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Next Article 
Volume 16, Number 12,
Issue of June 15, 1996
pp. 3791-3797
Copyright ©1996 Society for Neuroscience
Neural Agrin Activates a High-Affinity Receptor in C2 Muscle
Cells that Is Unresponsive to Muscle Agrin
David C. Bowen1,
Janice Sugiyama2,
Michael Ferns3, and
Zach W. Hall2
1 Regeneron Pharmaceuticals, Tarrytown, New York 10591, 2 National Institute of Mental Health, National Institutes
of Health, Bethesda, Maryland 20892, and 3 Department of
Neurosurgery, Montréal General Hospital Research Institute,
Montréal, Québec, Canada H3G 1A4
 ABSTRACT
- INTRODUCTION
- MATERIALS AND METHODS
- RESULTS
- DISCUSSION
- FOOTNOTES
- REFERENCES
ABSTRACT 
During synaptogenesis, agrin, released by motor nerves, causes the
clustering of acetylcholine receptors (AChRs) in the skeletal muscle
membrane. Although muscle  -dystroglycan has been postulated to be
the receptor for the activity of agrin, previous experiments have
revealed a discrepancy between the biological activity of soluble
fragments of two isoforms of agrin produced by nerves and muscles,
respectively, and their ability to bind -dystroglycan. We have
determined the specificity of the signaling receptor by investigating
whether muscle agrin can block the activity of neural agrin on intact
C2 myotubes. We find that a large excess of muscle agrin failed to
inhibit either the number of AChR clusters or the phosphorylation of
the AChR induced by picomolar concentrations of neural agrin. These
results indicate that neural, but not muscle, agrin interacts with the
signaling receptor. Muscle agrin did block the binding of neural agrin
to isolated -dystroglycan, however, suggesting either that
-dystroglycan is not the signaling receptor or that its properties
in the membrane are altered. Direct assay of the binding of muscle or
neural agrin to intact myotubes revealed only low-affinity binding. We
conclude that the signaling receptor for agrin is a high-affinity
receptor that is highly specific for the neural form.
Key words:
acetylcholine receptor;
agrin;
dystroglycan;
muscle;
receptor;
synaptogenesis
INTRODUCTION
Communication between skeletal muscle fibers and
the motor neurons that innervate them requires the development of
complex molecular machinery at the neuromuscular synapse. These events
are coordinated via local signaling cues derived from the neuron, the
muscle fiber, and the connecting basal lamina (Hall and Sanes, 1993 ).
One of the earliest events in synaptic differentiation is the
accumulation of acetylcholine receptors (AChRs) in the muscle membrane
underlying the nerve terminal. Initially, AChRs are found along the
length of the myotube surface, but when an ingrowing growth cone
contacts the muscle, the AChRs become concentrated at the site of
contact and extrasynaptic AChRs disappear (Salpeter et al., 1988). One
factor, agrin, has been isolated from the basal lamina and is able to
induce AChR clustering on cultured myotubes in the absence of
innervation (McMahan, 1990; Bowe and Fallon, 1995).
Agrin exists as several isoforms, generated by alternative splicing,
that differ in their biological activities and tissue localization
(Ferns et al., 1992; Ruegg et al., 1992; Rupp et al., 1992; Tsim et
al., 1992; Hoch et al., 1993). One of these forms is exclusively
localized in neurons (Hoch et al., 1993); it contains four and eight
amino acid inserts at the two most C-terminal splice sites,
respectively, and is designated Ag4,8. Assays of
cultured C2 myotubes have shown that Ag4,8 is the
most active isoform in causing AChRs to cluster (Ferns et al., 1992,
1993). The predominant form in muscle, Ag0,0,
which has inserts at neither of these sites, is significantly less
biologically active (Ferns et al., 1992, 1993). Investigation of
synapse formation in culture has shown that only neural-derived agrin
is responsible for initiating AChR clustering at sites of nerve-muscle
contact (Reist et al., 1992).
