Download Specific Associations of Neurosecretory or Neuromodulatory Axons

AMER. ZOOL., 35:566-577 (1995)
Specific Associations of Neurosecretory or Neuromodulatory Axons with
Insect Skeletal Muscles1
MARY B. RHEUBEN
Department of Anatomy, Michigan State University, East Lansing, Michigan 48824-1316
SYNOPSIS. The general process of neuromodulation in the skeletal muscles of various insects is accomplished via several different structural
forms. In addition to motor axons, which may contain a modulatory
substance as a co-transmitter, a second class of axons is found in close
association with insect skeletal muscles. These axons typically contain
dense cored vesicles. Some come directly into contact with the sarcolemma, but do not form a typical neuromuscular junction. Others have
finely distributed branches within the muscle but only directly contact
glial cells of the motor nerve branches. Immunocytochemistry has shown
that these nerve types contain one or more of several potential transmitters: glutamate, octopamine, serotonin, leucokinin, proctolin, or insulin.
While some of these substances are known to modulate muscle fiber
contractile abilities directly, or to affect the mechanisms of neuromuscular
transmission, others can be hypothesized to be involved in development,
respiration, and other undiscovered functions.
Nerve terminals containing large numbers of dense cored vesicles, or a mixture
of dense and clear cored vesicles, are also
seen in a variety of locations, including
adjacent to muscles. The dense cored vesicles vary in appearance in different terminal types, implying that their enclosed substances might differ. For example, five
distinct types of dense cored vesicles were
described in the peripheral neurosecretory
system in stick insect (Fifield and Finlayson,
1978), and Jia et al. (1993) report three morphologically distinct types of terminals that
contain dense cored vesicles in Drosophila
skeletal muscles.
Sometimes the terminals containing dense
cored vesicles differ from typical central,
motor or sensory synapses in that their postsynaptic target is either not evident at all
morphologically, being a distant target
reached by diffusion (neurohormonal association), or their structural associations with
the target cell are less well defined. A contact
region may partly resemble a synapse, but
often lacks obvious structural postsynaptic
specializations. Evidence for a specific region
1
From the Symposium Invertebrate Neuromuscular of the terminal dedicated to release of neuOrganization: Peripheral Contributions to Behavioral rosecretory material and reuptake of memVariability presented at the Annual Meeting of the brane can consist of a distinct cluster of clear
American Society of Zoologists, 27-30 December 1993,
cored vesicles resembling those found at
at Los Angeles, California.
INTRODUCTION
Skeletal muscles in most insect species
receive excitatory motor innervation from
one or more nerve terminals containing large
numbers of electron lucent vesicles, averaging 20-60 nm in diameter, with an occasional larger dense cored vesicle among them
(Osborne, 1975). These nerve terminals
typically produce excitatory junction potentials (EJPs) of two general types: either
"slow," in which the EJP is small, of longer
timecourse, and facilitates readily, or "fast"
in which the EJP is larger, usually above
threshold for an active membrane response,
and may depress on repeated stimulation.
Generally speaking, the structural differences between "slow" and "fast" nerve terminals are quantitative rather than qualitative (Rheuben, 1985;Titmus, 1981). Some
muscles are also innervated by inhibitory
motor neurones, and the vesicles of these
are also electron lucent or "clear cored"
(Titmus, 1981; Aizu, 1982).
566
NEUROMODULATORY AXONS IN INSECT MUSCLE
normal synapses ("synaptoid vesicles"), an
electron dense tuft or bar, resembling the
presynaptic active zone, and large dense
cored vesicles nearby. These regions of the
terminal or axon are often termed "synaptoid contacts" (Raabe, 1989). They may
occur in apposition to nothing more substantial than a layer of basal lamina.
The nature of the functional relationships
between the several types of nerve terminals
and their targets is something of a continuum. The distinction between neurotransmission, neuromodulation, and hormonal
effects is one of degree, and the same substance may function in all of the three ways.
In this paper the term neurotransmission is
used to mean the action of a substance which
is limited to a synaptic cleft; in the case of
insect muscle this refers either to excitatory
or inhibitory neuromuscular junctions which
produce an immediate electrical effect on
the muscle fiber. "Neuromodulation" refers
to the process by which a second nerve can
chemically modify synaptic transmission by
affecting either pre- or postsynaptic mechanisms. The term neurohormone refers to
substances which act after diffusing some
distance or after being transported to broadly
distributed sites of effect via the bloodstream. A neuromodulatory substance could
conceivably arrive at a muscle or neuromuscular junction either as a neurohormone or via a more closely applied nerve
process. See Raabe (1989) for a complete
description of the ways that these general
terms are applied to insects.
The most obvious function in skeletal
muscle to be modulated is contraction—its
amplitude and time course—and the mechanisms can range from direct and indirect
effects on muscle fiber membrane properties
to influences on cAMP levels. One of the
best known neuromodulatory substances to
be used in this fashion in insects is octopamine, which enhances the capabilities of
flight muscle via both pre- and postsynaptic
effects (reviewed by Orchard et al, 1993).
For example, octopamine increases miniature excitatory junction potential (MEJP)
frequency as well as the amplitude, the contraction rate, and the rate of relaxation of
the nerve-evoked twitch in the dorsal longitudinal flight muscles of cricket and locust
567
(Whim and Evans, 1988; O'Gara and
Drewes, 1990). In Manduca, octopamine
can decrease input resistance and hyperpolarize the muscle fiber membrane,
increase the amplitude and timecourse of
the excitatory junction potential, and
increase the amplitude and frequency of the
MEJP. The precise effects are dependent
upon the stage of development of the animal
(Klaassen and Kammer, 1985; Klaassen et
al, 1986; Fitch and Kammer, 1986). However other functions besides contraction,
such as those involving the metabolism and
respiration of the muscle fiber, its development, or the development of its innervation, should also be considered. The
diversity of structural types of neurosecretory axons in close association with skeletal
muscles support the likelihood of diverse
transmitters and diverse functions, and, as
will be described below, the structural
arrangements appear to be characteristic of
the particular insect group.
