Cellular distribution of a high-affinity glutamate transporter in the nervous system of the cabbage looper Trichoplusia ni
1 Southern Crop Protection and Food Research Centre, Agriculture and
Agri-Food Canada, London, Ontario, Canada N5V 4T3
2 Anatomical Institute, IMBA, University of Oslo, N-0317 Oslo,
Norway
3 Department of Zoology, The University of Western Ontario, London, Ontario,
Canada N6A 5B7
* Author for correspondence (e-mail: donlyc{at}agr.gc.ca)
Accepted 6 June 2002
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Summary |
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Key words: glutamate transporter, Trichoplusia ni, insect, anti-peptide antibody, immunohistochemistry, glutamate
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Introduction |
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Glutamate uptake has also been demonstrated in neural and neuromuscular
tissues in arthropods (Duce,
1988). In insects, glutamate has been shown to be a transmitter in
the CNS and at neuromuscular junctions
(Faeder and Salpeter, 1970
;
Usherwood and Machili, 1968
;
Beranek and Miller, 1968
;
Jan and Jan, 1976
). Glutamate
transporter cDNAs have been cloned from four orders of insects, namely
Lepidoptera (the cabbage looper Trichoplusia ni;
Donly et al., 1997
),
Hymenoptera (the honeybee Apis mellifera;
Kucharski et al., 2000
),
Diptera (the fruit fly Drosophila melanogaster;
Seal et al., 1998
;
Besson et al., 1999
) and
Dictyoptera (the cockroach Diploptera punctata;
Donly et al., 2000
). The
cabbage looper, cockroach and honeybee cDNAs were cloned from brain, and the
fly cDNA from embryonic tissue. All these proteins show a high degree of amino
acid identity and have comparable kinetic and pharmacological properties
(Caveney and Donly, 2002
).
Compared with mammalian EAATs, the peptide sequence of the honeybee EAAT is
most similar to mammalian EAAT2, while fruit fly EAAT1 is closest to mammalian
EAAT1 and cockroach EAAT closest to mammalian EAAT3. A second
Drosophila transporter has also been described
(Besson et al., 1999
),
suggesting that the insect CNS may have multiple forms as seen in mammals. The
T. ni transporter, designated TrnEAAT1 (T. ni excitatory
amino acid transporter 1), shows considerable sequence identity to human
EAAT-1 (40 %), -2 (37 %), -3 (42 %) and -4 (39 %), although it has the highest
identity (up to 53 %) to other cloned insect EAATs
(Donly et al., 2000
). Tissue
expression of TrnEAAT1 has been analyzed by northern blot analysis and the
highest levels were detected in extracts of RNA from brain, lower levels of
expression in the integument, and a weak expression in hindgut and rectum
(Donly et al., 1997
).
Since the cDNAs for most of these proteins were initially isolated using insect head or brain libraries, these transporters are most likely expressed in the CNS of these insects. Northern blot analysis has confirmed that the mRNAs for these proteins are synthesized in the CNS, optic lobes and possibly digestive tract of insects. However, these experiments do not give the precise localization of the protein, which may differ from the site of synthesis of its mRNA. Also, owing to the difficulty of isolating individual cell types in the preparation of cDNA libraries, the cells expressing the transporters have not been identified. Here we report on the cellular localization of the protein product of the TrnEAAT1 gene in tissues of the cabbage looper. Polyclonal antibodies were raised against peptide sequences in the C- and N-terminal regions of the TrnEAAT1 protein and used to localize the transporter. The transporter was found to be localized primarily at the neuromuscular junctions in skeletal muscle and in the ganglionic neuropile of the CNS.
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Materials and methods |
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Antibodies
Anti-peptide antibodies against TrnEAAT1 were prepared as described
(Lehre et al., 1995;
Danbolt et al., 1998b
) using
synthetic peptides corresponding to residues 2-15 (PLQIRR-NRCTSFLR) and
450-479 (LSQGDIDKSRALNEREAAPS-HELTELEKGDH) of the sequence
(Donly et al., 1997
) as
antigens. The former peptide was synthesized as a C-terminal amide, while the
latter was made with free carboxyl terminus. Both peptides were coupled to
keyhole limpet hemocyanin with glutaraldehyde and injected into New Zealand
White (female) rabbits. Rabbits 67JK, JP75, JQ19 and JQ31 were immunized with
the N-terminal peptide, while rabbits 8D0145 and 8D0268 were immunized with
the C-terminal peptide. The sera were affinity-purified on columns with
immobilized peptide as described (Danbolt, 1998b). This study is based on sera
from rabbits JQ31 and 8D0145, which yielded the best antibodies.