Despite its importance in synapse formation, the mechanism of agrin
action remains unknown. Recent experiments aimed at identifying the
receptor that mediates agrin-induced AChR clustering have identified
-dystroglycan as the major agrin-binding protein in muscle (Bowe et
al., 1994; Campanelli et al., 1994; Gee et al., 1994; Sugiyama et al.,
1994). -Dystroglycan is a component of the dystrophin-glycoprotein
complex that spans the muscle cell membrane and is thought to link the
cytoskeleton and the extracellular matrix (Ervasti and Campbell, 1991,
1993). Although -dystroglycan has been postulated to be the agrin
receptor responsible for AChR clustering (for review, see Bloch and
Randall, 1994; Fallon and Hall, 1994; Sealock and Froehner, 1994), one
troubling aspect of this hypothesis is that isolated -dystroglycan
binds soluble fragments of both Ag4,8 and
Ag0,0 with similar affinities (Sugiyama et al.,
1994), even though these fragments differ by 10,000-fold in their
biological activity (Ferns et al., 1993). We report here experiments
aimed at determining the specificity of the agrin signaling receptor
in situ. We find that the signaling receptor seems to be
highly specific, becoming activated by binding the neural, but not the
muscle, form of agrin with high affinity.
MATERIALS AND METHODS
Cell culture. C2 cells were cultured as described
previously (Gordon and Hall, 1989). Briefly, myoblasts were plated onto
tissue culture dishes or chamber slides (Nunc, Naperville, IL) and
maintained in C2 growth medium (DME-H21 containing 20% fetal calf
serum, 2 mM glutamine, 0.5% chick embryo extract
(Gibco, Gaithersburg, MD), and penicillin/streptomycin. At
confluence, the cells were induced to differentiate by switching the
medium to C2 fusion media (DME-H21 containing 5% horse serum and 2 mM glutamine).
Recombinant agrin production. Soluble agrin was produced by
transiently transfecting COS cells with rat agrin constructs containing
the C-terminal half of neural (C-Ag4,8) and
muscle (C-Ag0,0) agrin as described by Ferns et
al. (1993). COS cells were transfected using an adenovirus-mediated
DEAE-dextran method of transfection (Forsayeth and Garcia, 1994). The
conditioned medium containing the secreted agrin was collected, and the
concentration of agrin was measured by comparing the collected samples
with a known amount of agrin on Western blots. Purified, soluble agrin
was produced from HeLa suspension cultures infected with an agrin
recombinant vaccinia virus as described previously (Sugiyama et al.,
1994).
Antibodies. The rabbit serum antibody to purified rat agrin
was made commercially by Babco (Richmond, CA). The antibody against rat
agrin, mAb 131, was purchased from Stressgen Biotech (Victoria, BC,
Canada). The antibody against the AChR  subunit, mAb 124, was a gift
from Jon Lindstrom. 125I-labeled secondary
antibodies and HRP-conjugated antibodies were purchased from Amersham
(Arlington Heights, IL).
Agrin-induced AChR clustering assays. C2 myotube cultures
were grown on chamber slides as described above. After 2 d in C2 fusion
medium, soluble agrin (conditioned medium from agrin-transfected COS
cells) was added for 18 hr to induce AChR clustering as described
previously (Ferns et al., 1993). Excess muscle agrin (up to 1000-fold)
was added simultaneously with the neural agrin to determine whether
muscle agrin was able to inhibit neural agrin-induced clustering.
Myotubes with AChR clusters were visualized by staining the cultures
with rhodamine-conjugated -bungarotoxin for 1 hr at 37°C. The
cultures were then fixed in 2% paraformaldehyde for 20 min, mounted in
paraphenylenediamine, and visualized under fluorescence optics. AChR
clusters were quantified by counting 10 random visual fields under a
rhodamine filter with a 40× objective lens.
Agrin-induced phosphorylation of the AChR subunit. To
assay AChR-tyrosine phosphorylation, C2 myotube cultures were treated
without or with neural agrin (100 pM), muscle (10 nM) agrin, or both for 1 hr. Treated cells were
harvested in calcium/magnesium-free PBS containing 1 mM sodium orthovanadate, pelleted, and
resuspended in extraction buffer (25 mM Tris, pH
7.5, 25 mM glycine, 150 mM
NaCl, 5 mM each EDTA and EGTA, 50 mM sodium fluoride, 1 mM
sodium orthovanadate, and the protease inhibitors PMSF, benzamidine,
N-ethylmaleimide, and sodium tetrathionate) containing 1%
Triton X-100. Insoluble materials were removed by centrifugation for 4 min. Solubilized AChRs were isolated by incubating the extracts with
-bungarotoxin conjugated to Sepharose beads and then boiling the
beads in SDS-sample buffer to elute bound proteins. The proteins were
separated by SDS-PAGE and transferred to nitrocellulose membranes.
