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The Journal of Neuroscience, February 1, 2002, 22(3):931-945
Mechanisms of the Release of Anterogradely Transported
Neurotrophin-3 from Axon Terminals
XiaoXia
Wang1,
Rafal
Butowt1,
Michael R.
Vasko2, and
Christopher S.
von
Bartheld1
1 Department of Physiology and Cell Biology, University
of Nevada School of Medicine, Reno, Nevada 89557, and
2 Department of Pharmacology and Toxicology, University of
Indiana School of Medicine, Indianapolis, Indiana 46202-5120
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ABSTRACT |
Neurotrophins have profound effects on synaptic function and
structure. They can be derived from presynaptic, as well as
postsynaptic, sites. To date, it has not been possible to measure the
release of neurotrophins from axon terminals in intact tissue. We
implemented a novel, extremely sensitive assay for the release and
transfer of anterogradely transported neurotrophin-3 (NT-3) from a
presynaptic to a postsynaptic location that uses synaptosomal
fractionation after introduction of radiolabeled NT-3 into the
retinotectal projection of chick embryos. Release of the anterogradely
transported NT-3 in intact tissue was assessed by measuring the amount
remaining in synaptosomal preparations after treatment of whole tecta
with pharmacological agents. Use of this assay reveals that release of
NT-3 from axon terminals is increased by depolarization, calcium influx
via N-type calcium channels, and cAMP analogs, and release is most
profoundly increased by excitation with kainic acid or mobilization of
calcium from intracellular stores. NT-3 release depends on
extracellular sodium, CaM kinase II activity, and requires intact
microtubules and microfilaments. Dantrolene inhibits the high
potassium-induced release of NT-3, indicating that release of calcium
from intracellular stores is required. Tetanus toxin also inhibits NT-3
release, suggesting that intact synaptobrevin or synaptobrevin-like
molecules are required for exocytosis. Ultrastructural autoradiography and immunolabel indicate that NT-3 is packaged in
presumptive large dense-core vesicles. These data show that release of
NT-3 from axon terminals depends on multiple regulatory proteins and
ions, including the mobilization of local calcium. The data provide
insight in the mechanisms of anterograde neurotrophins as synaptic modulators.
Key words:
neurotrophic factors; secretion; presynaptic terminals; calcium; axonal transport; neurotrophin; synapse; synaptic
transmission
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INTRODUCTION |
Neurotrophins are important
regulators of neuronal differentiation and synaptic plasticity (Lohof
et al., 1993 ; Snider, 1994; Kang and Schuman, 1995; Thoenen, 1995;
McAllister et al., 1999; Poo, 2001). Traditionally, neurotrophins were
believed to be derived by retrograde axonal transport from target
cells, but recent studies revealed that neurotrophins are also
transported anterogradely along axons (von Bartheld et al., 1996; Zhou
and Rush, 1996; Altar et al., 1997; Conner et al., 1997; Heymach and
Barres, 1997; Smith et al., 1997; Yan et al., 1997; Fawcett et al.,
2000; von Bartheld and Butowt, 2000) and may act as extremely potent
excitatory neurotransmitters (Kafitz et al., 1999). Anterograde signals
provide trophic support from afferents (Linden, 1994; von Bartheld et
al., 1996; Altar et al., 1997), may mediate fast, local effects at
synaptic sites (Lohof et al., 1993; Kang and Schuman, 1995; Thoenen,
1995; Kafitz et al., 1999; Poo, 2001), and regulate dendritic growth,
neuronal cytoarchitecture, and phenotypes (Altar et al., 1997;
McAllister et al., 1999; Fawcett et al., 2000).
Neurotrophin 3 (NT-3) enhances synaptic transmission in neuromuscular
junctions (Lohof et al., 1993) and the brain (Kim et al., 1994), alters
short-term and long-term plasticity (Kang and Schuman, 1995; Kokaia et
al., 1998), and modifies the morphology of cerebellar and cortical
neurons (Lindholm et al., 1993; McAllister et al., 1999). In the
retinotectal system, endogenous and exogenous NT-3 are moved by
anterograde axonal transport (von Bartheld and Butowt, 2000), and
exogenous NT-3 is released from retinotectal terminals and taken up by
tectal dendrites, promoting the survival of postsynaptic neurons (von
Bartheld et al., 1996). Release mechanisms are crucial for an
understanding of the action of neurotrophins (Lo, 1995; Thoenen, 1995;
Fawcett et al., 1997; Aloyz et al., 1999; Berninger and Poo, 1999;
Schinder and Poo, 2000), but mechanisms of the release of anterogradely
transported neurotrophins are completely unknown.
Neuropeptides are released by a variety of mechanisms, presumably by
packaging into different types of vesicles that are released by
different mechanisms (Bartfai et al., 1988; Hille et al., 1999; Kasai,
1999). Therefore, release mechanisms of neurotrophins cannot be
predicted on the basis of what is known for other neuropeptides. The
role of anterogradely transported neurotrophins has been particularly difficult to assess because of the absence of an assay that
localizes the trophic factors within synapses and is sufficiently
sensitive to measure the excessively small amounts released from axon terminals.
We developed a technique that allows us to measure the release and
transynaptic transfer of picomolar amounts of anterogradely transported
NT-3. A novel combination of autoradiographic, biochemical, and
experimental pharmacological techniques in an in vivo model system revealed which molecules and events regulate the release of
anterogradely transported NT-3 from axon terminals. This technology will define the molecular mechanisms regulating neurotrophin release and help to better understand the role of neurotrophins in
neurotransmission and synaptic plasticity.
Parts of this work have been published previously in abstract form
(Wang et al., 1999).
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MATERIALS AND METHODS |
Sources of materials
4-Aminopyridine (4-AP), cadmium chloride, colchicine,
cytochalasin D, cytochrome c, kainic acid, KN-93,
strophanthidin, tetrodotoxin (TTX), and thapsigargin were from Sigma
(St. Louis, MO). 8-Br-cAMP, dantrolene, and forskolin were from ICN
Biomedicals (Cleveland, OH). BAPTA, BAPTA AM,  -conotoxin-GVIA,
KN-92, tetanus toxin, and tetanus toxin C fragment were from Calbiochem
(La Jolla, CA). Nerve growth factor (NGF) was from Mark Bothwell
(Seattle, WA) or Alomone Labs (Jerusalem, Israel). Brain-derived
neurotrophic factor (BDNF) and NT-3 were from Regeneron (Tarrytown,
NY). NT-3 antibody was from Robert Rush (Adelaide, Australia). Calcium
electrodes and calibration buffers were from World Precision
Instruments (Sarasota, FL). Fertilized chicken eggs were from
California Golden Eggs (Sacramento, CA). A total of 3800 chicken eggs
were used in this study; all embryos were staged (Hamburger and
Hamilton, 1951). Protocols were approved by the local animal care
committee, and surgical procedures were done after anesthesia with
sodium pentobarbital (Nembutal; 5 mg/embryo).
Injection of 125I-labeled proteins, trafficking,
and release
Neurotrophins, cytochrome c, and tetanus toxin were
radio-iodinated with lactoperoxidase (von Bartheld, 2001), and
substance P (SP) was iodinated with the Bolton-Hunter reagent (NEN Life Sciences, Boston, MA). Incorporation for NT-3 was ~90-95%, and specific activity was 90-150 cpm/pg. Specific activities for NGF were
75-189 cpm/pg, for cytochrome c were 36-72 cpm/pg, for
tetanus toxin were 40-61 cpm/pg, and for substance P were 18 cpm/pg.
