anterior to A3 [120,121] and several neurons that stain in a segment-specific manner such as the PM2 neurons in ganglia A5 and A7. These
particular neurons are therefore potential NO producers and responders. The
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
DNL
recording
A3
Muscle
recording
IN 703
MN1
DIM1,2
MN12
DEM
DEO
A4
Proleg
MNs
IN 505
DNL
recording
VNL recording
DIM
Figure 2 Identified NO-sensitive neurons in the ventral nerve cord of
M. sexta and configurations used to monitor their activity. Two ganglia
are shown from abdominal segments 3 and 4 (A3, A4). Motoneuron 1 (MN1)
has an axon in the contralateral dorsal nerve of the same segment.
Motoneuron 12 (MN12) and a pair of dorsal internal medial neurons (DIM1,2)
project into the connective and exit via the lateral branch of the dorsal nerve
(DNL) in the next posterior ganglion. MN12 innervates both the dorsal external medial (DEM) and the dorsal external oblique (DEO) muscles; DIM
innervates the dorsal internal muscle (DIM). The proleg motoneurons project
into the lateral branch of the ventral nerve (VNL). The targets of interneurons
(IN) 505 and 703 are not known.
PM2 neurons in ganglion A6 stain weakly if at all, suggesting a segmental difference in the responsiveness of PM2 neurons to NO.
In addition to neurons that accumulate cGMP in the presence of NO
donors many more NO-responsive neurons are detected using cGMP phosphodiesterase inhibitors such as zaprinast and sildenifil. These neurons accumulate
cGMP (as detected by immuno-staining) when they are exposed to NO donors,
even when action potentials are blocked using tetrodotoxin or low-sodium
saline, suggesting that they are direct targets for NO. They include some of the
motoneurons (MN1 and MN12) that control the body wall muscles [122].
© 2004 The Biochemical Society
Nitric oxide signalling: insect brains and photocytes
71
GCs and the effect of cGMP
Changes in cGMP levels in the CNS of Manduca are important during
metamorphosis and ecdysis (moulting or cuticle shedding). Thus increased
cGMP levels in specific cells activate motor patterns [123,124], co-ordinate
hormone cascades [118,125–129] and are associated with maturation and reconfiguration of the nervous system [76,117,130,131]. Although many of these
changes are NO-independent, cGMP changes associated with post-migratory
development of enteric neurons [132] and synaptogenesis [76] do appear to
involve NO. It has also been shown that some motoneuron axons and terminals produce large amounts of cGMP if stimulated by NO donors in the
presence of phosphodiesterase inhibitors [133]. Our own work demonstrates
that cholinergic agents can increase cGMP in the CNS and that most of this
change is mediated by a NO-sensitive sGC [47,134]. Four GCs have been
cloned from Manduca nerve tissue [35,135–139]. Sequence comparisons with
other family members suggest that two of these forms are NO-sensitive sGCs
(MsGC-1 and MsGC-1), and a third is an unusual isoform related to sGC-
but lacking the consensus cysteines for NO activation (MsGC-3). A fourth
gene is not identifiable as a receptor GC, a sGC or a member of the vertebrate
GC-A family, and has been named MsGC-I [140].
NO production and the cellular effects of NO
Biochemical responses to NO
The inhibition of cGMP accumulation by the sGC inhibitor ODQ
(1H-[1,2,4]oxadiazolo[4,3-a]-quinoxalin-1-one) can be used to assay NO activation in whole nerve cords. In Manduca the entire accumulation of cGMP
evoked by stimulation of metabotropic mAChRs is dependent on NO production [47,134] but this is small compared with the massive effect of activating the
ionotropic nAChRs (Figure 3) [50]. This response appears to be a direct effect
since it persists when spiking in the ganglion is blocked by low-sodium saline
and it is not affected by the Ca2-channel blocker, nickel [50]. Because the
Manduca nAChRs are permeable to Ca2 [141] it is possible that NO production is facilitated by a close association between NOS and nAChRs subunits
such as MARA1 and MARB [141,142]. Currently, it is unknown whether different transmitter systems have converge on the same NO/cGMP neurons, or
if different groups of NO/cGMP neurons are recruited by each transmitter
system.