The nitrocellulose was probed with a phosphotyrosine antibody, mAb
4G10, followed by HRP-conjugated sheep anti-mouse Ig, and then
visualized using the enhanced chemiluminescence method (Amersham). To
identify the phosphotyrosine-immunoreactive bands and to confirm that
equal amounts of AChR were present in all lanes, the blots were
stripped with acid treatment (200 mM glycine and
0.2% Tween 20, pH 2.5) for 20 min and reprobed with mAb 124, which
recognizes the AChR subunit.
Membrane extract preparation and agrin overlays. Membrane
extracts were prepared from C2 myotubes and Torpedo electric
organ as described previously (Sugiyama et al., 1994). Myotubes were
harvested from 15 cm dishes, pelleted, resuspended in a 20 mM HEPES buffer containing 250 mM sucrose, 2 mM each EGTA
and EDTA, 1 mM PMSF, and 20 µg/ml each
leupeptin and aprotinin, followed by cell lysis in a Dounce
homogenizer. After removing insoluble material by centrifugation,
membranes were isolated by high-speed centrifugation at 100,000 × g, the membrane pellet was extracted with 25 mM octylglucopyranoside in phosphate buffer, and
insoluble material was removed by centrifugation. Torpedo
membranes were prepared as described by Ma et al. (1993). Frozen tissue
was ground and homogenized in 50 mM Tris
containing 400 mM NaCl. The homogenate was
cleared by centrifuging twice at 6000 × g, followed by
centrifugation at 100,000 × g to pellet membranes.
Torpedo membranes were then solubilized as described for C2
myotubes.
Membrane proteins were separated on 6% SDS-PAGE and transferred to
nitrocellulose membranes. The membranes were first blocked with a
blocking buffer (10 mM Tris, pH 7.5, 150 mM NaCl, and 1 mM each
MgCl2 and CaCl2) containing
10% horse serum, 5% dry milk, and 0.1% Tween 20. Blocked membranes
were then incubated for 2 hr at room temperature with
125I-labeled neural agrin in the presence or
absence of a 100-fold excess of either muscle
(Ag0,0) or neural (Ag4,8)
agrin. Agrin-binding proteins were visualized by autoradiography.
Agrin radioimmunoassay. C2 myotubes were grown in 12-well
tissue culture dishes. The radioimmunoassay (RIA) used in these
experiments was adapted from Nastuk et al. (1991). Briefly, soluble
agrin was diluted with DME-H21 containing 1% BSA and added to the
myotubes for 30 min at 4°C. This was followed by 30 min incubations
at 4°C with a monoclonal antibody against rat agrin, mAb 131 (diluted
1:1000), and 125I-labeled sheep anti-mouse Ig
(diluted 1:400). The cells were then extracted with 0.1 N NaOH, and
agrin binding was measured by gamma counting.
RESULTS
Although soluble C-terminal fragments of neural and muscle agrin
differ from each other by only 12 amino acids, there is a dramatic
difference in their biological activities. The concentration required
for half-maximal activity of soluble neural agrin is ~50
pM. In contrast, soluble muscle agrin gives a
negligible response, even at concentrations that are 10,000-fold higher
(Fig. 1a). These results are consistent with
those previously reported (Ferns et al., 1993).
Fig. 1.
Excess muscle agrin does not inhibit AChR
clustering induced by neural agrin. Soluble agrin was added to C2
myotubes for 18 hr, and the cultures were stained with
rhodamine-conjugated -bugarotoxin to assay AChR clustering.
Top left, Concentration curves for neural and muscle agrin.
The results of two independent experiments are shown. Top
right, The concentration curve for neural agrin with ( ) and
without ( ) 1000-fold excess muscle agrin. As the neural agrin
concentration was increased, the muscle agrin concentration was
increased proportionally. The data for neural agrin alone are similar
to that shown in top left panel. Both curves (top
left, top right) were plotted on a log scale.
Bottom left, Two concentrations of neural agrin (10 and 100 pM) plus a 10-, 100-, or 1000-fold excess of
muscle agrin. In both experiments, an excess of muscle agrin failed to
decrease the number of AChR clusters induced by neural agrin.