Fertilized chicken eggs were incubated at 37°C. Embryos were windowed
on embryonic day 15 (E15) to E16, injected with 40-80 ng of
125I-labeled NT-3 in the eye in
ovo on E16-E17 (von Bartheld et al., 1996), and were perfused
intracardially 20 hr later (at E17-E18) with ice-cold nondepolarizing
buffer [low K+ Krebs' buffer (in
mM): 134 NaCl, 5 KCl, 1 MgSO4 · 7 H2O, 1.25 KH2PO4, 2.0 CaCl2 · 2 H2O, 16 NaHCO3, and 10 glucose, pH 7.5 (after gassing
with 95% O2, 5%CO2)] to
remove radioactivity that had leaked into systemic circulation. The
perfusion was also used to deliver pharmacological agents directly to
the brain and to improve penetration. The tectum was removed and placed
in cold low K+ buffer, and its
radioactivity was counted in a gamma counter. Only tecta containing
>6000 cpm (n = 694) were used for subsequent release
experiments, because such tecta typically generated >200 cpm in
synaptosomes and we found that ~200 cpm were necessary to reliably
quantify synaptosomal peaks. Each dissected tectum was incubated in 3 ml of oxygenated low K+ buffer (6 mM K+) or oxygenated
high K+ buffer (56 mM K+) (in
mM: 84 NaCl, 55 KCl, 1 MgSO4 · 7 H2O, 1.25 KH2PO4, 2 CaCl2 · 2 H2O, 16 NaHCO3, and 10 glucose) at room temperature with
or without experimental drugs for 60 min, unless indicated otherwise. Maximal transport capacity in the retinotectal projection was tested by
injection of 2-400 ng (~0.5-30 × 106 cpm in the eye at the time the animal
was killed) and gamma-counting of tecta after 20 hr. The amount
of injected 125I-labeled NT-3 in the eye
and the amount of anterogradely transported NT-3 in the tectum was
quantified; in addition, the amount of the radioactivity in the
synaptosomal peaks was taken as a measure for the amount and percentage
of 125I-labeled NT-3 within the
presynaptic terminals [excluding the amount in the general membrane
fraction containing the bulk of tectal radioactivity: axons and
dendrites (von Bartheld et al., 1996)].
Synaptosome preparation, data collection, and analysis
Synaptosomes were prepared to measure how much
125I-labeled NT-3 remained in presynaptic
terminals. The protocol of Viglietti et al. (1977) was used with slight
modifications. The incubation buffer was replaced with 5 ml of 0.32 M sucrose and 5 mM Tris, pH 7.4 (buffer B);
tecta were homogenized by trituration using a 1 ml syringe with a 20 G
1 1/2 gauge needle. The suspension was centrifuged at 4°C for 10 min
at 1500 × g to remove the crude nuclear fraction (P1).
The pellet was washed two times with 1 ml buffer B, and supernatants
were removed and centrifuged at 17,000 × g for 20 min
to pellet the membrane fraction (P2). Pellet 1 and supernatant (S2)
were counted in a gamma counter. P2 was resuspended in 10 ml of 0.32 M sucrose and loaded on a two-step discontinuous
ficoll-sucrose gradient in Quick-Seal tubes containing 15 ml of 13%
(w/v) ficoll in 0.32 M sucrose and overlaid with 15 ml of 7.5% (w/v) ficoll in 0.32 M sucrose.
The samples were centrifuged at 53,000 × g for 90 min
at 4°C. The ultracentrifuge tubes were fractionated (50 × 0.8 ml fractions), and each fraction was counted for 10 min in a gamma
counter. The radioactivity within the synaptosomal peak (defined by
visual inspection of the printed plots) was calculated as the ratio
(divided by the total radioactivity in the ultracentrifuge tube). The
average ratio of the synaptosomal peak in low
K+ buffer was defined as 100% unless
indicated otherwise. The ratio of the synaptosomal peak/tube in the
experimental condition was divided by the average ratio of the
synaptosomal peak/tube in the low K+
buffer to calculate the percentage of NT-3 released from retinotectal terminals. Statistical significance was determined by unpaired Student's t test or, for multiple comparisons, by one-way
ANOVA with Prism 2.01 software.
Proteins extracted from synaptosomal fractions derived from
125I-NT-3-loaded tecta were examined by
SDS-PAGE (15% gels). To examine whether release of NT-3 from
synaptosomal fractions (obtained from
125I-NT-3-loaded tecta) can be measured,
synaptosomes were centrifuged and resuspended in either high or low
K+ buffer, warmed to 37°C, gassed for 60 min, and centrifuged again in either a microcentrifuge or the
ultracentrifuge. The pellets and supernatants were gamma counted to
determine release of 125I-NT-3 into the buffer.
Pharmacological manipulations of release
A total of 694 successful release experiments were performed to
test molecules potentially involved in release of NT-3. Control experiments (low K+ or high
K+) were included in each experimental
series of up to eight centrifugation tubes for calibration of NT-3
release ("quality control"). For manipulation of release in
vitro, the dissected tecta were incubated with 3 ml of high
K+ buffer containing one of the following
agents: 4-aminopyridine (1 mM), 8-Br-cAMP (50 µM), cadmium chloride (100 µM), colchicine (175 µM), cytochalasin D (0.2 µM), dantrolene (50 µM), forskolin (10 µM),
KN-93 (10 µM), KN-92 (10 µM), nifedipine (10 µM), -conotoxin-GVIA (0.1 µM), strophanthidin (500 µM), tetrodotoxin (3 µM), or thapsigargin (5 µM). When drugs required dissolving in DMSO or
ethanol, control tecta were tested with identical vehicle solution.
BAPTA. Buffers were prepared as described, except that
CaCl2 was replaced with
MgCl2 at the same concentration. BAPTA AM (10 µM) or BAPTA (20 or 200 µM) was added to the calcium-free buffers overnight before use. Higher BAPTA concentrations were necessary because tecta were a source of calcium that leaked into the incubation solution. Calcium concentrations in the incubation buffer were measured
with and without tecta, and with normal calcium, or calcium-free buffers containing 20 µM BAPTA, 200 µM BAPTA, or 10 µM
BAPTA AM. Calcium electrodes and calibration buffers were used as
described previously (Baudet et al., 1994).
N-Methyl-D-glucamine.
N-Methyl-D-glucamine (NMG) was used to
replace extracellular sodium in high K+
buffer (in mM: 100 NMG-HCl, 39 KCl, 1 MgSO4, 1.25 KH2PO4, 2 CaCl2, 16 KHCO3, and 10 glucose). Chick embryos were perfused with high K+ buffer containing NMG, dissected optic
tecta were incubated in NMG buffer for 60 min, and the tissue was
further processed as described. Some NMG-treated tecta were incubated
in normal (Na-containing) high K+ buffer
to determine whether the NMG effect was reversible.
Tetanus toxin. Tetanus toxin (1-4 µg) was injected into
the eyes 1 d before NT-3 injection to target the toxin by
anterograde axonal transport to the retinotectal nerve terminals.
Whereas 4 µg was lethal, 1-2 µg was tolerated. The tetanus toxin C
fragment (responsible for trafficking of the molecule) was
radio-iodinated with lactoperoxidase to a specific activity of 40-61
cpm/pg. Free iodide was removed by membrane centrifugation (von
Bartheld, 2001). Radiolabeled toxin C fragment (300 ng) was injected
into the eye of chick embryos, and the tecta were processed for
autoradiography to confirm that the toxin was transported anterogradely
by retinal ganglion cell (RGC) axons to the optic tectum.
125I-NT-3 was coinjected with (cold)
tetanus toxin to test whether the toxin interfered with the normal
anterograde transport of NT-3. The tecta, loaded with tetanus toxin and
125I-NT-3, were treated with high
K+ buffer and processed as described.