Protein isolation
Virus-infected cells
Total protein extracts were prepared from High Five cells (BTI-TN-5B1-4,
Invitrogen) infected with recombinant baculovirus expressing the TrnEAAT1 gene
(Donly et al., 1997) and from
uninfected control cells. Cells were harvested from the growth medium,
centrifuged and resuspended in 10 mmol l-1 sodium phosphate buffer
and immediately frozen. Thawed cells were homogenized in cold phosphate buffer
and centrifuged at 16,000 g for 40 min at 4 °C. The
supernatant (water-soluble fraction) was collected and frozen and the pellet
solubilized at room temperature in buffer containing 1 % SDS. The sample was
centrifuged, the supernatant (`membrane' fraction) collected and frozen, and
the pellet discarded.
Insect tissues
T. ni caterpillars were anaesthetized in 75 % ethanol, and quickly
dissected to isolate tissue samples. Caterpillar integument and associated
skeletal muscle was briefly rinsed in buffer and then immediately frozen in
liquid nitrogen. Adults were anaesthetized at 4 °C prior to isolation of
flight muscle and heads, which were immediately frozen in liquid nitrogen.
Thawed tissue samples were quickly homogenized in 10 volumes of phosphate
buffer using a Brinkmann Polytron Homogenizer. Samples were then processed as
described for High Five cells.
Western blot analysis
Frozen samples containing 30 µg of protein per well were thawed, diluted
in SDS-gel electrophoresis sample buffer [60 mmol l-1 Tris base, pH
6.8, 25 % glycerol, 2 % sodium dodecyl sulfate (Sigma), 14.4 mmol
l-1 2-mercaptoethanol (Sigma), 0.1 % Bromophenol Blue (Sigma)] and
incubated at room temperature for 15 min. SDS-PAGE was performed as described
by Laemmli (1970) on
separating gels consisting of 8 % acrylamide. Protein molecular mass standards
were Bio-Rad prestained SDS-PAGE broad range marker proteins. Gels were then
electroblotted to nitrocellulose (Bio-Rad) membranes
(Towbin et al., 1979
).
Membranes were blocked with 5 % nonfat milk in TBS (10 mmol l-1
Tris base, 150 mmol l-1 NaCl, pH 7.5) with 0.05 % Triton X-100
(Sigma) for 1 h at room temperature, transferred to primary antibodies
(diluted to 1 µg/ml anti-C antibody and 30 µg/ml anti-N antibody) in the
blocking buffer, and incubated for 24 h. The membrane was then washed in TBS
(10 mmol l-1) for 1 h with changes, then incubated in anti-rabbit
IgG (whole molecule)-alkaline phosphatase conjugate (1:30,000; Sigma) in
blocking buffer for 4-24 h at 4 °C on a shaker. The membrane was washed
three times in TBS, and then incubated for 15 min in alkaline phosphatase
buffer (0.1 mol l-1 Tris base, pH 9.5, 0.1 mol l-1 NaCl,
5 mmol l-1 MgCl2). Blots were developed in BCIP/NBT
(5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium) liquid substrate
(Sigma) to visualize the labelled bands. The reaction was stopped in 20 mmol
l-1 EDTA in TBS and the blots photographed.
Isolation of tissues for light and electron microscopy
Third or fourth instar caterpillars were anesthetized for 30 s in 75 %
ethanol, then dissected in cold 4 % paraformaldehyde/0.5 % glutaraldehyde in
Tris (25 mmol l-1) buffered saline (TBS), pH 7.4. Adults were
immobilized in the cold (4 °C) for 30 min then dissected in fixative as
per caterpillars. Tissue samples were removed after 5 min and placed in fresh
fixative at 4 °C for 4 h. The fixative was removed and the samples washed
in TBS for 1 h at 4 °C.
Light microscopy of nerve tissue
Caterpillars were chilled at 4 °C for at least 1 h and then injected
with 1 % Methylene Blue in normal saline
(Plotnikova and Nevmyvaka,
1980). The insects were left at room temperature for 4 h and then
dissected in a fixative of 12% ammonium molybdate (aqueous). Tissue samples
were excised and immediately examined in the light microscope.