In 1971 Osborne (Osborne et al, 1971)
described neurosecretory type endings in
association with striated muscles from three
insect species and from frog. At the end of
the discussion, those authors commented
"Thus somatic muscles in general are innervated by a variety of axons, some of which
appear to be neurosecretory. The physiological significance of these endings clearly
remains to be discovered." In 1994, while
the number of structural examples have
increased, and while the potential identities
of their transmitters have been deduced in
some cases using immunocytochemistry,
little is known about their normal uses. The
objective of this article is to summarize what
is known about the subset of neurosecretory
or neuromodulatory axons that specifically
associate with skeletal muscles, to add some
recent work from our laboratory, and to suggest some future directions for study.
PERIPHERAL NEUROSECRETORY
STRUCTURES IN INSECT
SKELETAL MUSCLE
The neurosecretory or neuromodulatory
endings found in close association with skeletal muscle fibers may be categorized into
three broad structural types for the purposes
of discussion:
568
MARY B. RHEUBEN
1. The neuromodulatory function is
combined directly with that of synaptic
transmission, so that the neuromodulatory
substance is actually a co-transmitter in a
"normal" motor neuron.
2. The neuromodulatory ending is a separate structure from the motor neuron. Its
terminal contains numerous dense cored
vesicles, and is in direct contact with the
muscle fiber membrane. In some there may
be, in addition to the dense cored vesicles,
a "synaptoid" or synapse-like structure, with
its cluster of clear vesicles surrounding an
electron dense bar. A distinct postsynaptic
specialization may be lacking.
3. The neuromodulatory ending accompanies the fine motor nerve branches within
the muscle, but does not directly contact the
muscle fiber. Apparent synaptoid release
sites may occur in juxtaposition with the
glial cells ensheathing the neurosecretory
axon or only with basal lamina. There is no
significant structural barrier between a
released substance and the hemolymph.
Each of these structural designs may be
used in different parts of an insect's nervous
system, depending on the species.
/. The neuromodulatory substance is a
co-transmitter with the excitatory
transmitter in the motor neurones
The best investigated example of an insect
skeletal muscle receiving innervation from
a motor neuron having two transmitters with
differing functions is found in the cockroach. The peptide proctolin has been identified in a slow motor neuron (Ds) that
innervates the coxal depressor muscle
(O'Shea and Bishop, 1982; Adams and
O'Shea, 1983). Both the cell body and its
fine branches within the depressor muscle
have been shown to contain a proctolin-like
substance using both single cell neurochemical methods and immunocytochemistry
(O'Shea et al, 1982). Application of proctolin to the muscle directly produces a slow
rise in tension, without membrane depolarization. Tonic high frequency stimulation of the nerve also causes a slow increase
in tension superimposed upon the nerve
evoked twitches. The single evoked excitatory junction potential was similar to that
seen from non-proctolin containing motor
neurones, so it has been suggested that proctolin is present in conjunction with another
excitatory transmitter, presumably glutamate (Adams and O'Shea, 1983). Only particular divisions of muscle 177 in the metathoracic segment received innervation from
Ds.
The tonic bundle of the locust extensor
tibia muscle exhibits great sensitivity to
proctolin, which enhances the force of the
spontaneous myogenic contractions at 5 x
10-" M (May et al, 1979) and proctolin
has been identified in its slow motor neuron
(Worden et al, 1985). Unspecified body wall
muscles in the cockroach abdomen (O'Shea
et al, 1982) also receive branches from a
small number of proctolin containing neurones (lateral white cells) that can be identified within the abdominal ganglia. Proctolin-containing beaded endings are seen at
the light microscope level on the mandibular closer muscle in locusts, and proctolin
enhances the neurally evoked contractions
as well as elevating inositol trisphosphate
(IP3) (Baines et al, 1990a). From a survey
of skeletal muscles for proctolin-like immunoreactivity it has been suggested that proctolin may be contained in some but not all
of the slow motor neurones associated with
tonic (slow) muscles (Witten and O'Shea,
1985). In the skeletal muscles of insects and
other arthropods that have been studied
physiologically, proctolin enhances contraction or generates a slow rise in tension
similar to a "catch" state (O'Shea and Shaffer, 1985), so it may often have a role in
maintaining resting tension. Octopamine
and serotonin have also been associated with
sustained tension or postural muscle control
in insects and Crustacea (Hoyle, 1984; review
by Kravitz, 1988).
Besides forming endings in close association with skeletal muscles, neurones containing proctolin or a proctolin-like substance also extensively innervate the heart,
digestive tract, oviduct musculature, and
neurohaemal regions (Nassel and O'Shea,
1987, blowfly; Witten and O'Shea, 1985,
cockroach; Veenstra et al, 1985, Colorado
potato beetle; Davis et al, 1989, Lepidoptera) so that proctolin may commonly operate widely as a neurohormone as well as a
neuromodulator, perhaps with general
NEUROMODULATORY AXONS rN INSECT MUSCLE
functions in feeding and digestion. In the
visceral organs the structural specializations
associated with the proctolin-containing
terminals include both direct synaptoid
contacts with muscle fibers and neurohemal
type release sites (Klemm et al, 1986).
569
fibers directly. At least four types of axon
have been observed so far to innervate particular muscles of this group. These axons
differ characteristically in size of bouton and
type of vesicle.
One category, "Type I," corresponds best
to a motor nerve function. It has been
divided into two subtypes, Is and Ib (Atwood
et al., 1993) or CV and CVo (Jia et ai, 1993)
for muscle fibers 6 and 7, and is characterized by relatively large boutons, clear cored
vesicles, and the presence of a subsynaptic
reticulum formed by the muscle fiber at the
synaptic sites. Occasional dense cored vesicles have been seen in Type Is terminals.