Physiological responses to NO
NO donors have marked effects upon the activity of Manduca neurons.
For example, application of 0.5 mM sodium nitroprusside or S-nitroso-N-acetylD,L-pencillamine to an isolated abdominal ganglion doubles the spike rates in the
proleg motor nerve, the VNL (lateral branch of the ventral nerve), a change that
can be blocked by ODQ [47]. Application of NO donors also changes the activity of motor neurons supplying the dorsal muscles. These changes are
neuron-specific: hence, motor neurons controlling the large longitudinal dorsal
internal muscle (Figure 2) [143] increase their spike activity in the presence of
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
1800
0.5 mM Zaprinast NMDG
10 M ODQ
cGMP (fmol/nerve cord)
1600
800
600
400
200
0.5
mM DM
Za
SO
Z
pr
ina apr
st ina
st
0.1
N
mM MDG
Ox
0.1
omM
Ox M
o
Pil
oc -M
Ca
arp
0
P
.1
rb
ine
ilo
ac
ho mM carp
Ca
l
ine
rb
0.0
Ca
ac
rb
ac Carb 1 mM hol
ho
l acho sco
p.
l
0.
Ca 01 m sc
o
rb
ac M m p.
ho
ec
l
a
0.1 m .
mM eca
.
GA
0.1
BA
Ni mM GAB
co
Ni
tin
c A
e otin
Ox e
oM
0
Figure 3 Nicotine evokes a sGC-dependent increase in neuronal
cGMP The levels of cGMP in isolated Manduca ventral nerve cords were
measured by enzyme immunoassay. Inhibition of cGMP-phosphodiesterase by
0.50 mM zaprinast significantly increased cGMP levels both in normal saline
(P0.0001) and when most depolarizing activity was blocked using low-sodium
saline (replacing sodium with 126 mM N-methyl-D-glucamine, NMDG), suggesting a basal turnover of cGMP. Additional treatments (in the presence of
zaprinast) with cholinergic agonists [oxotremorine-M (oxo-M), pilocarpine and
carbachol], cholinergic antagonists [scopolamine (scop) and mecamylamine
(meca)], or with -aminobutyric acid (GABA), did not increase cGMP significantly. However, an exception was nicotine (0.1 mM), which evoked a 4-fold
increase in cGMP (P0.0001), both alone, and in combination with
oxotremorine-M (0.1 mM). Accumulation of cGMP was blocked by pre-treating
nerve cords with the sGC inhibitor ODQ (10 mM, P0.0001). The sample
number for each mean value is shown above the bars.
sodium nitroprusside or spermine-NONOate. These responses have been
analysed in most detail for MN12. NO mediates a conductance-increase depolarization that is mimicked by the cell-permeant cGMP analogue 8-bromo-cGMP
but not by 8-bromo-cAMP and is blocked by the sGC inhibitor ODQ (R.M.
Zayas and B.A. Trimmer, unpublished work; [122]). Intracellular recordings
from the ventral internal medial motoneuron show that similar inhibition of
© 2004 The Biochemical Society
Nitric oxide signalling: insect brains and photocytes
73
spontaneous spiking is mediated by a dramatic hyperpolarization that cannot be
overridden even by the injection of large depolarizing current.
Additional studies show that spontaneous production of NO by ganglia
can regulate basal and stimulated spike rates. For example, the resting spike rate
in the VNL, and that stimulated by oxotremorine-M, are reduced 50–70% by
incubation with the NOS inhibitor NG-nitro-L-arginine and this suppression
can be prevented by co-incubation with the NOS substrate arginine [47]. It is
also noteworthy that the dorsal internal muscle motor neuron (which is stimulated by exogenous NO) is inhibited by application of NG-nitro-L-arginine to
the isolated nerve cord.