Control experiments
To determine whether NT-3 was released constitutively during the
incubation in low K+ buffer, the amount of
NT-3 in retinotectal terminals was compared between tecta processed
immediately after killing, and tecta were incubated for 60 min in
oxygenated low K+ buffer. To determine
whether NT-3 or other neurotrophins or peptides bound to terminals
during the preparation of synaptosomes, the tectal homogenates of
normal (not injected) chick embryos were incubated with 0.1-0.3 ng/ml
radio-iodinated NT-3, BDNF, NGF, SP, or cytochrome c and
subjected to ficoll-sucrose gradient centrifugation, and gamma counts
were plotted after fractionation. Some NT-3 preparations were incubated
with 500-fold excess cold NT-3 to determine specific, receptor-mediated
binding. Total protein content of synaptosomal peaks was measured by UV
absorption [Bio-Rad (Hercules, CA) standard protein assay].
Anterograde transport and release of other proteins
To compare the transport and release of NT-3 with that of other
proteins, we measured NGF, tetanus toxin C fragment, and cytochrome c. These proteins were chosen because, unlike substance P,
they can be transported in the retinotectal system after intraocular injection. The amount of radiolabeled protein injected in the eye was
measured 20 hr later to assess retention in the eye. The transport
efficiency was measured by comparing the amount in the eye at the time
the animal was killed with the amount contained in the contralateral
tectum. By competing with excess cold protein (20- to 50-fold), it was
examined whether anterograde transport was receptor mediated.
Accumulation of the transported protein in retinotectal terminals was
assessed by measuring the radioactivity in the synaptosomal peak and
comparing it with the total radioactivity in the contralateral tectum.
In vivo release of NT-3
Kainic acid (8 µl, 5 mM) was injected into the eye
(estimated concentration in the eye, 0.625 mM) of
125I-NT-3 treated chick embryos 45 min
before dissection of the tecta. To determine whether the effect of the
kainic acid could be blocked by TTX, 5 µl of TTX (100 µM) or PBS (as a control for the TTX application) was
injected into the eyes, followed by 8 µl of kainic acid (5 mM) 15 min later. The embryos were anesthetized 45 min later and perfused intracardially, and the tecta were processed as described.
Release of substance P from tectal terminals
Radio-iodinated NT-3 was injected in the eye 20 hr before
homogenization of the tecta to identify the fractions containing the
synaptosomal peak. Tecta were treated with either low or high K+ buffer as described. Fractions were
counted in the gamma counter for 1 min. HCl was added to 3 ml fractions
containing synaptosomes, and samples were stored immediately at
 80°C. Samples of 500 µl were analyzed by radioimmunoassay for the
content of SP (Chen et al., 1996).
Vesicle purification
Tecta from E17 chick embryos loaded with
125I-labeled NT-3 were dissected and
homogenized at 4°C in 15 ml of buffered sucrose (4 mM
HEPES and 320 mM sucrose, adjusted to pH 7.3 with NaOH). Vesicles were purified according to Huttner et al. (1983) and Han and
Fischbach (1999). After centrifugations, resuspended vesicles (6.7 ml)
were loaded on sucrose step gradients in 40 ml Quick-Seal centrifuge
tubes, loaded with 6.75 ml each of 50, 200, 400, 550, and 800 mM sucrose (from bottom to top), and centrifuged at 24,500 rpm (42.1 rotor; 65,000 × g) for 5 hr. Fractions were
collected, counted, and plotted as described for synaptosome fractionations.
Electron microscopic autoradiography and immunocytochemistry
Localization and quantification of anterogradely transported
125I-NT-3 in tectal tissue was performed
by autoradiography as described previously (von Bartheld et al., 1996;
von Bartheld, 2001). Five hundred thirty silver grains from two E17
tecta with 125I-NT-3-loaded retinotectal
terminals were analyzed. Silver grains overlying presynaptic or
postsynaptic compartments were quantified, and the percentages of
silver grains were calculated. In addition, 87 silver grains within the
stratum opticum were analyzed for association with either endosomes or
large dense-cored vesicles (LDCVs) within axons. Synaptosomes from
125I-NT-3-loaded tecta were fixed in 2.5%
glutaraldehyde and centrifuged at 10,000 × g at 4°C,
embedded in Spurr, and sectioned at 80-90 nm. Sections were coated
with a monolayer of Ilford L4 emulsion and exposed for 8-10 weeks at
4°C. Developed sections were examined on a Philips CM10 transmission
electron microscope. The distribution of silver grains was
photographed, and 94 random silver grains were quantified at a
magnification of 15,000-20,000×. For immunolabeling, tectal tissue
from E19-E20 embryos was fixed in 4% paraformaldehyde and 0.1%
glutaraldehyde by intracardial perfusion. Standard pre-embedding immunolabeling techniques were used. Tecta were sectioned with a
vibratome at 100-150 µm. Endogenous peroxidase activity was quenched
by pretreatment with 3%
H2O2. The sections were
incubated for 24 hr at 4°C with 20 µg/ml purified polyclonal
antibodies to NT-3 (Zhou and Rush, 1993) (courtesy of R. Rush, Flinders
University, Adelaide, Australia) and processed with the ABC kit
(Vector Laboratories, Burlingame, CA). Sections were embedded in Spurr
and thin sectioned at 80-90 nm. In control sections, the primary
antibody was omitted.
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RESULTS |
Anterogradely transported 125I-NT-3 in tectal terminals
can be quantified
Injections of 40-80 ng of 125I-NT-3
in the eye resulted in a total of ~100 pg of
125I-NT-3 in the contralateral tectum. The
synaptosomal peaks contained 5.94 ± 0.36% (SEM;
n = 19) of the total radioactivity measured in the
contralateral tectum, equivalent to 15-20% of the radioactivity in
the ultracentrifugation tube (Fig.
1A). Much of the
radioactivity was found in the general membrane fraction, containing
125I-NT-3 in retinal axons en route to
their terminals in the tectum (von Bartheld et al., 1996). There was no
radioactive peak in the synaptosomes from the ipsilateral (control)
tecta, indicating that accumulation of
125I-NT-3 in tectal synaptosomes was not
attributable to diffusion or uptake after leakage into systemic
circulation, and 125I-NT-3 arrived in the
tectum intact. The localization of anterogradely transported
125I-NT-3 in synaptosomes (after
fractionation) was confirmed by quantitative autoradiography at the
ultrastructural level. Synaptosomes were identified in expected quality
and quantity ( 60-70% purity) (Viglietti et al., 1977), and >60%
of silver grains were located over clearly identified synaptosomes
(Fig. 1B,C). The identity of
synaptosomal fractions was verified by increased levels of synaptic
vesicle protein (SV2) in Western blots (data not shown). The
association of radiolabeled NT-3 in synaptosomal peaks with retinotectal terminals (rather than other tectal terminals) can be
concluded because of the fact that the highest level of
autoradiographic labeling in the tectum in situ occurs in
the layer known to contain the retinotectal terminals (von Bartheld et
al., 1996). To determine whether 125I-NT-3
in multivesicular bodies (MVBs) of postsynaptic dendrites may
contribute to the radioactivity in the synaptosomal peak, silver grains
over MVBs were quantified. Only ~2% of silver grains labeled MVBs
(Fig. 1C), ruling out significant contamination by these and
other postsynaptic elements. These data confirm that synaptosomal
fractionation measures the content of anterogradely transported
125I-NT-3 in retinotectal terminals.
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Figure 1.
Quantification of radiolabeled NT-3 in
retinotectal terminals after anterograde axonal transport.