The Ib and Is terminals appear to give rise
to slow and fast synaptic potentials respectively, (Atwood et al., 1993; Kurdyak et al.,
1993) and are described more fully elsewhere in this volume (Atwood and Cooper,
1995).
The second axon type, "Type II" (Johansen et al., 19896; Budnik and Gorczyka,
1992; Monastirioti et al., 1995) is characterized at the light microscope level by substantially smaller (less than 2 pm) rounded
boutons and a wider, more irregular distribution of its terminals over the muscle fibers.
Type II terminals are immunoreactive for
the Small Synaptic Bouton antigen "SSB,"
whereas the Type I motor nerve terminals
are not (Budnik and Gorczyka, 1992). An
ultrastructural comparison of muscle fibers
6 and 7 (which do not usually have type II
endings) and muscle fibers 12 and 13 (which
do) suggests that at least some of the type
II endings have elliptical dense cored vesicles as well as clear cored vesicles (Jia et
al., 1993). These are referred to as "mixed
vesicle", MV, type axons. In subsequent
studies, Monastirioti et al. (1995) have found
that two octopamine immunoreactive neurones supply nearly all of the body wall musculature via these characteristic type II endings.
2. A separate neurosecretory type axon
forms a direct contact with skeletal
muscle fibers
In Dipterans many muscles receive direct
innervation from axons containing large
numbers of dense cored vesicles as well as
from more conventionally structured motor
axons. Because their peripheral axons and
nerve terminals are less well covered by glial
cells than in other types of insects, they are
amenable subjects for immunocytochemical studies. The larval abdominal musculature has several neurosecretory type axons
which have been shown by electron microscopy to make direct contact with the sarcolemma:
For example, muscle fiber " 8 " (the
abdominal muscles in Dipterans are
described anatomically by number) in larval Drosophila, Calliphora, and Phormia is
innervated by three axons, two of which
have terminals containing dense cored vesicles and which occupy shallow depressions
in the muscle fiber membrane. One of the
terminals having dense cored vesicles is
immunoreactive to antiserum to leucokinin
I (LKIR). A single LKIR neuron innervates
each of 7 abdominal hemisegments, makes
intimate contact only with muscle fiber 8 in
each of these segments, has an extension to
a trachea, and has superficially placed varicosities along part of its axon which could
release material into the hemolymph. In the
adult the LKIR reactive axons innervate
abdominal spiracles. Consequently LKIR
axons could have not only a specific and
direct effect on muscle 8, but also be
involved in respiration and the function of
adjacent muscles which are influenced by
A third structural type of axon, "Type
LKIR indirectly through the hemolymph III," was first identified from nerve termi(Cantera and Nassel, 1992).
nals immunoreactive for an insulin-like
Muscle fibers 6, 7, 12, and 13 in Dro- peptide. These terminals exclusively innersophila have been studied extensively. Some vate muscle fiber 12 in segments 2-5, and
of these fibers have, in addition to motor have boutons intermediate in size between
innervation, two or more kinds of neurose- those of typical type I and type II endings,
cretory type axons which contact the muscle and which are more elongated or oval in
570
MARY B. RHEUBEN
shape (Gorczyca et al, 1993). They were
immunologically distinct from the type of
terminal stained by the SSB (small synaptic
bouton) antibody, associated with type II
terminals (Budnik and Gorczyca, 1992). Jia
et al, (1993), after comparing terminal types
at the electron microscopic level on muscles
6, 7, 12, and 13, suggest that a set of terminals containing spherical dense cored
vesicles may correspond to the type III insulin-containing terminals. These axons also
contained clusters of clear cored vesicles in
association with presynaptic densities,
forming a synaptoid; each of the latter structures was found in apposition to the muscle
fiber in depressions of the sarcolemma near
to junctions made by type I terminals
(Atwood et al, 1993; Jia et al., 1993).
Even though the insulin-containing terminals are restricted to a particular muscle,
virtually all body wall neuromuscular junctions exhibit localized immunoreactivity for
insulin receptors, which appeared to be in
the postsynaptic muscle membrane. This
suggests both immediate and long distance
effects for an insulin-like peptide.
Immunoreactivity for proctolin (or proctolin-like substances) has also been seen in
terminals supplying muscle fibers 12 and 13
and selected other fibers in the abdominal
musculature (Anderson et al., 1988; Keshishian et al, 1993). It is not yet certain which
type of terminal houses the proctolin, or
whether it is co-localized to motor nerve
type I terminals containing glutamate, or to
neurosecretory type terminals also containing insulin or octopamine. Furthermore,
glutamate immunoreactivity has been
observed in most if not all type I and type
II fibers in Drosophila (Johansen et al,
1989a, b), with anti-HRP and anti-glutamate antibodies identifying the same terminals in double label experiments done on
muscle fibers 2, 4, 6, 7, 12, and 13. Therefore, in the case of the specialized, type II,
dense cored vesicle-containing terminals,
glutamate may be a co-transmitter to the
octopamine housed in the dense cored vesicles!
When all these points are considered, it
seems likely that some dense cored vesiclecontaining axons of either Type II or Type
III morphologies may well contain different
and possibly multiple transmitter types. If
this is the case then one must define motor
axons and neuromodulatory axons very
carefully, since their functions will be
dependent on the presence and responses of
receptors to their transmitters in the postsynaptic membranes and in adjacent regions.
Further physiological and ultrastructural
double labelling studies will be necessary to
clarify the situation.
In summary, in Drosophila and other
Dipterans, there are at least four or five types
of axon which make direct contact with the
skeletal muscle fibers. Two or three of these
may be excitatory or inhibitory motor axons,
but the other endings could be responsible
for supplying leucokinin I, octopamine,
proctolin, insulin, or other modulatory substances to specific locations on specific muscle fibers. While these latter endings often
closely parallel the motor endings, their separate nature would allow an independent,
more versatile action on the muscle fiber
when compared to the arrangement where
the modulatory substance is a co-transmitter in the motor nerve terminal, and where
release is still presumably linked to motor
axon activity.