Immunohistochemistry
Tissues were incubated in TBS/0.1% Triton X-100 (TTBS) containing 1 %
bovine serum albumin (BSA) for 4 h at 4 °C, then treated with primary
antibody in TTBS/BSA for 8-24 h at 4 °C on a shaker. After primary
incubation, the samples were washed in TTBS for 4 h and then incubated with
anti-rabbit IgG alkaline phosphatase conjugate (Sigma; diluted 1:400 in
TTBS/BSA) for 4 h to overnight at 4 °C. The tissues were washed in TTBS
for 4 h at room temperature. Visualization of the probe was achieved with the
indirect chromogenic method using one of the following substrates: Vector Red,
Vector Blue, Vector Black (Vector Laboratories) and BCIP/NBT (Sigma). Although
blocking for endogenous alkaline phosphatase in the insect tissues was not
needed, the resultant reaction was checked by comparing it to the results
obtained with secondary antibodies linked to rhodamine (Cappel Laboratories)
or gold (British BioCell). Both of these non-enzymatic systems produced the
same results as the alkaline phosphatase, confirming its reliability in this
study. After the substrate produced a suitable colour reaction, the tissue was
washed in water, mounted in Geltol Mounting Media (Immunon) and photographed
with an Axiovert 35 photomicroscope (Zeiss) on Elite Chrome 160 T slide film
(Eastman Kodak).
Silver enhancement of gold for light microscopy was performed by the method
of Danscher (1981).
Gold labelling
Fixed muscle tissues were washed in TBS at 4 °C for 1-2 h, followed by
a treatment in TTBS/1 % BSA for 4 h. Specimens were incubated overnight in
primary antibody (C-peptide antibody) diluted in TTBS/BSA, then washed in TBS
for 5 h with solution changes, followed by incubation in goat antirabbit IgG 1
nm or 10 nm gold (British BioCell) diluted 1:100 (manufacturer's
recommendations) in buffer for 24 h. After washing overnight in buffer,
samples were re-fixed in 2 % glutaraldehyde for 10 min and washed again in
buffer. Tissues were then treated with 0.5 % osmium tetroxide/TBS for 30 min
(room temperature), washed with double-distilled water for 20 min and stained
with 2 % aqueous uranyl acetate for 30 min. Samples were washed in
double-distilled water and dehydrated through a graded acetone series and
embedded in Epon/Araldite. Sections were cut with a diamond knife on a
Reichert ultramicrotome and mounted on nickel grids. Grids were examined in a
Philips CM10 electron microscope.
Ganglia were fixed, dehydrated and embedded in epon as described above, except that the osmium tetroxide was omitted. Sections mounted on nickel grids were floated on a drop of saturated sodium metaperiodate for 10 min (room temperature), and then washed three times in distilled water, followed by a treatment in 0.1 mol l-1 HCl for 5 min. The grids were washed in PTBN buffer (8.5 g NaCl, 40 µl 0.5 mol l-1 NaPO4, pH 7.4, 500 µl Tween 20, 1 g BSA 1-1 ddH2O) for 4x10 min), incubated in primary antibody (1/100-1/500) diluted in PTBN (overnight at room temperature), washed in buffer (4x 1 min), and then treated with anti-rabbit immunoglobulin coupled to 15 nm gold (British BioCell) in PBTN (1/75) for 30 min. Finally, grids were washed in double distilled water and stained with 3 % aqueous uranyl acetate for 30 min prior to examination in the electron microscope.
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Results |
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The labelling patterns obtained with the two antibodies were the same, but
the C-terminal antibody consistently exhibited the highest affinity for the
TrnEAAT1 protein and as a result the N-terminal antibody had to be used at a
higher concentration. The three main bands that were consistently visualized
included two at 40-50 kDa and one at 80-100 kDa. The predicted molecular mass
of the TrnEAAT1 protein is 52 kDa, so it is probable that the strongest band
(nearer 50 kDa) represents the TrnEAAT1 monomer. The reason for this anomalous
mobility on SDS-PAGE is not clear, although the redox state of the markers can
affect apparent mobility relative to other transporters
(Haugeto et al., 1996). This
band also matches that found in insect tissues
(Fig. 2). Because these kinds
of protein easily form SDS-resistant complexes
(Haugeto et al., 1996
;
Danbolt, 2001
), the largest
band at 80-100 kDa is most likely a TrnEAAT1 dimer
(Fig. 1). The third and fastest
migrating band may represent unglycosylated TrnEAAT1 since its mobility is
greater than the product observed in insect tissue (which would be expected to
be fully processed). A fourth faint product migrating between the TrnEAAT1
monomer and dimer was not consistently observed. Despite the heterogeneity of
the products, these blots demonstrate specificity for the TrnEAAT1 protein at
two levels. First, the only reactivity occurs in cells infected with the
TrnEAAT1-expressing virus. Second, the same bands are recognized by two
separate antibodies raised in different rabbits against two different
peptides, one from each terminus of the T. ni glutamate
transporter.