[View Larger Version of this Image (28K GIF file)]
Muscle agrin does not competitively inhibit neural agrin-induced
AChR clustering
The large differences in biological activities between muscle and
neural agrin could reflect a large difference in affinity of the two
forms of agrin for the receptor that mediates their effect on
clustering activity; alternatively, the two forms could have a similar
affinity for the postulated receptor but differ in their ability to
activate the receptor and stimulate downstream events. To test these
two possibilities, we have investigated the ability of muscle agrin to
compete with the neural form of agrin. If muscle agrin has an affinity
for the receptor that is comparable to that of neural agrin but fails
to activate the receptor, it is, in effect, a competitive inhibitor of
neural agrin and should block its activity. Alternatively, if the two
have affinities that differ widely, muscle agrin should be ineffective
in blocking the induction of AChR clusters by neural agrin.
We initially constructed a concentration curve measuring the ability of
neural agrin to induce AChR clustering in the presence or absence of
1000-fold excess muscle agrin by increasing the amount of muscle agrin
added with each concentration of neural agrin tested (Fig.
1b). At all concentrations of neural agrin tested, the
addition of 1000-fold excess muscle agrin did not affect the number of
AChR clusters. The highest concentration of muscle agrin used
competitively to inhibit neural agrin activity, 1500 pM, does not on its own induce AChR clustering
above background (see Fig. 1a). The presence of muscle agrin
also had no apparent effect on the size or shape of AChR clusters
induced by neural agrin (data not shown). As a further test, we chose
two concentrations of neural agrin (10 and 100 pM) in the lower range of the concentration curve
and tested the effect of varying excess amounts of muscle agrin (10×,
100×, and 1000×) on the ability of neural agrin to induce AChR
clusters (Fig. 1c). Again, an excess of muscle agrin failed
to inhibit the activity of neural agrin, similar to the results
obtained in Hoch et al. (1994). The results of our experiments thus
show that neural agrin induces AChR clusters by activating a receptor
that binds muscle agrin poorly, if at all.
Muscle agrin does not inhibit neural agrin-induced phosphorylation
of the AChR subunit
We also tested whether muscle agrin could block downstream events
induced by neural agrin. Although the signaling pathway by which agrin
induces the formation of AChR clusters is unknown, agrin has been shown
to induce phosphorylation of the subunit of the AChR, both in chick
(Wallace et al., 1991; Qu and Huganir, 1994) and mouse muscle (Ferns et
al., 1996). In C2 myotubes, agrin produces a rapid, but transient,
tyrosine phosphorylation of the subunit (Ferns et al., 1996).
Tyrosine kinase inhibitors that block AChR phosphorylation also block
agrin-induced AChR clustering under the same conditions (Wallace, 1994;
Ferns et al., 1996). We tested, therefore, whether this early event in
the signaling pathway for neural agrin can be blocked by muscle agrin.
Neural agrin was added to myotube cultures at approximately the same
concentration as that needed to induce AChR clustering (see Fig.
1a). When C2 myotubes were treated for 1 hr (a time point
that correlates to peak phosphorylation of the subunit) (Ferns et
al., 1996) with neural agrin at 100 pM, a
concentration that induces approximately half-maximal clustering, there
was a strong induction of tyrosine phosphorylation of the subunit
(Fig. 2). In contrast, muscle agrin failed to induce
tyrosine phosphorylation of the subunit even at a concentration of
10 nM. The immunoblots were reprobed with an
antibody to the AChR subunit to confirm the identity of the
phosphotyrosine band and to ensure that equal amounts of AChR were
present for each sample. When a 100-fold excess of muscle agrin was
added to 100 pM neural agrin, there was no
detectable effect on the level of -subunit phosphorylation (Fig. 2).
The failure of muscle agrin to block this aspect of neural agrin
signaling suggests that neural and muscle agrin differ significantly in
their affinities for the signaling receptor. Thus, as with AChR
clustering, neural agrin seems to stimulate tyrosine phosphorylation of
the AChR subunit via a receptor that is specific for the neural
isoform.
Fig. 2.