The CNS can also be stimulated in intact larvae by placing them into a
low concentration (70 p.p.m.) of NO gas. After a 5 min exposure to NO the
pattern of cGMP-stained neurons resembles that seen after applying NO
donors to the isolated nerve cord and this accumulation can be prevented by
pre-injecting larvae with ODQ [144]. Presumably NO quickly penetrates the
tracheal system and directly activates sGC in target neurons. Within seconds of
exposure to NO, larvae begin to twitch and undergo exaggerated but reasonably co-ordinated movements including crawling and repeated cycles of proleg
retraction and adduction. Although it is unlikely that NO is the main regulator
of such behaviours in normal animals these results suggest that NO is able to
modulate a variety of constitutive motor systems.
Range and specificity of NO in the CNS
Neurons in culture
Manduca neurons can be maintained in culture [145,146] and loaded with
the NO indicator DAF-2-DA (4,5-diaminofluorescein diacetate). Most have
low baseline fluorescence that is evenly distributed throughout the cytoplasm.
The basal fluorescence of cultured neurons (mean pixel intensity of 9
fields = 1167.1 arbitrary units) can be reduced by the NOS inhibitor L-NAME
(NG-nitro L-arginine methyl ester; 1 mM; pixel intensity, 73.66.5 arbitrary
units) and this inhibition is prevented by co-incubating in arginine (10 mM;
pixel intensity, 136.37.4 arbitrary units), suggesting that the accumulated
fluorescence is generated by the endogenous production of NO. Some neurons
have particularly high baseline fluorescence and these are presumed to be
endogenous NO producers. Interestingly, neurons growing nearby (a few cell
diameters or less) are often quite dark, suggesting that basal NO production
does not spread far in the culture medium. These dark cells are loaded with
DAF-2 (4,5-diaminofluorescein) as their fluorescence can be increased two
orders of magnitude by the addition of an NO donor to the culture medium.
The rate of fluorescence accumulation can be increased transiently and repeatedly by applying puffs of ACh (200 ms, 1 mM; Figure 4). These responses are
blocked by bath-applied L-NAME (1–10 mM) showing that they result from
increases in NO synthesis. The same puffs of ACh produce increases in intracellular Ca2 of a similar duration [50,141].
© 2004 The Biochemical Society
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
Rate of change in
fluorescence (dF/dt)
Fluorescence intensity
(A) 90
DAF-2-DAloaded neuron
80
70
60
ACh pipette
50
40
1
0
0
100
200
300
Response magnitude (total change in rate)
(B) 60
400
L-NAME
500
(1 mM)
50
40
30
200 s
20
10
0
1000
2000
3000
4000
Figure 4 ACh stimulates NO release from Manduca neurons in culture.
Neurons cultured for 2 days were incubated in DAF-2-DA (60 min, 25 M). The
accumulated DAF-2 fluorescence of a single neuron (A, upper trace) was monitored during the application of ACh from a pressure pipette (2–5 pulses, 200 ms,
filled boxes). Each application caused a transient increase in the rate of fluorescence accumulation. This is most easily seen in the lower trace plotting the rate of
fluorescence change after removing fast noise transients with a low-pass two-pole
Butterworth filter at 0.05 Hz. These increases in fluorescence presumably represent pulses of increased NO production. This interpretation is supported by the
results shown in (B). A measure of the increased production of NO over basal
levels was made by integrating the total change in fluorescence over the duration
of each response (the response magnitude). Repeated pulses of ACh evoked reasonably constant changes in fluorescence for 40 min. However, after bath
application of the NOS inhibitor L-NAME (1 mM, dark bar), the responses to continued pulses of ACh declined rapidly. The inset trace shows the last nine
responses during L-NAME treatment. The sensitivity of the ACh response to
L-NAME suggests that DAF-2 fluorescence in this neuron is attributable to
NOS activity.