|
Having concluded that the antibodies recognize the TrnEAAT1 protein, the specificity of the antibodies was tested on immunoblots of crude preparations of T. ni tissues. Both caterpillar skeletal muscle and adult head tissue probed with either antibody recognized a band between the 36 and 52 kDa markers (Fig. 2A,B). A high molecular mass oligomeric band was also present in the head tissue lanes, as seen in the immunoblots of extracts of virally infected High Five cells. Also consistent with the High Five blots, the major bands are detected only in the lanes with membrane proteins (although one very weak band can be seen in the adult head lane of a water soluble fraction; Fig. 2A, lane 3). Fig. 2A,B shows that the two antibodies specifically stain the TrnEAAT1 protein in both the head and body of T. ni. However, the anti-N serum, which had to be used at a higher concentration than the anti-C serum in order to visualize the bands, gave a less intense signal and higher background. Because of this all tissue staining experiments were performed with the anti-C serum, as it exhibited no crossreactivity in the tissue extracts.
Tissue distribution of TrnEAAT1
A whole-mount view of a caterpillar abdominal segment stained with the
TrnEAAT1 C-terminal antibody is shown in
Fig. 3. The branching pattern
seen on the surface of the segmental skeletal muscle fibres was identified as
a network of axoglial projections formed by motor neurons and associated glial
cells (details below). The expression pattern of TrnEAAT1 shown in
Fig. 3 shows that these stained
projections are segmentally compartmented. The projections do not cross the
segment border but instead terminate where the muscle inserts into the body
wall (Fig. 3). This segmentally
recurring and restricted pattern of antibody staining of skeletal muscle is
found along the caterpillar's body. The ventral nerve cord in whole-mount
preparations also stained with the TrnEAAT1 C-terminal antibody, but only the
ganglia were seen to stain above background levels. The connectives and the
nerve branches radiating out from the ganglia did not stain
(Fig. 3). Residual tissues left
after the dissection, such as fat body, trachea, and integument also did not
stain.
|
Table 1 lists the
distribution of the transporter protein in caterpillar tissues detected by the
C-terminal antibody. Strong positive reactions were seen in skeletal muscle,
flight muscle and the neuropile of ganglia. Some fibres in intra-oesophageal
muscle stained in a pattern similar to that of the skeletal muscle. Midgut and
hindgut tested negative. Isolated silk glands, gonads, fat bodies, trachea and
integument did not react detectably to the antibody, and the muscles of the
dorsal vessel and adjoining diaphragm also failed to react. Northern blot
analysis had previously suggested that mRNA for TrnEAAT1 was expressed weakly
in the integument, hindgut and rectum
(Donly et al., 1997). The
previous detection of TrnEAAT1 mRNA in the isolated integument
(Donly et al., 1997
) most
likely resulted from contamination by muscle tissue.
|
No reaction was seen in tissues treated with either preimmune serum or when the primary antibody was omitted. The staining reaction was prevented when the antibody was treated with the peptide against which it was raised before use on insect tissues. A commercial anti-glutamate transporter antibody (Chemicon International), raised against the unique amino acid sequence of the C terminus of cloned rat EAAC1 (504-523) also failed to stain insect tissues. Finally, despite its lower affinity for TrnEAAT1 protein, treatment with the N-terminal antibody produced the same tissue staining patterns as those produced with the C-terminal antibody.
Muscle staining
TrnEAAT1 C-terminal antibody staining revealed a branched network of nerves
(motor neurons and associated glial cells) that completely covered the surface
of caterpillar skeletal muscle (Fig.