Muscle agrin does not competitively inhibit
phosphorylation of the AChR subunit induced by neural agrin. C2
myotubes were treated with neural agrin (C-Ag4,8)
for 1 hr in the presence or absence of a 100-fold excess of muscle
agrin (C-Ag0,0). AChRs were isolated from cell
extracts using -bugarotoxin-conjugated Sepharose beads and immuno-
blotted with a phosphotyrosine antibody as described in Materials
and Methods. Lane 1, Blank
beads+; lane 2, no agrin;
lane 3, 10 nM muscle agrin;
lane 4, 100 pM neural agrin;
lane 5, 100 pM neural agrin + 10 nM muscle agrin. A 100-fold excess of muscle
agrin did not alter the level of neural agrin-induced AChR phosphorylation. The two faint tyrosine-phosphorylated bands of ~106
kDa in the muscle agrin-treated extracts appeared inconsistently and
were not further characterized.
[View Larger Version of this Image (25K GIF file)]
Muscle agrin inhibits binding of neural agrin
to -dystroglycan on nitrocellulose membranes
Because -dystroglycan has been postulated to be the agrin
receptor (for review, see Bloch and Randall, 1994; Fallon and Hall,
1994; Sealock and Froehner, 1994), we further investigated the binding
of the agrin isoforms to -dystroglycan in vitro. If
-dystroglycan were the receptor, a possible explanation for the
discrepancy between our previous results in vitro (Sugiyama
et al., 1994) and our present results is that neural and muscle agrin
bind to two different, noncompeting sites on -dystroglycan. We
therefore wished to examine competition between binding of the two
isoforms to -dystroglycan in vitro. We extracted crude
membrane preparations from Torpedo electric organ and C2
myotubes with detergent and subjected the proteins in the extract to
SDS-PAGE. The separated proteins were then transferred to
nitrocellulose blots and incubated with 0.5 nM
125I-labeled C-terminal, soluble neural agrin. In
agreement with our previous results, a single band of 200 or 150 kDa,
corresponding to -dystroglycan, was visualized in blots of
Torpedo electric organ (Fig. 3a) and C2
muscle extracts (Fig. 3b), respectively. To determine
whether muscle agrin could inhibit the binding of neural agrin to
-dystroglycan, we added a 100-fold excess of either unlabeled muscle
or unlabeled neural agrin simultaneously with the radiolabeled neural
form. In both cases, radiolabeled neural agrin binding to
-dystroglycan in the extracts was inhibited by excess unlabeled
agrin. The ability of muscle agrin to block the binding of neural agrin
to -dystroglycan on nitrocellulose membranes indicates that muscle
and neural agrin bind to isolated -dystroglycan in a competitive
manner.
Fig. 3.
Muscle agrin competitively inhibits neural agrin
binding to -dystroglycan on nitrocellulose blots. Membrane extracts
from (A) Torpedo electric organ (T)
and (B) C2 myotubes (C) were isolated by
SDS-PAGE, transferred to nitrocellulose membranes, and incubated with
125I-labeled neural agrin (0.5 and 0.8 nM, respectively) without and with 50 nM neural (C-Ag4,8) and
muscle (C-Ag0,0) agrin. Radiolabeled agrin bound
to a protein of 200 kDa (A) and 150 kDa (B) was
identified as -dystroglycan. This binding was inhibited by an excess
of either unlabeled neural or muscle agrin.
[View Larger Version of this Image (30K GIF file)]
Neural and muscle agrin binding to cultured myotubes
To investigate the discrepancy between the specificity of
biological activity and the nonspecificity displayed by neural and
muscle agrin binding to -dystroglycan, we examined the binding of
agrin to intact C2 myotube cultures. C2 myotubes were incubated with
soluble rat agrin, and binding was measured by an RIA using mAb 131, a
mouse antibody specific for rat agrin, followed by
125I-labeled sheep antibody to mouse total Ig.
mAb 131 recognizes rat but not mouse agrin, thus differentiating
between endogenous and exogenous agrin. Agrin binding to cultured C2
myotubes required calcium (Fig. 4a), and over
the concentration range used, the half-maximal binding in different
experiments ranged from 50 to 200 nM (Fig.
4b). The high concentrations of agrin required to see
binding is in general agreement with experiments by Campanelli et al.
(1994), who estimated a binding constant of 10-30
nM, based on a fluorescence immunoassay and flow
cytometry of C2 myotubes in suspension. In contrast, Nastuk et al.
(1991) estimated a value of 100-500 pM for the
binding of agrin to chick myotubes, using an RIA similar to ours.
Fig. 4.