A, Quantification of 125I-NT-3 in the
synaptosomal peak after centrifugation on ficoll-sucrose gradients and
fractionation. The synaptosomal peak (SY) and the general
membrane fraction (GM) are indicated. B,
Synaptosome shows a silver grain (arrowhead) next to a large
dense core vesicle (arrow). C, Quantification of
silver grains over synaptosomes shows majority of grains over
synaptosomes. Silver grains (n = 94) were randomly
analyzed. D, Synapse before release of NT-3. Cluster of
silver grains over a presynaptic terminal in layer f of the stratum
griseum et fibrosum superficiale (SGFSf) of the optic tectum after
anterograde transport. Note the synaptic profile
(S). E, Synapse in the process of
releasing NT-3. Silver grains over a presynaptic profile
(S) containing a large dense core vesicle
(arrow) and over a postsynaptic organelle reminiscent of a
multivesicular body (arrowhead). F, Synapse after
release of NT-3. Silver grain on the postsynaptic side of the synapse
(S) overlying a multivesicular body
(MVB). Scale bars, 200 nm.
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125I-NT-3 in presynaptic terminals in
situ: quantitative electron microscopy
To verify the amount of 125I-NT-3 in
the presynaptic retinotectal terminals in situ, quantitative
autoradiography was performed at the ultrastructural level. Typical
examples of silver grains over presynaptic and postsynaptic profiles
are shown in Figure 1D-F. A total of 530 silver
grains from two chick tecta were sampled randomly, and their location
was analyzed at a magnification of 24,400×. In many (42.5%) but not
all of the labeled terminals, LDCVs were visible in close vicinity
(within 100 nm) of the silver grains (Fig. 1E),
consistent with the notion that anterograde neurotrophins localize in
LDCVs (Fawcett et al., 1997; Michael et al., 1997; Holstege et al.,
1999). The percentage of silver grains over presynaptic terminals was
5.48% (range, 4.72-6.17%) of the total 530 grains, and 3.15% were
over postsynaptic terminals (Fig.
1E,F). The percentage for
presynaptic localization of 125I-NT-3
(5.48%) is remarkably similar to that obtained by synaptosomal fractionation (5.94%). Although we cannot rule out coincidence, this
similarity supports the validity of our assay for the measurement of
exogenous NT-3 in presynaptic axon terminals.
125I-NT-3 in synaptosomes is intact by SDS-PAGE
At least some of the NT-3 that arrives in the tectum after
anterograde axonal transport from the eye is intact (von Bartheld et
al., 1996), but it was not known whether this included the 125I-NT-3 within synaptosomes. Protein
extracted from synaptosomal fractions was examined by SDS-PAGE. The
large majority of 125I-NT-3 recovered from
synaptosomes was intact compared with native 125I-NT-3 (Fig.
2A). When treated with
detergents and trypsin, a major fraction of NT-3 in the synaptosomal
fraction was degraded, indicating that NT-3 was located within a
membrane-confined compartment (data not shown). These results indicate
that anterogradely transported NT-3 within synaptosomes remains
intact.
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Figure 2.
NT-3 in synaptosomes:
K+-induced-release of NT-3 and other proteins from
axon terminals and binding of neurotrophins and neuropeptides to
synaptosomes. A, SDS gel shows that the large majority of
NT-3 recovered from synaptosomes (Synapt.) migrates
identically to native NT-3 at ~14 kDa. The arrow indicates
minor degradation products. Three month exposure. B, Plot of
anterogradely transported 125I-NT-3 in the contralateral
tectum (cpm) as a function of the amount of 125I-NT-3
remaining in the injected eye. C, Plot of the
125I-NT-3 in the synaptosomal fraction from the
contralateral tectum (filled circles) as a function of the
amount of 125I-NT-3 remaining in the injected eye. When the
125I-NT-3 in the eye was competed with 20-50-fold excess
cold NT-3, only background cpm were found in the synaptosomes
(open circles). Note that the amounts of NT-3 in the tectum
and in the synaptosomal peak saturate at ~10 × 106-15 × 106 cpm in the eye.
D, Representative examples show decreased size of
synaptosomal peaks (SY) from tectum depolarized in high
K+ buffer (right panel) compared with low
K+ buffer (left panel). E,
Quantification of D reveals a 15-20% decrease
in NT-3 content in depolarized terminals (HiK, high
K+ buffer; ***p  0.001, ANOVA,
error bars indicate SEM). The average of the values for low
K+ buffer (LoK, 60 min) was taken as 100%.
Comparison of incubation for 60 min versus 0 min in low
K+ shows lack of basal (constitutive) release.
Numbers of experiments are indicated on bars. F, Release of
radiolabeled tetanus toxin C fragment (Tet tox) was not
statistically significant in high K+ buffer.
G, Radiolabeled cytochrome c
(Cyt. C) was not released in high
K+ buffer compared with low K+
buffer. H, Depolarized tecta showed a 25-50% release of
substance P (SP) from tectal terminals, measured by
radioimmunoassay in 500 µl samples (1:6 dilution).
*p 0.05, t test, HiK versus LoK,
60 min, error bars indicate SEM. I, When homogenized
tecta were incubated with radiolabeled NT-3, NGF, or cytochrome
c (Cyt C), NT-3 bound to tectal terminals
which was competed by excess cold NT-3, whereas NGF and cytochrome
c showed very little binding. SY,
Synaptosomal peak; GM, general membrane fraction.
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Direct release from synaptosomes using the conventional assay
We attempted to measure release of anterogradely transported
125I-NT-3 directly from synaptosomes by
using a conventional static release protocol (Androutsellis-Theotokis
et al., 1996), with the modification that
125I-NT-3 was loaded into tectal
synaptosomes via anterograde transport from the retina in
vivo. After injections of 50-100 ng of
125I-NT-3 into the eye, ~3-4 pg
(450-600 cpm) of 125I-NT-3 was present in
the synaptosomal fraction, but only a fraction of this amount (~1 pg
100-200 cpm) was recovered per tectum after washes, resuspension,
and warming to 37°C. The radioactivity in the supernatant after
centrifugation ranged from 30 to 60%, and this amount was not
increased when synaptosomes were resuspended in high
K+ buffer instead of low
K+ buffer (data not shown). The
radioactivity in the supernatant likely represents
125I-NT-3 that leaked out of damaged
immature synaptosomes, thus yielding a relatively high and variable
background that made it impossible to measure secreted
125I-NT-3. The radioactivity levels in
this prep were approximately three orders of magnitude lower than what
is usually used for synaptosomal release studies
(Androutsellis-Theotokis et al., 1996), and it was impossible to
increase the amount of anterogradely transported
125I-NT-3 per tectum, because its
transport into synapses saturated at ~10 × 106 cpm/eye (Fig.
2B,C). We conclude that the
measurement of anterogradely transported
125I-NT-3 release directly from
synaptosomes is impossible with the conventional technique.
Depolarization of tectal terminals induces release of NT-3
Next we tested whether release of NT-3 from terminals in intact
tissue can be measured by quantifying the amount remaining within terminals rather than measuring the amount that is released. Tecta whose retinotectal terminals had been loaded with anterogradely transported 125I-NT-3 were incubated in
oxygenated buffer containing high K+ (56 mM), whereas control tecta were incubated in low
K+ (5 mM), and the amount of
125I-NT-3 remaining in terminals was
quantified by measuring the amount in the synaptosomal fraction
relative to the total amount in the tectum or the total amount in the
ficoll-sucrose gradient tube. Depolarization resulted in smaller
synaptosomal peaks of 125I-NT-3 (Fig.