In addition, the finding of an insulin-like
peptide and insulin receptors that increase
in distribution during development, and by
analogy with the vertebrate system, has suggested that growth and development of
muscle and neuromuscular junctions are
phenomena in insects that could be regulated by the neurosecretory type axons
(Gorczyca et al, 1993).
3. The neuromodulatory or neurosecretory
axons are in close association with the
motor nerve branches within the
muscle but do not directly contact the
muscle fibers
Insect neurohormones are released from
a variety of specialized organs such as the
corpora cardiaca, which is a glandular structure near the brain, the perisympathetic
organs which are segmentally arranged in
association with the median and transverse
nerves, and various neurohemal release sites
along peripheral nerves (Raabe, 1989). The
single neurosecretory type axons which have
release sites along the fine motor nerve
NEUROMODULATORY AXONS IN INSECT MUSCLE
branches might be viewed as a special case
of neurohemal organs or might represent a
distinctly different mechanism depending
upon the actual target.
For example, serotonin immunoreactive
neurites have been found to form a meshwork in the outer layers of the glial sheath
surrounding motor nerves. In Periplaneta,
a small number of serotonergic cells sends
processes by a very circuitous route from
the subesophageal ganglion to the nerves
supplying the muscles of mastication. When
the serotonergic axons reach the regions of
the nerve adjacent to the muscles, they
divide to form many fine branches along
the surface of the motor nerve trunk (Davis,
1987). Fine beaded branches are also seen
to run in close proximity to muscle fibers
of the mandibular closer muscle (locust,
Baines et al., 19906). In Calliphora, the
homologous nerves contain serotonergic
profiles in the outer neural sheath that are
seen at the electron microscope level to be
separated from the hemolymph only by a
layer of basal lamina. These 5-HT immunoreactive axons contain both clear and
dense cored vesicles (Nassel and Elekes,
1984). Their precise function relative to the
skeletal muscles is unknown; in visceral
muscles of several species exogenously
applied 5-HT results in enhancement of
contraction or increase in frequency of contraction (Nassel, 1988).
Another neuromodulatory cell type whose
terminals form a distant association with
muscle fibers is the octopamine-containing
DUM (dorsal unpaired median) neurones
of the thoracic and abdominal ganglia in
orthopterans, lepidopterans, and hemipterans. DUM cells, unlike most other central
neurones, are capable of generating an action
potential in their somata. Each sends out a
single primary axon which then bifurcates
to provide mirror image branches to the
right and left halves of the thorax (reviewed
by Evans, 1980).
Two DUM neurones have been investigated in particular, the DUMETi (dorsal
unpaired median cell to the extensor tibia
muscle) and DUMDL (dorsal unpaired
median cell to the dorsal longitudinal muscle). Hoyle et al. (1980) traced the axon of
DUMETi to the periphery using a combi-
571
nation of electrophysiological and ultrastructural methods. It was observed to
accompany the fast motor neuron, initially
within the common glial sheath. In branches
of the nerve within the muscle itself, the
presumed DUMETi axon lay in or on the
outermost glial layer, paralleled the fast
motor axon when it branched, formed
"swellings," but was not seen to come into
direct contact with the muscle membrane.
The DUMDL, although not continuously
traced from ganglion to muscle, seemed to
have a similar structure. These DUM cell
axons branch to associate with specific muscles, but yet appear not to form a structural
specialization with the fibers directly. This
suggests a mechanism of modulation that
might differ from those used by the previous
two types of neurosecretory innervation. In
the case of the extensor tibia, one known
effect of octopamine and/or stimulation of
the DUMETi neuron is slowing of a myogenic rhythm present in one of the muscle
bundles. For the dorsal longitudinal flight
muscles both octopamine and activation of
the DUM neuron is associated with a complex enhancement of muscle function and
stretch receptor function at the initiation of
flight (Orchard et al, 1993), although distinguishing the effects of circulating octopamine from those supplied by the DUM
cells directly to the muscles is difficult.
In Manduca, we have investigated a similar type of neurosecretory axon in the larval
and adult mesothoracic muscles. The association of neurosecretory type axons with
individual muscle fibers was examined in
detail using extensive serial sectioning
(Rheuben and Autio, in preparation). It was
found for larval dorsal longitudinal muscles
"A," "B," and "C" that contribute to the
adult dorsal longitudinal flight muscles, that
a single neurosecretory axon followed and
branched with the single motor axon along
the surface of the muscle fiber. The neurosecretory twig was not ensheathed completely by glial cell processes as was the
motor axon, but rather lay loosely wrapped
in the outermost layer of the glial sheath.
In several long sets of serial sections, the
neurosecretory twig was found to enlarge to
form varicosities at intervals. These varicosities contained both clear and dense cored
572
MARY B. RHEUBEN
vesicles. Occasionally clusters of clear cored
vesicles were found in conjuction to a structure comparable to a presynaptic density,
forming a "synaptoid" contact. This contact
was often in direct apposition to a glial cell
membrane or occasionally to the thick layer
of basal lamina that surrounded the outside
of the nerve as a whole (Figs. 1, 2). Even
though the motor axon branched off to form
neuromuscular junctions at intervals, the
neurosecretory twig did not. No direct contacts were seen between the neurosecretory
axon and the muscle fiber, either within or
adjacent to the neuromuscular junctions.
Virtually all larval terminal motor nerve
branches were found to have an associated
neurosecretory twig in the outer part of the
glial sheath. This type of synaptoid contact
with glial processes also occurs in the dorsal
longitudinal nerve in flightless grasshoppers
even though the muscle is greatly reduced
or absent (Arbas and Tolbert, 1986).