Binding characteristics of neural and muscle agrin
to cultured C2 myotubes. Agrin binding to intact C2 myotubes was
measured by RIA as described in Materials and Methods. Top
left, C2 myotubes were incubated with increasing concentrations of
muscle agrin in the presence () or absence () of calcium (2 mM EDTA). Top right, Neural () and
muscle () agrin-binding curves were constructed with C2 myotubes.
Both curves (top left, top right) were plotted on
a log scale. Bottom left, C2 myotubes were incubated with
500 µM muscle agrin and increasing
concentrations of heparin. The low-affinity agrin binding to intact
myotubes is dependent on the presence of calcium and is relatively
insensitive to heparin.
[View Larger Version of this Image (23K GIF file)]
The binding characteristics of both neural and muscle agrin isoforms to
intact C2 myotubes indicate the presence of very abundant, low-affinity
agrin-binding sites. This is in contrast to the nanomolar
concentrations required to observe binding of agrin to solubilized
-dystroglycan from C2 myotube extracts on nitrocellulose blots
(Sugiyama et al., 1994) and to the picomolar concentrations required
for agrin-induced AChR-clustering activity (Ferns et al., 1993). In
addition, at least 1000-fold more heparin was needed to inhibit agrin
binding to intact myotubes (Fig. 4c) than to inhibit agrin
binding to -dystroglycan in vitro (Gee et al., 1994;
Sugiyama et al., 1994) or to inhibit agrin-induced AChR clustering
(Hirano and Kidokoro, 1989; Wallace, 1990; Saito et al., 1993). Because
of these differences, it seems unlikely that the binding site that we
are measuring corresponds to -dystroglycan or to the receptor that
induces AChR clustering.
DISCUSSION
Understanding the mechanism by which neurally derived agrin acts
requires the characterization and identification of the signaling
receptor responsible for transducing information across the
postsynaptic membrane. A prominent feature of the ability of agrin to
stimulate AChR clustering is the remarkable specificity of the spliced
forms. The addition of two inserts of four and eight amino acids,
respectively, to a soluble, C-terminal fragment of agrin results in a
dramatic increase in biological activity (Ferns et al., 1993). The
relative activities of the cell-attached, full-length agrin forms are
more difficult to estimate quantitatively but, even in this context,
the neural form of agrin is clearly more active than the muscle form
(Ferns et al., 1992; Ruegg et al., 1992).
The large difference between neural and muscle agrin activities raises
the question of whether the two isoforms have different affinities for
the physiologically active agrin receptor or whether they differ in
their ability to activate downstream events. If the muscle isoform
binds the signaling receptor with an affinity similar to that of neural
agrin but is unable to activate the signaling pathway, it should act as
an antagonist for neural agrin activity. When an excess of muscle agrin
(up to 1000-fold) was added along with the neural form, however, it was
unable to inhibit neural agrin-induced AChR clustering on C2 myotube
cultures (Fig. 1b,c) competitively. These results
are consistent with those of Hoch et al. (1994), who found that a
fragment of soluble muscle agrin did not inhibit the activity of the
soluble neural form.
Further evidence that muscle agrin does not act as an antagonist for
neural agrin comes from tyrosine phosphorylation assays. Neural agrin
induces subunit phosphorylation at concentrations corresponding to
its clustering activity (Ferns et al., 1996). Furthermore, tyrosine
kinase inhibitors that prevent phosphorylation of the subunit also
inhibit agrin-induced AChR clustering (Wallace, 1994; Ferns et al.,
1996), suggesting that tyrosine phosphorylation of the AChR subunit
or other proteins is involved in the agrin-mediated clustering process.
Muscle agrin, on the other hand, neither induces phosphorylation of the
AChR subunit at the concentrations examined (Ferns et al., 1996)
nor clusters AChRs at the corresponding concentrations (Fig.
1a). When we investigated whether the presence of excess
muscle agrin affects neural agrin-induced tyrosine phosphorylation of
the AChR subunit, we found that the neural agrin retained its
activity (Fig. 2). Thus, the results from experiments assaying
agrin-induced AChR clustering and AChR phosphorylation show that muscle
agrin does not act as a competitive inhibitor for the neural agrin
signaling pathway. The differences between the two agrin isoforms must
therefore arise from their differing affinities for the signaling
receptor rather than through a downstream effect.