2D), equivalent to a loss of 15-20% of the
radioactivity in the terminals after depolarization
(p 0.001; ANOVA; multiple comparisons). These
data were normalized to the total amount of radioactivity in the
gradient ultracentrifugation tube. With an input of 40-80 ng of
125I-NT-3 in the eye, we found that
~8000-12,000 cpm was transported to the tectum, and 6% (~500 cpm)
of this amount was reliably measured in the synaptosomal peak (Table
1). No basal release of NT-3 was detected
during the 60 min incubation step in low
K+ buffer (Fig. 2E). To
determine whether the release of 125I-NT-3
was specific or merely reflected the loss of 10-15% of presynaptic
protein attributable to stimulated release, the total protein content
of synaptosomal preparations was compared between depolarized and
control tecta. There was no significant difference between the two
groups (data not shown), indicating that NT-3 was specifically
released. These data show that the novel release assay has significant
advantages over the conventional assay in terms of recovery,
reliability, and sensitivity, primarily because the synaptosomes are
used merely to determine the amount of radiolabeled NT-3. Because the
release has occurred in primarily intact tissue (the tectum),
the assay does not require the synaptosomes to be functionally intact,
which is a major challenge for immature synaptosomes prone to leakage
at 37°C.
Control experiments with other transported proteins: limitations of
the new assay
To determine whether the release of
125I-NT-3 was specific or whether other
proteins were released similarly when measured with the same assay, we
compared the transport, synaptic accumulation, and release of NT-3,
NGF, tetanus toxin, and cytochrome c. The novel assay
requires that the protein is retained in the eye (to be internalized by
retinal ganglion cells), that it be transported to the terminals in the
tectum with sufficient efficiency, and that it accumulate in the
retinotectal terminals. The data showing how these three requirements
are met for five different proteins are summarized in Table 1.
Cytochrome c showed the highest retention in the eye (47%)
after 20 hr, and HSP showed the lowest (2%). The transport
efficiencies of NT-3, NGF, and cytochrome c were similar
(1-2 pg/ng in the tectum retained in the eye) and was significantly
more efficient for tetanus toxin (4 pg/ng in the eye). The greatest
differences between the proteins were found with synaptic accumulation:
NT-3 accumulated in terminals almost six times more than NGF, whereas
tetanus toxin and cytochrome c accumulated only slightly
more than NGF (Table 1). These data provide evidence for some
remarkable differences in the trafficking of these proteins in the
retinotectal system. When we attempted to determine release of NGF in
high K+ buffer, it turned out that the
counts per minutes amounts in the synaptosomal fractions were too low
to be reliably quantified (Table 1). Release of tetanus toxin could be
quantified in this system; although there was a trend for lower peaks
with depolarization, the difference did not reach statistical
significance (Fig. 2F). Release of
125I-cytochrome c could also be
quantified, and there was no release of this protein with high
K+ buffer (Fig. 2G). NGF
transport appeared to be more efficient than in a previous study (von
Bartheld et al., 1996), probably because here we measured radioactivity
exclusively in the tectum, and free iodine and degradation products
were not removed by dehydration of the tissue. Together, these data
show that 125I-NT-3 is specifically stored
in retinotectal terminals, and this capacity makes it possible to
quantify release of NT-3, which is released during depolarization,
unlike other proteins that can also be transported to retinotectal
terminals but do not accumulate there. Requirements and limitations of
our novel release assay thus include the following: (1) internalization
by projection neurons; (2) anterograde axonal transport; (3)
accumulation of the transported protein in presynaptic terminals; and
(4) release of the protein from the terminal. The utility of this assay
may be useful for any protein and neuronal projection system in which sufficient amounts of protein can be quantified, and the novel assay
thus may not be restricted to NT-3 and the retinotectal system of chick embryos.
Control experiments with substance P
A fraction of retinotectal projections in chicken use SP as a
neurotransmitter (Yamagata and Sanes, 1995), and release of SP from
synaptosomes can be induced by treatment with high
K+ (Lembeck et al., 1977). Radiolabeled SP
was very poorly transported anterogradely by RGCs after injection in
the eye (Table 1), presumably attributable to the lack of SP receptors
on RGCs that transport endogenous SP, and therefore it was not possible
to measure release of (exogenous) SP from retinotectal terminals. To
determine whether depolarization of tecta induces release of endogenous
SP from tectal terminals, similar to that of anterogradely transported NT-3, synaptosomal fractions were prepared from
125I-NT-3-loaded tecta (to identify the
synaptosomal peaks), and the amount of SP in the synaptosomal peak
after depolarization (or maintenance of tecta in low
K+ buffer as controls) was measured by
radioimmunoassay (Chen et al., 1996). Approximately 25-50% of SP was
released by depolarization (Fig. 2H), which is
similar to amounts of other neuropeptides released from synaptosomes
(25%) (Verhage et al., 1991b). These data confirm the validity of our
release protocol by comparison with a classical neuropeptide.
Control experiments with added neurotrophins or
cytochrome c
Release of neuropeptides exclusively from terminals is
conventionally examined by measuring the release from synaptosomes (Androutsellis-Theotokis et al., 1996) rather than examining the release in intact tissue and then preparing synaptosomes to measure the
amount remaining. To determine whether neurotrophins and other small
peptides bind to synaptosomes prepared from E17 chick tecta, homogenized tecta were incubated for 15-20 min with radiolabeled neurotrophins or radiolabeled cytochrome c, and the tissue
was processed as described. Significant amounts of BDNF and NT-3
accumulated in the synaposomal peak, but very little NGF or cytochrome
c accumulated (Fig. 2I). To test whether
binding of BDNF and NT-3 to synaptosomes was receptor mediated,
radiolabeled BDNF and NT-3 were incubated in the presence of 500-fold
excess cold same trophic factor. Much of the BDNF (data not
shown) and NT-3 (Fig. 2I) bound to synaptosomes in a
receptor-mediated manner. These data indicate that NT-3 receptors [presumably tyrosine kinase receptor C (trkC)] are present on tectal
terminals (not necessarily retinotectal terminals). These data also
show that release of NT-3 from synaptosomes in a static release system
likely would fail or grossly underestimate release, because the
released NT-3 would bind to receptors present on synaptosomal membranes, thus confounding the results.
Calcium dependence of NT-3 release
To determine whether the depolarization-induced release of NT-3
required physiological concentrations of extracellular calcium, 125I-NT-3-loaded tecta were incubated in
calcium-free buffer containing the calcium chelator BAPTA. High
concentrations of BAPTA (200 µM) abolished the high
K+-induced release of NT-3 (Fig.
3A), whereas lower
concentrations (20 µM) did not (Table
2), indicating that the lower
concentration of BAPTA was not sufficient to chelate the calcium that
may leak from the dissected tecta into the buffer (volume of 3 ml). To determine whether this was indeed the case, calcium was measured in
buffers with and without tecta in four conditions (0 µM, 20 µM, and 200 µM BAPTA and 10 µM
BAPTA AM) in calcium-free buffers and in addition in normal calcium
buffers as shown in Table 2. Calcium leakage from tecta increased the
calcium concentration in calcium-free buffers from ~3 to ~6
µM, and this increase was not significantly
attenuated by 20 µM BAPTA. BAPTA at 200 µM reliably reduced the calcium concentration
in these buffers to <1 µM calcium. BAPTA AM at
10 µM did not reduce the extracellular calcium
concentration significantly. We will refer to the 2 mM calcium concentration as "normal calcium,"
to the 2-10 µM calcium concentration as "low calcium," and to buffers with 1 µM
calcium concentration as "calcium-free" buffers. It should be noted
that the effective calcium concentration at the nerve terminals within
the tectum likely differs from the calcium concentration that was
measured in the incubation buffer.
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Figure 3.
Manipulation of NT-3 release from axon terminals
by modulation of calcium, other ions, or second messengers.
A, High K+-induced release of NT-3 was
abolished by BAPTA (BAP), BAPTA/AM (B/AM),
cadmium chloride (CAD), -conotoxin GVIA (CON),
dantrolene (DAN), but not nifedipine (NIF).
Release of NT-3 was elicited by thapsigargin (THA), both in
high and low K+, and to a lesser extent by
4-aminopyridine (4AP). B, Release of NT-3 was
induced by 8-bromo-cAMP (8BR) or forskolin (FOR).