4). This is of special interest since previous work has shown that
glial (sheath) cells at neuromuscular junctions in insects take up glutamate
(Faeder and Salpeter, 1970;
Faeder et al., 1974
). The
nerve branches imaged by the C-terminal antibody run alongside the trachea in
some instances, or pass over or under them. In some preparations, the nerves
were seen to run the length of the muscle, giving off projections along the
muscle surface. A similar pattern was reported in the larva of the waxmoth
Galleria mellonella by Belton
(1969
). Intra vitam
Methylene Blue staining was used to compare the antibody labelling pattern
with that of previously published patterns of nerve distribution on skeletal
muscle. The pattern of staining produced by Methylene Blue
(Fig. 5A) closely matched the
immunohistochemical staining pattern (Fig.
5B). Breaks in the pattern occur along the nerve indicating areas
where TrnEAAT1 expression is either obscured by trachea or sites where it is
not expressed. The TrnEAAT1 antibodies also stained nerves associated with
adult flight- and inter-segmental muscle (not shown). The fine skeletal muscle
bundles that run between segments in the adult abdomen exhibited a pattern of
innervation similar to that in the caterpillar.
|
|
On examining the larval motor neuron terminals at higher magnification, distinctive features of the antibody labelling pattern became evident (Fig. 6). The individual neurons did not appear to be labeled by antibody, but instead their axons were silhouetted as unstained channels surrounded by strongly immunoreactive glial cells. The inner surface of the glial sheath (immediately adjacent to the axons) stained most intensely. Small circular dark plaques, thought to be the sites of the neuromuscular synapses, were distributed along the tracts formed by the axons and associated glial cells. The outer surface of the glial sheath, as well as the glial cytoplasm was only weakly immunopositive, staining at an intensity barely detectable over the background (muscle) staining (Fig. 6). Except where they form neuromuscular synapses, axon terminals traversing the surface of muscle fibres are wrapped in an immunopositive glial cell sheath. This was seen in cross-sections of muscle in which the transporter had been labelled with a secondary antibody linked to gold and intensified with silver (Fig. 7, inset). All the antibody-labelled glial components are restricted to the outer surface of the muscle, penetrating only a short distance into it. In some cases, glial cells were wrapped over axons that lay in grooves on the muscle surface.
|
|
Electron microscopy was used to determine on which membrane surface the
glial cells and/or axons expressed EAAT1. Labelling was seen to occur at
regular areas over the surface of the muscle in thick sections enhanced with
silver (Fig. 7, inset). When
viewed in the electron microscope, abundant gold label was seen on the inner
surface of the glial plasma membranes (Fig.
7). Labelling occurred only at or near neuromuscular junctions, as
seen in Fig. 7 by the presence
of a rete synapticum, a specialized membranous network in the muscle that lies
between the synapse and the contractile elements of the muscle
(Edwards et al., 1958).
Fig. 8 shows a glial cell
expressing transporter protein on the cytoplasmic face of its plasma membrane
where it envelopes an adjacent axon. The axonal membrane appears to be
unlabelled.
|
Nervous system staining
The neuropiles of the ganglia in the larval CNS stained strongly when
probed with TrnEAAT1 C-terminal antibody
(Fig. 9). The cortex of the
ganglion, in which the neuronal cell bodies reside, and the perineurial sheath
did not react with the antibody. The neuropile region is composed of finely
interwoven nerve terminals interspersed with glial processes. Because synaptic
contacts are largely restricted to the ganglionic neuropile in the CNS, a need
for glutamate removal from the extraneuronal space in this region might be
presumed. The nerve connectives between the ventral ganglia and the nerves
branching from them were not stained by TrnEAAT1 antibody. The pattern of EAAT
staining was quite different from that seen when anti-FMRFamide antibody was
used as a control probe for the ganglion. This antibody labelled only cell
bodies in the ganglion.
|
Ultrastructural examination of the ganglion revealed that the transporter was found in the glial cell membranes that insulate the nerve processes in the neuropile (Fig. 10). Gold particles were distributed randomly along the glial membranes and not clustered. No staining was evident in the perineurium, or the ganglionic cortex. This distribution matches the staining pattern seen in the light microscope after chromogenic labelling (Fig. 9). This cellular pattern of label is in agreement with that seen at the neuromuscular junction, where only the glial cell processes were decorated (Figs 7, 8).
|
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Discussion |
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The distribution of TrnEAAT1 at the neuromuscular synapse, and its specific
expression by glial cells confirms earlier physiological studies on glutamate
uptake by nervous tissue in insects. Faeder and Salpeter
(1970) and Faeder et al.