Nitric oxide signalling: insect brains and photocytes
75
Neurons in situ
Experiments in Manduca suggest that NO cannot reach all regions of the
intact CNS. Hence when nerve cords are incubated in the ACh esterase
inhibitor neostigmine it increases basal ACh levels and evokes a large increase in
both the number and intensity of cGMP-immunoreactive neurons. However,
despite the known presence of NO-responsive neurons in all regions of the ganglion [119,144], neostigmine only increased cGMP accumulation in ventral
neurons. This region coincides with the ventral ramifications of most cholinergic sensory neurons and is inconsistent with a rapid and uniform diffusion of
NO throughout the ganglion. Instead, it suggests that the actions of NO can be
spatially restricted even within the small confines of an insect ganglion.
Furthermore, when ganglia are incubated overnight in DAF-2-DA,
numerous cellular ‘hot spots’ are visible. Because DAF-2 becomes irreversibly
fluorescent after exposure to NO these bright cells probably represent locally
high concentrations of NO. Subsequent addition of excess NOC-7 [3-(2hydroxy-1-methyl-2 nitrosohydrazino)-N-methylpropanamine; an NO donor]
turns the entire ganglion bright, suggesting that DAF-2 is distributed throughout the tissue. Other explanations are possible. In some tissues DAF-2 is
sensitive to the concentration of ascorbic acid [147,148] or to Ca2 and illumination [149,150]. DAF-2 does not react with NO directly but with one of its
rapidly generated oxidation products [151]. Therefore the supply of oxygen
within the nervous system also determines the development of DAF-2 fluorescence even if NO can diffuse freely. In many insect tissues the tracheolar system
supplies oxygen to individual cells thereby eliminating diffusive uptake from the
extracellular space [152]. Because cells are also sites of oxygen consumption
the extracellular space in very dense active tissues, such as the neuropile, is
potentially anoxic.
Recent experiments on Manduca also suggest that the outer sheath and
perineural cell layers have a major impact on the reactivity of NO. When cells
in intact abdominal ganglia are loaded with DAF-2 and exposed to NO donors
they do not fluoresce strongly until the perineural sheath and the underlying
cells are disrupted. The sheath region constitutes a powerful protective barrier
for neurons with the P-glycoprotein enzyme system [153] and cytochrome
P450-dependent polysubstrate mono-oxygenase (Figure 1) [154]; hence it
could chemically exclude NO. However, it is more likely that removal of the
barrier layer changes the extracellular milieu and promotes formation of fluorescent triazolofluorescein from DAF-2. This tight regulation of the oxidative
conditions inside the neuropil could have important repercussions for NO signalling in insects.
NO in the firefly lantern
Firefly courtship depends on a remarkable flash communication system
involving precisely timed, rapid bursts of bioluminescence that vary in duration and flash patterns among firefly species [155,156]. The biochemical
reactions directly responsible for bioluminescence are well understood and it is
generally accepted that the limiting step for light production is the supply of
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
oxygen to the activated luciferin complex [157]. However, the mechanism of
oxygen ‘gating’ during firefly flashes has yet to be determined. Recent reports
implicate NO as the key mediator of on/off flashing in the photocytes of adult
Photuris sp. fireflies [100,158].
NO regulation of firefly flash control
Each firefly flash is initiated by a burst of neural activity that stimulates
the release of octopamine from terminals in the lantern [159–161]. These nerve
endings do not synapse on to the light-producing photocytes but are found
instead on tracheolar cells surrounding terminal branch points of the tracheolar
air supply. The chemical production of light by the ATP- and oxygen-dependent luciferin/luciferase reaction takes place in the peroxisomes located deep
inside the large photocytes. What has long been a mystery is how the neural
signal is communicated to the photocytes and how it regulates a change in oxygen access to the peroxisomes.