High K+-induced release of NT-3 was reduced by the
CaM kinase II inhibitor KN-93 (KN93), but not by the
inactive analog KN-92 (KN92). Replacing extracellular sodium
with N-methyl glucamine (NMG) significantly
reduced release of NT-3 induced by high K+. High
K+-induced release of NT-3 was unaffected by
tetrodotoxin (TTX) treatment. The Na/K ATPase inhibitor
strophanthidin (STR) induced release in low
K+ buffer. The number of independent experiments is
indicated on bars. Error bars indicate SEM. All experimental data
points were calibrated in the same batch by obtaining control data
without the pharmacological agent. r., release;
n.r., no release. Statistical significance compared with LoK
( ) or HiK (*) was determined by ANOVA with p 0.05 (* or ), p 0.01 (** or ), and
p 0.001 (*** or ).
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Table 2.
Calcium concentrations in incubation buffers with (+ tect)
and without ( tect) a chick optic tectum as a function of the
concentration of the calcium chelator BAPTA and resulting release of
NT-3 from retinotectal terminals
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To examine whether functional voltage-gated calcium channels were
required for depolarization-induced NT-3 release, tecta were incubated
in normal calcium buffer containing 100 µM cadmium chloride, a nonselective blocker of voltage-gated
Ca2+ channels. Cadmium significantly
reduced the K+-evoked NT-3 release (Fig.
3A). To determine whether functional N-type calcium channels
were required for NT-3 release, tecta were incubated in buffer
containing 0.1 µM -conotoxin, an N-type Ca2+ channel blocker. -Conotoxin
significantly reduced NT-3 release (Fig. 3A), consistent
with the localization of N-type calcium channels on presynaptic
terminals (Westenbroek et al., 1992). Nifedipine (10 µM), a blocker of L-type
Ca2+ channels, showed no significant
effect on release (Fig. 3A). These data demonstrate that
high K+-induced release of NT-3 depends on
extracellular calcium, which, during depolarization, enters the synapse
via N-type calcium channels.
Release of neurotrophins from brain slices and cultured neurons depends
on mobilization of calcium from intracellular stores (Blöchl and
Thoenen, 1995; Thoenen, 1995; Griesbeck et al., 1999). To test whether
NT-3 release from axon terminals also required intracellular calcium
release, tecta were incubated in low calcium buffer containing
the membrane-permeable calcium chelator BAPTA AM at 10 µM
(calcium concentrations of 4-6 µM) (Table 2). BAPTA AM
effectively chelates calcium only after membrane penetration, cleavage
by intracellular esterases, and intracellular accumulation (Tsien,
1981). The amount of extracellular calcium (4-6 µM)
remaining in the BAPTA AM-treated solution would have been sufficient
to allow for the release of NT-3 (compare with 20 µM
BAPTA in Table 2). Addition of the relatively small amount of 10 µM BAPTA AM significantly attenuated the release of NT-3
(Fig. 3A). To verify that mobilization of calcium from
intracellular stores affects NT-3 release, three experiments were
performed. Dantrolene, an inhibitor of calcium release from
caffeine-sensitive intracellular stores (Blöchl and Thoenen,
1995), was used at 50 µM in both low calcium
and normal calcium high K+ buffer to
determine whether it would prevent the release of NT-3. In both low
calcium and in normal calcium buffer (as defined in Table 2),
dantrolene completely prevented the release of NT-3 (data shown for low
calcium in Fig. 3A). These experiments indicate that high
K+-induced NT-3 release requires the
release of calcium from intracellular stores (calcium-induced calcium
release). The role of intracellular calcium was further tested by
incubation in 1 mM 4-aminopyridine (4-AP) in
normal calcium buffer, which increases intracellular (cytosolic)
calcium and has been suggested to cause the release of trophic factors
(Grimaldi et al., 2001). 4-AP caused significant NT-3 release (Fig.
3A). Finally, to verify that mobilization of calcium from
intracellular stores and subsequent increase in intracellular calcium
is sufficient to induce release of NT-3, tecta loaded with
125I-NT-3 were incubated in high
K+ buffer containing 5 µM thapsigargin. Thapsigargin induced a greater release of NT-3 than depolarization alone (Fig. 3A). When
thapsigargin was tested in low K+ buffer,
the evoked NT-3 release was of a similar magnitude as seen with
thapsigargin in high K+ buffer (Fig.
3A). It should be noted that thapsigargin acutely induces a
transient calcium spike by mobilizing calcium from intracellular stores. This acute effect is distinct from the chronic effects of
thapsigargin because of the depletion of intracellular calcium stores
that can be seen in dynamic (superfusion) release assays. Such
long-term effects of thapsigargin would not be seen in our release
assay, because NT-3 is not replenished as shown by colchicine experiments (see below). Such considerations have to be kept in mind
when interpreting data obtained with our novel release assay. To verify
that the mobilization of calcium from intracellular stores by
thapsigargin is immediate and does not require 60 min of incubation, we
tested whether NT-3 was released within <10 min incubation. As
expected, the effect of thapsigargin on NT-3 release was
virtually identical within 10 min (data not shown) and 60 min.
Together, these data indicate that mobilization of calcium from
intracellular stores is sufficient to trigger release of NT-3 from terminals.
Role of cAMP and CaM kinase II
cAMP has been implicated in the regulation of release of
transmitters and neurotrophins (Edwards et al., 1988; Thoenen, 1995; Goodman et al., 1996). To determine whether cAMP is involved in NT-3
release, we used the cAMP analog 8-Br-cAMP and forskolin. Tecta were
incubated in low K+ buffer containing 50 µM 8-Br-cAMP. This drug induced NT-3 release (Fig.
3B), consistent with the notion that cAMP is involved in the
release mechanism. NT-3 release was also induced by 10 µM forskolin, an adenylate cyclase activator
(Fig. 3B). Thus, two distinct ways to elevate cAMP levels
showed the same result, namely release of NT-3, further supporting the
notion that cAMP indeed is involved in the regulation of NT-3 release.
Protein kinases regulate synaptic transmission, with the most direct
evidence for Ca2+/calmodulin-dependent
protein kinase II (CaM kinase II) (Llinas et al., 1991; Kelly, 1993).
To test whether CaM kinase II affects NT-3 release, we used the
selective inhibitor, KN-93, at doses which primarily abolish the
activity of CaM kinase II, but not of other kinases (10 µM) (Sumi et al., 1991). As a control, we used
the inactive analog, KN-92 (10 µM). KN-93
significantly reduced high K+-induced NT-3
release, whereas KN-92 had no effect (Fig. 3B). These data
indicate that cAMP and CaM kinase II may act downstream of the
depolarization induced by high K+.
Role of extracellular sodium
The role of extracellular sodium for the release of neurotrophins
is controversial. Initial studies on the release of neurotrophins concluded that extracellular sodium is required for regulated release
of neurotrophins from dendritic and somal compartments (Blöchl
and Thoenen, 1995), but this may have been attributable to an
independent blocking effect of the sodium substitute,
N-methyl-D-glucamine (NMG) (Hoener, 2000).
Release of neuropeptides from terminals (rather than whole cells) does
seem to require sodium influx (Stuenkel and Nordmann, 1993; Thirion et
al., 1999). To determine whether low extracellular sodium may attenuate
NT-3 release from axon terminals,
125I-NT-3 loaded tecta were incubated in
buffer containing high K+, and NMG to
replace > 90% of the extracellular sodium. As seen in Figure
3B, replacement of sodium reduced the release of NT-3. When
NMG was washed out and the buffer replaced with normal sodium, high
K+ buffer induced NT-3 release (data not
shown), indicating that the effect of NMG was reversible. To determine
whether the opposite treatment, increased influx of sodium rather than
decreased influx as with NMG, will have the opposite effect on NT-3
release, we used strophanthidin, a sodium-potassium ATPase inhibitor
(Meyer and Cooper, 1981). As expected, incubation of tecta with 500 µM strophanthidin induced release of NT-3 (Fig.