In contrast, in the adult dorsal longitudinal muscles innervated by some of the
same larval motor axons (Rheuben and
Kammer, 1980) the neurosecretory type
axon has been seen within the muscle in
affiliation with the terminal branches of the
motor axon only very rarely. Axons containing similar types of dense cored vesicles
have been seen in the outer sheaths of main
nerve branches close to the third axillary
muscles (Rheuben and Kammer, 1983) in
adults, and are frequent at the bases of the
nerves supplying the dorsal longitudinal
muscles, near the neurohemal organs at each
segmental ganglion (Wasserman, 1985). The
reasons for this developmental change in
apparent distribution for one particular
nerve type are not known, nor is the transmitter. The functions and fiber types of larval and adult muscles are quite different, as
are some of the adult muscles from each
other, so the presence or absence of a modulatory neurosecretory axon could relate
either to developmental changes or to not
yet understood differences in function.
FUNCTIONS OF THE NEUROSECRETORY
TYPE AXONS SUGGESTED BY THEIR
STRUCTURE
In both the structural type in which a
modulatory substance is carried as a co-
transmitter, and that in which a separate
neurosecretory type axon makes direct contact with the muscle sarcolemma, it would
seem that the most immediate target is the
postsynaptic muscle fiber and the most
immediate function is modulation of contraction. However, the morphology does not
exclude secondary targets such as adjacent
glial cells, tracheoblasts, or adjacent muscles and their innervation. The existence of
the third structural type, in which the neurosecretory axon only comes as close as the
tertiary branches of the motor axon, further
suggests that other targets and other functions may be important. It might be useful
to speculate on some of these.
1. Modulation of glial function
In Manduca (Rheuben and Autio, in
preparation) and in locusts (Hoyle et ah,
1980) neurosecretory type axons form varicosities and synaptoid structures with or
near the glial cells which enshroud the motor
axon. This could be happenstance or it could
suggest that the glial cell is an immediate
target. If the glial cell is the immediate target, what functions might be modulated? In
locust several layers of glial cells form a
"blood-brain" barrier in central and peripheral nerves, with a voltage being developed
across it. Applied octopamine modulates the
ability of this potential to change in response
to alterations in extracellular potassium
concentrations, with one possible explanation being a fall in K + permeability of the
basolateral membranes (Schofield and
Treherne, 1986). Octopamine also directly
hyperpolarizes Schwann cell membranes in
squid (Reale et al, 1986). DUM neurites
form structures similar to presynaptic densities in apposition to glial cells within the
ganglia of locusts (Watson, 1984). In subsequent studies, the DUM cell processes in
the same regions were shown to be immunoreactive to octopamine (Stevenson et al,
1992). Taken together these findings suggest
the hypothesis that octopaminergic neurosecretory type neurons, particularly the
DUM neurons, may have a direct effect on
glial cells that they "innervate." Similarly
the observations that exogenous 5-HT
affects the membrane potentials of leech glial
cells (Walz and Schlue, 1982) and seroto-
FIG. 1. Tertiary motor nerve branch in the vicinity of a neuromuscularjunction. An oblique section of the
nerve is shown in the lower right comer of the micrograph. A varicosity of the neurosecretory axon (NS) lies
in an indentation of the outermost glial process, and a small portion of the motor axon (A) is at the edge of the
section. The profile of the neurosecretory axon contains a single dense cored vesicle and a cluster of clear cored
vesicles. Between varicosities, the neurosecretory axons are much smaller in cross-sectional area, as little as 0.2
iim. The neuromuscularjunction and its associated glia are on the left. The neurosecretory axons were not found
any closer to neuromuscular junctions or to muscle fibers than as shown here. The motor axon typically sends
a terminal branch into a junction from the tertiary nerve branch, but the terminal branch is not accompanied
by a process from the neurosecretory axon. G, glial process; Tr, tracheole; F, fibroblast. 51,300 x.
574
MARY B. RHEUBEN
FIG. 2. Synaptoid of a neurosecretory type axon. A grazing section of a varicosity from a tertiary motor branch
from a 5th instar Manduca larva is completely surrounded by a glial process in this region, with the motor axon
and the rest of the glial wrappers being out of the photograph. Clear cored vesicles predominate in a synaptoid.
The presynaptic electron dense specialization (arrow) is shown here in direct apposition to the glial process.
Several microtubules pass through the varicosity. Between varicosities the axon contains only electron lucent
cytoplasm and the continuing microtubules. Dense cored vesicles are sparsely present, with possibly one shown
to the left of the synaptoid. In extensive serial sections this axon type was not found to directly contact the
muscle fiber. 73,600 x.
nergic neurosecretory axons are found commonly in close apposition to the glial sheath
cells of the mandibular nerves evokes speculation of a possible functional relationship.
The idea that glial cells might be targets
of traditional neurotransmitters is not new:
for example, the relationship between glial
cell glycogenolysis and noradrenergic compounds has been explored extensively (Stone
and Ariano, 1989). At the neuromuscular
junction, Schwann cells respond to a lack
of neuronal activity by upregulating glial
fibrillary acidic protein. This effect appeared
to be mediated via nerve terminal transmitter release (Georgiou et al., 1994). The
morphology of astroglial cells is controlled
by beta-adrenergic receptors (Shain et al,
1987). A trophic relationship between glia
and axon in which the glial cells provide
nutritive substances or transmitter precursors has long been suspected. For example,
glia contain the preponderance of glycogen
in the honeybee retina (Tsacopoulos et al.,
1987), and use either the glycogen or glucose
to make alanine which is exported to the
photoreceptors for use as a mitochondrial
energy source (Tsacopoulos et al., 1994). It
would not be surprising if, in some systems,
regulation of the rate of synthesis of these
energy sources were synchronized to
expected needs of the axon via a direct CNS
link.