Photocyte
Trachea
Nerve
Flash
NO
Light
O2
Luciferin*
O2
ATP
Quiescent
Luciferin
Tracheolar
end cells
Peroxisome
Mitochondria
Figure 5 A model for the control of firefly lantern flash by NO. When
the lantern (photocyte) is dark (quiescent; below the horizontal line) oxygen is
delivered by the trachea and tracheoles and is consumed through respiration in
mitochondria packed into the peripheral cytoplasm of the photocytes. Little
oxygen reaches peroxisomes that contain the light-producing reactions of the
luciferin/luciferase pathway. In this quiescent state, ATP produced by oxidative
phosphorylation promotes formation and accumulation of the activated
luciferin-adenylate intermediate (denoted as luciferin*) by luciferase. In flash
mode (above the horizontal line) nerve activity causes octopamine release that
transiently activates lantern NOS. NO diffuses rapidly and binds to COX,
where it inhibits oxygen use by the photocyte mitochondria. Now oxygen
delivered by the tracheal system diffuses through photocytes to the peroxisomes where it triggers the light-producing reaction. The light flash itself might
help to terminate respiratory inhibition by promoting NO release from COX
(thin arrows).
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Nitric oxide signalling: insect brains and photocytes
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Recently a model has been proposed (Figure 5) [100,158] in which
octopamine transiently activates NOS to produce a burst of NO. The NO
rapidly diffuses into the peripheral cytoplasm of local photocytes where mitochondria are densely packed. Here the NO binds to COX (cytochrome c
oxidase) and inhibits mitochondrial oxygen consumption. The oxygen arriving
at the tracheoles thus is free to diffuse past the mitochondria to the more centrally located peroxisomes, which otherwise are essentially anoxic due to
mitochondrial oxygen consumption. The resulting increase in oxygen availability to the peroxisomes triggers light production by reaction with the
accumulated luciferyl-adenylate intermediate.
Three major findings support this model [100]. First, adult fireflies
(Photuris sp.) maintained in an atmosphere of 80% nitrogen/20% oxygen commenced light production when NO (70 p.p.m.) was introduced, and ceased light
production when NO was removed. Secondly, partially dissected lantern organs
(with neural inputs removed but the tracheal system intact) produced light in
response to NO, and to the neural transmitter octopamine, and this response was
inhibited by the NO scavenger carboxy-PTIO [2-(4-carboxyphenyl)-4,4,5,5tetramethylimidazoline-1-oxyl-3-oxide]. Thirdly, immmunocytochemistry and
NADPH-diaphorase staining showed that NOS is located in cells between the
ends of fine tracheoles and the photocytes.
NO-mediated inhibition of COX
This model is consistent with the known NO-mediated reversible inhibition of COX that occurs at physiological concentrations of NO. (It is
important to distinguish this reversible inhibition from the irreversible and
indiscriminate inhibition of many enzymes that occurs at higher concentrations of NO due to chemical modification, catalysed by products of NO
metabolism such as peroxynitrite [162].) COX is a multi-subunit respiratory
enzyme complex located in the inner mitochondrial membrane. It is the terminal electron acceptor of the mitochondrial electron-transfer chain, and
catalyses the reaction that converts molecular oxygen to water. Nanomolar
concentrations of externally added NO inhibit mitochondrial COX in isolated
preparations of the enzyme, and in isolated mitochondria, brain nerve terminals and a variety of cultured mammalian cell types [162,163]. Oxygen
consumption in mitochondria isolated from firefly lantern organs has also been
shown to be completely inhibited in the presence of the NO donor NOC-7
[158]. NO inhibition of mitochondrial COX has a rapid onset and occurs via
NO binding to one or more enzyme sites in competition with oxygen [164].