3B). Release was not attenuated by incubation with
tetrodotoxin (TTX, 3 µM) (Fig. 3B),
indicating that TTX-sensitive sodium channels are not required for the
release process when induced by high K+.
Together, these results may suggest that an unconventional
sodium-dependence of neurotrophin release may be a general feature for
neuropeptide release from axon terminals (Stuenkel and Nordmann, 1993;
Thirion et al., 1999).
Structural proteins involved in NT-3 release
Microtubules and microfilaments have been implicated in the
movement of vesicles from the axon shaft to the membrane at nerve terminals for exocytosis (Westrum et al., 1983; Hirokawa et al., 1989;
Ashton and Dolly, 1991). Microfilaments are thought to act as a barrier
to small synaptic vesicle (SSV) fusion, but they may aid in the
translocation of LDCVs (Doussau and Augustine, 2000) and LDCVs are
known to concentrate outside the active zone (Verhage et al., 1991a).
To determine whether microfilaments are required for NT-3 release,
tecta were incubated with 0.2 µM cytochalasin D. This
drug significantly reduced NT-3 release (Fig.
4A). Microtubules are
present in tectal nerve terminals (Bird, 1989). To determine whether
intact microtubules are required for evoked NT-3 release, 125I-NT-3 loaded tecta were incubated in
high K+ buffer containing 175 µM colchicine. Colchicine abolished the release
of NT-3 (Fig. 4A), indicating that intact
microtubules are necessary for evoked NT-3 release from axon terminals.
The colchicine experiments also show that there is no significant replenishment of NT-3 from axons into terminals during the 60 min
release period. If the released NT-3 was rapidly replenished by
anterograde flow from the axon, one would not expect a net loss of NT-3
in terminals. Because colchicine blocks all anterograde axonal
transport of NT-3 (von Bartheld et al., 1996), we can exclude this
possibility.
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Figure 4.
Manipulation of NT-3 release from axon terminals
with toxins to structural proteins and examination of upstream and
downstream events. A, High K+-induced
release of NT-3 (HiK) was abolished by cytochalasin D
(CYD), colchicine (COL), and pretreatment with
tetanus toxin (TET). Tetanus toxin was targeted specifically
to retinotectal terminals by injection of 1 µg of tetanus toxin in
the eye 24 hr before NT-3 injection. B, Autoradiography
shows that radiolabeled tetanus toxin (125I-TET) was
transported anterogradely by retinal ganglion cells to the optic
tectum. Scale bar, 500 µm. C, Anterograde transport of
125I-NT-3 was not reduced by doses of TET (1-2 µg) in
the eye that were used to target the toxin to retinotectal terminals.
CTL, control. D, Simultaneous treatment with
N-methyl-glucamine (NMG) and thapsigargin
(THA) did not reduce the amount of NT-3 that was released,
indicating that sodium influx may be upstream of THA action.
Cotreatment with BAPTA (BAP) in calcium-free buffer did not
attenuate NT-3 release. Simultaneous treatment with colchicine
(COL) and THA significantly reduced the release of NT-3,
indicating that COL was acting downstream of THA. The number of
independent experiments is indicated on each bar. Error bars indicate
SEM. Statistical significance compared with LoK (), HiK (*) THA only
(§, D) was determined by ANOVA with p 0.05 (*, §), p 0.01 (**), and
p 0.001 (*** or ).
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Role of synaptobrevin/synaptobrevin analogs in NT-3 release
Synaptobrevin and synaptobrevin analogs are required for release
of neurotrophins from cell bodies and/or dendrites (Blöchl, 1998). Synaptobrevin can be cleaved by tetanus toxin (Schiavo et al.,
2000). To determine whether tetanus toxin treatment affects NT-3
release from terminals, tecta were pretreated by intraocular injection
of the whole tetanus toxin (1-2 µg) which is subsequently transported anterogradely to the retinotectal terminals (Manning et
al., 1990). To verify this mode of drug delivery, 300 ng of radiolabeled tetanus toxin C-fragment was injected into the eye of
chick embryos. Within 24-48 hr, 1 ng of the toxin accumulated in the
retinotectal axons and terminals (Fig. 4B).
Subsequent anterograde transport of
125I-NT-3 to the tectum was not affected
by intraocular injection of 1-2 µg tetanus toxin (Fig.
4C). Tetanus toxin in the retinotectal terminals (estimated
5 ng/tectum) reduced the release of NT-3 from tectal terminals (Fig.
4A). This suggests that intact synaptobrevin (or
synaptobrevin analogs) are required for the release of NT-3 from axon
terminals, and that at least some of the release is accomplished by
synaptobrevin-mediated vesicle trafficking and/or exocytosis.
Requirements for the effect of thapsigargin on NT-3 release
Are extracellular sodium, calcium, or intact microtubules required
for the effect of thapsigargin? When tecta were cotreated with NMG and
thapsigargin, there was no significant reduction in the amount of NT-3
that was released (Fig. 4D), indicating that the
effect of thapsigargin did not require extracellular sodium.
Thapsigargin did not require the influx of extracellular calcium for
triggering release, because incubation in calcium-free solution
containing 200 µM BAPTA did not reduce the
magnitude of release (Fig. 4D). When colchicine
treatment was combined with thapsigargin, there was a significant
reduction in the amount of NT-3 that was released (Fig.
4D), indicating that colchicine may act downstream of
the mobilization of calcium from internal stores. These data indicate
that mobilization of calcium from intracellular stores is sufficient to
trigger NT-3 release, but requires intact microtubules, presumably for
vesicle translocation (Doussau and Augustine, 2000).
Kainic acid induces NT-3 release in vivo
Kainic acid and tetrodotoxin increase and decrease, respectively,
the electrical activity of RGCs (Catsicas and Clarke, 1987; Karlsson
and Hallböök, 1998). To determine whether stimulation of
RGCs in vivo induces release of NT-3 similar to the
depolarization of retinotectal terminals in vitro, 1 µg
kainic acid or PBS vehicle control was injected in the eyes of chick
embryos in ovo, 19 hr after intraocular injection of
125I-NT-3. The final concentration of
kainic acid in the eye ( 80 µl volume) was estimated to be 62 µM. Animals were killed 45 min after the
intraocular injection of kainic acid, and their tecta were processed
immediately for synaptosomal fractionation. Kainic acid induced the
release of ~30% of the exogenous NT-3 present in the terminals (Fig.
5). To determine whether this effect of kainic acid could be blocked by treatment with tetrodotoxin (TTX), the
eyes of embryos were pretreated with TTX (0.16 µg injected, final
concentration estimated at 6.25 µM), 20 min
before the injection of kainic acid. TTX treatment abolished the effect
of kainic acid (Fig. 5B). These data confirm in an in
vivo experiment the results from our exposure to drugs by bath
application of whole tecta.
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Figure 5.
Kainic acid induced release of NT-3 from axon
terminals in vivo. Injection of kainic acid
(KA) in the eye increased the amount of NT-3 that was
released from retinotectal terminals compared with PBS-treated controls
(+PBS). The KA-induced release of NT-3 was prevented by
injection of tetrodotoxin (TTX) in the eye 15 min before KA
injection when compared with PBS injections as a control for TTX. The
number of independent experiments is indicated on each bar. Error bars
indicate SEM. Statistical significance compared with +PBS
() or the KA+PBS condition (§§) was determined by ANOVA with
p 0.01 (, §§).
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Is NT-3 packaged in large dense core vesicles?