NEUROMODULATORY AXONS IN INSECT MUSCLE
2. Developmental roles
All insects undergo growth in a series of
molts, and many aspects of this are under
the control of circulating hormones. In
addition, it is possible that some aspects of
this process are regulated locally. Insulin and
insulin-like growth factors may be important in synaptogenesis and maintenance of
synapses, as has been shown for neurones
in culture (Seecof and Dewhurst, 1974;
Vanhems et al, 1990). An insulin-like compound is produced in the corpora cardiaca
of locust (Hetru et al, 1991) presumably for
hormonal type release. In Drosophila a single insulin containing terminal was found
in certain segments at m.f. 12, but insulin
receptors were found at the bases of virtually all neuromuscular junctions in the body
wall musculature (Gorczyca et al., 1993).
Immunoreactivity for both the insulin like
compound and the insulin receptors was not
detectable until late first or early second
instars, with an increase thereafter. These
authors speculate that it might be involved
in expansion of the neuromuscular junctions as the larva grows.
Glia are important to early development
in some areas of the insect nervous system,
forming a scaffold for transverse nerve formation in Manduca (Taghert et al., 1988),
and Drosophila (Gorczyka et al, 1994), and
for central nervous system tracts in Drosophila (Jacobs and Goodman, 1989). Later
in development, in Manduca, the peripheral
glia, both intramuscular and those surrounding main nerve branches, and which
are the apparent targets for the neurosecretory type axons, undergo quite marked morphological changes during metamorphosis
(Rheuben, 1992). This change in morphology of the intramuscular glial cells occurs at
the same time as motor axons are withdrawing from their targets. Later, the same
or similar glia are present, cradling the
growing nerve tip, during the formation of
the adult neuromuscular junctions (Rheuben and Kammer, 1981). Direct application
of a regulatory substance by a neurosecretory axon would allow specific glial cells to
respond to the changing needs of their axons
in a timely fashion during such sequences
of complex developmental changes. Since
various muscles do not develop or degen-
575
erate in synchrony in insects, a specific control mechanism could be useful.
An interesting function for a serotonin
containing neurosecretory type axon has
been proposed for an eclosion muscle of the
tsetse fly. In this muscle a single "immunocyte" or macrophage-like cell is activated
and sends out multiple processes in and
amongst the muscle fibers when degeneration begins. A serotonergic axon terminates
in the vicinity of the immunocyte, and cutting it stops process outgrowth of the immunocyte and concurrent muscle degeneration
(Miyan and Tyrer, 1993). Consequently
these authors postulate that activity of the
immunocyte is controlled in some way by
the neurosecretory type axon, thus indirectly regulating the timing of muscle degeneration.
In summary, we have considered only a
very few of the ways that neurosecretion and
neuromodulation are important. However
these examples suggest the need to consider
diverse roles for the neurosecretory type
axons in association with the skeletal muscles of insects. Although many, such as the
octopaminergic axons in locust, may have
primary effects on muscle contraction, others may be more important in developmental processes, in regulation of non-neuronal
cells, or in respiration. Very little is concretely known about these cells at present
and they offer the opportunity to add to our
understanding of developmental and functional processes in insects in general.
ACKNOWLEDGMENTS
Support for the work in progress reported
here was from NIH Grant NS 17132. We
thank the Insect Physiology Laboratory,
Department of Agriculture, Beltsville, MD
for providing the Manduca eggs. The author
is grateful to Ms. Dawn Autio for her excellent technical assistance, and to Drs. Harold
Atwood, Yoshi Kidokoro, Motojiro Yoshihara, and Michael Gorczyka for their helpful discussions.
REFERENCES
Adams, M. E. and M. O'Shea. 1983. Peptide cotransmitter at a neuromuscular junction. Science 221:
286-289.
Aizu, S. 1982. Morphological differences between
576
MARY B. RHEUBEN
excitatory and inhibitory nerve terminals in cockroach coxal muscles. Tissue Cell 14:329-339.
Anderson, M. S., M. E. Halpern, and H. Keshishian.
1988. Identification of the neuropeptide transmitter proctolin in Drosophila larvae: Characterization of muscle fiber-specific neuromuscular
endings. J. Neurosci. 8:242-255.
Arbas, E. A. and L. P. Tolbert. 1986. Presynaptic
terminals persist following degeneration of "flight"
muscle during development of a flightless grasshopper. J. Neurobiol. 17:627-636.
Atwood, H. L. and R. L. Cooper. 1995. Functional
and structural parallels in crustacean and Drosophila neuromuscular systems. Amer. Zool. 35:
000-000.
Atwood, H. L., C. K. Govind, and C.-F. Wu. 1993.
Differential ultrastructure of synaptic terminals on
ventral longitudinal abdominal muscles in Drosophila larvae. J. Neurobiol. 24:1008-1024.
Baines, R. A., A. B. Lange, and R. G. H. Downer.
1990a. Proctolin in the innervation of the locust
mandibular closer muscle modulates contractions
through the elevation of inositol trisphosphate. J.
Comp. Neurol. 297:479^86.
Baines, R. A., N. M. Tyrer, and R. G. H. Downer.
1990b. Serotoninergic innervation of the locust
mandibular closer muscle modulates contractions
through the elevation of cyclic adenosine monophosphate. J. Comp. Neurol. 294:623-632.
Budnik, V. and M. Gorczyca. 1992. SSB, an antigen
that selectively labels morphologically distinct
synaptic boutons at the Drosophila larval neuromuscular junction. J. Neurobiol. 23:1054-1066.
Cantera, R. and D. R. Nassel. 1992. Segmental peptidergic innervation of abdominal targets in larval
and adult dipteran insects revealed with an antiserum against leucokinin I. Cell Tiss. Res. 269:
459-471.
Davis, N. T. 1987. Neurosecretory neurons and their
projections to the serotonin neurohemal system of
the cockroach Periplaneta americana (L), and
identification of mandibular and maxillary motor
neurons associated with this system. J. Comp.