The results described above provide a strong criterion for
identification of the agrin receptor or receptor complex that mediates
its biological activity: it must show a much higher affinity for neural
agrin than muscle agrin. Previous work has identified -dystroglycan
as an agrin-binding, membrane-associated glycoprotein (Bowe et al.,
1994; Campanelli et al., 1994; Gee et al., 1994; Sugiyama et al., 1994)
and has led to the suggestion by some that it is the agrin receptor
(for review, see Bloch and Randall, 1994; Fallon and Hall, 1994;
Sealock and Froehner, 1994). No other agrin-binding protein in muscle
or Torpedo electric organ has been identified. Some of the
binding characteristics of -dystroglycan correlate to the
characteristics of the biological activity of agrin; for example, both
the binding and the AChR-clustering activity are calcium-dependent and
inhibited by polyanions such as heparin. Moreover, agrin binding to
-dystroglycan is diminished in muscle cell variants that respond
poorly to agrin (Campanelli et al., 1994; Gee et al., 1994; Sugiyama et
al., 1994). Several features of the binding of agrin to
-dystroglycan, however, are not consistent with its biological
activity: isolated -dystroglycan binds both neural and muscle agrin
with a binding affinity that is lower than that required for biological
activity, and it shows no apparent specificity for the neural form
(Sugiyama et al., 1994). We now have extended these results to show
that excess muscle agrin can block the binding of neural agrin to
-dystroglycan on nitrocellulose blots (Fig.
3a,b), thus ruling out the possibility that the
two forms are binding to nonoverlapping or noninteracting sites on
-dystroglycan.
The binding properties of -dystroglycan in situ could
differ from those of the isolated protein, either because denaturation
and refolding on the nitrocellulose blots changed its original
conformation or because specific interactions with neighboring proteins
in its native environment were lost. We attempted to resolve these
difficulties by measuring the binding of agrin to -dystroglycan in
intact muscle cells. Unfortunately, in C2 myotubes, the binding that we
detected lacked both the characteristics of the agrin receptor, as
defined by its biological activity, and the characteristics of binding
to -dystroglycan, as defined on nitrocellulose blots. Agrin binding
to C2 myotubes (Fig. 4b) was detectable only at much higher
concentrations than those required either for binding to
-dystroglycan on nitrocellulose blots or for biological activity,
and the binding was relatively insensitive to heparin (Fig.
4c). In fact, in comparison, the binding of muscle agrin
seems to be higher than that of neural agrin (Fig. 4b);
thus, it may also bind to a second component on C2 myotubes that does
not detect neural agrin. Ma et al. (1993) have found a binding site in
chick muscle and in Torpedo membranes, which they believe to
be -dystroglycan, that has a higher affinity (~0.1
nM) than that described here. The affinities of
these binding sites for the two forms of agrin, however, have not been
tested. In C2 myotubes, the high-abundance, low-affinity sites that we
see may obscure the binding to -dystroglycan or to any other
high-affinity binding site that presumably occurs. In any case, the
low-affinity binding that we observe, whatever its identity, does not
have the specificity of the biological response.
Because we are unable to detect the binding of agrin to
-dystroglycan in intact myotubes, our experiments do not yield
decisive information about whether it is the agrin receptor that
activates the pathway leading to AChR clustering. In situ,
-dystroglycan is part of a complex of six proteins associated with
dystrophin and is believed to be tightly associated to one of them, the
integral membrane protein -dystroglycan (Ervasti et al., 1990;
Yoshida and Ozawa, 1990; Ervasti and Campbell, 1991). This association,
or others, could change the specificity of agrin binding exhibited by
-dystroglycan. Such a change must be a large one, because the
isolated protein exhibits roughly equal affinities for the two forms of
agrin, whereas the biological receptor shows a preference of over
1000-fold for the neural form, which differs from the muscle form by
only 12 amino acids. Although the role of -dystroglycan thus remains
unsettled, the high degree of specificity that we demonstrate for the
agrin-signaling receptor in situ provides an important
criterion for its ultimate identification.
FOOTNOTES
Received Jan. 10, 1996; revised March 18, 1996; accepted March 25, 1996.
This work was supported by grants from National Institutes of
Health.
D. C. B. and J. S. contributed equally to this manuscript.
Correspondence should be addressed to Dr. Zach W. Hall, Office of the
Director, National Institute of Neurological Disorders and Stroke,
National Institutes of Health, Bethesda, MD 20892.
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