Our ultrastructural autoradiography localized
125I-NT-3 in the vicinity of large dense
core vesicles (LDCVs) (Fig. 1E), but the resolution
of the 125I-autoradiography was not
sufficient for unambiguous identification of the types of vesicles
which contain NT-3 in presynaptic terminals. Therefore, vesicles of
different sizes were purified by differential sedimentation in sucrose
gradients (Huttner et al., 1983; Han and Fischbach, 1999). Vesicle
purification showed that only the anterogradely transported
125I-NT-3 accumulated in fractions known
to contain LDCVs (Han and Fischbach, 1999), but not the
125I-NT-3 that was incubated with
synaptosomes (Fig. 6A).
After incubation, 125I-NT-3 was marginally
increased in fractions containing SSVs. Because the LDCVs are
heterogeneous in size and the fractions containing LDCVs also contain
endosomes, these data do not prove that the anterogradely transported
NT-3 is exclusively associated with LDCVs, but our data are consistent
with this notion. When the superficial layer of the optic tectum of E17
chick embryos was immunolabeled for endogenous NT-3 and examined at the
electron microscopic level, immunolabel was detected in LDCVs (Fig.
6B). We estimate that ~25% of terminals in
retinorecipient layers of the tectum contained LDCVs, and ~5-6% of
such terminals contained LDCVs with NT-3-like immunoreactivity. No
labeled LDCVs were detected when the primary antibody was omitted (data
not shown). If 125I-NT-3 was present in
LDCVs within retinal axons, one would expect to see silver grains over
LDCVs in the stratum opticum (SO) within the tectum. Indeed, we found
silver grains over and adjacent to dense-cored vesicles within axons of
the SO (Fig. 6C). Because of the relatively small size of
the LDCVs and the fact that many silver grains may obscure them, we did
not attempt to quantify the correlation of silver grains and LDCVs.
Taken together, these results further support the idea that
anterogradely transported neurotrophins are packaged in LDCVs.
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Figure 6.
Association of NT-3 with large dense core
vesicles (LDCV) rather than small synaptic vesicles
(SSV) in retinotectal terminals. A, Vesicle
purification of tecta containing anterogradely transported
125I-NT-3 shows higher levels of radioactivity in
sedimentation fractions known to contain LDCVs in chick embryo brains
(Han and Fischbach, 1999), but not in fractions known to contain SSVs
(top panel). When homogenized tecta were incubated with
125I-NT-3 (bottom panel), radioactivity was seen
in sedimentation fractions known to accumulate SSVs (Han and Fischbach,
1999). B, Section through a retinotectal terminal from a
17-d-old chick embryo immunolabeled with an antibody to NT-3, processed
with horseradish-peroxidase, and visualized with diaminobenzidine. Note
reaction product in a large dense core vesicle (arrow).
Scale bar, 100 nm. C, Section through the stratum opticum
(SO) of the optic tectum after injection of
125I-NT-3 in the eye. Note a cluster of silver grains in
the vicinity of a large dense core vesicle (arrow) in
sectioned retinotectal fibers. Scale bar, 500 nm.
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DISCUSSION |
Our study provides first direct evidence for mechanisms of release
of anterogradely transported neurotrophins from axon terminals. We
found that NT-3 is released by depolarization, that this release depends on intracellular and extracellular calcium, and can be evoked
by mobilization of calcium from intracellular stores. Release is
further regulated by cAMP and CaM kinase II. We show that release occurs by exocytosis of presumptive large dense-core vesicles (LDCVs).
Anterogradely transported and presynaptically located neurotrophins are
released by changes in local, intracellular calcium concentrations.
Technical considerations
Previous studies used immunological techniques to measure the
amounts of neurotrophins released from entire cells (cell bodies, dendrites and axons) into medium or perfusate (Blöchl and
Thoenen, 1995; Goodman et al., 1996), but this technology was not
sufficiently sensitive to measure the excessively small amounts
(molecules per synapse) exclusively released from axon terminals.
Therefore, it is important to consider whether we accurately measured
the amount of 125I-NT-3 in retinotectal
terminals by synaptosomal fractionation and gamma counting. When
synaptosomal measurements were compared with quantitative analysis of
the in situ distribution of silver grains after
autoradiography at the ultrastructural level, the morphological and the
biochemical/pharmacological data showed a remarkable agreement for the
estimates of exogenous NT-3 in axon terminals (5.48% vs. 5.94%).
Another important issue is whether anterogradely transported exogenous
NT-3 is released by the same mechanisms as anterogradely transported
endogenous NT-3. Do endogenous and exogenous neurotrophins pass through the same organelles? RGCs in chicks express NT-3, and
endogenous NT-3 is anterogradely transported to the terminals in the
optic tectum (von Bartheld and Butowt, 2000). Proteins destined for
anterograde transport have to be processed in the Golgi system
(Hammerschlag et al., 1982). Exogenous NT-3 (but very little BDNF)
accumulates in the Golgi system of RGCs (Butowt and von Bartheld, 2001;
von Bartheld et al., 2001) and transcytoses RGCs in a receptor-mediated
manner, possibly by sequential binding to trkC and p75 receptors
(Butowt and von Bartheld, 2001). Therefore, it is likely that exogenous
NT-3 joins the pathway normally taken by endogenous NT-3.
Alternatively, there may be parallel pathways of release, one for newly
synthesized NT-3, and one for transcytosed NT-3 (von Bartheld et al.,
2001).
Our release assay has significant advantages (sensitivity and synapse
specificity) as well as limitations compared with conventional release
assays. It requires sufficient internalization, anterograde axonal
transport, and accumulation of the protein in presynaptic terminals,
and it is not a dynamic assay with replenishment of the source.
Nevertheless, its utility is not necessarily restricted to the
retinotectal system or to neurotrophins.
Molecular determinants of NT-3 release
Our results discern 3 major steps that lead to the release of NT-3
from axon terminals (Fig. 7). These
conclusions are based on data obtained with a variety of
pharmacological agents at concentrations commonly used for tissue
slices. The first step involves depolarization of the axon terminus
with influx of calcium through N-type calcium channels. The second step
may be induced by calcium influx and/or mobilization of calcium from
intracellular stores. Increased calcium concentrations in the
presynaptic compartment (Verhage et al., 1991a), as well as kinase
activation (cyclic AMP-dependent kinases and CaM kinase II), may
provide signals for vesicle translocation via microtubules to the
active zone at the synaptic cleft. The vesicle then fuses with the
membrane, presumably mediated by synaptobrevin, because release is
tetanus-toxin-sensitive. The neurotrophin is released and may diffuse
in the synaptic cleft to bind to presynaptic and/or postsynaptic
neurotrophin receptors (Levine et al., 1995; Wu et al., 1996; Kafitz et
al., 1999; von Bartheld et al., 2001). The role of sodium influx for
neurotrophin release from whole cells is controversial (Blöchl
and Thoenen, 1995; Hoener, 2000). We find that NT-3 release from
terminals requires sodium influx, apparently via TTX-insensitive sodium
channels, consistent with reports of TTX-insensitive, sodium-evoked
release of neuropeptides from axon terminals (Stuenkel and Nordmann,
1993) and sodium influx into docked vesicles (Thirion et al., 1999).
Sodium influx may also interact with mobilization of calcium from
internal stores (Berridge, 1998).
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Figure 7.
Diagram summarizes the proposed mechanisms of
release of anterogradely transported NT-3 from axon terminals.
Depolarization by high potassium (K+) opens N-type
calcium channels. Calcium influx induces calcium release from internal
stores. Increased intraterminal calcium concentrations are sufficient
to induce release of NT-3 containing vesicles. Release can also be
induced by cAMP and requires CaM kinase II activity. Second messengers
and kinases may act directly on vesicle release or indirectly via
calcium mobilization from internal stores (Hille et al., 1999).
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ABSTRACT
INTRODUCTION