Neurol. 259:604-621.
Davis, N. T., S. G. Velleman, T. G. Kingan, and H.
Keshishian. 1989. Identification and distribution of a proctolin-like neuropeptide in the nervous system of the gypsy moth, Lymantria dispar,
and in other Lepidoptera. J. Comp. Neurol. 283:
71-85.
Evans, P. D. 1980. Biogenic amines in the insect
nervous sytem. Adv. Insect Physiol. 15:317-473.
Fifield, S. M. and L. H. Finlayson. 1978. Peripheral
neurons and peripheral neurosecretion in the stick
insect, Carausius morosus. Proc. R. Soc. London
B. 200:63-85.
Fitch, G. K. and A. E. Kammer. 1986. Effects of
octopamine and forskolin on excitatory junction
potentials of developing and adult moth muscle.
J. Neurobiol. 17:303-316.
Georgiou, J., R. Robitaille, W. S. Trimble, and M. P.
Charlton. 1994. Synaptic regulation of glial protein expression in vivo. Neuron 12:443-455.
Gorczyca, M. G., R. W. Phillis, and V. Budnik. 1994.
The role of tinman, a mesodermal cell fate gene,
in axon pathfinding during the development of the
transverse nerve in Drosophila. Development 120:
1-10.
Gorczyca, M., C. Augart, and V. Budnik. 1993. Insulin-like receptor and insulin-like peptide are localized at neuromuscular junctions in Drosophila. J.
Neurosci. 13:3692-3704.
Hetru, C, K. W. Li, P. Bulet, M. Lagueux, and J. A.
Hoffmann. 1991. Isolation and structural characterization of an insulin-related molecule, a predominant neuropeptide from Locusta migratoria.
Eur. J. Biochem. 201:495-499.
Hoyle, G. 1984. Neuromuscular transmission in a
primitive insect: Modulation of octopamine, and
catch-like tension. Comp. Biochem. Physiol. 77:
219-232.
Hoyle, G , W. Colquhoun, and M. Williams. 1980.
Fine structure of an octopaminergic neuron and
its terminals. J. Neurobiol. 11:103-126.
Jacobs, J. R. and C. S. Goodman. 1989. Embryonic
development of axon pathways in the Drosophila
CNS: I. A glial scaffold appears before the first
growth cones. J. Neurosci. 9:2402-2411.
Jia,X.-X.,M. Gorczyca, and V. Budnik. 1993. Ultrastructure of neuromuscular junctions in Drosophila: Comparison of wild type and mutants with
increased excitability. J. Neurobiol. 24:1025-1044.
Johansen,J.,M.E.Halpern,andH. Keshishian. 1989a.
Axonal guidance and the development of muscle
fiber-specific innervation in Drosophila embryos.
J. Neurosci. 9:4318^1332.
Johansen, J., M. E. Halpern, K. M. Johansen, and H.
Keshishian. 1989b. Stereotypic morphology of
glutamatergic synapses on identified muscle cells
of Drosophila larvae. J. Neurosci. 9:710-725.
Keshishian, H., A. Chiba, T. N. Chang, M. S. Halfon,
E. W. Harkins, J. Jarecki, L. Wang, M. Anderson,
S. Cash, M. E. Halpern, and J. Johansen. 1993.
Cellular mechanisms governing synaptic development in Drosophila melanogaster. J. Neurobiol.
24:757-787.
Klaassen, L. W. and A. E. Kammer. 1985. Octopamine enhances neuromuscular transmission in
developing and adult moths, Manduca sexta. J.
Neurobiol. 16:227-243.
Klaassen, L. W., A. E. Kammer, and G. K. Fitch.
1986. Effects of octopamine on miniature excitatory junction potentials from developing and adult
moth muscle. J. Neurobiol. 17:291-302.
Klemm, N., R. Hustert, R. Cantera, and D. R. Nassel.
1986. Neurons reactive to antibodies against
serotonin in the stomatogastric nervous system
and in the alimentary canal of locust and crickets
(Orthoptera, Insecta). Neuroscience 17:247-261.
Kravitz, E. A. 1988. Hormonal control of behavior:
Amines and the biasing of behavioral output in
lobsters. Science 241:1775-1781.
Kurdyak, P., H. L. Atwood, B. A. Stewart, and C.-F.
Wu. 1993. Differential physiology and morphology of motor axons to ventral longitudinal
muscles in larval Drosophila. Neurosci. Abstr. 19:
116.51.
May, T. E., the late B. E. Brown, and A. N. Clements.
1979. Experimental studies upon a bundle of tonic
fibres in the locust extensor tibialis muscle. J. Insect.
Physiol. 25:169-181.
Miyan, J. A. and N. M. Tyrer. 1993. Innervation of
NEUROMODULATORY AXONS IN INSECT MUSCLE
dipteran eclosion muscles: Ultrastructure, immunohistochemistry, physiology and death. Phil.
Trans. R. Soc. London B. 341:361-374.
Monastirioti, M., M. Gorczyca, J. Rapus, M. Eckert,
K. White, and V. Budnik. 1995. Octopamine in
the fruit fly Drosophila melanogaster. J. Comp.
Neurol. (In press).
Nassel, D. R. 1988. Serotonin and serotonin-immunoreactive neurons in the nervous system of insects.
Prog. Neurobiol. 30:1-85.
Nassel, D. R. and K. Elekes. 1984. Ultrastructural
demonstration of serotonin-immunoreactivity in
the nervous system of an insect (Calliphora erythrocephala). Neurosci. Letts. 48:203-210.
Nassel, D. R. and M. O'Shea. 1987. Proctolin-like
immunoreactive neurons in the blowfly central
nervous system. J. Comp. Neurol. 265:437-454.