Competition by NO binding to COX raises the apparent Km for oxygen, thus
dampening the rate of oxygen utilization [165]. Concentrations of NO measured in a range of biological systems are similar to those shown to reversibly
inhibit COX and mitochondrial respiration, suggesting that NO inhibition of
COX may be a involved in the physiological regulation of mitochondrial respiratory rate [162,166]. In fact, constitutively expressed, endogenously released
NO has been shown to regulate mitochondrial respiration through modulation
of COX in vascular endothelial cells [167].
© 2004 The Biochemical Society
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B.A. Trimmer, J. Aprille and J. Modica-Napolitano
Flash termination
The distinctive spatial arrangement of NOS-containing cells in the firefly
lantern, the rapid synthesis and high diffusion rate of NO, the ability of NO to
permeate the plasma membrane and membrane-bound cytoplasmic compartments and the reversible inhibition of COX by NO are all essential elements in
the putative role of this molecule as a signal for flash initiation. Additional features of the interaction of NO with COX may be of importance in the
termination of firefly flashing. For example, NO-induced inhibition of mitochondrial respiration is reversible and therefore is relieved by NO degradation
[12] or by increasing concentrations of oxygen [168]. In the firefly lantern, flash
termination could occur as NO is rapidly degraded following cessation of the
CNS signal that stimulates NO production, or by competition between NO
and locally increased oxygen for binding to COX. Furthermore, the inhibition
of COX by NO is rapidly and completely reversible by light in mammalian
cells [169] and in mitochondria isolated from firefly lantern organs [158]. These
results support the hypothesis that in the firefly lantern, the flash itself may
contribute to rapid flash termination [100].
Additional mechanisms for flash control
There may be other mechanisms that contribute to the control of oxygen
access to the firefly photocyte peroxisomes. It has been postulated that rapid
gating of oxygen to light-emitting lantern photocytes is modulated by tracheolar fluid levels [157]. According to this mechanism, CNS stimulation of tracheal
end cells leads somehow to a transient increase in intracellular osmotic potential and the absorption of tracheolar fluid by these cells. The decrease in fluid
levels and resultant decreased diffusion barrier increases the rate of oxygen
supply from tracheoles to the photocyte peroxisomes, thus stimulating the
oxygen-dependent light-emitting reaction.
Because oxygen availability is so important for regulation of firefly flashing, more than one mechanism for controlling oxygen delivery is likely to be
involved. Different mechanisms may dominate during the day when flashing
does not occur, compared with night-time, when precise flashing is critical for
mating. A complete understanding of flash control will surely require an analysis of tracheole fluid control and oxygen-diffusion kinetics, as well as the
further elucidation of the role of NO.
Closing comments
Because insect tissues are small, highly structured and contain relatively
few cells they are very good models for understanding the relationship between
tissue morphology and NO function. Hence in the CNS, individual NO-producing and -responsive neurons can be identified and used to estimate the range
and specificity of NO/cGMP signalling. Initial evidence suggests that NO does
not permeate an entire ganglion but instead might be restricted to fairly small
regions of the neuropil. The mechanisms controlling this range are unknown
but are expected to involve the unique oxygen supply and oxidative protective
mechanisms used by insects. Similarly, the firefly lantern provides clues about
© 2004 The Biochemical Society
Nitric oxide signalling: insect brains and photocytes
79
how NO can co-ordinate the activity of groups of cells (the photocytes) and help
them to respond synchronously. This tissue also suggests ways in which NO can
evoke rapid behavioural effects through the regulation of mitochondrial oxygen
consumption. The remaining challenges require a better understanding of
the redox conditions in the insect CNS and a more through examination of the
kinetics and magnitude of NO production and its biological effects.
We thank Dr Ricardo Zayas for permission to use some of his unpublished results
(including Figure 3) and Dr Jonathon Issberner for his observations on the distribution of NO in the CNS. Much of the work reported in this chapter was sponsored
by a grant from the NSF (IBN 0077812) to B.A.T.
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