(1974
) showed that radioactive
glutamate was taken up preferentially by sheath cells surrounding stimulated
nerves in a cockroach, with the highest levels of uptake at neuromuscular
junctions, rather than in the nerve branches running to the muscle fibres.
This observation agrees with the immunohistochemical data reported here.
Faeder's observations were later extended to locusts by Botham et al.
(1978
) and Van Marle et al.
(1985
), confirming that in
insects glutamate is a major neurotransmitter at neuromuscular junctions, and
that glial cells play a major role in glutamate recycling. In the caterpillar,
only nerves directly in contact with skeletal muscle and not the nerve fibres
running from the ganglion to the muscles stained positively for EAAT. Our data
also indicate that the transporter protein is localized to glial membranes and
not to axonal membranes. Since axons may travel deep into the muscle and
beyond their glial sheaths, it has been proposed that muscle glutamine
synthetase may be responsible for glutamate inactivation in areas where glial
cells are not present (Dowton et al.,
1988
). Medium-affinity glutamate transporters are present in
epidermal cells in many insects, where they help suppress the levels of
glutamate in the haemoplasm (Tomlin et
al., 1993
). However, local glutamate uptake by glial cells is
likely the main regulator of the transmitter glutamate levels at the
synapse.
In insects, there are many points of contact between the axons and the
muscle fibres. Most insect neuromuscular junctions occur at or near the
surface of the muscle (reviewed by
Osborne, 1970). This is the
case with T. ni, where stained nerve processes were seen only running
over the surface of the skeletal muscles and not plunging deep within them. In
many instances, nerve processes also appear to run along grooves in the muscle
fibre. Insect axons are multi-terminal and may make contact with the fibre
thousands of times (Osborne,
1975
). Judging by the extensive networks of innervating processes
that are stained, even on smaller muscles, it would seem that T. ni
larval skeletal muscles also exhibit this attribute. In our preparations,
numerous circular darkly staining bodies are seen running alongside the axon
in the glial sheath. We interpret these to be areas of high expression of the
transporter by the glial cells at individual neuromuscular synapses. A similar
pattern has been shown in the hornworm Manduca sexta
(Rheuben and Reese, 1978
;
Rheuben, 1985
). Rheuben
described circular profiles of glial processes that alternate with muscle
extensions contacting the nerve where synapses occur. The insect glutamate
transporter appears to be most highly expressed in glial processes between
these synapses. The synaptic distribution of this protein in glial cells
agrees with findings on some mammalian glutamate transporters. The absence of
overall labelling of the glial cell surface indicates that following its
synthesis the transporter protein is rapidly directed to the cell surface
nearest sites of transmitter release. The mammalian GABA transporter, is also
known to be targeted to sub-domains of the cell surface
(Pietrini et al., 1994
).
Considering that the caterpillar glutamate transporter was cloned from a
head cDNA library, it is hardly surprising that the transporter was
subsequently found to be expressed in the CNS of this insect. TrnEAAT1 was
detected in the central neuropile only, and not in the connectives or nerve
branches that communicate from the ganglion to other parts of the insect's
body. Individual glial cell bodies were not stained as was seen when FMRFamide
was localized in this insect's CNS (R.B.G., unpublished). Instead a diffuse
glial distribution in the neuropile was demonstrated. The neuropile contains
numerous axonal connections that require the removal of glutamate after
signaling occurs. This is not the case in connectives and communicating
nerves, which form synapses only when they terminate at a muscle or another
nerve(s). Our findings on the localization of the transporter in the insect
CNS are consistent with the reported localization of glutamate transporters in
the mammalian CNS (for a review, see
Danbolt, 2001). Although some
of these transporters are expressed by neurons (EAAT3), the transporters most
important for glutamate removal are expressed by glial cells (EAAT-1 and
-2).
This is the first description of the cellular pattern of localization of a glutamate transporter protein at the neuromuscular junction and in the CNS of an insect. It will prove interesting to see whether other glutamate transporters are localized in a similar fashion in excitable tissues of other insects. Currently we are examining other insect transporters to determine their localization and co-expression in glial or neuronal cells. These studies should help delineate the roles of these essential proteins in synaptic signaling in insects.
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Acknowledgments |
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