O'Gara, B. A. and C. D. Drewes. 1990. Modulation
of tension production by octopamine in the metathoracic dorsal longitudinal muscle of the cricket
Teleogryllusoceanicus. J. Exp. Biol. 149:161-176.
O'Shea, M. and C. A. Bishop. 1982. Neuropeptide
proctolin associated with an identified skeletal
motoneuron. J. Neurosci. 2:1242-1251.
O'Shea, M. and M. Schafter. 1985. Neuropeptide
function: The invertebrate contribution. Annu.
Rev. Neurosci. 8:171-198.
O'Shea, M., M. E. Adams, and C. A. Bishop. 1982.
Identification of proctolin-containing neurons.
F.A.S.E.B. 41:2940-2947.
Orchard, I., J.-M. Ramirez, and A. B. Lange. 1993.
A multifunctional role for octopamine in locust
flight. Annu. Rev. Entomol. 38:227-249.
Osborne, M. P. 1975. The ultrastructure of nervemuscle synapses. In P. N. R. Usherwood (ed.),
Insect muscle, pp. 151-205. Academic Press Inc.,
New York.
Osborne, M. P., L. H. Finlayson, and M. J. Rice. 1971.
Neurosecretory endings associated with striated
muscles in three insects (Schistocerca, Carausius,
and Phormia) and a frog (Rand). Z. Zellforsch.
116:391-404.
Raabe, M. 1989. Recent developments in insect neurohormones. Plenum Press, New York.
Reale, V., P. D. Evans, and J. Villegas. 1986. Octopaminergic modulation of the membrane potential of the Schwann cell of the squid giant nerve
fibre. J. Exp. Biol. 121:421-443.
Rheuben, M. B. 1985. Quantitative comparison of
the structural features of slow and fast neuromuscular junctions in Manduca. J. Neurosci. 7:17041716.
Rheuben, M. B. 1992. Degenerative changes in the
structure of neuromuscular junctions of Manduca
sexta during metamorphosis. J. Exp. Biol. 167:
119-154.
Rheuben, M. B. and A. E. Kammer. 1980. Comparison of slow larval and fast adult muscle innervated by the same motor neurone. J. Exp. Biol.
84:103-118.
Rheuben, M.B. and A. E. Kammer. 1981. Membrane
structures and physiology of an immature synapse.
J. Neurocytol. 10:557-575.
Rheuben, M. B. and A. E. Kammer. 1983. Mechanisms influencing the amplitude and time course
of the excitatory junction potential. In A. D. Grin-
577
nell and W. Moody (ed.), The Physiology of excitable cells, pp. 393-409. Alan R. Liss, Inc., New
York.
Schofield, P. K. and J. E. Treherne. 1986. Octopamine sensitivity of the blood-brain barrier of an
insect. J. Exp. Biol. 123:423-439.
Shain, W., D. S. Forman, V. Madelian, and J. N. Turner. 1987. Morphology of astrogial cells is controlled by beta-adrenergic receptors. J. Cell Biol.
105:2307-2314.
Seecof, R. L. and S. Dewhurst. 1974. Insulin is a
Drosophila hormone and acts to enhance the differentiation of embryonic Drosophila cells. Differentiation 3:63-70.
Stevenson, P. A., H.-J. Pfluger, M. Eckert, and J. Rapus.
1992. Octopamine immunoreactive cell populations in the locust thoracic-abdominal nervous
system. J. Comp. Neurol. 315:382-397.
Stone, E. A. and M. A. Ariano. 1989. Are glial cells
targets of the central noradrenergic system? A
review of the evidence. Brain Res. Rev. 14:297—
309.
Taghert, P. H., J. N. Carr, and J. B. Wall. 1988. The
formation of a neurohaemal organ during insect
embryogenesis. Adv. Insect Physiol. 20:87-117.
Titmus, M. J. 1981. Ultrastructure of identified fast
excitatory, slow excitatory, and inhibitory neuromuscular junctions in the locust. J. Neurocytol.
10:363-385.
Tsacopoulos, M., J. A. Coles, and G. Van De Werve.
1987. The supply of metabolic substrate from glia
to photoreceptors in the retina of the honeybee
drone. J. Physiol., Paris 82:279-287.
Tsacopoulos, M., A.-L. Veuthey, S. G. Saravelos, P.
Perrottet, and G. Tsoupras. 1994. Glial cells
transform glucose to alanine, which fuels the neurons in the honeybee retina. J. Neurosci. 14:1339—
1351.
Vanhems, E., E. Delbos, and J. Girardie. 1990. Insulin and neuroparsin promote neurite outgrowth in
cultured locust CNS. Eur. J. Neurosci. 2:776-782.
Veenstra, J. A., H. M. Romberg-Privee, and H. Schooneveld. 1985. A proctolin-like peptide and its
immunocytochemical localization in the Colorado
potato beetle, Leptinotarsa decemlineata. Cell Tiss.
Res. 240:535-540.
Walz, W. and W. R. Schlue. 1982. Ionic mechanisms
of a hyperpolarizing 5-hydroxytryptamine effect
on leech neuropile glial cells. Brain Res. 250:111121.
Wasserman, A. J. 1985. Central and peripheral neurosecretory pathways to an insect flight motor
nerve. J. Neurobiol. 16:329-345.
Watson, A. H. D. 1984. The dorsal unpaired median
neurons of the locus metathoracic ganglion: Neuronal structure and diversity, and synapse distribution. J. Neurocytol. 13:303-327.
Whim, M. D. and P. D. Evans. 1988. Octopaminergic
modulation of flight muscle in the locust. J. Exp.
Biol. 134:247-266.
Witten, J. L. and M. O'Shea. 1985. Peptidergic innervation of insect skeletal muscle: Immunochemical
observations. J. Comp. Neurol. 242:93-101.
Worden, M. K., J. L. Witten, and M. O'Shea. 1985.
Proctolin is a co-transmitter for the SETi motoneuron. Neurosci. Abstr. 11:327.